Download Progressive Transverse Microtubule Array

This article is a Plant Cell Advance Online Publication. The date of its first appearance online is the official date of publication. The article has been
edited and the authors have corrected proofs, but minor changes could be made before the final version is published. Posting this version online
reduces the time to publication by several weeks.
Progressive Transverse Microtubule Array Organization in
Hormone-Induced Arabidopsis Hypocotyl Cells
W
Laura Vineyard, Andrew Elliott, Sonia Dhingra, Jessica R. Lucas,2 and Sidney L. Shaw1
Department of Biology, Indiana University, Bloomington, Indiana 47405
The acentriolar cortical microtubule arrays in dark-grown hypocotyl cells organize into a transverse coaligned pattern that is
critical for axial plant growth. In light-grown Arabidopsis thaliana seedlings, the cortical array on the outer (periclinal) cell face
creates a variety of array patterns with a significant bias (>3:1) for microtubules polymerizing edge-ward and into the side
(anticlinal) faces of the cell. To study the mechanisms required for creating the transverse coalignment, we developed a dualhormone protocol that synchronously induces ;80% of the light-grown hypocotyl cells to form transverse arrays over a 2-h
period. Repatterning occurred in two phases, beginning with an initial 30 to 40% decrease in polymerizing plus ends prior to
visible changes in the array pattern. Transverse organization initiated at the cell’s midzone by 45 min after induction and
progressed bidirectionally toward the apical and basal ends of the cell. Reorganization corrected the edge-ward bias in
polymerization and proceeded without transiting through an obligate intermediate pattern. Quantitative comparisons of
uninduced and induced microtubule arrays showed a limited deconstruction of the initial periclinal array followed by
a progressive array reorganization to transverse coordinated between the anticlinal and periclinal cell faces.
INTRODUCTION
Microtubules are cellular polymers composed of repeating a-b
tubulin subunits assembled in a head-to-tail orientation to form
hollow tubes (Ledbetter and Porter, 1964; Howard and Hyman,
2003). In most interphase animal cells, microtubules nucleate from
g-tubulin–containing complexes that are localized to a discrete
organelle called the centrosome (Mazia, 1987; Doxsey, 2001;
Bornens, 2002). By collecting the microtubule minus ends to the
centrosome, animal cells create a radial array having the dynamic
plus ends projecting outward toward the cell periphery. Plant microtubules also nucleate on g-tubulin complexes (Liu et al., 1993,
1994; McDonald et al., 1993; Murata et al., 2005; Pastuglia et al.,
2006), but unlike animals, flowering plants do not make centrioles
or a centralized microtubule-organizing center (Newcomb, 1969).
Interphase plant microtubules nucleate from distributed sites
throughout the cell cortex, giving rise to a dynamic polymer network closely associated with the plasma membrane (Shaw et al.,
2003; Murata et al., 2005; Nakamura and Hashimoto, 2009).
The organizational state of the plant cortical microtubule array
influences the direction of plant cell expansion (Baskin, 2001;
Lloyd, 2011). Hypocotyl and root cells extend axially to push the
chlorophyll-bearing cotyledons into the sunlight and the primary
root into the soil, respectively. The cortical microtubules in both
cases organize into coaligned patterns that are transverse to the
plant growth axis (Baskin, 2001). These microtubules, in turn,
1 Address
correspondence to [email protected].
Address: Santa Clara University, Department of Biology, 500 El
Camino Real, Santa Clara, California, 95053
The author responsible for distribution of materials integral to the findings
presented in this article in accordance with the policy described in the
Instructions for Authors (www.plantcell.org) is: Sidney L. Shaw (sishaw@
indiana.edu).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.112.107326
2 Current
pattern cellulose deposition into the cell wall by guiding the
plasma membrane–bound cellulose synthase complexes (Green,
1962; Giddings et al., 1980; Paredez et al., 2006; Chan et al.,
2010). The transversely oriented cellulose fibers are proposed
to retard turgor-driven radial cell swelling in favor of axial cell
extension (Baskin, 2005). In the case of the dark-grown seedling
hypocotyl, cells can extend to ;500 times their original length by
this mechanism in the absence of new cell divisions (Gendreau
et al., 1997; Le et al., 2005). Exposing etiolated hypocotyl cells
to light rapidly reorganizes the microtubule arrays, leading to
a new pattern of cellulose deposition accompanying the switch
from axial growth to radial thickening (Refrégier et al., 2004;
Vandenbussche et al., 2005).
The mechanisms by which the cortical microtubules coalign and
orient to the cell growth axis have been the subject of considerable
speculation (Green, 1962; Ledbetter, 1982; Lloyd and Chan, 2002;
Dixit and Cyr, 2004a; Ehrhardt and Shaw, 2006; Hashimoto and
Kato, 2006; Baulin et al., 2007; Lucas and Shaw, 2008; Sedbrook
and Kaloriti, 2008; Allard et al., 2010b; Eren et al., 2010; Ambrose
et al., 2011). Early proposals, based on electron and immunofluorescence microscopy of fixed cells, focused on lateral sliding of
the microtubules, possibly using motors to power the interactions
between microtubules on the cell cortex (Hardham and Gunning,
1978; Lloyd and Wells, 1985; Cyr, 1994; Cyr and Palevitz, 1995;
Wymer and Lloyd, 1996; Lloyd and Chan, 2002). Observations
made in live cells found no evidence for lateral microtubule sliding,
finding instead that polymers in hypocotyl cortical arrays remained
attached to the cell cortex and exhibited a form of polymer
treadmilling (Shaw et al., 2003; Ehrhardt and Shaw, 2006). The
treadmilling motility facilitates cortical polymer interactions that
often result in microtubule bundling (Shaw et al., 2003; Dixit and
Cyr, 2004b) or changes to the polymerization state of the microtubule (Wightman and Turner, 2007). Plant microtubules were
observed to nucleate at the cell cortex (Shaw et al., 2003; Murata
The Plant Cell Preview, www.aspb.org ã 2013 American Society of Plant Biologists. All rights reserved.
1 of 15
2 of 15
The Plant Cell
et al., 2005) and either coalign to form a bundle or branch from
an existing microtubule (Shaw et al., 2003; Murata et al., 2005;
Nakamura et al., 2010). These live-cell observations of plant
cortical microtubules in hypocotyl cells have provided a number
of possible mechanisms that could contribute to transverse array
organization (Ehrhardt, 2008; Lloyd and Chan, 2008; Lucas and
Shaw, 2008).
Contemporary models for transverse microtubule array organization have focused on cell geometry and the potential
self-organizing properties of the treadmilling microtubules. Selforganizing properties have generally been taken to be those that
are a state property of all the microtubules in the system that
can affect some change in the orientation of the microtubules
within an array. Computational modeling was used to explore
the proposal that angle-dependent association of treadmilling
microtubules into bundles drives array coalignment (Wasteneys
and Ambrose, 2009). These studies suggest that angledependent bundling plays a significant role but does not constitute a complete self-organizing system with reference to the
cell axis (Baulin et al., 2007; Allard et al., 2010b; Eren et al.,
2010). The addition of other phenomena, such as new microtubules entering from the cell’s longitudinal side walls or an increased probability of switching from growth to shortening (i.e.,
catastrophe) at microtubule intersections or at the apical or basal
ends of the cell, have been introduced to improve the likelihood of
convergence to a transverse coalignment (Baulin et al., 2007;
Allard et al., 2010b; Eren et al., 2010; Sambade et al., 2012).
Recent experimental evidence for localized catastrophes at apical
and basal edges of the cell supports a model where longitudinally
oriented microtubules are preferentially removed to leave a predominantly transverse array (Ambrose et al., 2011). In sum, the
leading model to explain how plant microtubules become coaligned to a transverse orientation relies on a mechanism for actively removing the longitudinally oriented polymers going into, or
emerging from, the apical and basal ends of the cell and selforganizing properties, such as bundling interactions between the
remaining polymers (Ehrhardt and Shaw, 2006; Ambrose et al.,
2011; Sambade et al., 2012).
A major impediment to testing these models and for determining the molecular and biophysical contributions to array
organization has been the inability to routinely observe the array
organizing into a transverse pattern. Light in general, and specifically from the microscope, actively signals the hypocotyl cells
to form nontransverse arrays, making high-resolution imaging
studies of cells organizing into a transverse coalignment
somewhat inaccessible. Both auxin and gibberellic acid affect
hypocotyl cell elongation, and both hormones alter microtubule
array organization when exogenously applied (Shibaoka, 1974,
1993, 1994; Roberts et al., 1985; Ishida and Katsumi, 1991;
Lloyd et al., 1995; Shibaoka and Takesue, 1999; Wenzel et al.,
2000; Folta et al., 2003). Prior studies have clearly established
that either hormone can promote microtubule array organization, though high-resolution temporal and spatial data have not
been available from these studies for application to models for
array organization (Takesue and Shibaoka, 1999; Wenzel et al.,
2000). Using a combination of these hormones to synchronously
induce light-grown Arabidopsis thaliana seedlings, we captured
extensive time-lapse observations of the temporal and spatial
changes in array organization leading to transverse array patterning in this important model system. Our quantitative analysis
of these data provides a novel cell biological model for transverse patterning with array organization initiating as a band of
transverse polymers at the cell midzone and progressively reorganizing toward the apical and basal cell faces.
RESULTS
Gibberellic Acid/Indole-3-Acetic Acid Induce Robust
Transverse Cortical Microtubule Array Organization
Our first goal was to establish a reproducible method for synchronously inducing light-grown Arabidopsis hypocotyl cells to
form transverse coaligned microtubule arrays. We began by
classifying the array patterns in untreated cells and determining
the distribution of pattern classes in 6-d-old light-grown seedlings expressing a tubulin to green fluorescent protein (GFP):
TuA6 transgene. Using reconstructed image composites of the
entire hypocotyl length from confocal fluorescence microscopy,
we visually classified the microtubule array organization for each
cell from two adjacent hypocotyl cell files as one of four array
patterns (Figures 1A to 1D). Arrays without a dominant coalignment typically had microtubules radiating from the interior of
the outer (periclinal) cell face down the (anticlinal) sides of the
cell, forming an inverted basket pattern (Figure 1A). Arrays with
predominantly coaligned microtubules were classified as longitudinal (Figure 1B), oblique (Figure 1C), or transverse (Figure 1D)
patterns, where oblique was 15° to 75° relative to the cell growth
axis. Using the percentage of cells in each hypocotyl as a
quantitative measure of array organization states (Figure 1E), we
found that ;60% of the cells in light-grown control plants
showed a strong degree of microtubule coalignment at this
developmental stage with ;40% showing a less coaligned (i.e.,
basket) organization. We found fewer than 20% of the cells
exhibited a transverse coalignment when taken directly from
growth plates to the microscope (untreated in Figure 1F).
Using this quantitative assay to determine the percentage of
each array class, we examined the effects of auxin (indole-3acetic acid [IAA]) and gibberellic acid (GA4) on array organization. Transfer of seedlings from agar growth plates to agar
treatment plates was inconsistent and damaging to plants. We
therefore transferred seedlings from agar plates into liquid culture containing hormones or solvent controls and incubated the
seedlings for 2 h in the light at 22°C prior to imaging. For each
treatment, we tabulated the total percentage of arrays falling into
each pattern class (Figure 1E).
Transfer of seedlings into a liquid medium, with or without
solvents, led to a small but significant (P < 0.01 comparing 22
untreated to 14 Murashige and Skoog [MS] plants) increase in
the percentage of cells with transversely aligned microtubule
arrays when compared with controls taken directly from growth
plates (Figure 1F). Solvent treatment trended slightly higher for
transverse arrays but was not significantly different (P > 0.05
comparing 13 solvent to 14 MS plants) from liquid medium alone
(Figure 1F). These experiments establish that placing seedlings
into a liquid medium will alter the distribution of microtubule
Transverse Microtubule Array Organization
3 of 15
array patterns but does not induce a single or predominant class
of array pattern.
Treatment with 10 µM GA4 increased the relative percentage of
cells with transverse arrays to near 50% at 2 h (Figure 1E). GA4
treatment appeared to reduce the percentage of basket-patterned
arrays but had little effect on the total percentage of oblique arrays
at 2 h. Treatment with 0.5 µM IAA resulted in hypocotyls having
slightly more than 50% of cells exhibiting transversely coaligned
microtubule arrays within 2 h (Figure 1E). All other array organizations were proportionately reduced. Both the GA4 (P = 0.01
comparing 18 GA4 to 14 solvent-treated plants) and IAA (P < 0.01
comparing 18 IAA to 14 solvent-treated plants) treatments were
significant relative to solvent controls (Figure 1F).
The relatively modest increase in cells with transverse microtubule arrays and the variability between plants observed with GA4
or IAA used alone was not sufficient for robust quantitative assays
or routine imaging of individual cells. We therefore tested induction
using both hormones in combination and observed 75 to 85% of
the epidermal hypocotyl cells in each plant to organize transverse
array patterns by 2 h (Figures 1E and 1F). The effect of combining
both hormones was significant (P < 0.01 comparing 15 GA4/IAA
plants) relative to the addition of either hormone alone. Plants
observed at 6 h had 75 to 85% transverse microtubule arrays,
suggesting that this may be an upper limit for this treatment
regime. These experiments establish a robust and repeatable
method for synchronously inducing the microtubule arrays in wildtype light-grown Arabidopsis seedlings at 6 d after germination to
organize from a variety of array patterns into a transverse coaligned array over a 2-h time course.
GA4/IAA Induce Significant Axial Extension of the
Light-Grown Hypocotyl
Figure 1. GA4/IAA-Induced Cortical Microtubule Array Reorganization.
(A) to (D) Confocal microscopy images of 6-d-old light-grown epidermal
hypocotyl cells showing basket (A), longitudinal (B), oblique (C), and
transverse (D) microtubule array organization in GFP:TUA6-expressing
plants. Bars = 10 µm.
(E) The percentage of epidermal hypocotyl cells exhibiting each array
organization class for seedlings imaged directly from germination plates
(untreated) or after 2 h in liquid culture with the associated treatment
condition (bar pattern designations for array class displayed in [A] to [D]).
(F) Box and whisker quartile plots for the percentage of cells with
transverse arrays at 2 h including the mean value (filled circle) and outlying data (open circles) for each treatment. Number of observations for
(E) and (F), as plants/cells, were as follows: untreated (26/827), halfstrength MS (15/376), solvent (13/397), GA4 (19/477), IAA (19/478), GA4 +
IAA (15/476).
(G) The effect of all treatments on hypocotyl length was assayed by direct
measurement of seedling hypocotyls after 72 h of continuous treatment.
Bars represent mean values from >45 total plants pooled from three experiments for each treatment with variation displayed as the pooled SD.
Both GA4 and IAA have varying effects on hypocotyl growth,
possibly related to the developmental stage and growth environment of the seedling (Shibaoka, 1974, 1994; Huang and
Lloyd, 1999; Shibaoka and Takesue, 1999). The effect of single
and combination treatments with GA4 and IAA on hypocotyl
growth on 6-d-old light-grown Arabidopsis seedlings was determined by measuring hypocotyl length 72 h after hormone
treatment. Light-grown seedlings were treated with 10 µM GA4,
0.5 µM IAA, or both hormones in combination starting at 6 d
after germination. Hypocotyl length was longer for plants
transferred to liquid culture with and without the solvent control (Figure 1G). Treatment of light-grown seedlings at this
developmental stage with GA4 did not produce a substantive
increase in hypocotyl length over solvent controls (P > 0.05
for 36 GA4 and 48 solvent-treated plants), but IAA treatment
resulted in a significant change (P < 0.01 for 36 IAA-treated
plants) with hypocotyls ;130% as long as solvent control
treated plants. Treatment with both hormones resulted in
hypocotyls that were ;200% as long as solvent controls,
significantly longer (P < 0.01 for 48 GA4/IAA-treated plants)
than control and IAA-treated seedlings (Figure 1G). These data
indicate that the combined hormone treatment leads to a substantially higher percentage of hypocotyl cells with transverse
coaligned microtubule arrays correlated with a substantial increase
in axial hypocotyl extension.
4 of 15
The Plant Cell
Timing of Hormone-Induced Transverse Array Organization
We examined the relative timing of microtubule array reorganization
from each starting array pattern to a transverse coaligned pattern.
We placed seedlings into liquid culture on slides containing
a combination of 10 µM IAA and 0.5 µM GA4 and imaged hypocotyls every 15 min for 2 h. The percentage of hypocotyl cells
observed in each array pattern at each time point was tabulated for
;40 cells on the imaged side of each seedling from the time course
(Figure 2A). The compiled data for cells with transverse arrays
suggest that the induced reorganization occurs in two phases.
There is an initial lag phase from 0 to 45 min where the number of
cells classed as transverse increases relatively slowly. The lag was
followed by a second, faster phase, established by 45 to 60 min
after induction and continuing until 75 to 80% of the cells were
transverse at 120 min (Figure 2A). The starting percentage of hypocotyl cells exhibiting a longitudinal array pattern (;20%) showed
an initial drop during the lag phase followed by a slow dissipation to
zero by 2 h (Figure 2A). Although we observed a small initial rise in
arrays classed as baskets, we found no obligate transition to an
intermediate array pattern (e.g., oblique to basket) prior to achieving
a transverse array organization.
To determine if any starting array pattern was more rapidly
reorganized to a transverse pattern, we examined the relative
timing of cells transitioning from each starting array pattern to
a final transverse array orientation (Figure 2B). Cells that started
as oblique arrays reorganized to transverse at approximately the
same rate as cells that started with basket-patterned arrays.
These data indicate that the coalignment of microtubules found
for the oblique arrays does not predispose them to a more rapid
transition to transverse coalignment. Cells having initially longitudinal arrays were somewhat slower to be classed as transverse (15 min delay relative to other patterns) but ultimately
became transverse in the same 2-h period as the other arrays
(Figure 2B).
Transverse Array Organization Initiates in the
Midzone of the Cell
To determine how the pattern of microtubules changes over
time to produce a transverse coalignment, we imaged individual
hypocotyl cells expressing a GFP:TuA6 microtubule marker at
7.5- or 15-min intervals for 2.5 h using confocal laser scanning
microscopy (Figure 3). Solvent-treated control cells showed
occasional rearrangements of the cortical microtubules over the
120- to 150-min time period but showed no distinct organizational changes into a defined pattern on this time scale (Figure
3A). We examined the progression of changes to microtubule
array organization in control cells (Figure 3A) and in hormonetreated cells that originated in basket (Figure 3B), oblique (Figure
3C), or longitudinal (Figure 3D) array patterns. Hormone-treated
cells originating in a transverse coalignment simply maintained
a transverse coalignment.
Microtubule array organization to a transverse coalignment
exhibited three consistent features that were independent of the
starting array pattern. Single-cell imaging confirmed that no
significant changes in array pattern appeared in the initial 30 to
45 min after hormone induction (Figures 3B to 3D). Transverse
Figure 2. Timing of Hormone-Induced Microtubule Array Reorganization.
Epidermal hypocotyl cells (n = 196 cells across six plants) were imaged after
induction with 10 µM IAA and 0.5 µM GA4 at 15-min intervals for 120 min.
(A) The percentage of all cells exhibiting each array pattern class is
plotted at each time interval. The percentage of cells with arrays classed
as transverse shows an initially slow rise followed by a more rapid phase.
(B) The relative percentage of each array class that has reorganized to
a transverse orientation was plotted for arrays starting as oblique, longitudinal, or basket patterns. Cells with initially oblique (;40% of cells)
arrays transited to transverse slightly ahead of cells with basket (;32%
of cells) array forms and longitudinal arrays (;18% of cells) were slightly
delayed when compared with both basket and oblique.
coalignment was initially observed around the midzone of the
elongated hypocotyl cells and progressed in both an apical and
basal direction. Finally, we observed that the apical and basal
regions of the outer cell face were the last regions to reorganize
into a transverse pattern. Persistent longitudinal polymers at the
cell’s apical and basal ends often interacted with the advancing
front of transverse microtubules, yielding curved polymers in the
corners of the cell (e.g., Figure 3D, 82.5 to 97.5 min).
Cells with an initially oblique microtubule array progressed
through a series of oblique organizational states, giving the
appearance of a rotating array orientation (Figure 3C) as previously described (Roberts et al., 1985; Yuan et al., 1995; Wymer
and Lloyd, 1996; Wymer et al., 1996). Using time-lapse image
data taken at 3-min intervals, we observed new microtubule
polymer appearing between the existing oblique microtubules
and a progressive loss of the original oblique polymers (Figure
Transverse Microtubule Array Organization
5 of 15
Figure 3. Cortical Microtubule Reorganization from Basket, Oblique, or Longitudinal Patterns to Transverse over Time.
Confocal image projections of GFP:TUA6-labeled microtubules in epidermal hypocotyl cells grown for 6 d in continuous light. Time in minutes. Bars =
10 µm in (A) to (D).
(A) Selected images from a solvent control cell over 120 min of imaging. The array pattern changes over the course of observation but achieves no
defined pattern of organization.
(B) to (D) Plants treated with GA4 and IAA.
(B) Cells with basket formed arrays are characterized by microtubules oriented perpendicular to the cell perimeter around the entire edge of the outer
periclinal cell face. Transversely aligned microtubules appear across the middle of the cell between 45 and 60 min and progress toward the cell apex
and base. Note the persistence of longitudinal microtubules at the cell apex and base.
(C) Oblique arrays appeared to rotate into a transverse position, progressively creating arrays of shallower angle relative to the cell growth axis. The
middle zone of the cell becomes transverse ahead of the apex and base (67.5 min), creating a transient sigmoidal shape for the array polymers prior to
becoming transverse.
(D) Reorganization of longitudinal arrays was characterized by the invasion of new microtubules from the lateral side faces of the cell (90 to 97.5 min)
instead of the progressive shift to shallower angles.
3C; see Supplemental Movie 1 online). We interpret the observations as showing new transversely oriented microtubules
interacting with (i.e., bundling) the existing oblique microtubules
at progressively shallower angles and therefore find that the
microtubules are not sliding or rotating. Consistent with the
other starting array patterns, we observed that the microtubules
toward the midzone of the cell were initially more transversely
aligned than the polymers toward the apical and basal ends of
the cell. This gave the microtubules on the outer periclinal cell
face a transiently sigmoidal pattern (Figure 3C, 52.5 to 67.5 min).
We found that oblique arrays with a left-handed pitch reorganized
to the right, and arrays with a right-handed pitch reorganized to
the left, suggesting no structural bias in the organizing mechanism for these wild-type plants.
Cells with an initially longitudinal array organization developed
transverse microtubules first at the lateral edges of the periclinal
6 of 15
The Plant Cell
cell surface toward the center of the cell (Figure 3D). These
microtubules polymerized transversely across the outer periclinal array and occasionally created a basket or focused to form
a star-shaped array (Sambade et al., 2012) before establishing
a transverse coalignment, consistent with the slight initial increase we observed in basket shaped arrays (Figure 2A). Longitudinally oriented microtubules toward the apical and basal
cell faces persisted until the rest of the array had formed
a transverse pattern (Figure 3D, 97.5 and 135 min). Longitudinal
arrays differed from oblique arrays in that they did not typically
show a progression of oblique arrays (i.e., apparent rotation)
prior to be becoming transversely aligned. This was apparently
due to the microtubules interacting with the longitudinal polymers at an angle that was too high to redirect microtubule polymerization into a bundle.
In sum, for arrays starting in a basket, oblique, or longitudinal
pattern, no obvious changes in array pattern were observed
during the first 30 to 45 min after induction (Figure 3), corresponding to the initial lag phase found when quantifying the
percentage of cells in each array pattern over time (Figure 2). For
each starting array pattern, we consistently found that transverse coalignment initiated at the midzone of the cell 30 to 45
min after induction and progressed to the apical and basal ends
of the cell (Figures 3B to 3D). In the cases of the longitudinal and
basket patterned arrays, the longitudinally oriented polymers at
the apical and basal ends of the cell were clearly maintained
throughout the first 90 to 120 min of induction and were the last
elements of array pattern to change (Figures 3B to 3D).
Microtubule Plus Ends Reveal Structural
Properties of the Array
To investigate the underlying structural changes associated with
the microtubule array reorganization into a transverse coalignment,
we imaged cells expressing a GFP:AtEb1 probe (Figure 4) that
labels polymerizing microtubule plus ends (Mathur et al., 2003). We
imaged single hypocotyl cells every 15 min for 5-min durations
over a time course of 2.5 h. To capture the entire curved surface of
the outer periclinal cell face, we captured seven axial sections at
0.34-µm increments every 6 s and projected the information to
a single image (Figures 4A and 4C). The pattern of microtubule
array organization at each 15-min time point was evaluated by
summing all of the projected frames together from each 5-min
time-lapse subsegment (Figures 4B and 4D) to visualize the GFP:
AtEb1 tracks formed by the microtubule polymerization trajectories
(Figures 4A to 4D; see Supplemental Movies 2 to 4 online).
A visual evaluation of the time-lapse image data revealed that
the cortical microtubule arrays in all uninduced cells are structured with the majority of plus ends polymerizing toward the cell
perimeter (see Supplemental Movies 2 to 4 online). Hormoneinduced cells with transversely coaligned arrays (Figure 4D)
appeared to have lost this bias for edge-directed polymerization
in the outer periclinal arrays (see Supplemental Movies 2 to 4
online), suggesting a significant change in the structure of the
cortical array concurrent with the change in array organization.
To quantify the changes in underlying array structure over the
course of array organization to transverse, we identified all GFP:
AtEb1-labeled microtubule ends that were within 2 µm of the
perimeter of the outer periclinal cell face (Figure 4A, cell perimeter and dashed contour lines) over sequential 30-s time intervals for each 5-min data set (seven intervals per 5-min time
lapse; see Methods) at each 15-min interval. Using the timelapse data, we marked each plus end as polymerizing outward
from the periclinal cell surface to the anticlinal side faces, inward
from the anticlinal cell faces to the outer periclinal array (Figure
4A, In or Out), or parallel to the adjacent cell face. The extracted
data were plotted as the average number of plus ends (mean of
seven 30-s intervals per 5-min time lapse) moving either in or out
of the periclinal array per 1 (lateral) µm of cell perimeter at each
15-min time point (Figures 4E and 4F). Since the timing of reorganization differed slightly between different cells with different starting patterns (Figure 2), we present examples of the
aggregate numerical averages of the plus-end densities and
directions at the cell perimeter plotted for a single cell (Figures
4G to 4J) and as the average of multiple cells (Figures 4K to 4M).
Consistent with visual inspection of the time-lapse data from
untreated control cells, our quantitative analysis revealed that 70
to 75% of the plus ends at the cell periphery were polymerizing
out of the periclinal cell face with <20% of the plus ends coming
into the periclinal face from the anticlinal side faces (Figures 4E,
4H, 4L, and 4M). The balance of microtubule ends (5 to 10%)
were moving parallel to a perimeter side face and were counted
for total density. The bias in polymerization trajectories was
observed for all array pattern classes, including cells that were
initially transverse. Microtubules polymerizing into the periclinal
cell face were found in a few concentrated regions around the
cell perimeter and rarely originated from the cell’s apical or basal
ends (Figure 4E). We observed that the apical and basal ends of
the cell maintained a relatively high density of polymerizing plus
ends when compared with the side faces (Figures 4E, shaded
regions, 4G, 4K, and 4M). These data show a substantial
asymmetry (;3:1) in the number of microtubule plus ends polymerizing out of the outer cell face relative to those entering the
outer cell face, and, related to constructing models for array
organization, we find relatively few microtubules originating from
the apical and basal regions of the cell in untreated control
plants, independent of the array pattern.
The addition of GA4/IAA to the GFP:Eb1-expressing plants
resulted in reorganization of the array to a transverse coalignment (Figure 4D). We found that the density of microtubule plus
ends measured at the cell perimeter declined by ;30% within
the first 5 min of hormone treatment and continued to decrease
to <40% by 60 min after induction when compared with solventtreated control cells (Figure 4F). The decrease in GFP:Eb1labeled ends therefore occurred during the early lag phase of
induction and prior to any substantial pattern changes observed
with the GFP:TUA6 microtubule marker. The loss of polymerizing plus ends was most apparent at the apical and basal ends of
the cell (Figure 4F, shaded regions), mostly due to the continued
absence of microtubules entering the outer periclinal cell face
from these positions. However, we observed that cells retained
the longitudinally aligned polymers, likely to be microtubule
bundles, at the apical and basal portions of the cell until the
remainder of the cell became transverse.
While the total density of GFP:Eb1-labeled plus ends measured at the cell periphery decreased, we observed that the
Figure 4. Time-Lapse Imaging of GFP:AtEb1 in Hormone-Induced Cells Organizing into a Transverse Coalignment.
8 of 15
The Plant Cell
density of plus ends polymerizing into the periclinal cell face
from the anticlinal side faces increased slightly over time with
the hormone induction (Figures 4I, 4K, and 4M). Coupled with
the loss of outward polymerizing plus ends, we found that the
relative percentage of all plus ends polymerizing into the periclinal cell face shows an average net increase from ;15 to
;35% of total density after hormone induction (Figures 4J and
4L). These quantitative data support the initial visual inspection,
suggesting that the balance of microtubule plus ends moving
into and out of the outer cortical array moves closer to a 1:1 ratio
before the array becomes transversely coaligned. We observe
that the majority of effect on the ratio of microtubules polymerizing into versus out of the array came from the reduction in
polymerizing plus ends in the periclinal array and not through an
increase in the anticlinal faces.
The microtubule plus ends polymerizing into the outer periclinal array prior to induction were typically concentrated into
clumps and rarely originated from apical or basal cell faces
(Figure 4). We found that the distribution of microtubule plus
ends moving into the outer periclinal array changed dramatically
after induction from concentrated clumps to a uniformly spaced
distribution along the lateral sides of the cell (Figure 4E). The
sites were further characterized by having microtubules polymerizing both into and out of the outer periclinal array. The total
density of microtubule plus ends at the midzone did not appear
to increase during array reorganization, but the spatial distribution of microtubules moving into the outer periclinal array had
a tendency to spread bidirectionally from the cell’s midzone with
an advancing front of transversely oriented polymers (Figure 4F).
These data suggest that it is not a significant increase in the
number or density of microtubules at the cell midzone that
correlates with the appearance of the initial transverse microtubules, but rather the origin and orientation of these polymers
relative to the cell axis. While hormones induced a redistribution
of sites around the cell perimeter where microtubules entered
the periclinal array, we found that the apical and basal ends of
the cell were still excluded (Figure 4F).
Microtubules Polymerize across Cell Face Junctions
To determine if the redistribution of microtubule plus-end trajectories was occurring due to microtubules either preferentially
initiating from, or terminating at, the edges or junctions between
cell faces, we examined the GFP:Eb1 plus-end trajectories in
the trans-facial regions (Figures 5A and 5B). Using threedimensional (3D) confocal time-lapse microscopy, we created
data sets that isolated the side faces of the cell (Figures 5C to
5E). When observed in cross section (Figure 5F), the hypocotyl
cells did not show a sharp transition or edge between the outer
periclinal and anticlinal cell faces. We therefore bracketed the
region where adjacent cells form a junction and imaged to
a position >3 µm below that position. We observed the GFP:Eb1
probe in cells 60 min after hormone induction traveling either
from the outer periclinal cell face down the anticlinal side of the
cell or from the anticlinal side face and into the outer periclinal
array. It was not possible to unambiguously track individual
GFP:Eb1-labeled microtubule ends due to the density of ends
and bleed-through of GFP:Eb1 signal from the adjacent cell.
Summing the 3D time-lapse data to visualize the GFP:Eb1 trajectory paths from the lateral face (Figure 5G) and the apical/
basal face, we found that there were no apparent barriers to
microtubule growth at the junction between adjacent cells and
no obvious increase in the density of GFP:Eb1 origins (Figures
5C to 5E; see Supplemental Movie 5 online). We observed that
many of the plus-end trajectories appear to follow the same
tracks, implying the formation of microtubule bundles extending from the periclinal surface array to the anticlinal side
face of the cell.
DISCUSSION
The formation of coaligned microtubule arrays organized into
a transverse pattern relative to the plant growth axis has long
been associated with the axial growth of plant cells (Green,
1962; Hepler and Newcomb, 1964; Cyr, 1994; Baskin, 2001;
Figure 4. (continued).
Confocal imaging of control ([A] and [B]) and hormone-induced ([C] and [D]) hypocotyl cells expressing a GFP:AtEb1 transgene marking the polymerizing microtubule plus ends. A single time point (projection of 5-µm depth) from 0 and 120 min ([A] and [C]) is paired with a summation image (sum
of 50 projected frames taken at 6-s intervals) from 0 and 120 min ([B] and [D]) to show the trajectory of the microtubule plus ends and the representative
pattern of microtubule array organization. Bars = 3 µm in (A) to (D). The density and orientation of plus-end polymerization was measured at the
perimeter of the outer periclinal array (outline and dashed contour in [A]) at 15-min intervals ([E] to [M]). A 5-min time-lapse image series was acquired at
5-s intervals every 15 min for 120 min where the position and number of GFP:AtEb1 spots entering (In), exiting (out), or moving parallel to a cell edge
(Side) was recorded within the ;3-µm space delimited by the dashed contour line. The mean number of GFP:AtEb1 spots (i.e., microtubule plus ends)
per micrometer of cell perimeter per 30-s time interval moving out of (green) or into (red) the outer periclinal array in a control (E) and an induced (F) cell
are plotted for each 15-min time point in the 2-h time-lapse series (cells in [A] to [D] depicted in [E] and [F]). The cell perimeter was made linear in (E)
and (F) with the cell’s apex and base shaded (gray). Note the spreading of incoming plus ends (red) on the lateral sides of the cell and the loss of plus
ends at the apical and basal ends for the induced cell (F). Data from a single control (E) or treated (F) cell plotted numerically to show the change in the
mean density ([G] and [I]) of plus ends polymerizing out of (green) or into (red) the outer periclinal array at the lateral sides (o) or ends (x) of the cell. The
percentage of all plus ends in each orientation through time counted per time point in a single control (H) and treated (J) cell. The mean number of plus
ends moving out of (green) or into (red) the periclinal array per 30 s 6 SD (K) aggregated from six control (solid lines) and six treated (dashed lines) cells.
The percentage of all plus ends in each orientation through time (L) aggregated from six control ([L], solid lines) and six hormone-induced ([L], dashed
lines) cells. Combined density of polymerizing plus ends (In + Out) for six control ([M], solid lines) and six treated ([M], dashed lines) at the cell’s lateral
sides (blue) and ends (red).
Transverse Microtubule Array Organization
9 of 15
Figure 5. Microtubule Polymerization Is Continuous between Periclinal and Anticlinal Hypocotyl Faces during Array Reorganization.
(A) Schematic of two adjacent hypocotyl cells viewed end on (yz projection) indicating the cross section of the imaged region of the lateral cell junction
(dashed box).
(B) Rotation and cutaway of the schematic (xz projection) indicating the projected image volume used to assess the continuity of the microtubule
polymerization trajectories in the lateral side faces of the cell.
(C) to (E) Confocal images of GFP:Eb1 taken at sequential axial (z axis) positions were combined and reprojected from the side (xz projections as in [B])
to create time-lapse images of microtubule plus-end trajectories.
(F) A rotation of the image data (yz projection as in [A]) indicating the position of the cell junction. Reference lines (dashed lines in [C] to [K]) bracket the
cell junction.
(G) A summation of 100 consecutive side projected images (xz projections as in [B]) taken at 3-s intervals indicating the track or trajectory for the
microtubule plus ends polymerizing across the cell junction.
(H) to (K) Summation images from time-lapse GFP:Eb1 data at the apical/basal cell face (schematic in [H]) for three cells at 60 min after hormone
induction indicating continuous polymerization trajectories across trans-facial boundaries ([I] to [K]).
Wasteneys and Galway, 2003; Ehrhardt and Shaw, 2006;
Hashimoto and Kato, 2006; Lloyd and Chan, 2008; Lloyd, 2011).
Several models have been proposed to explain the molecular
and biophysical factors required to achieve this special state of
microtubule array organization. A significant limitation for these
models has been the lack of time-resolved observations from
cells that are transitioning into a transverse coaligned pattern.
We developed a robust and reproducible protocol for synchronously inducing the cortical microtubules in light-grown hypocotyl cells to reorganize from their preexisting array patterns into
a transverse coalignment. The induction method, combining two
plant hormones previously shown to affect microtubule array
organization (Shibaoka, 1974, 1993; Ishida and Katsumi, 1991;
Lloyd et al., 1995; Shibaoka and Takesue, 1999), permits a direct
quantitative comparison of treated and untreated plants under
identical circumstances and at a temporal and spatial scale
commensurate with microtubule turnover in the array. Using this
approach, we evaluated the changes in microtubule array structure
and pattern leading to transverse microtubule coalignment.
Our data reveal a progressive change in organization occurring
in two principal phases (Figure 6). The initial response (Figures 6A
and 6B) to hormones was a significant decrease in the number of
GFP:Eb1-labeled microtubule plus ends occurring prior to any
substantive change in the array pattern. A second phase (Figures
6C to 6E) was characterized by the appearance of transversely
oriented microtubules around the midzone of the cell where
transverse organization proceeded bidirectionally over a 30- to 60min time course toward the apical and basal regions of the cell.
The repatterning to transverse coalignment was marked by a notable transformation of the outer periclinal microtubule array
structure. Specifically, we found that control cells had approximately three microtubules polymerizing toward the perimeter of
that cell face for every one microtubule polymerizing toward the
interior of the outer periclinal array. This remarkable bias in microtubule polymerization trajectory was progressively corrected
after induction as the array reorganized from the cell’s midzone to
become transverse (Figure 6B).
We propose from our observations that the well-studied
microtubule arrays on the outer periclinal cell face of these
light-grown hypocotyl cells create patterns somewhat independently from the arrays on the other cell faces (Chan
et al., 2011). We speculate that these array patterns contribute
to cell wall construction that is required for radial hypocotyl
thickening (Chan et al., 2007, 2010; Lloyd, 2011). Hormone
induction altered the array structure from a system having 70
to 80% of the growing plus ends exiting the outer periclinal
10 of 15
The Plant Cell
Figure 6. Depiction of Hormone-Induced Cortical Array Reorganization to a Transverse Pattern.
Hypothetical hypocotyl cell with cortical microtubule array displayed (black lines) with polymerizing plus ends (arrowheads).
(A) Initial array organization with 3:1 ratio of outward (red arrowheads) to inward (green arrowheads) polymerizing plus ends.
(B) Hormone treatment induces a change in array structure observed as a decrease in the number of microtubule ends (mottled surface) on the outer
periclinal cell face prior to significant changes to array pattern.
(C) to (E) Transverse microtubules initially appear at the cell midzone ([C]; yellow surface) and organize progressively ([D] and [E]) from the midzone to
the apex and base of the cell.
array, independent of array pattern, to a system with more
balanced numbers of microtubules entering and exiting the
outer periclinal array at the cell’s lateral side faces. We infer
from our data that creation of a transverse coaligned microtubule array that is functional for axial cell growth requires
coordination across multiple cell faces. We speculate that
these globally coordinated transverse microtubule arrays are
critical for producing a transverse cellulose microfibril alignment around the cell for axial growth (Baskin, 2001, 2005;
Ehrhardt and Shaw, 2006; Lloyd, 2011).
Mechanistic Models for Transverse
Microtubule Coalignment
Earlier observations of microtubules in hormone-treated cells,
using immunofluorescence or microinjected fluorescent tubulins, proposed that array organization occurred through movements of existing polymers through intermediate patterns rather
than through direct polymerization into a new pattern (Roberts
et al., 1985; Yuan et al., 1995; Wymer and Lloyd, 1996; Wymer
et al., 1996). More recent models for transverse array patterning,
drawing from observations in live cells, propose that organization is driven by preferential loss of longitudinally oriented microtubules at the cell’s apical and basal ends (Ehrhardt and
Shaw, 2006; Baulin et al., 2007; Allard et al., 2010a, 2010b; Eren
et al., 2010; Ambrose et al., 2011; Sambade et al., 2012). These
models propose that if microtubules at the cell’s apical and
basal ends are disallowed from entering the outer periclinal array
or are selectively destroyed through preferential catastrophes at
cell edges (Ambrose et al., 2011), then the remaining microtubules will passively form a transverse coalignment. The degree
of microtubule coalignment for the remaining microtubules is
hypothesized to improve through the action of angle-dependent
bundling (Allard et al., 2010b; Eren et al., 2010), intrabundle
versus branched microtubule nucleation (Nakamura et al., 2010;
Kirik et al., 2012), or through an increase in the number of microtubules entering from the cell’s lateral side faces (Sambade
et al., 2012). Computer simulations have been developed to show
the plausibility of these mechanisms for organizing an existing
microtubule population into a transverse coalignment (Baulin
et al., 2007; Allard et al., 2010b; Eren et al., 2010; Ambrose et al.,
2011; Sambade et al., 2012).
Our repeated observations of hormone-induced microtubule
array transitions from a variety of initial patterns to a transverse
coalignment shed significant light on what mechanisms create
a global transverse pattern. We observed a progressive change
in the array pattern, somewhat consistent with previous findings
from GA4- or IAA-treated cells (Roberts et al., 1985; Ishida and
Katsumi, 1991; Lloyd et al., 1995; Yuan et al., 1995; Wymer and
Lloyd, 1996; Wymer et al., 1996; Shibaoka and Takesue, 1999).
However, we demonstrate that the reorganization does not occur through reshuffling or sliding of existing microtubules. For
example, we clearly show that the previously posited rotation of
cortical array microtubules (Roberts et al., 1985; Yuan et al.,
1995; Wymer and Lloyd, 1996; Wymer et al., 1996) corresponds
to a situation where newly appearing transverse microtubules
are interacting with an initially oblique array to form microtubule
bundles of increasingly shallow transverse orientation. The relatively long time period (;60 min) required for transverse patterning, relative to polymer turnover (Shaw et al., 2003; Shaw
and Lucas, 2011), indicates that the transitional patterns observed after induction likely arise through creation of new
microtubules and not through reorganization of the existing
polymers.
Our observations in hypocotyl cells do not support current
models for transverse patterning that rely substantively on the
targeted destruction of longitudinal microtubules at the cell’s
apical and basal ends. Our naive expectation was that if the
microtubules were preferentially removed at the apical and basal
ends of the cell, hormone treatment would induce a loss of
longitudinal microtubules at the cell’s ends, leading to a transverse reorganization initiating from the ends of the cell. Contrary
to this expectation, our data show that the longitudinal microtubules at the cell’s apex and base were consistently the last
Transverse Microtubule Array Organization
elements of the original array pattern to disappear (Figures 6D
and 6E). Furthermore, we consistently observed that transverse
array organization initiated at the midzone of the cell and
progressed to the apex and base, independent of the original
array pattern (Figures 6C to 6E). These observations were most
clearly observed for the ;20% of cells that started with
a predominantly longitudinal microtubule coalignment. New
microtubules emerging from the lateral anticlinal sides of the
cell invaded the central midzone portion of the outer periclinal
array before any changes were observed in the pattern of
longitudinal microtubules running the length of the cell (Figures
3 and 6C to 6E). Based on these observations, we conclude
that transverse array organization is driven primarily by an
active process that creates new transverse microtubules rather
than by passive retention of transverse microtubules in cells
where the longitudinal microtubules are directly targeted for
destruction (Figure 6).
Our findings suggest that the apical and basal ends of these
axially extended hypocotyl cells have distinct properties for
creating or maintaining microtubules. We found relatively few
GFP:Eb1-labeled ends entering the outer periclinal array from
the apical and basal cell faces. Critically, this phenomenon was
observed in both the control cells and in hormone-induced cells.
This suggests that the apical and basal cell faces, at this developmental stage, are not contributing significantly to the
population of longitudinal microtubules in the outer periclinal
array. Nearly all of the longitudinally oriented microtubules toward the apex and base of the outer periclinal cell face were
polymerizing toward the cell edges. We infer from our quantitative observations that the majority of these longitudinal
microtubules must then be originating from positions on the
outer periclinal cell face. Given our observation that hormone
induction rapidly depleted the total number of plus ends on the
outer periclinal cell face, we speculate that the cell is ultimately
cutting off the supply of these longitudinal microtubules rather
than targeting the existing polymers for destruction. We found
that the reorganization to transverse was marked by the progressive elimination of the edge-ward polymerization bias along
the lateral sides of the cell (Figures 6C to 6E). We infer that the
apical and basal cell faces have important properties because
this progressive correction stops and does not extend into those
cell faces to produce microtubules that polymerize into the outer
periclinal array.
Our discovery that the cell progressively organizes from the
midzone to the ends of the cell, regardless of initial array pattern,
suggests a mechanism that is spatially regulated by the cell. We
therefore propose that a global transverse coalignment does not
arise strictly due to self-organizing properties imposed globally
on the microtubules. We suspect that changing some of the
known microtubule behaviors, such as branched nucleation or
severing at microtubule crossover positions, should have important consequences for creating or maintaining an array pattern. But we propose that some of these phenomena are likely to
be deployed in a spatially distinct manner to affect transverse
array patterning and not simply changed on a global scale. How
different microtubule-related activities are spatially controlled,
and precisely which activities are required for transverse patterning, remain to be determined.
11 of 15
Working Model for Transverse Array Organization
Keeping in mind that exogenously applied hormone is not
a naturally occurring event for the plant, we propose that the
temporal sequence of events leading to transverse coalignment
shows us one pathway among several possible pathways to the
transverse coaligned state. Comparing the induced transverse
arrays to the four general pattern classes in the untreated cells,
we point to the distribution of positions along the cell’s lateral
sides where microtubules are polymerizing both into and out of
the outer periclinal array as the key difference in array structure
(Figure 6). Our quantitative evaluation of GFP:Eb1-labeled
polymer ends suggests that this change in array structure arises
not through a significant rise in the total number of plus ends
emerging from the side faces, but more directly from their redistribution along the lateral cell faces. Therefore, our data are in
partial agreement with conclusions drawn from prior work suggesting that new microtubules coming from the side faces of the
cells would help drive transverse coalignment (Sambade et al.,
2012). That work looked almost exclusively at arrays that began
in a longitudinal coalignment and concluded that all arrays form
an obligate star pattern prior to cell growth. We interpret our
data, examining a full spectrum of starting array patterns, as
indicating that all cells initiate transverse patterning toward the
midzone of the cell before that patterning progresses to the
cell’s ends. In the specific case of cells with initially longitudinal
arrays (;20% of population), the early part of the reorganization
at the midzone likely corresponds to the previously characterized star pattern. However, in the more general case of cells
that do not start with a longitudinal array (;80% of population),
those cells do not reorganize into a star or basket pattern as
an obligate intermediate step toward becoming transverse.
The new microtubules creating the transverse pattern interact
with the existing array pattern and eventually dominate the
organization.
We speculate that the hormone-induced reorganization of
arrays into a transverse coalignment is primarily driven by a redistribution of microtubule nucleation sites. The 3:1 bias in
microtubules polymerizing edge-ward in the outer periclinal
arrays strongly suggests that microtubule nucleations are more
prominent on this cell face than on the anticlinal cell faces in
untreated cells. This proposal is consistent with, and may possibly explain, prior microtubule dynamics measurements indicating more total polymerization (;2:1) than depolymerization
(Shaw et al., 2003; Shaw and Lucas, 2011). A strong bias for
microtubule nucleation in the outer periclinal cell face, with
concomitant microtubule extinction of the treadmilling polymers
more prominent on other cell faces, could explain both the observed asymmetry in plus-end trajectories and the apparent
non-steady state microtubule dynamics measurements. We
note that this positive drift coefficient (Maly, 2002), and nucleation rates in general, are dealt with somewhat arbitrarily in
current computer simulations for array organization (Baulin et al.,
2007; Allard et al., 2010b; Eren et al., 2010; Hawkins et al., 2010;
Tindemans et al., 2010; Deinum et al., 2011; Sambade et al.,
2012).
Alternatively, the bias for edge-ward polymerization could be
due to cell edge effects (Ambrose et al., 2011; Zhang et al.,
12 of 15
The Plant Cell
2011), such as an upregulation of catastrophe or microtubule
severing at the perimeter of these untreated cells (Burk et al.,
2001; Wasteneys and Ambrose, 2009; Nakamura et al., 2010). If
microtubules underwent a cycle of preferential catastrophe or
severing at the edge of the periclinal/anticlinal cell face, followed
by a high probability of rescue to polymerization at points within
the periclinal array, we would expect to count more apparent
plus ends polymerizing toward the cell edge than coming from
the cell edge. While our 3D time-lapse observations of the GFP:
EB1 probe at the cell’s side and end faces revealed no visible
interruption of plus-end polymerization trajectory at cell edges,
we cannot discount the possibility of an increased catastrophe
frequency within the anticlinal side faces of the cell (Oda and
Fukuda, 2012).
In summary, we propose that transverse coalignment of the
microtubules occurs in these cells primarily owing to an influx
of transversely oriented microtubules from the lateral side faces
of the cell initiating at the cell midzone and progressing bidirectionally to the apex and base (Figure 6). How the transverse
polymerization trajectory is set for these microtubules is not
known, but we speculate that they are following the relatively
transverse orientation of the anticlinal side facing arrays (Crowell
et al., 2011) and eventually bridge across the outer periclinal
array to form transverse antiparallel bundles. We propose that
an increase or redistribution of the nucleation sites for these
polymers, at the expense of microtubules in all other orientations
(i.e., not just transverse) originating on the outer periclinal cell
face, promotes the antiparallel microtubule bundles transverse
to the cell’s axis (Figures 6C to 6E). We speculate that the
progression of transverse array organization from the middle of
the cell to the ends may be due to a gradient in signaling for
microtubule nucleation in the lateral anticlinal cell faces and
influenced by a lag in the disassembly of the longitudinal polymer bundles in the apical and basal regions of the cell.
with 3;0.5 mL of medium and sealed between slide and cover glass using
silicon vacuum grease (Dow Corning) to form a chamber.
Confocal Microscopy
METHODS
Hypocotyl cells were imaged using a Leica SP5 confocal laser scanning
microscope with either a 340 1.2–numerical aperture (NA) or a 363 1.2NA water emersion objective lens. Confocal volume imaging of the apical
and basal side faces of the cell was accomplished with a Nikon A1
confocal laser scanning microscope using a 340 1.2-NA water immersion
lens. The GFP-based probes were excited using a 488-nm laser line, and
excitation was collected using the spectrophotometric detector.
To capture array organization in whole hypocotyls, the hypocotyls
were imaged in sections (typically four) beginning with the root/hypocotyl
junction and progressing apically to the nascent meristem. Optical
sections for reconstructed image projections were taken at 0.25- to
0.34-mm steps. Two-dimensional projections of GFP:TUA6 data sets
were created by converting Leica files into tiff format (Imaris; Bitplane) and
subsequently creating a summation or standard deviation projection
using ImageJ (NIH, Wayne Rasband). Photoshop (Adobe Software) was
then used to linearly scale the contrast for presentation. Images showing
the entire hypocotyl were composited by hand in Illustrator (Adobe
Software).
To determine the number and direction of polymerizing microtubule
plus ends at time intervals during array organization, Eb1:GFP in control
and GA4 and IAA cotreated seedlings were imaged. Data were collected at
15-min intervals starting at the time when plants were placed into the
liquid inducing media on the microscope slide and terminating 2.5 h later.
Plants saw exposure to hormone for no more than 3 min before the first
time point. The seedlings were continuously imaged for the first 5 min of
every 15-min interval. A series of seven frames, taken at 0.5-µm steps
along the microscope’s optical axis (upper 3 to 4 µm of the cell), was taken
every 6 s to produce the 3D time-lapse data sets.
For evaluation of microtubule ends polymerizing in the lateral side and
end faces of the cell, we used the Leica confocal laser scanning microscope to collect 2- to 5-min duration time-lapse movies of 6-d-old
GFP:Eb1-expressing plants at 3.1- to 5.9-s intervals. A narrow XY field of
view was used to capture only the side or end face of interest from the
outer to the inner periclinal cell faces. Images were taken every 0.2 µm to
a minimum depth of 10 µm from the top surface of the cell.
Plant Materials
Data Analysis
Seed was sterilized in 100% ethanol for 10 s before planting on 0.3% agar
(Sigma-Aldrich) plates containing half-strength MS medium and no sugar
(Sigma-Aldrich). We exposed the plated seed to 2 h of light, wrapped the
plates in foil, and stored them for 2 d at 4°C to synchronize germination.
The foil was removed and the plates were incubated vertically at 22°C
under continuous light for 6 d prior to imaging. Wild-type Arabidopsis
thaliana (Columbia-0) plants expressing a 35S-GFP:TUA6 transgene were
used for visualizing microtubules and a 35S-Eb1:GFP expressing line for
visualizing polymerizing microtubule plus ends (Mathur et al., 2003).
For assaying the percentage of cells with transverse cortical arrays, the
two-dimensional projections were given to two individuals for single blind
evaluation using the same classing key (Figure 1) for scoring array types.
Array organization was scored in two adjacent cell files along the entire
hypocotyl length (;28.0 6 3.1 cells) to avoid any bias in organization for
cell file. Counts for each array class from different counters were averaged
for final reporting.
A series of MATLAB (The Mathworks) scripts was developed and
implemented as a graphical user interface for assessing the GFP:Eb1
trajectories and for computing summary statistics. Prior to analysis, each
of the 3D data sets was projected to a single image creating 50 or more
time-lapse image frames at 6-s intervals for each 5-min image series.
Each 50-frame data set of the nine data sets comprising the 2-h time lapse
was used separately for analysis. Our first goal was to measure the
average number of microtubule plus ends polymerizing into or out of the
outer periclinal array during each 15-min interval. Our second goal was to
relate any change in number or ratio of trajectories to the spatial position
on the cell perimeter. To accomplish these goals, a line was traced around
the perimeter of the cell using the computer pointing device and a contour
was automatically generated inside the cell creating a zone where GFP:
Eb1 spots were counted one time. Each 5-min time-lapse sequence was
Hormone Treatments
Seedlings were taken from agar germination plates after 6 d at 22°C and
placed into liquid half-strength MS medium for treatment. A 10 mM GA4
(Sigma-Aldrich) stock dissolved in 1:1 (v/v) of ethanol and distilled water
was made and plants were treated with a 10 mM final concentration. A 0.1
mM IAA (Sigma-Aldrich) stock dissolved in 100% DMSO (Sigma-Aldrich)
was made and plants were treated with a 0.5 mM final concentration. For
observations made at 2 h, plants were placed in 5 mL of liquid and moved
to a cover glass with the same medium immediately prior to imaging. For
time-lapse experiments, plants were positioned onto a glass slide
Transverse Microtubule Array Organization
broken into seven sequential time periods with five images per period.
Using the computer pointing device, each GFP:Eb1 spot in the perimeter
delimited zone was marked only once by hand, and the trajectory of the
spot was recorded as either moving “in” to the periclinal array, “out” of the
periclinal array, or “sideways” relative to the cell edge. The trajectory
assignment was made by looping the five images in the time period to
visualize polymerization direction. The width of the counting zone contour
was set to ;3 µm such that GFP:Eb1 spots were not double counted
across the seven time periods. Summary statistics for each 5-min time
period were therefore calculated as the average number of microtubule
plus ends per 1-µm linear distance of cell perimeter that were traversing
the cell perimeter per 30 s with a number of seven time periods per 5 min.
To validate the data transcription, one control and one experimental cell
were counted for all nine time points by two different individuals, where
the resulting differences in the number and direction of GFP:Eb1 spots
were found to be negligible.
For determining the spatial distributions of each spot, the marked GFP:
Eb1 position was projected onto the hand-drawn perimeter line for each
cell and the measured pixels/micrometer value was used to convert and
scale the data. The apical and basal regions of each cell were marked by
hand in the MATLAB graphical user interface from known orientations of
the cells in the images.
For side and end face dynamics, the data were projected side-on (xz or
yz projections) using 2.1- to 2.5-µm thick slices of the 3D time-lapse data
set to evaluate the GFP:Eb1 trajectory. For examining tracks, data sets
from sequential 3D projections were summed to a single image (Imaris).
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Movie 1. Short Interval Imaging of an Oblique Array
Transitioning to Transverse.
Supplemental Movie 2. Microtubule Plus-End Trajectories Indicate
Dramatic Changes to the Array Architecture after Hormone Induction.
Supplemental Movie 3. Microtubule Plus-End Trajectories Indicate
Dramatic Changes to the Array Architecture after Hormone Induction.
Supplemental Movie 4. Microtubule Plus-End Trajectories Indicate
Dramatic Changes to the Array Architecture after Hormone Induction.
Supplemental Movie 5. Microtubules Polymerize Continuously between the Periclinal and Anticlinal Cell Faces.
ACKNOWLEDGMENTS
We thank Roger Hangarter for advice regarding the hormone treatments
and growth conditions and Jim Powers at the Indiana University Light
Microscopy Imaging Center for help with the microscopy. We acknowledge support for this work from the National Science Foundation (MCB0920555 and MCB-1157982 to S.L.S.).
AUTHOR CONTRIBUTIONS
L.V. designed and executed the majority of experiments. J.R.L.
performed the original hormone induction experiments. A.E. measured
and correlated the array orientation assays. S.D. performed the hypocotyl length assays. S.L.S. developed the project and wrote the article.
Received November 10, 2012; revised January 31, 2013; accepted
February 8, 2013; published February 26, 2013.
13 of 15
REFERENCES
Allard, J.F., Ambrose, J.C., Wasteneys, G.O., and Cytrynbaum,
E.N. (2010a). A mechanochemical model explains interactions between cortical microtubules in plants. Biophys. J. 99: 1082–1090.
Allard, J.F., Wasteneys, G.O., and Cytrynbaum, E.N. (2010b).
Mechanisms of self-organization of cortical microtubules in plants
revealed by computational simulations. Mol. Biol. Cell 21: 278–286.
Ambrose, C., Allard, J.F., Cytrynbaum, E.N., and Wasteneys, G.O.
(2011). A CLASP-modulated cell edge barrier mechanism drives
cell-wide cortical microtubule organization in Arabidopsis. Nat. Commun.
2: 430.
Baskin, T.I. (2001). On the alignment of cellulose microfibrils by cortical
microtubules: A review and a model. Protoplasma 215: 150–171.
Baskin, T.I. (2005). Anisotropic expansion of the plant cell wall. Annu.
Rev. Cell Dev. Biol. 21: 203–222.
Baulin, V.A., Marques, C.M., and Thalmann, F. (2007). Collision induced spatial organization of microtubules. Biophys. Chem. 128:
231–244.
Bornens, M. (2002). Centrosome composition and microtubule anchoring mechanisms. Curr. Opin. Cell Biol. 14: 25–34.
Burk, D.H., Liu, B., Zhong, R., Morrison, W.H., and Ye, Z.H. (2001).
A katanin-like protein regulates normal cell wall biosynthesis and
cell elongation. Plant Cell 13: 807–827.
Chan, J., Calder, G., Fox, S., and Lloyd, C. (2007). Cortical microtubule arrays undergo rotary movements in Arabidopsis hypocotyl
epidermal cells. Nat. Cell Biol. 9: 171–175.
Chan, J., Crowell, E., Eder, M., Calder, G., Bunnewell, S., Findlay,
K., Vernhettes, S., Höfte, H., and Lloyd, C. (2010). The rotation of
cellulose synthase trajectories is microtubule dependent and influences the texture of epidermal cell walls in Arabidopsis hypocotyls.
J. Cell Sci. 123: 3490–3495.
Chan, J., Eder, M., Crowell, E.F., Hampson, J., Calder, G., and
Lloyd, C. (2011). Microtubules and CESA tracks at the inner epidermal wall align independently of those on the outer wall of lightgrown Arabidopsis hypocotyls. J. Cell Sci. 124: 1088–1094.
Crowell, E.F., Timpano, H., Desprez, T., Franssen-Verheijen, T.,
Emons, A.-M., Höfte, H., and Vernhettes, S. (2011). Differential Regulation of Cellulose Orientation at the Inner and Outer Face of Epidermal
Cells in the Arabidopsis Hypocotyl. Plant Cell 23: 2592–2605.
Cyr, R.J. (1994). Microtubules in plant morphogenesis: Role of the
cortical array. Annu. Rev. Cell Biol. 10: 153–180.
Cyr, R.J., and Palevitz, B.A. (1995). Organization of cortical microtubules in plant cells. Curr. Opin. Cell Biol. 7: 65–71.
Deinum, E.E., Tindemans, S.H., and Mulder, B.M. (2011). Taking
directions: The role of microtubule-bound nucleation in the selforganization of the plant cortical array. Phys. Biol. 8: 056002.
Dixit, R., and Cyr, R. (2004a). The cortical microtubule array: From
dynamics to organization. Plant Cell 16: 2546–2552.
Dixit, R., and Cyr, R. (2004b). Encounters between dynamic cortical
microtubules promote ordering of the cortical array through angledependent modifications of microtubule behavior. Plant Cell 16:
3274–3284.
Doxsey, S. (2001). Re-evaluating centrosome function. Nat. Rev. Mol.
Cell Biol. 2: 688–698.
Ehrhardt, D.W. (2008). Straighten up and fly right: Microtubule dynamics and organization of non-centrosomal arrays in higher plants.
Curr. Opin. Cell Biol. 20: 107–116.
Ehrhardt, D.W., and Shaw, S.L. (2006). Microtubule dynamics and organization in the plant cortical array. Annu. Rev. Plant Biol. 57: 859–875.
Eren, E.C., Dixit, R., and Gautam, N. (2010). A three-dimensional computer simulation model reveals the mechanisms for self-organization
of plant cortical microtubules into oblique arrays. Mol. Biol. Cell 21:
2674–2684.
14 of 15
The Plant Cell
Folta, K.M., Pontin, M.A., Karlin-Neumann, G., Bottini, R., and
Spalding, E.P. (2003). Genomic and physiological studies of early
cryptochrome 1 action demonstrate roles for auxin and gibberellin in the
control of hypocotyl growth by blue light. Plant J. 36: 203–214.
Gendreau, E., Traas, J., Desnos, T., Grandjean, O., Caboche, M.,
and Höfte, H. (1997). Cellular basis of hypocotyl growth in Arabidopsis thaliana. Plant Physiol. 114: 295–305.
Giddings, T.H., Jr., Brower, D.L., and Staehelin, L.A. (1980). Visualization of particle complexes in the plasma membrane of Micrasterias denticulata associated with the formation of cellulose
fibrils in primary and secondary cell walls. J. Cell Biol. 84: 327–339.
Green, P.B. (1962). Mechanism for plant cell morphogensis. Science
138: 1404–1405.
Hardham, A.R., and Gunning, B.E. (1978). Structure of cortical microtubule arrays in plant cells. J. Cell Biol. 77: 14–34.
Hashimoto, T., and Kato, T. (2006). Cortical control of plant microtubules. Curr. Opin. Plant Biol. 9: 5–11.
Hawkins, R.J., Tindemans, S.H., and Mulder, B.M. (2010). Model for
the orientational ordering of the plant microtubule cortical array.
Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 82: 011911.
Hepler, P.K., and Newcomb, E.H. (1964). Microtubules and fibrils in
cytoplasm of coleus cells undergoing secondary wall deposition. J.
Cell Biol. 20: 529–532.
Howard, J., and Hyman, A.A. (2003). Dynamics and mechanics of the
microtubule plus end. Nature 422: 753–758.
Huang, R.F., and Lloyd, C.W. (1999). Gibberellic acid stabilises microtubules in maize suspension cells to cold and stimulates acetylation of alpha-tubulin. FEBS Lett. 443: 317–320.
Ishida, K., and Katsumi, M. (1991). Immunofluorescence microscopical observation of cortical microtubule arrangement by gibberellin in d5 mutant of Zea mays. Plant Cell Physiol. 32: 409–417.
Kirik, A., Ehrhardt, D.W., and Kirik, V. (2012). TONNEAU2/FASS regulates the geometry of microtubule nucleation and cortical array organization in interphase Arabidopsis cells. Plant Cell 24: 1158–1170.
Le, J., Vandenbussche, F., De Cnodder, T., Van Der Straeten, D.,
and Verbelen, J.P. (2005). Cell elongation and microtubule behavior in the Arabidopsis hypocotyl: Responses to ethylene and
auxin. J. Plant Growth Regul. 24: 166–178.
Ledbetter, M.C. (1982). The role of microtubules in plant cell wall
growth. Recent Adv. Phytochem. 16: 125–150.
Ledbetter, M.C., and Porter, K.R. (1964). Morphology of microtubules of plant cell. Science 144: 872–874.
Liu, B., Joshi, H.C., Wilson, T.J., Silflow, C.D., Palevitz, B.A., and
Snustad, D.P. (1994). g-Tubulin in Arabidopsis: Gene sequence,
immunoblot, and immunofluorescence studies. Plant Cell 6: 303–314.
Liu, B., Marc, J., Joshi, H.C., and Palevitz, B.A. (1993). A gammatubulin-related protein associated with the microtubule arrays of
higher plants in a cell cycle-dependent manner. J. Cell Sci. 104:
1217–1228.
Lloyd, C. (2011). Dynamic microtubules and the texture of plant cell
walls. Int. Rev. Cell Mol. Biol. 287: 287–329.
Lloyd, C., and Chan, J. (2002). Helical microtubule arrays and spiral
growth. Plant Cell 14: 2319–2324.
Lloyd, C., and Chan, J. (2008). The parallel lives of microtubules and
cellulose microfibrils. Curr. Opin. Plant Biol. 11: 641–646.
Lloyd, C., Shaw, P.J., Warn, R.M., and Yuan, M. (1995). Gibberellicacid-induced reorientation of cortical microtubules in living plant
cells. J. Microsc. 181: 140–144.
Lloyd, C.W., and Wells, B. (1985). Microtubules are at the tips of root
hairs and form helical patterns corresponding to inner wall fibrils. J.
Cell Sci. 75: 225–238.
Lucas, J., and Shaw, S.L. (2008). Cortical microtubule arrays in the
Arabidopsis seedling. Curr. Opin. Plant Biol. 11: 94–98.
Maly, I.V. (2002). Diffusion approximation of the stochastic process of
microtubule assembly. Bull. Math. Biol. 64: 213–238.
Mathur, J., Mathur, N., Kernebeck, B., Srinivas, B.P., and
Hülskamp, M. (2003). A novel localization pattern for an EB1-like
protein links microtubule dynamics to endomembrane organization.
Curr. Biol. 13: 1991–1997.
Mazia, D. (1987). The chromosome cycle and the centrosome cycle in
the mitotic cycle. Int. Rev. Cytol. 100: 49–92.
McDonald, A.R., Liu, B., Joshi, H.C., and Palevitz, B.A. (1993). Gammatubulin is associated with a cortical-microtubule-organizing zone in the
developing guard cells of Allium cepa L. Planta 191: 357–361.
Murata, T., Sonobe, S., Baskin, T.I., Hyodo, S., Hasezawa, S.,
Nagata, T., Horio, T., and Hasebe, M. (2005). Microtubuledependent microtubule nucleation based on recruitment of gammatubulin in higher plants. Nat. Cell Biol. 7: 961–968.
Nakamura, M., Ehrhardt, D.W., and Hashimoto, T. (2010). Microtubule and katanin-dependent dynamics of microtubule nucleation
complexes in the acentrosomal Arabidopsis cortical array. Nat. Cell
Biol. 12: 1064–1070.
Nakamura, M., and Hashimoto, T. (2009). A mutation in the Arabidopsis gamma-tubulin-containing complex causes helical growth
and abnormal microtubule branching. J. Cell Sci. 122: 2208–2217.
Newcomb, E.H. (1969). Plant microtubules. Annu. Rev. Plant Physiol.
20: 253–288.
Oda, Y., and Fukuda, H. (2012). Initiation of cell wall pattern by a Rho- and
microtubule-driven symmetry breaking. Science 337: 1333–1336.
Paredez, A.R., Somerville, C.R., and Ehrhardt, D.W. (2006). Visualization of cellulose synthase demonstrates functional association
with microtubules. Science 312: 1491–1495.
Pastuglia, M., Azimzadeh, J., Goussot, M., Camilleri, C., Belcram,
K., Evrard, J.L., Schmit, A.C., Guerche, P., and Bouchez, D.
(2006). Gamma-tubulin is essential for microtubule organization and
development in Arabidopsis. Plant Cell 18: 1412–1425.
Refrégier, G., Pelletier, S., Jaillard, D., and Höfte, H. (2004). Interaction between wall deposition and cell elongation in dark-grown
hypocotyl cells in Arabidopsis. Plant Physiol. 135: 959–968.
Roberts, I.N., Lloyd, C.W., and Roberts, K. (1985). Ethylene-induced
microtubule reorientations - Mediation by helical arrays. Planta 164:
439–447.
Sambade, A., Pratap, A., Buschmann, H., Morris, R.J., and Lloyd,
C. (2012). The influence of light on microtubule dynamics and
alignment in the Arabidopsis hypocotyl. Plant Cell 24: 192–201.
Sedbrook, J.C., and Kaloriti, D. (2008). Microtubules, MAPs and
plant directional cell expansion. Trends Plant Sci. 13: 303–310.
Shaw, S.L., Kamyar, R., and Ehrhardt, D.W. (2003). Sustained microtubule treadmilling in Arabidopsis cortical arrays. Science 300: 1715–1718.
Shaw, S.L., and Lucas, J. (2011). Intrabundle microtubule dynamics
in the Arabidopsis cortical array. Cytoskeleton (Hoboken) 68: 56–67.
Shibaoka, H. (1974). Involvement of wall microtubules in gibberellin
promotion and kinetin inhibition of stem elongation. Plant Cell
Physiol. 15: 255–263.
Shibaoka, H. (1993). Regulation of gibberellins of the orientation of cortical
microtubules in plant cells. Aust. J. Plant Physiol. 20: 461–470.
Shibaoka, H. (1994). Plant hormone-induced changes in the orientation of cortical microtubules: Alterations in the cross-linking between microtubules and the plasma membrane. Annu. Rev. Plant
Physiol. 45: 527–544.
Shibaoka, H., and Takesue, K. (1999). Auxin-induced longitudinal-totransverse reorientation of cortical microtubules in nonelongating
epidermal cells of azuki bean epicotyls. Protoplasma 206: 27–30.
Takesue, K., and Shibaoka, H. (1999). Auxin-induced longitudinal-totransverse reorientation of cortical microtubules in nonelongating
epidermal cells of azuki bean epicotyls. Protoplasma 206: 27–30.
Transverse Microtubule Array Organization
Tindemans, S.H., Hawkins, R.J., and Mulder, B.M. (2010). Survival
of the aligned: Ordering of the plant cortical microtubule array.
Phys. Rev. Lett. 104: 058103.
Vandenbussche, F., Verbelen, J.P., and Van Der Straeten, D.
(2005). Of light and length: Regulation of hypocotyl growth in
Arabidopsis. Bioessays 27: 275–284.
Wasteneys, G.O., and Ambrose, J.C. (2009). Spatial organization of
plant cortical microtubules: Close encounters of the 2D kind. Trends
Cell Biol. 19: 62–71.
Wasteneys, G.O., and Galway, M.E. (2003). Remodeling the cytoskeleton for growth and form: An overview with some new views.
Annu. Rev. Plant Biol. 54: 691–722.
Wenzel, C.L., Williamson, R.E., and Wasteneys, G.O. (2000).
Gibberellin-induced changes in growth anisotropy precede gibberellindependent changes in cortical microtubule orientation in developing
epidermal cells of barley leaves. Kinematic and cytological studies on
15 of 15
a gibberellin-responsive dwarf mutant, M489. Plant Physiol. 124:
813–822.
Wightman, R., and Turner, S.R. (2007). Severing at sites of microtubule crossover contributes to microtubule alignment in cortical
arrays. Plant J. 52: 742–751.
Wymer, C., and Lloyd, C. (1996). Dynamic microtubules: Implications
for cell wall patterns. Trends Plant Sci. 1: 222–228.
Wymer, C.L., Wymer, S.A., Cosgrove, D.J., and Cyr, R.J. (1996).
Plant cell growth responds to external forces and the response
requires intact microtubules. Plant Physiol. 110: 425–430.
Yuan, M., Warn, R.M., Shaw, P.J., and Lloyd, C.W. (1995). Dynamic microtubules under the radial and outer tangential walls of microinjected pea
epidermal cells observed by computer reconstruction. Plant J. 7: 17–23.
Zhang, D., et al. (2011). Drosophila katanin is a microtubule depolymerase that regulates cortical-microtubule plus-end interactions
and cell migration. Nat. Cell Biol. 13: 361–370.
Progressive Transverse Microtubule Array Organization in Hormone-Induced Arabidopsis
Hypocotyl Cells
Laura Vineyard, Andrew Elliott, Sonia Dhingra, Jessica R. Lucas and Sidney L. Shaw
Plant Cell; originally published online February 26, 2013;
DOI 10.1105/tpc.112.107326
This information is current as of June 18, 2017
Supplemental Data
/content/suppl/2013/02/20/tpc.112.107326.DC1.html
Permissions
https://www.copyright.com/ccc/openurl.do?sid=pd_hw1532298X&issn=1532298X&WT.mc_id=pd_hw1532298X
eTOCs
Sign up for eTOCs at:
http://www.plantcell.org/cgi/alerts/ctmain
CiteTrack Alerts
Sign up for CiteTrack Alerts at:
http://www.plantcell.org/cgi/alerts/ctmain
Subscription Information
Subscription Information for The Plant Cell and Plant Physiology is available at:
http://www.aspb.org/publications/subscriptions.cfm
© American Society of Plant Biologists
ADVANCING THE SCIENCE OF PLANT BIOLOGY