Download Microtubule cortical array organization and plant cell morphogenesis

Microtubule cortical array organization and plant cell
morphogenesis
Alex Paradez1,2, Amanda Wright3 and David W Ehrhardt1
Plant cell cortical microtubule arrays attain a high degree of
order without the benefit of an organizing center such as a
centrosome. New assays for molecular behaviors in living cells
and gene discovery are yielding insight into the mechanisms by
which acentrosomal microtubule arrays are created and
organized, and how microtubule organization functions to
modify cell form by regulating cellulose deposition. Surprising
and potentially important behaviors of cortical microtubules
include nucleation from the walls of established microtubules,
and treadmilling-driven motility leading to polymer interaction,
reorientation, and microtubule bundling. These behaviors
suggest activities that can act to increase or decrease the local
level of order in the array. The SPIRAL1 (SPR1) and SPR2
microtubule-localized proteins and the radial swollen 6 (rsw-6)
locus are examples of new molecules and genes that affect
both microtubule array organization and cell growth pattern.
Functional tagging of cellulose synthase has now allowed the
dynamic relationship between cortical microtubules and the
cell-wall-synthesizing machinery to be visualized, providing
direct evidence that cortical microtubules can organize
cellulose synthase complexes and guide their movement
through the plasma membrane as they create the cell wall.
Addresses
1
Department of Plant Biology, Carnegie Institution, 260 Panama Street,
Stanford, California 94305, USA
2
Department of Biology, Stanford University, Stanford, California 94305,
USA
3
Section of Cell and Developmental Biology, University of California San
Diego, 9500 Gilman Drive, La Jolla, California 92093-0116, USA
Corresponding author: Ehrhardt, David W ([email protected])
Current Opinion in Plant Biology 2006, 9:571–578
This review comes from a themed issue on
Cell biology
Edited by Laurie G Smith and Ulrike Mayer
Available online 28th September 2006
1369-5266/$ – see front matter
# 2006 Elsevier Ltd. All rights reserved.
DOI 10.1016/j.pbi.2006.09.005
Introduction
Plant cell growth is achieved by cell wall expansion that is
driven by high internal pressure, or turgor. To acquire
specific shapes that are important for cell function and
organized multicellular development, the cell wall has to
yield to uniformly applied internal pressure in a nonwww.sciencedirect.com
uniform, or anisotropic, pattern. Plant cell morphogenesis
is influenced by both the microtubule and actin cytoskeletal networks and the signaling mechanisms that control
their organization. Interphase microtubule cortical arrays
assume a variety of configurations that vary by cell type
and shape. In cells that are destined to undergo rapid axial
elongation, such as those in the root axis or the etiolated
hypocotyl, the cortical array assumes a high degree of
order, with polymers lying roughly in parallel to each
other and oriented transversely or obliquely relative to
the cell axis [1]. By contrast, in highly lobed pavement
cells, there is no global orientation of microtubules but
rather local and periodic patches of parallel polymers that
are correlated with the sinuses of the undulating cell
perimeter [2]. It is likely that basic mechanisms for
the creation of cortical array organization apply in all
cell types, and that modifications and variations of these
mechanisms operate in cells that have specialized
shape. The molecular mechanisms by which cortical
microtubule patterns are established and maintained
are not yet known, but new insights are arriving from
a combination of genetic, biochemical, and live-cellimaging studies.
What are the functions of the plant cortical microtubule
array? In 1962, Paul Green [3] reported that colchicine, a
drug known to disrupt the fibers in mitotic spindles,
caused uniform swelling of algal cells and loss of cell wall
birefringence as measured by polarization microscopy.
He hypothesized that colchicine-sensitive fibers were
somehow responsible for organizing the direction in
which the major structural polymers in the cell wall were
deposited, the orientation of these wall fibers being the
basis of the material anisotropy responsible for the direction of cell wall expansion. A year later, Ledbetter and
Porter [4] observed the first cortical microtubules in plant
cells, noting that they lay just under the plasma membrane and were often parallel to each other, and coining
the name ‘microtubule’ because of their annular appearance in cross section. These authors and others observed
that microtubules were often parallel to fibers in the cell
wall [5,6], supporting Green’s original idea. Many studies
with both drugs and mutants have supported the microfibril guidance hypothesis (reviewed in [7]), but microtubule orientation and cellulose orientation can become
uncoupled [7,8], and cellulose microfibrils can be laid
down in a parallel fashion without an intact cortical
array [9]. These observations suggest that the function
of microtubules in cell wall organization might be
more complicated than simple one-on-one guidance of
Current Opinion in Plant Biology 2006, 9:571–578
572 Cell biology
cellulose orientation. Here, we review recent progress in
our understanding of interphase cortical microtubule
organization and the function of this array in building
the cell wall and regulating cell wall expansion pattern.
Cortical array creation and organization
Cortical microtubule nucleation
Most current evidence suggests that interphase microtubules are first polymerized then organized into the
cortical array. In the course of normal root axis development, microtubules appear at the cortex of post-mitotic
cells in random orientations before the array attains a high
degree of order. Likewise, when cortical microtubules are
depolymerized with drugs then allowed to recover, the
array is initially disorganized and gradually regains an
ordered appearance, showing that microtubules are not
polymerized into their final position [10–12]. Plant cells
lack an obvious central microtubule nucleating center,
such as a centrosome or basal body. Instead, nucleation
activity appears to be distributed in the plant cell [13–
15,16], with cortical polymers originating at multiple
sites on the cortex itself [17,18,19]. Plant microtubule
nucleation activity was recently confirmed to require
g-tubulin [16], which also appears to be distributed
widely at the cell cortex, being prevalent along the walls
of existing microtubules [13,15,16]. In a breakthrough
study, Murata et al. [19] demonstrated that new microtubules can be nucleated from sites along microtubule
walls that are marked by g-tubulin. Remarkably, these
microtubules arise at a fixed angle of about 40 degrees to
the wall of the existing polymer, both in vitro and in vivo
[19]. These observations demonstrated that new
microtubules can be created at specific angles, but in
relation to other polymers rather than the cell axis. As
microtubules have also been observed to arise within
bundles [20], it will be interesting to investigate whether
the angle of new polymerization with respect to the
subtending polymer is a point of regulation. New polymers arising at zero degrees would tend to maintain
current array structure whereas nucleation at 40 degrees
would be expected to disrupt current array organization, a
potentially useful property for making array transitions
in response to signals or for remodeling arrays that continuously change their orientation over the course of
cell growth.
Polymer treadmilling, bundling and self organization
If microtubules are not generally polymerized into an
ordered array [21–24], how is interphase organization
created? The prevalent alternative theories have been
that microtubules of diverse orientations and positions are
selectively stabilized or that polymers are moved by
motor activities from one location and orientation to
another [6,21]. Live cell-imaging studies have effectively
ruled out the latter hypothesis, provided support for
selective stabilization in some cells, and have revealed
a third possible mechanism for array remodeling.
Current Opinion in Plant Biology 2006, 9:571–578
Once initiated, many cortical microtubules do not remain
attached to their presumed sites of nucleation, instead
they are often observed to detach from these locations
and to move across the cell cortex [17]. Photobleaching
experiments showed that this migration is the result of a
hybrid polymer treadmilling mechanism. The leading
end of the microtubule alternates between episodes of
growth, shrinking and pause, a pattern called dynamic
instability, with subunit gain being greater over time than
subunit loss. The lagging end of the microtubule mainly
shows episodes of shrinking and pause [17]. The net
result is apparent translocation of the microtubule, not
by motor activity but by biased polymerization activity.
These treadmilling polymers appear be tightly associated
with the cell membrane, as judged by a lack of lateral
translocation in cells that have rapidly streaming cytosol
[17]. Together, these results suggest that the translocation of microtubules by motor activity is not a significant
mechanism in cortical array organization [25].
Microtubule attachment to the plasma membrane
[17,18,26,27], possibly mediated by a phospholipase
D-dependent mechanism [28], confines migrating microtubules to a two-dimensional space where growing plus
ends can encounter and interact with other microtubules
[25]. Dixit and Cyr [29] showed that one consequence of
these interactions in tobacco tissue culture cells is that
when growing microtubule ends encounter other polymers at steep angles, complete microtubule depolymerization (i.e. catastrophe) often results. These authors
predicted that, over time, this behavior selects against
polymers of discordant orientation, producing a more
uniformly aligned array. Similar behavior is not obvious
in the microtubule arrays of Arabidopsis hypocotyl cells
(SL Shaw, DW Ehrhardt, unpublished), but interactionsensitive catastrophe might play a role in the arrays of
other cell types in Arabidopsis.
Microtubule interactions that are driven by treadmilling
result in a second important outcome: reorientation of
polymer growth and apparent bundling with the encountered polymer [17,29]. Interestingly, this bundling interaction shows a strong dependence on the angle of polymer
encounter, with bundling being very efficient at angles
below 30 degrees and rare at angles steeper than 40
degrees ([29]; SL Shaw, DW Ehrhardt, unpublished).
Once assimilated into a bundle, single polymers tend
to remain associated with that bundle [17,25], and thus
bundles seem to act as positional and orientational traps.
Together, polymer migration and angle-dependent interaction that leads to bundling have properties of a selforganizing behavior that might increase order in the array
at a local level [25,29]. Candidates for proteins that might
participate in or facilitate bundling interactions include
proteins that reside at the growing polymer end, such as
SPIRAL1 (SPR1) [30,31] and the Arabidopsis homologs of
the plus-end-associated proteins EB1/BIM1 [18,32].
www.sciencedirect.com
Microtubule cortical array organization and plant cell morphogenesis Paradez, Wright and Ehrhardt 573
Additional candidates are proteins that form cross bridges
between polymer walls, such as members of the microtubule-associated family MAP65, which have been shown
to promote bundling of microtubules in vitro and in vivo
[20,33–36,37,38].
variable and independent orientations from cell to
cell [45]. Identification of the product of rsw-6 should
provide an interesting part of the microtubule cortical
array story.
Local control of cortical array organization
Orientation of cortical microtubule arrays
Although potentially important, the significance of treadmilling-mediated bundling in the creation and maintenance of cortical array organization remains to be
determined. On their own, these activities cannot account
for how the net orientation of the cortical microtubule
array is selected. Other inputs must feed into the control
of array orientation, which appears to be a dynamic
process in many cells. Array orientation is known to
change steadily along a developmental gradient in the
root axis [7]. Moreover, recent studies in sunflower [39]
suggest that cortical arrays in hypocotyls cells undergo
continuous rotation or oscillation, patterns also recently
observed by Clive Lloyd and colleagues in observations of
live Arabidopsis hypocotyl cells (C Lloyd, pers. comm.).
Cortical array orientation is also well known to respond to
extrinsic cues such as light and hormones [7].
Insight into the molecular mechanisms that control the
orientation of cortical microtubule arrays has begun to
emerge from genetic studies. Mutations in the plantspecific plus-end-localized protein SPR1 [30,31,40],
and in another novel protein SPR2 [40,41,42], cause
cortical array orientation in root cells to become pitched
in a left-hand helix [41]. By contrast, suppressing mutations at the a–b dimer interface of a-tubulin drives the
orientation of the same arrays in the opposite direction,
toward a right-handed helix [43]. Likewise, modification
of a-tubulin at its carboxyl terminus with either green
fluorescent protein (GFP) or a short epitope tag, or
mutation of residues in a-tubulin that are hypothesized
to promote GTP hydrolysis, caused arrays to pitch to the
left [44]. These studies suggest that there is a dynamic
balance between mechanisms that drive cortical arrays to
either a left- or right-handed pitch [41]. Do these mutations affect array formation primarily by acting on microtubule dynamics, thus affecting the outcome of possible
self-organizing mechanisms, or do they modify the ability
of microtubules to perceive or respond to cellular cues
that direct array orientation? Analysis of how some of
these mutations affect basic microtubules properties,
such as polymerization and depolymerization rates
[44], has begun but it remains to be determined if other
potentially important polymer behaviors, such as membrane association and bundling interactions, are changed.
A mutant reported by Baskin and colleagues [45], radial
swollen 6 (rsw-6), might provide a new tool for teasing
apart the mechanism of array orientation. rsw-6 mutants
are able to maintain arrays of parallel microtubules in
root epidermal cells, but the orientation of these arrays is
no longer coordinated with the cell axis and displays
www.sciencedirect.com
Cortical microtubules in cells that have complex shapes,
such as pavement cells in the leaf epidermis, are arranged
in more complicated patterns that suggest regional control
of cytoskeletal organization within the cell. An exciting
study by Fu and colleagues [2] shed new light on the
mechanisms of regional regulation of microtubule organization in pavement cells. In lobed pavement cells,
microtubule organization has a periodic pattern, only
achieving marked co-alignment in the sinuses. Yang’s
group [2] found that gain- and loss-of-function mutations in the small G-protein ROP2 (Rho of plants 2), the
wildtype version of which localizes to cell lobes, both
created a more uniform transverse array and reduced the
amplitude of cell lobes. In addition, overexpression of
RIC1, a ROP2-interacting protein that binds microtubules in a ROP2-dependent manner, virtually eliminated
cell lobing while promoting a highly organized transverse
microtubule array [2]. Together, these result suggest
that ROP2 modifies microtubule organization through the
local modification of RIC1 activity [2]. As with a host of
other mutated proteins and molecular manipulations that
affect cytoskeletal organization, it will be informative
to determine what specific microtubule behaviors are
targeted by RIC1.
Interaction of the cortical microtubule array
with cell wall biosynthetic machinery
Visualization of dynamic cellulose synthase
As described in the introduction to this review, it is
proposed that cortical array organization is important
because it guides the deposition of cell wall cellulose
microfibrils, thus generating material anisotropy in the
cell wall that is the basis for directional cell expansion
during turgor-driven growth. Although parallelism
between microtubules and cellulose has long been noted
[7], the uncoupling of these polymer arrays has also been
observed [9]. A limitation to understanding the true
relationship between the cortical cytoskeleton and cellulose deposition has been the ability to observe cellulose
synthase itself. Cellulose is not secreted but is synthesized from a large protein complex in the plasma membrane. Freeze-fracture scanning electron microscopy
(SEM) has revealed that the cellulose synthase complex
is a hexagonal rosette 25 nm in diameter [46]. These
rosettes are thought to be comprised of about 36 catalytic
subunits of the CESA protein, of which at least three
isoforms appear to be required for activity [46]. Although
SEM and transmission electron microscopy have revealed
details of cellulose synthase and microtubule organization
in plant cells, they are limited in their ability to reveal the
dynamic relationships among molecules.
Current Opinion in Plant Biology 2006, 9:571–578
574 Cell biology
Imaging of a functional yellow fluorescent protein (YFP)
fusion to CESA6 (YFP<CESA6) has allowed the
dynamic relationship between cortical microtubules
and cellulose synthase itself to be observed in living cells
[47]. As revealed by spinning disk confocal microscopy,
YFP<CESA6 localized to the plasma membrane of etiolated hypocotyl cells in linear arrays of distinct particles
(Figure 1). These particles translocated along linear paths
Figure 1
with steady velocities (averaging 330 nm/min) and were
sensitive to the cellulose synthesis inhibitor isoxaben,
suggesting that the particles were active cellulose
synthase rosettes or collections of rosettes. Consistent
with the alignment hypothesis, the trajectories of the
particles traced the paths of microtubules labeled
with co-expressed CFP<TUBULIN (CFP<TUA1).
YFP<CESA6 particles followed microtubules even along
curved polymers. Furthermore, when microtubules underwent rapid rearrangement, the pattern of YFP<CESA6
was coordinately rearranged, with new microtubule
polymerization preceding new arrangements and trajectories of particles. The colocalization of YFP<CESA6
complexes and microtubules was often tightly coordinated but was not absolute: approximately 60% of the
YFP<CESA6 label overlapped with CFP<TUA1 label.
Considering the large difference in the dynamic properties of these two systems, it is not surprising that CESA
and microtubule localization patterns are not completely
coupled. Elements of the cortical microtubule array were
frequently observed to depolymerize or translocate by
treadmilling, leaving much slower CESA6 complexes
behind. CESA6 complexes remained mobile after abandonment by microtubules, consistent with earlier studies
that indicated that cellulose production itself does not
require microtubules [48].
Observation of YFP<CESA6 complexes suggests an
important role for the polymer bundles that are created
by treadmilling microtubules. Treadmilling allows polymers to be repositioned, giving the array organizational
flexibility, but individual microtubules tend not to remain
in a single position long enough to serve as guides for
slowly translocating CESA6 complexes [47]. Microtubule bundles, on the other hand, have much longer lifetimes [25], providing the positional stability that is
needed to guide cellulose synthase.
Co-localization of YFP<CESA6 and CFP<TUA1 in etiolated hypocotyl
cells of Arabidopsis. Images were acquired every 10 s on a spinning
disk confocal microscope system. The top row of images are the
average of three image frames, showing particulate YFP<CESA6
localization along cortical microtubules. The bottom row shows an
average of 60 frames to visualize the trajectories of YFP<CESA6
complexes as they move through the cell membrane [47].
Current Opinion in Plant Biology 2006, 9:571–578
It is likely that the relationship between the cortical
microtubule array and cellulose synthase is under developmental control, changing over the life of a cell and
varying among different cell types. For example, the
correlation between microtubule and cellulose microfibril
orientation breaks down near the end of the expansion
zone in Arabidopsis roots, with cellulose deposition patterns remaining roughly transverse whereas microtubules
become longitudinally arranged [8]. Moreover, microtubules are not parallel to the observed cellulose deposition
pattern in root hairs [49], suggesting that cellulose
synthase complexes in these cells do not associate with
cortical microtubules. There are ten CESA genes in
Arabidopsis and active cellulose synthase complexes are
predicted to contain at least three cellulose isoforms
[50,51]. Functional tagging of additional CESA proteins
should reveal if complexes of different subunit composition have different abilities to associate with the cortical
cytoskeleton.
www.sciencedirect.com
Microtubule cortical array organization and plant cell morphogenesis Paradez, Wright and Ehrhardt 575
What is the mechanism of CESA guidance?
Models for the guidance of cellulose synthase by cortical
microtubules fall into two general categories: those that
postulate direct or indirect molecular binding of the
cellulose synthase complex to microtubules, and those
that postulate that microtubules simply act as passive
barriers that constrain the trajectories of translocating
complexes. Any model needs to take into account the
observation that YFP<CESA6 particles were observed to
move bi-directionally along every measured track defined
by a single microtubule bundle. Pauses in particle movement were not observed, indicating that particles moving
in opposite directions might be efficiently segregated to
avoid collision.
If CESA complexes are associated directly with microtubules by molecular linkers, uninterrupted bi-directional
movement could be accommodated by lateral interactions
between microtubule bundles and CESA complexes,
with one set of complexes moving ‘north’ on one side
of a microtubule bundle and one set moving ‘south’ on
the other side. Motor proteins, such as kinesins, would be
well suited to the task of a linker, as they would allow the
constant association of moving CESA complexes with the
microtubule wall through coordinated cycles of attachment and de-attachment by the motor domains. The
Arabidopsis genome is also blessed with a plethora of
kinesin motors, with at least 61 identified by molecular
homology to date [52]. Kinesins and other microtubule
motors are directional: capable of translocating towards
one pole of the microtubule or the other. If all microtubule bundles consisted of microtubules that have
anti-parallel polarity, a motor linker would account
nicely for the segregation of bidirectional movement
by CESA complexes. Cortical bundles have been
observed to have both parallel and anti-parallel polarity
[17,53], however, raising a challenge to the development of a simple model for bidirectional movement
involving motor-based linkers. Models in which a nondirectional linker protein is postulated to provide lateral
stabilization of CESA-complexes with microtubules
face a similar challenge. It should be emphasized that
the translocation of the CESA complex itself does not
require a motor protein. Cellulose synthesis and deposition proceed in the absence of microtubules [48], and
YFP<CESA6 complexes continue to move through the
cell membrane when microtubules are depolymerized
[47]. The motive force for complex movement is most
likely provided by cellulose synthesis and microfibril
crystallization [46,48].
Given the observation that YFP<CESA6 complexes can
easily track curved microtubules, passive constraint models that involve more than one microtubule must be
applicable to narrow channels between polymers in
microtubule bundles or bundles that are spaced below
the optical resolution limit of the light microscope.
www.sciencedirect.com
Polymers within bundles have been observed to be
associated with each other by 25 nm cross bridges, probably provided by MAP65 proteins [33], and so the channels formed between adjacent polymers would be only
just large enough to accommodate a single, unadorned
25 nm cellulose synthase rosette. The possibility that the
rosette complex might extend further into the cytosol
because of the association of accessory proteins ([46]; M
Brown, pers. comm.) posses a challenge to this model, as
does the observation of uninterrupted bidirectional
movement of complexes. The possibility that sets of
parallel bundles below the optical resolution limit might
guide CESA complexes cannot be ruled out, but these
polymer arrangements would need to be created at a high
efficiency to account for both the high correlation between
YFP<CESA6 tracks and resolved microtubule bundles
and the concerted re-orientation of YFP<CESA6 tracks
with rapid cortical array re-orientation [47].
Another class of passive restraint model is one in which
there is an inherent curvature to the direction of unrestrained complex movement. If complexes follow a curved
path with a constant handedness, to the left for example,
they will tend to be pressed up against a microtubule fence
when they travel in one direction along the barrier, but will
curve away from the fence when travelling in the opposite
direction. The net result would be the accumulation of
complexes moving in one direction on one side of a single
microtubule bundle and complexes moving in the opposite direction on the other side of the barrier, just as
observed. While curved microfibrils have often been
observed [54], it is not known whether this is the natural
path of an unrestrained complex. At present, neither class
of model, passive constraint or molecular linker, can be
ruled out and further work is needed to distinguish among
these hypotheses.
YFP<CES6 organization in the absence of
microtubules and hypotheses for the function of
microtubule guidance
Near-complete disassembly of microtubules did not
cause a randomization of YFP<CESA6 organization.
Indeed, in the absence of microtubules, YFP:CESA6
was observed to trace roughly parallel trajectories at
oblique angles to the cell axis. Similarly, microfibrils have
been observed to remain transverse in elongating root
cells after disruption of the cortical cytoskeleton by
mutation or drug treatment [9]. These results suggest
that there is a default pattern for CESA6 organization in
the absence of microtubules. This pattern might depend
on a self-organizing mechanism similar to that proposed
by Emons and colleagues [49] or alternatively, might
be the result of interaction with a backup guidance
mechanism. These results also raise the question of
whether microtubules have functions in cellulose or
cell wall biosynthesis besides the tuning of microfibril
deposition pattern. There are at least four alternative
Current Opinion in Plant Biology 2006, 9:571–578
576 Cell biology
Figure 2
that are required to perform work required for cell morphogenesis, such as localization and guidance of cellulose
synthase. The mechanisms by which particular array
structures are created, how the microtubules and actin
cytoskeletons interact and are coordinated, the means by
which cell signaling pathways feed into and reorganize
these structures at both a cellular and tissue-wide scale,
and how intracellular organization is converted into cell
shape remain challenging and exciting problems. The
next few years should prove to be exciting ones for
exploring the molecular mechanisms by which plant cells
create form as new tools for live-cell observation and
experimentation are created, new mutants are discovered,
and genomic and proteomic analyses yield a growing list
of possible molecular players [57–59].
Acknowledgements
The authors would like to thank Jordi Chan and Clive Lloyd and R
Malcolm Brown Jr for sharing data before publication, and Clive Lloyd,
Malcom Brown, Herman Höfte, Andrew Staehelin, Sid Shaw, Tim Stearns,
Chris Somerville and John Sedbrook for stimulating discussions about
microtubule organization and cellulose biosynthesis.
References and recommended reading
Papers of particular interest, published within the annual period of
review, have been highlighted as:
of special interest
of outstanding interest
1.
Schematic of alternative hypotheses for the function of cellulose
synthase guidance by the microtubule cortical array.
hypotheses (Figure 2). First, microtubules might be
required for synthase processivity. In the absence of
microtubules, cellulose synthase complexes might have
shorter lives and produce shorter microfibrils that fail to
regulate wall expansion normally [55]. Second, microtubules might be required to coordinate cellulose deposition with the delivery of other proteins or molecules that
are required for cell wall function [56]. Third, microtubules might be required to concentrate and organize
synthetic complexes so that their products can more
readily assemble into higher order fibers or clusters of
fibers. These macro-fibers or clusters might have mechanical or organizational properties that are important for cell
wall growth. Last, it is possible that dynamic remodeling
of the cellulose deposition pattern is important for wall
expansion and that microtubule guidance allows the cell
greater control in creating these changing patterns.
2.
Fu Y, Gu Y, Zheng Z, Wasteneys G, Yang Z: Arabidopsis
interdigitating cell growth requires two antagonistic pathways
with opposing action on cell morphogenesis. Cell 2005,
120:687-700.
Evidence for a mechanism that exerts local control over cytoskeletal
organization in plant cells. Evidence is presented that the small G-protein
ROP2 and a ROP2-interacting protein, RIC1, regulate microtubule organization and cell lobing in pavement cells. A model is suggested by which
ROP2 negatively regulates RIC1 in cell lobes, and RIC1 actively promotes
microtubule assembly or stabilization in cell sinuses.
3.
Green PB: Mechanism for plant cell morphogenesis.
Science 1962, 138:1404.
4.
Ledbetter MC, Porter KR: A ‘microtubule’ in plant cell fine
structure. J Cell Biol 1963, 19:239-250.
5.
Lloyd CW, Clayton L, Dawson PJ, Doonan JH, Hulme JS,
Roberts IN, Wells B: The cytoskeleton underlying side walls and
cross walls in plants: molecules and macromolecular
assemblies. J Cell Sci Suppl 1985, 2:143-155.
6.
Palevitz B: Potential significance of microtubule
rearrangement, translocation and reutilization in plant cells. In
The Cytoskeletal Basis of Plant Growth and Form. Edited by Lloyd
C. Academic Press; 1991:45–55.
7.
Baskin TI: On the alignment of cellulose microfibrils by cortical
microtubules: a review and a model. Protoplasma 2001,
215:150-171.
8.
Sugimoto K, Williamson RE, Wasteneys GO: New techniques
enable comparative analysis of microtubule orientation; wall
texture; and growth rate in intact roots of Arabidopsis.
Plant Physiol 2000, 124:1493-1506.
9.
Sugimoto K, Himmelspach R, Williamson RE, Wasteneys GO:
Mutation or drug-dependent microtubule disruption causes
radial swelling without altering parallel cellulose microfibril
deposition in Arabidopsis root cells. Plant Cell 2003,
15:1414-1429.
Conclusions
The plant cortical microtubule array is emerging as a
dynamic structure in which individual polymers are constantly being destroyed, rebuilt and repositioned by polymer treadmilling. Dynamic polymers can assemble into
bundles to form more stable elements of the cortical array
Current Opinion in Plant Biology 2006, 9:571–578
Lloyd C, Seagull RW: A new spring for plant cell biology:
microtubules as dynamic helices. Trends Biochem Sci 1985,
10:476-478.
www.sciencedirect.com
Microtubule cortical array organization and plant cell morphogenesis Paradez, Wright and Ehrhardt 577
10. Cyr R: Microtubules in plant morphogenesis: role of the
cortical array. Annu Rev Cell Biol 1994, 10:153-180.
11. Cyr RJ, Palevitz BA: Organization of cortical microtubules in
plant cells. Curr Opin Cell Biol 1995, 7:65-71.
12. Lambert A: Microtubule-organizing centers in higher plants:
evolving concepts. Bot Acta 1995, 108:535-537.
13. Liu B, Joshi HC, Wilson TJ, Silflow CD, Palevitz BA, Snustad DP:
g-Tubulin in Arabidopsis: gene sequence, immunoblot, and
immunofluorescence studies. Plant Cell 1994, 6:303-314.
14. Erhardt M, Stoppin-Mellet V, Campagne S, Canaday J, Mutterer J,
Fabian T, Sauter M, Muller T, Peter C, Lambert AM et al.: The plant
Spc98p homologue colocalizes with g-tubulin at microtubule
nucleation sites and is required for microtubule nucleation.
J Cell Sci 2002, 115:2423-2431.
15. Seltzer V, Pawlowski T, Campagne S, Canaday J, Erhardt M,
Evrard JL, Herzog E, Schmit AC: Multiple microtubule
nucleation sites in higher plants. Cell Biol Int 2003, 27:267-269.
16. Pastuglia M, Azimzadeh J, Goussot M, Camilleri C, Belcram K,
Evrard JL, Schmit AC, Guerche P, Bouchez D: Gamma-tubulin is
essential for microtubule organization and development in
Arabidopsis. Plant Cell 2006, 18:1412-1425.
Genetic analysis demonstrates that g-tubulin function is crucial for microtubule organization throughout the cell cycle.
29. Dixit R, Cyr R: Encounters between dynamic cortical
microtubules promote ordering of the cortical array through
angle-dependent modifications of microtubule behavior.
Plant Cell 2004, 16:3274-3284.
30. Sedbrook JC, Ehrhardt DW, Fisher SE, Scheible WR,
Somerville CR: The Arabidopsis SKU6/SPIRAL1 gene encodes
a plus end-localized microtubule-interacting protein involved
in directional cell expansion. Plant Cell 2004, 16:1506-1520.
31. Nakajima K, Furutani I, Tachimoto H, Matsubara H, Hashimoto T:
SPIRAL1 encodes a plant-specific microtubule-localized
protein required for directional control of rapidly expanding
Arabidopsis cells. Plant Cell 2004, 16:1178-1190.
32. Mathur J, Mathur N, Kernebeck B, Srinivas BP, Hülskamp M:
A novel localization pattern for an EB1-like protein links
microtubules dynamics to endomembrane organization.
Curr Biol 2003, 13:1991-1997.
33. Chan J, Jensen CG, Jensen LC, Bush M, Lloyd CW: The 65-kDa
carrot microtubule-associated protein forms regularly
arranged filamentous cross-bridges between microtubules.
Proc Natl Acad Sci USA 1999, 96:14931-14936.
34. Smertenko A, Saleh N, Igarashi H, Mori H, Hauser-Hahn I,
Jiang CJ, Sonobe S, Lloyd CW, Hussey PJ: A new class of
microtubule-associated proteins in plants. Nat Cell Biol 2000,
2:750-753.
17. Shaw S, Kamyar R, Ehrhardt D: Sustained microtubule
treadmilling in Arabidopsis cortical arrays. Science 2003,
300:1715-1718.
35. Wicker-Planquart C, Stoppin-Mellet V, Blanchoin L, Vantard M:
Interactions of tobacco microtubule-associated protein
MAP65-1b with microtubules. Plant J 2004, 39:126-134.
18. Chan J, Calder G, Doonan J, Lloyd C: EB1 reveals mobile
microtubule nucleation sites in Arabidopsis. Nat Cell Biol 2003,
5:967-971.
36. Smertenko AP, Chang HY, Wagner V, Kaloriti D, Fenyk S,
Sonobe S, Lloyd C, Hauser MT, Hussey PJ: The Arabidopsis
microtubule-associated protein AtMAP65-1: molecular
analysis of its microtubule bundling activity. Plant Cell 2004,
16:2035-2047.
19. Murata M, Sonobe S, Baskin TI, Hyodo S, Hasezawa S, Nagata T,
Horio T, Hasebe M: Microtubule-dependent microtubule
nucleation based on recruitment of g-tubulin in higher plants.
Nat Cell Biol 2005, 7:961-968.
In vitro and in vivo assays demonstrate g-tubulin-dependent microtubule
nucleation from the walls of existing microtubules. New polymers grow
within a narrow range of angles, around 408, relative to the existing
microtubule.
20. Van Damme D, Van Poucke K, Boutant E, Ritzenthaler C, Inzé D,
Geelen D: In vivo dynamics and differential microtubulebinding activities of MAP65 proteins. Plant Physiol 2004,
136:3956-3967.
21. Wasteneys G, Williamson R: Microtubule resassembly in
internodal cell of Nitella tasmanica: assembly of cortical
microtubules in branching clusters and its relevance to steady
state microtubule assembly. J Cell Sci 1989, 93:705-714.
22. Wasteneys G, Gunning B, Hepler P: Microinjection of
fluorescent brain tubulin reveals dynamic properties of
cortical microtubules in living plant cells. Cell Motil Cytoskel
1993, 24:205-213.
23. Yuan M, Shaw P, Warn R, Lloyd C: Dynamic reorientation of
cortical microtubules, from transverse to longitudinal, in living
plant cells. Proc Natl Acad Sci USA 1994, 91:6050-6053.
24. Wymer CL, Fisher DD, Moore RC, Cyr RJ: Elucidating the
mechanism of cortical microtubule reorientation in plant cells.
Cell Motil Cytoskeleton 1996, 35:162-173.
25. Ehrhardt DW, Shaw SL: Microtubule dynamics and organization
in the plant cortical array. Annu Rev Plant Biol 2006, 57:859-875.
26. Dhonukshe P, Gadella TW Jr: Alteration of microtubule
dynamic instability during preprophase band formation
revealed by yellow fluorescent protein-CLIP170 microtubule
plus-end labeling. Plant Cell 2003, 15:597-611.
27. Vos J, Dogterom M, Emons A: Microtubules become more
dynamic but not shorter during preprophase band formation: a
possible ‘search-and-capture’ mechanism for microtubule
translocation. Cell Motil Cytoskeleton 2004, 57:246-258.
28. Gardiner J, Harper J, Weerakoon N, Collings D, Ritchie S, Gilroy S,
Cyr R, Marc J: A 90-kD phospholipase D from tobacco binds to
microtubules and the plasma membrane. Plant Cell 2001,
13:2143-2158.
www.sciencedirect.com
37. Chang HY, Smertenko AP, Igarashi H, Dixon DP, Hussey PJ:
Dynamic interaction of NtMAP65-1a with microtubules in vivo.
J Cell Sci 2005, 118:3195-3201.
FRAP experiments show that MAP65-1 protein on microtubule bundles
turns over much faster than microtubule treadmilling. This is an important
property for a cross-linking protein that is found in bundles composed of
highly dynamic constituent polymers.
38. Smertenko AP, Chang HY, Sonobe S, Fenyk SI, Weingartner M,
Bögre L, Hussey PJ: Control of the AtMAP65-1 interaction
with microtubules through the cell cycle. J Cell Sci 2006,
119:3227-3237.
MAP65-1 localization and function in the mitotic spindle is shown to be
regulated by CDK- and MAPK-dependent phosphorylation.
39. Hejnowicz Z: Autonomous changes in the orientation of
cortical microtubules underlying the helicoidal cell wall of the
sunflower hypocotyl epidermis: spatial variation translated
into temporal changes. Protoplasma 2005, 225:243-256.
Analysis of microtubule orientation in hundreds of sunflower hypocotyl
cells, together with mathematical modeling, suggests that the observed
distributions of array orientations are consistent with a model in which
array orientation in these cells is constantly rotating or oscillating.
40. Nakajima K, Kawamura T, Hashimoto T: Role of the SPIRAL1
gene family in anisotropic growth of Arabidopsis thaliana.
Plant Cell Physiol 2006, 47:513-522.
Mutations in the SPR1 gene family are shown to be additive, resulting in
pronounced loss of cell expansion anisotropy.
41. Furutani I, Watanabe Y, Prieto R, Masukawa M, Suzuki K, Naoi K,
Thitamadee S, Shikanai T, Hashimoto T: The SPIRAL genes are
required for directional control of cell elongation in
Arabidopsis thaliana. Development 2000, 127:4443-4453.
42. Buschmann H, Fabri CO, Hauptmann M, Hutzler P, Laux T,
Lloyd CW, Schaffner AR: Helical growth of the Arabidopsis
mutant tortifolia1 reveals a plant-specific microtubuleassociated protein. Curr Biol 2004, 14:1515-1521.
43. Abe T, Thitamadee S, Hashimoto T: Microtubule defects and cell
morphogenesis in the lefty1 lefty2 tubulin mutant of
Arabidopsis thaliana. Plant Cell Physiol 2004, 45:211-220.
44. Abe T, Hashimoto T: Altered microtubule dynamics by
expression of modified a-tubulin protein causes right-handed
Current Opinion in Plant Biology 2006, 9:571–578
578 Cell biology
helical growth in transgenic Arabidopsis plants. Plant J 2005,
43:191-204.
Modifications of the C-terminus of a-tubulin act dominantly to reduce
microtubule dynamicity and cause right-handed twisting of organ growth.
The authors present evidence that these alterations might reduce the rate
of GTP hydrolysis, stabilizing and extending the G-cap on microtubule
plus ends.
45. Bannigan A, Wiedemeier AM, Williamson RE, Overall RL,
Baskin TI: Cortical microtubule arrays lose uniform alignment
between cells and are oryzalin resistant in the Arabidopsis
mutant, radially swollen 6. Plant Cell Physiol 2006,
47:949-958.
In the rsw-6 mutant, microtubule arrays in root cells are randomized with
respect to axial orientation, but retain parallel architecture. rsw6 might be
defective in a function that links microtubule orientation within the cellular
reference frame, or in coordinating the orientation of arrays across
neighboring cells.
46. Somerville C: Cellulose synthesis in higher plants. Annu Rev Cell
Dev Biol 2006, in press.
47. Paredez AR, Somerville CR, Ehrhardt DW: Visualization of
cellulose synthase demonstrates functional association with
microtubules. Science 2006, 312:1491-1495.
Spinning disk confocal microscopy of a functional YFP<CESA6 fusion
protein allows the observation of dynamic cellulose synthase complexes
in the plant cell membrane for the first time. Membrane protein complexes
that contain CESA6 are shown to track the positions of cortical microtubules and to re-organize their positions and trajectories as microtubules
are reorganized.
48. Heath IB: A unified hypothesis for the role of membrane bound
enzyme complexes and microtubules in plant cell wall
synthesis. J Theor Biol 1974, 48:445-449.
49. Mulder B, Schel J, Emons AM: How the geometrical model for
plant cell wall formation enables the production of a random
texture. Cellulose 2004, 11:395-401.
50. Taylor NG, Howells RM, Huttly AK, Vickers K, Turner SR:
Interactions among three distinct CesA proteins essential for
cellulose synthesis. Proc Natl Acad Sci USA 2003, 100:14501455.
51. Sonobe S, Yamamoto S, Motomura M, Shimmen T: Isolation of
cortical MTs from tobacco BY-2 cells. Plant Cell Physiol 2001,
42:162-169.
52. Lee YR, Liu B: Cytoskeletal motors in Arabidopsis. Sixty-one
kinesins and seventeen myosins. Plant Physiol 2004, 136:38773883.
53. Dixit R, Chang E, Cyr R: Establishment of polarity during
organization of the acentrosomal plant cortical microtubule
array. Mol Biol Cell 2006, 17:1298-1305.
Imaging of EB1 in tobacco tissue culture cells and in transformed
Arabidopsis reveals that many cells display regional and global patterns
of preferred co-orientation of cortical microtubules with respect to
polymer polarity.
54. Giddings TH, Staehelin LA: Spatial relationship between
micotubules and plasma-membrane rosettes during the
deposition of primary wall microfibrils in Closterium sp.
Planta 1988, 173:22-30.
55. Wasteneys GO, Fujita M: Establishing and maintaining axial
growth: wall mechanical properties and the cytoskeleton.
J Plant Res 2006, 119:5-10.
56. Robert S, Bichet A, Grandjean O, Kierzkowski D, Satiat
Jeunemaitre B, Pelletier S, Hauser MT, Höfte H, Vernhettes S:
An Arabidopsis endo-1,4-beta-D-glucanase involved in
cellulose synthesis undergoes regulated intracellular cycling.
Plant Cell 2005, 17:3378-3389.
KORRIGAN protein is shown to localize to a variety of intracellular compartments including Golgi, an early endosomal compartment, the tonoplast, and a fourth compartment that is distinct from the others. This fourth
compartment appears to be localized at the cell cortex and accumulates in
response to treatment with the cellulose synthase inhibitor isoxoben. The
compartment exhibits microtubule-dependent motility. It is proposed that
KORRIGAN activity might be regulated by intracellular recycling.
57. Persson S, Wei H, Milne J, Page GP, Somerville CR: Identification
of genes required for cellulose synthesis by regression
analysis of public microarray data sets. Proc Natl Acad Sci USA
2005, 102:8633-8638.
58. Muench DG, Chuong SD, Franceschi VR, Okita TW: Developing
prolamine protein bodies are associated with the cortical
cytoskeleton in rice endosperm cells. Planta 2000,
211:227-238.
59. Persson S, Wei H, Milne J, Page GP, Somerville CR: Identification
of genes required for cellulose synthesis by regression
analysis of public microarray data sets. Proc Natl Acad Sci USA
2005, 102:8633-8638.
How to re-use Elsevier journal figures in multimedia presentations
It’s easy to incorporate figures published in Trends, Current Opinion or Drug Discovery Today journals
into your multimedia presentations or other image-display programs.
1.
2.
3.
Locate the article with the required figure on ScienceDirect and click on the ‘Full text + links’
hyperlink
Click on the thumbnail of the required figure to enlarge the image
Copy the image and paste it into an image-display program
Permission of the publisher is required to re-use any materials from Trends, Current Opinion or Drug
Discovery Today journals or from any other works published by Elsevier. Elsevier authors can obtain
permission by completing the online form available through the Copyright Information section of
Elsevier’s Author Gateway at http://authors.elsevier.com. Alternatively, readers can access the
request form through Elsevier’s main website at:
www.elsevier.com/locate/permissions
Current Opinion in Plant Biology 2006, 9:571–578
www.sciencedirect.com