Download Progress in understanding the role of microtubules in plant cells

Progress in understanding the role of microtubules in plant cells
Geoffrey O Wasteneys
Microtubules have long been known to play a key role in plant
cell morphogenesis, but just how they fulfill this function is
unclear. Transverse microtubules have been thought to
constrain the movement of cellulose synthase complexes in
order to generate transverse microfibrils that are essential
for elongation growth. Surprisingly, some recent studies
demonstrate that organized cortical microtubules are not
essential for maintaining or re-establishing transversely
oriented cellulose microfibrils in expanding cells. At the same
time, however, there is strong evidence that microtubules are
intimately associated with cellulose synthesis activity,
especially during secondary wall deposition. These apparently
conflicting results provide important clues as to what
microtubules do at the interface between the cell and its wall.
I hypothesize that cellulose microfibril length is an important
parameter of wall mechanics and suggest ways in which
microtubule organization may influence microfibril length. This
concept is in line with current evidence that links cellulose
synthesis levels and microfibril orientation. Furthermore, in
light of new evidence showing that a wide variety of proteins
bind to microtubules, I raise the broader question of whether
a major function of plant microtubules is in modulating signaling
pathways as plants respond to sensory inputs from the
environment.
Addresses
Department of Botany, University of British Columbia, 3529-6270
University Boulevard, Vancouver, British Columbia, Canada V6T 1Z4
e-mail: [email protected]
Current Opinion in Plant Biology 2004, 7:651–660
This review comes from a themed issue on
Cell biology
Edited by Martin Hülskamp and Yasunori Machida
Available online 25th September 2004
1369-5266/$ – see front matter
# 2004 Elsevier Ltd. All rights reserved.
DOI 10.1016/j.pbi.2004.09.008
Abbreviations
AtKSS
Arabidopsis thaliana KATANIN SMALL SUBUNIT
DCB
2,6-dichlorobenzonitrile
fra2
fragile fiber2
KCBP
kinesin-like calmodulin binding protein
mor1-1 microtubule organization1-1
spr1
spiral1
XTH
xyloglucan endotransglucosylase/hydrolase
Introduction
Just when the advances in microscopy, molecular biology
and genomics converged to make work easier for biolwww.sciencedirect.com
ogists, biological systems seem to have become more
complex, and long-held concepts have begun to crumble.
This is very well illustrated in the study of microtubule
function in plant cells. Since their first clear description in
1963 in a publication by Ledbetter and Porter [1], in
which the term ‘microtubule’ was first coined, the question of how and not whether microtubules control the
orientation of cellulose microfibrils has been the central
focus of most studies on cortical microtubules in plants.
Ledbetter and Porter [1] noted that these fine tubular
structures mirrored the orientation of cellulose microfibrils, whose function as the main load-bearing and
growth-axis-determining component of cell walls had
already been established. A year before, Paul Green [2]
speculated that colchicine-sensitive ‘proteins of a spindle
fiber nature’ at the plasma membrane would be active in
the control of wall texture and cell form. Indeed, it was
eventually proven that spindle fibers were also microtubules, and colchicine-binding protein was renamed tubulin to reflect the fact that it was the primary component of
microtubules [3]. In plant cells, microtubules were soon
found to be a major part of the previously described
phragmoplast, which builds the cell plate during telophase, and to form a cortical band before prophase, which
somehow marks the site at which the cell plate will
eventually connect with the parent cell membrane (for
review see [4,5]).
During the two decades that followed the discovery of
cortical microtubules, several research teams equipped
with transmission electron microscopes put forward models to explain how cortical microtubules could control
the orientation of cellulose microfibrils. The most widely
accepted model, the cellulose-synthase-constraint hypothesis, is summarized in a 1991 article by Giddings and
Staehelin [6]. This model is shown here in a model
diagram produced by Brian Gunning (Figure 1), which
incorporates xyloglucan tethers. According to this theory,
microtubules, through their close interaction with the
plasma membrane, form barriers that constrain the paths
of cellulose synthase complexes as they deposit cellulose
chains in the cell wall.
Lack of direct evidence has always marred the synthaseconstraint model. In 2000, I highlighted some studies that
brought the obligate nature of microtubule–microfibril
co-alignment into question [7]. I speculated that combined genetic and cytological approaches would soon
yield new models to describe how the microtubule cytoskeleton regulates the anisotropic properties of the cell
wall [7]. A comprehensive survey of pre-2001 research
by Tobias Baskin [8] also reminded us of the many
Current Opinion in Plant Biology 2004, 7:651–660
652 Cell biology
Figure 1
Microfibrils embedded
in wall matrix
β (1-4) glucan chains in a
cellulose microfibril
all
Microfibrils linked
by xyloglucans
ll w
Ce
PF
Microfibril
emerging through
plasma mambrane
Lipid bilayer of
plasma mambrane
F
P
PF
Cellulose microfibril
emerging from rosette,
parallel to microtubule
Intermicrotubule
bridge
Microtubule
Microtubule bridged
to plasma membrane
(and cell wall?)
Relationships between cellulose-synthesizing complexes (rosette type), wall microfibrils, plasma membrane and microtubules. Diagram
provided by Brian Gunning.
experimental systems in which well-organized microfibrils could be produced in the absence of microtubules.
Baskin also articulated the ‘templated-incorporation’
model in which cellulose microfibrils adhere to a scaffold
deposited by microtubules, but in which established
microfibrils can continue to be deposited in the direction
of this scaffold regardless of whether cortical microtubules remain in place [8].
One key to understanding how microtubules work is to
recognize that they are far more complex than the clean,
green structures we usually see labeled with fluorescent
probes. Microtubule surfaces are heavily congested
places, the landing platforms for all sorts of proteins,
membranous inclusions, nucleotides and ions. Some residents, the microtubule-associated proteins and their regulatory elements, co-ordinate the assembly, bundling and
stability of the microtubules. Others are motor proteins
that shuttle cargo or participate in microtubule organization. Perhaps hundreds of other elements bind to microtubules, either because their function is somehow
regulated by microtubules or simply because microtubules provide a convenient surface on which they can
perform their functions. Amidst emerging evidence of
numerous microtubule-binding elements, I now wonder
what the real functions of microtubules in plant cells
include. Are microtubules simply rigid structural elements that provide useful mechanical support, barriers
and rapid transport routes? Or, can they also act as
repositories for signaling molecules to modulate, by their
Current Opinion in Plant Biology 2004, 7:651–660
changeable polymer status, metabolic responses to external cues?
Cortical microtubules and cell-wall
mechanics
Microtubule function has been studied using drugs that
target tubulin and interfere with microtubule assembly.
This approach is useful but limited by variable drug
accessibility to cells. Moreover, these drugs do not discriminate between microtubules that are involved in cell
division and those in interphase. If cell division planes
are disturbed or cell division arrested altogether, consequent disturbance to cell axis formation, including
disturbance to the polar distribution of auxin efflux
transporters, can help to explain why cellulose microfibrils lose global transverse orientation after drug
treatments [9]. Mutations that affect microtubule organization provide an alternative means of assessing cortical microtubule function. Mutants, however, are not
without their own drawbacks. Those with constitutive
phenotypes may be embryo-lethal or accumulate defects
as they develop. The temperature-sensitive microtubule
organization1-1 (mor1-1) mutant is unique to date
because its growth and development is normally indistinguishable from that of wildtype plants, yet shifting
the culture temperature by a few degrees generates
microtubule disorganization, left-handed organ twisting,
and loss of growth anisotropy in these mutants [10].
Culturing mor1-1 mutants at the threshold of the restrictive temperature range disorganizes cortical arrays
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Microtubules in plant cells Wasteneys 653
almost immediately but has relatively little effect on
microtubules in dividing cells [10].
Microtubule disruption generates radial
swelling without altering the orientation of
cellulose microfibrils
Using a method developed to compare the patterns of
microtubule and cellulose-microfibril orientation in
equivalent locations of roots of known growth status
[11], we found that in mor1-1 roots cells, cellulose microfibrils continued to be deposited transverse to the long
axis of the root even after prolonged disruption of cortical
microtubule arrays and the onset of radial expansion
[12]. We also examined microfibril patterns after depolymerizing cortical microtubules with the drug oryzalin or
stabilizing them with taxol. We detected no changes to the
transverse order of cellulose microfibrils in cells that had
entered the elongation zone before the drug treatment.
Longer treatments with these drugs generated variable
patterns of microfibril orientation in cells whose division
had been perturbed, but in these cases, the microfibrils
remained locally parallel [12]. Baskin and colleagues
[13] recently described similar findings in Arabidopsis
roots treated with much lower, non-saturating concentrations of the drug oryzalin. Using birefringence assays in
addition to field-emission scanning electron microscopy,
they confirmed that microfibrils remain parallel, if not
better aligned, when microtubule organization is disturbed. Nevertheless, they contend that microtubules
remain in global control of microfibril alignment [13].
Figure 2
(a)
(b)
(c)
(d)
(e)
(f)
Neither of the studies described above ruled out the
possibility that if a transverse template of cellulose microfibrils is established before microtubule disruption, then
new microfibrils may continue to be deposited transversely even after microtubules are lost. To check this
possibility, we designed an experiment in which the drug
2,6-dichlorobenzonitrile (DCB) was used to randomize
the orientation patterns of cellulose microfibrils in the
mor1-1 mutant (Figure 2a,b; [14]). Once cellulose
microfibril texture was disturbed, we shifted plants to
mor1-1’s restrictive temperature (Figure 2c,d) and, finally,
allowed cellulose synthesis to recover by washing out the
DCB. The working hypothesis was that creating disordered microtubules together with a disordered microfibril
template should prevent the recovery of well-ordered
cellulose microfibrils. By contrast, our experiments
revealed that cellulose microfibrils could not only recover
parallel order as cells underwent radial swelling but also
that the net orientation of the microfibrils was transverse
to the root long axis (Figure 2e–g). These results throw
out a strong challenge to the idea that microtubules orient
cellulose microfibrils.
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Frequency (%)
(g) 25
Microfibrils can establish transverse, parallel
order in the absence of both transverse
microtubules and a well-ordered template of
microfibrils
Microfibrils
Microtubules
20
15
10
5
0
0
20
40
60
80 100 120
Orientation (degrees)
140 160
180
An experiment to test whether microfibrils can re-establish parallel
order when the existing microfibril template is disordered and cortical
microtubules are disorganized. After DCB treatment for 4 h,
(a) cortical microtubules remain transversely oriented in mor1-1
root cells, whereas (b) the formerly transverse orientation of cellulose
microfibrils has been disrupted by inhibiting cellulose synthesis.
Shifting the temperature from 21 8C to 31 8C (c) disorganizes the
microtubules while (d) the disordered microfibril texture is maintained.
Maintaining the 31 8C temperature leads to significant lateral
expansion of cells and (e) continued disorganization of the cortical
microtubules, but 18 h after DCB’s removal, (f) microfibrils are
organized in parallel arrays. (g) At the time point shown in (e) and (f),
microtubule orientation is widely dispersed about the transverse axis
(blue bars), while microfibrils (red bars) show relatively tight
distribution about the transverse axis. Microtubules were examined by
immunofluorescence and cellulose microfibrils by field-emission
scanning electron microscopy. Scale bars are 50 mm for (a), (c) and
(e), and 200 nm for (b), (d) and (f). This figure is modified from
Figures 2 and 3 of [14].
Current Opinion in Plant Biology 2004, 7:651–660
654 Cell biology
Microfibril alignment depends on sufficient
levels of cellulose synthesis
An important clue to understanding the results of the
above studies is that, at its restrictive temperature, cellulose content is not reduced in mor1-1 [12]. Several
studies provide evidence of a correlation between the
level of cellulose synthesis and the pattern of microfibril
orientation in cellulose-deficient mutants [15–17],
although this correlation may not be universal [18]. Thus,
the apparently unperturbed orientation of microfibrils in
mor1-1 reflects normal patterns of deposition that are
consistent with normal levels of cellulose synthesis. This
correlation between microfibril orientation and cellulose
production is equally important for interpreting the aberrant microfibril orientation reported in the mutant fragile
fiber2 (fra2) [19], a null allele of the katanin p60 ‘small’
subunit (AtKSS) [20]. AtKSS’s demonstrable microtubule-severing activity [21] plays an important role in
cortical microtubule organization [4,22], and numerous
mutants in which AtKSS is affected have now been
identified in screens for altered morphology [23], stem
strength [23,24], trichoblast cell-fate specification [25],
responses to gibberellin [26] and tubulin-targeted drug
sensitivity [27]. These mutants are unable to establish
transverse cortical microtubule arrays after cytokinesis. In
fra2, cellulose levels are reduced and microfibril patterns
are moderately disturbed, although not in the same
pattern as cortical microtubules [19,26]. Thus, the disordered microfibril orientation patterns in fra2 may be a
consequence of reduced cellulose synthesis.
Do cortical microtubules regulate the length
of cellulose microfibrils?
Anisotropic cell expansion requires, as a minimum, first,
sufficient levels of cellulose synthesis to achieve parallel
microfibril order and, second, microtubules that align in
the same direction as nascent microfibrils, especially
during rapid organ growth. Other factors are also required
[28]. This close correspondence between microtubule
and cellulose-microfibril orientation should, therefore,
remain a central feature of any alternative to the cellulose-synthase-constraint or templated-incorporation
hypotheses. I propose a model in which microtubules
ensure the synthesis and integrity of long microfibrils by
forming parallel cortical arrays in the same direction as
cellulose-microfibril deposition. I call this the microfibrillength-regulation hypothesis.
This hypothesis is based on a premise that intimate
connections between matrix polysaccharides and cellulose microfibrils are central to wall loosening [29]. Wall
loosening is achieved through the activity of wall proteins
such as expansins [30], which are believed to act by
breaking hydrogen bonds that link wall polymers, along
with the activities of wall-modifying enzymes, especially
the xyloglucan endotransglucosylase/hydrolases (XTHs),
which can both cut and ligate xyloglucans [31]. Matrix
Current Opinion in Plant Biology 2004, 7:651–660
breakage and annealing to allow the controlled slippage
of the load-bearing microfibrils elegantly explains wall
extension in one direction (elongation) but it does not
explain how walls expand laterally when microfibrils
remain abundant and transversely oriented. I suggest that
microtubule disorganization results in relatively short
microfibrils or induces periodic weaknesses in cellulose
chains. This allows turgor-driven, enzyme-assisted separation of microfibrils in two directions, while maintaining
microfibrils in a transverse orientation.
Two variations of this model are illustrated in Figure 3.
For simplicity, the matrix, comprising xyloglucan tethers
and pectic polysaccharides, is depicted as a sheet-like
structure, although in reality it is inhomogeneous.
Under conditions in which microtubule arrays are intact
(Figure 3a), cellulose microfibrils are long and only move
apart from one another with no lateral displacement. This
is perfect anisotropy. In the first variation of the microfibril-length-regulation model (Figure 3b), close linkage
of cortical microtubules to the plasma membrane stabilizes newly synthesized microfibrils against mechanical
weakness. Imperfections in microfibrils, which are indicated by dark regions on the newly formed microfibrils
(Figure 3b), might arise before the cellulose chains are
consolidated into paracrystalline fibrils, and would be
vulnerable to breakage when stressed. The fragmented
microfibrils would then move apart from each other, both
at right angles and parallel to their direction of deposition,
generating isotropic wall expansion. This model emphasizes a role for cortical microtubules in maintaining
consistent microfibril strength through mechanical interactions with the plasma membrane. Consistent with
experimental evidence, partial disruption of the microtubule array or aberrant orientation will generate partial
loss of anisotropy. Complete loss of cortical microtubules,
either through depolymerization or disconnection from
the plasma membrane, will lead to isotropic expansion.
The second variation (Figure 3c) invokes modulation of
cellulose-synthase-complex longevity and/or activity by
altering microtubule polymer status, resulting in the
production of short but otherwise normal microfibrils.
The activity and/or life-span of the cellulose-synthase
complex directly controls microfibril length. Unfortunately, there is currently almost no information available
on the longevity of cellulose-synthase complexes,
although one earlier study estimated a life-span of
20 min in moss protonemata [32]. A role for microtubules
in maintaining synthase activity or in stimulating the
insertion of synthase complexes is consistent with the
close association of microtubules with secondary wall
ingrowths, where massive cellulose deposition takes place
[33,34,35]. A recent article demonstrates a close association of microtubules, sucrose synthase and cellulose biosynthesis in Zinnia tracheary elements [36], reminding us
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Microtubules in plant cells Wasteneys 655
Figure 3
(a) Microtubules intact, microfibrils long: anisotropic expansion
(b) Microtubules disorganized, microfibrils fragmented: isotropic expansion
(c) Microtubules disorganized, microfibrils short: isotropic expansion
Cellulose microfibril
Xyloglucan/pectic polysaccharide matrix
Current Opinion in Plant Biology
The microfibril-length-regulation hypothesis. Microtubule activity at the plasma membrane influences the length of cellulose microfibrils, whose
separation, which is mediated by the activity of enzymes on inter-microfibril polysaccharide tethers, determines the direction of cell-surface
expansion (dark blue arrows). (a) Under normal conditions, in which microtubules (not shown) are plentiful and oriented in the direction of
microfibril synthesis, long microfibrils are produced and their separation is only at right angles to their orientation. (b) Loss of cortical
microtubules or their mis-orientation generates periodic weaknesses in microfibrils, which are prone to breakage, allowing separation of
microfibrils in the lateral as well as longitudinal direction. (c) Alternatively, the loss of well-organized microtubules affects the longevity or
activity of cellulose synthase complexes, resulting in relatively short microfibrils. This model explains how cells can undergo radial expansion
while microfibril orientation remains transverse.
that the availability of UDP-glucose precursors, which is
determined by sucrose synthase activity, is yet another
requirement of cellulose synthesis that may be microtubule-dependent.
The best way to pack microtubules may be
transversely
The microfibril-length-regulation hypothesis still requires cells to have transverse microtubules for optimal
cell elongation. An earlier article that suggested that
microtubule orientation can derive information from cellulose microfibrils [37] is supported by two recent studies
that employed inhibitors of cellulose synthesis [14,38].
Microtubule organization in pollen tubes is altered by
isoxaben treatments [38] and cortical microtubules
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become more dispersed about the transverse axis when
root epidermal cells are treated with DCB [14]. Perhaps
transverse orientation is the best way to pack large numbers of microtubules into a cylinder, in which case polymer status may be more important for microtubule
function than their orientation. Transverse cortical arrays
appear to be more heavily populated than the righthanded oblique arrays that are found in Arabidopsis
root cells during growth cessation [11]. Ordering large
populations of microtubules into transverse arrays may
occur, by default, through self-assembly mechanisms [4].
Recent observations of microtubule activity in living cells
[39–41,42,43] support this view. The role that microtubule-associated proteins play in this activity is described in detail elsewhere [4,22,44,45].
Current Opinion in Plant Biology 2004, 7:651–660
656 Cell biology
Microtubule control of directional
handedness?
Organ twisting is an important feature of bending
responses and twining habits. Mutants that have organtwisting phenotypes have been identified, and most have
defects in microtubule organization. Early predictions
that the right-twisting (Z-form) mutant spiral1 (spr1) is
defective in a microtubule-related process [7,46,47] have
proven to be correct. SPR1/SKU6 has been recently
identified as a novel microtubule-associated protein of
tiny size that preferentially associates with the plus ends
of microtubules [48,49]. Treatments with low concentrations of microtubule-targeted drugs [46], the
temperature-sensitive alleles of MOR1 [10,50] and the
semi-dominant tubulin mutants lefty1 and lefty2 [51], all
generate left-handed (S-form) twisting in most organs.
Curiously, genetic or pharmacological left-handed twisting always seems to override or suppress right-handed
twisting. Left-handed twisting in mor1 mutants occurs
early and at the lower end of the restrictive temperature,
whereas the lefty1 lefty2 double mutant has moredisrupted microtubules and radial swelling in addition
to strong twisting [52]. Attributing the handedness of
organ twisting to the handedness of helical cortical microtubule arrays [46,47,51] is therefore tenuous. Twisting
occurs in the mor1-1 mutants, which do not have biased
microtubule orientation [10]; and the right-handed helical
cortical microtubule arrays in the lefty single mutants [51]
follow the default loss of transverse orientation seen
normally in cells as growth rates and microtubule numbers decline [11]. An alternative interpretation is that
non-uniform loss of anisotropy along radial tissue gradients in concert with inherent torsional handedness may
explain twisting handedness [9].
Does microtubule polymer status regulate
signaling and transcriptional events?
A large-scale proteomics project identified a staggering
122 proteins from a tubulin affinity chromatography column [53]. Such rich diversity of microtubule-binding
proteins suggests that microtubules fulfill numerous roles
beyond coordinating cell division and morphogenesis.
Several recent articles (described below) provide evidence for such functions, especially in relation to adaptive
responses.
Here, the idea is that signaling molecules, bound to
microtubules either directly or indirectly through protein
complexes, may be released to the cytoplasm and become
active when microtubules are depolymerized [54]. Many
environmental triggers, including cold [55], osmotic
stress [56], exposure to heavy metals [57–59,60,61]
and pathogens [62–66], are associated with reorganization
of the cytoskeleton. This may involve transient microtubule disruption, which is likely to be associated with a
sudden increase in cytosolic calcium, or be stimulated by
kinase activity [27,67,68]. In these situations, proteins
Current Opinion in Plant Biology 2004, 7:651–660
that are sequestered on microtubule surfaces will be
released and possibly activated or mobilized to new
targets. Depending on the intensity of the signal, the
activation of signal transducers or repressors may be
sufficient to elicit an appropriate metabolic response.
Recent progress in understanding how microtubules are
linked to the plasma membrane has come from the discovery that phospholipase D associates with microtubules
[69]. Compounds that activate phospholipase D, and signals associated with osmotic stress, result in microtubule
disorganization and detachment from the plasma membrane [70], inhibiting normal seedling development [71].
Two examples of how microtubules might gauge ambient
conditions include cold acclimation and the response to
aluminum in the soil. Freeze-tolerance generally requires
a period of cool temperatures to stimulate metabolic
changes, and freeze-tolerant wheat varieties consistently
respond to cold treatments with a transient microtubulereorganization event [55]. Remarkably, freeze-sensitive
wheat cultivars can be rendered freeze-tolerant by briefly
treating them with a herbicide that disassembles their
microtubules. Cross-resistance to microtubule-specific
herbicides and chilling stress has been found in activation-tagged lines of tobacco [72]. A recent study suggests
that aluminum exposure stimulates glutamate efflux,
which, via the activation of glutamate receptors, generates Ca2+ influx [60]. The result is transient microtubule
disassembly, which may integrate with the signaling
cascade that eventually protects the plant by the production and secretion of organic acids.
The kinesin-like calmodulin-binding protein (KCBP)
and the complexes that it forms provide an intriguing
example of a link between Ca2+-dependent microtubule
activity and transcriptional regulation. Although the zwichel mutant phenotype focuses attention on KCBP’s role
in trichome morphogenesis through its ability to bind and
possibly bundle microtubules in a Ca2+-dependent manner [73], there are tantalizing hints that KCBP is part of a
bigger network of regulatory mechanisms. Recent studies
demonstrate that KCBP interacts directly with the plant
homologue of CtBP/BARS (carboxy-terminal binding
protein/brefeldin A ADP-ribosylated substrate) transcriptional repressors, ANGUSTIFOLIA [74,75]. Loss of
ANGUSTIFOLIA function through mutation causes
the overexpression of numerous genes, including one
that encodes the wall-modifying XTH enzyme [75].
Thus, a tantalizing loop emerges that involves Ca2+
signaling, microtubule-bundling activity, and the sequestration by KCBP of a transcriptional regulator that regulates general morphological characteristics that may
involve microtubule activity [76].
Finally, microtubules may play a crucial role in remodeling the actin cytoskeleton [76], and this in turn, may
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Microtubules in plant cells Wasteneys 657
explain how microtubules regulate morphogenesis, possibly by controlling the length of cellulose microfibrils
across the growth continuum. The small GTPase molecular switches known as ROPs (Rho of plants) regulate
both the assembly of the fine networks of microfilaments
that are associated with tip growth and the interdigitating
growth of leaf pavement epidermal cells [77]. A regulatory
role for microtubules in ROP signaling to the actin
cytoskeleton may explain why microtubule disruption
affects morphogenetic processes that are thought to be
largely under the control of actin microfilaments. Understanding how microtubule organization affects the activity and function of actin microfilaments will yield
important knowledge about morphogenesis and the
workings of plant cells.
Conclusions
The availability of conditional microtubule disruption in
mor1-1 mutants, along with improvements in scanning
electron microscopy, has enabled the testing and questioning of the existing synthase-constraint and templatedincorporation models. Although additional independent
testing is warranted before all aspects of the old models
are rejected, it is not too early to use these findings to
think more flexibly about cell-wall expansion and the
specific role that microtubules play. In this article, I have
introduced the concept that microtubule-dependent
microfibril length may contribute, along with microfibril
orientation, to the mechanical properties of the plant cell
wall, a concept we are now testing experimentally. I have
also suggested, in light of the burgeoning number of
microtubule-binding proteins and some intriguing recent
experimental evidence, that microtubule polymer status
may modulate responses to many environmental signals.
Whether or not ‘microtubulomics’ becomes a hot new
area of cell and developmental biology, I hope that these
new ideas provoke a rethinking of the role that microtubules play in all aspects of plant cell function.
Acknowledgements
I acknowledge Keiko Sugimoto and Regina Himmelspach for their
careful work, Richard Williamson for helpful discussions, Tsubasa
Shoji and David Collings for comments on this manuscript, and other
members of my laboratory, whose research efforts and enthusiasm have
helped to stimulate new ideas. I also thank Brian Gunning for
generously providing Figure 1.
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.
Ledbetter MC, Porter KR: A ‘‘microtubule’’ in plant cell fine
structure. J Cell Biol 1963, 19:239-250.
2.
Green PB: Mechanism for plant cellular morphogenesis.
Science 1962, 138:1404-1405.
3.
Weisenberg RC, Borisy GG, Taylor EW: The colchicine-binding
protein of mammalian brain and its relation to microtubules.
Biochemistry 1968, 7:4466-4479.
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4.
Wasteneys GO: Microtubule organization in the green
kingdom: chaos or self-order? J Cell Sci 2002, 115:1345-1354.
5.
Hussey PJ, Hawkins TJ, Igarashi H, Kaloriti D, Smertenko A:
The plant cytoskeleton: recent advances in the study of the
plant microtubule-associated proteins MAP-65, MAP-190 and
the Xenopus MAP215-like protein, MOR1. Plant Mol Biol 2002,
50:915-924.
6.
Giddings TH, Staehelin LA: Microtubule-mediated control of
microfibril deposition; a re-examination of the hypothesis. In
The Cytoskeletal Basis of Plant Growth and Form. Edited by Lloyd
CW. London: Academic Press; 1991:85-100.
7.
Wasteneys GO: The cytoskeleton and growth polarity.
Curr Opin Plant Biol 2000, 3:503-511.
8.
Baskin TI: On the alignment of cellulose microfibrils by cortical
microtubules: a review and a model. Protoplasma 2001,
215:150-171.
9.
Wasteneys GO, Collings DA: Expanding beyond the great divide:
the cytoskeleton and axial growth. In The Plant Cytoskeleton in
Cell Differentiation and Development. Edited by Hussey PJ.
Oxford: Blackwell; 2004:83-115.
Check out this review for a more comprehensive analysis of cytoskeletal
function in growth-axis establishment, tropisms and twisting.
10. Whittington AT, Vugrek O, Wei KJ, Hasenbein NG, Sugimoto K,
Rashbrooke MC, Wasteneys GO: MOR1 is essential for
organizing cortical microtubules in plants. Nature 2001,
411:610-613.
11. 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.
12. 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.
The work described in this paper demonstrates that microfibrils retain
transverse parallel order when lateral growth is induced the root epidermal cells of Arabidopsis thaliana by the temperature-dependent mor1-1
mutation or by oryzalin or taxol treatments. The additive mor1-1 radial
swelling1-1 (rsw1-1) double-mutant phenotype and the unperturbed
cellulose content of mor1-1 mutants suggest that microtubules control
the mechanical properties of the wall by some means other than cellulose
synthesis or microfibril orientation.
13. Baskin TI, Beemster GTS, Judy-March JE, Marga F:
Disorganization of cortical microtubules stimulates tangential
expansion and reduces the uniformity of cellulose microfibril
alignment among cells in the root of Arabidopsis. Plant Physiol,
135:2279-2290.
Drug concentrations that are insufficient to depolymerize microtubules
completely generate radial swelling. However, the orientation of cellulose
microfibrils is unperturbed both by this treatment and in the mor1-1
mutant [12]. This is possibly the first study to directly measure lateral
wall growth on the surface of root epidermal cells. The authors also use
birefringence retardation measurements to investigate microfibril angle.
14. Himmelspach R, Williamson RE, Wasteneys GO: Cellulose
microfibril alignment recovers from DCB-induced disruption
despite microtubule disorganization. Plant J 2003, 36:565-575.
This work reveals that cellulose microfibrils can be oriented precisely
without an organized cortical microtubule array or a pre-existing template
of well-ordered microfibrils. Combining exposure to the cellulose-synthesis-perturbing drug DCB with temperature-dependent microtubule disruption in the mor1-1 mutant provided a rigorous test of existing models
of how microtubules control the mechanical properties of the wall. The
work also suggests that microtubule orientation may be influenced by
cellulose microfibril patterns.
15. Sugimoto K, Williamson RE, Wasteneys GO: Wall architecture in
the cellulose-deficient rsw1 mutant of Arabidopsis thaliana:
microfibrils but not microtubules lose their transverse
alignment before microfibrils become unrecognizable in the
mitotic and elongation zones of roots. Protoplasma 2001,
215:172-183.
16. Sato S, Kato T, Kakegawa K, Ishii T, Liu YG, Awano T, Takabe K,
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Current Opinion in Plant Biology 2004, 7:651–660
658 Cell biology
membrane-bound endo-1,4-beta-glucanase KORRIGAN in
cell elongation and cellulose synthesis in Arabidopsis
thaliana. Plant Cell Physiol 2001, 42:251-263.
17. Pagant S, Bichet A, Sugimoto K, Lerouxel O, Desprez T,
McCann M, Lerouge P, Vernhettes S, Höfte H: KOBITO1 encodes
a novel plasma membrane protein necessary for normal
synthesis of cellulose during cell expansion in Arabidopsis.
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18. Zhong R, Burk DH, Morrison WH III, Ye ZH: A kinesin-like
protein is essential for oriented deposition of cellulose
microfibrils and cell wall strength. Plant Cell 2002,
14:3101-3117.
19. Burk DH, Ye ZH: Alteration of oriented deposition of cellulose
microfibrils by mutation of a katanin-like microtubulesevering protein. Plant Cell 2002, 14:2145-2160.
20. McClinton RS, Chandler JS, Callis J: cDNA isolation,
characterization, and protein intracellular localization of a
katanin-like p60 subunit from Arabidopsis thaliana.
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21. Stoppin-Mellet V, Gaillard J, Vantard M: Functional evidence for
in vitro microtubule severing by the plant katanin homologue.
Biochem J 2002, 365:337-342.
22. Hashimoto T: Dynamics and regulation of plant interphase
microtubules: a comparative view. Curr Opin Plant Biol 2003,
6:568-576.
23. Bichet A, Desnos T, Turner S, Grandjean O, Höfte H: BOTERO1 is
required for normal orientation of cortical microtubules and
anisotropic cell expansion in Arabidopsis. Plant J 2001,
25:137-148.
24. Burk DH, Liu B, Zhong R, Morrison WH, Ye ZH: A katanin-like
protein regulates normal cell wall biosynthesis and cell
elongation. Plant Cell 2001, 13:807-827.
25. Webb M, Jouannic S, Foreman J, Linstead P, Dolan L: Cell
specification in the Arabidopsis root epidermis requires the
activity of ECTOPIC ROOT HAIR 3 — a katanin-p60 protein.
Development 2002, 129:123-131.
26. Bouquin T, Mattsson O, Naested H, Foster R, Mundy J: The
Arabidopsis lue1 mutant defines a katanin p60 ortholog
involved in hormonal control of microtubule orientation during
cell growth. J Cell Sci 2003, 116:791-801.
A screen for altered regulation by gibberellin of a luciferase reporter
construct identified an AtKSS allele that encodes a truncated protein. A
yeast two-hybrid screen demonstrated that AtKSS may interact with a
katanin p80 homologue, as well as with a kinesin-related protein. This is
the first evidence that the larger regulatory katanin subunit may be
functional in plants.
27. Naoi K, Hashimoto T: A semidominant mutation in an
Arabidopsis mitogen-activated protein kinase phosphataselike gene compromises cortical microtubule organization.
Plant Cell 2004, 16:1841-1853.
A tantalizing connection between mitogen-activated protein kinase activity and microtubule-associated protein function is made in this study. The
authors also report that a screen for hypersensitivity to propyzamide
identified many new AtKSS and tubulin mutations.
28. Wiedemeier AM, Judy-March JE, Hocart CH, Wasteneys GO,
Williamson RE, Baskin TI: Mutant alleles of Arabidopsis
RADIALLY SWOLLEN 4 and 7 reduce growth anisotropy
without altering the transverse orientation of cortical
microtubules or cellulose microfibrils. Development 2002,
129:4821-4830.
29. Cosgrove DJ: Wall structure and wall loosening. A look
backwards and forwards. Plant Physiol 2001, 125:131-134.
30. Cosgrove DJ: Loosening of plant cell walls by expansins.
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31. Rose JK, Saladie M, Catala C: The plot thickens: new
perspectives of primary cell wall modification. Curr Opin Plant
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32. Rudolph U, Schnepf E: Investigations of the turnover of the
putative cellulose-synthesizing particle ‘rosettes’ within the
plasma membrane of Funaria hygrometrica protonema cells. I.
Current Opinion in Plant Biology 2004, 7:651–660
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33. Chaffey N, Barlow P, Sundberg B: Understanding the role of the
cytoskeleton in wood formation in angiosperm trees: hybrid
aspen (Populus tremula x P. tremuloides) as the model
species. Tree Physiol 2002, 22:239-249.
34. Gardiner JC, Taylor NG, Turner SR: Control of cellulose synthase
complex localization in developing xylem. Plant Cell 2003,
15:1740-1748.
This article verifies the close association between microtubules and the
specific cellulose synthase (CesA) enzymes that are involved in the
secondary wall thickenings of xylem elements.
35. Dejardin A, Leple JC, Lesage-Descauses MC, Costa G, Pilate G:
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comparative analysis from multiple libraries. Plant Biol (Stuttg)
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36. Salnikov VV, Grimson MJ, Delmer DP, Haigler CH: Sucrose
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37. Fisher DD, Cyr RJ: Extending the microtubule/microfibril
paradigm — cellulose synthesis is required for normal cortical
microtubule alignment in elongating cells. Plant Physiol 1998,
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38. Lazzaro MD, Donohue JM, Soodavar FM: Disruption of cellulose
synthesis by isoxaben causes tip swelling and disorganizes
cortical microtubules in elongating conifer pollen tubes.
Protoplasma 2003, 220:201-207.
Pollen-tube growth in conifers is dramatically slower than that in angiosperms, but this allows some interesting analysis of tip growth. In this
study, the inhibition of cellulose synthesis generates aberrant microtubule
organization, supporting the idea that microtubules gain information for
their organization from the cell wall.
39. Chan J, Calder GM, Doonan JH, Lloyd CW: EB1 reveals mobile
microtubule nucleation sites in Arabidopsis. Nat Cell Biol 2003,
5:967-971.
This study, along with that described in [41], demonstrates that the End
Binding1 (EB1) proteins bind to the plus ends of cortical microtubules in
plant cells. Both studies use high expression levels of EB1, so some
caution is warranted in interpreting the unusual localizations documented. Follow-up studies using immunolocalization and mutational analysis
are required.
40. 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.
A heterologous fusion protein construct is used to great effect to identify
microtubule plus ends in this analysis of microtubule dynamics.
41. Mathur J, Mathur N, Kernebeck B, Srinivas BP, Hülskamp M:
A novel localization pattern for an EB1-like protein links
microtubule dynamics to endomembrane organization.
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42. Shaw SL, Kamyar R, Ehrhardt DW: Sustained microtubule
treadmilling in Arabidopsis cortical arrays. Science 2003,
300:1715-1718.
A modified form of treadmilling is detected for the first time in living plant
cells. In this study, the authors reveal that both the leading and lagging
ends of microtubules can undergo intermittent growth and shortening.
43. Vos JW, Dogterom M, Emons AM: 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.
A careful comparative analysis of microtubule behavior reveals the
reorganization of the cortical array in readiness for cell division. This
analysis suggests that microtubule turnover is an effective remodeling
mechanism in the formation of parallel cortical microtubule arrays.
44. Mathur J, Hülskamp M: Microtubules and microfilaments in cell
morphogenesis in higher plants. Curr Biol 2002, 12:R669-R676.
45. Mayer U, Jürgens G: Microtubule cytoskeleton: a track record.
Curr Opin Plant Biol 2002, 5:494-501.
46. Furutani I, Watanabe Y, Prieto R, Masukawa M, Suzuki K, Naoi K,
Thitamadee S, Shikanai T, Hashimoto T: The SPIRAL genes are
www.sciencedirect.com
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Arabidopsis thaliana. Development 2000, 127:4443-4453.
47. Hashimoto T: Molecular genetic analysis of left–right
handedness in plants. Philos Trans R Soc Lond B Biol Sci 2002,
357:799-808.
48. 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.
49. 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.
These studies reveal that SPR1/SKU6 is a 12-kDa plant-specific protein
that, according to SPR1–green fluorescent protein (GFP) fusion expression, associates with the plus ends of microtubules at all stages of the cell
cycle. spr1/sku6 mutants have slightly abnormal microtubule orientation
patterns in root epidermal cells. The association of SPR1 with microtubules may be indirect, however, as SPR1 fails to co-sediment with
microtubules.
50. Konishi M, Sugiyama M: Genetic analysis of adventitious root
formation with a novel series of temperature-sensitive
mutants of Arabidopsis thaliana. Development 2003,
130:5637-5647.
51. Thitamadee S, Tuchihara K, Hashimoto T: Microtubule basis for
left-handed helical growth in Arabidopsis. Nature 2002,
417:193-196.
52. 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.
This study demonstrates that the left-twisting phenotypes previously
described for single lefty mutants represent a partial loss of growth
anisotropy. The more severe phenotypes of the lefty1 lefty2 double
mutant demonstrate how altered microtubule dynamics is an important
regulator of wall mechanical properties.
53. Chuong SD, Good AG, Taylor GJ, Freeman MC, Moorhead GB,
Muench DG: Large-scale identification of tubulin binding
proteins provides insight on subcellular trafficking, metabolic
channeling, and signaling in plant cells. Mol Cell Proteomics
2004, in press.
This premier, large-scale proteomic search for tubulin-binding proteins
identified an astonishing 122 proteins from cultured cells of Arabidopsis
thaliana. Importantly, this work verifies just how congested the microtubule surface is. It also suggests regulatory functions for microtubules in
diverse processes, including gene expression, protein translation, signaling, metabolism and cell-wall modification.
54. Wasteneys GO: Microtubules show their sensitive nature.
Plant Cell Physiol 2003, 44:653-654.
55. Abdrakhamanova A, Wang QY, Khokhlova L, Nick P: Is
microtubule disassembly a trigger for cold acclimation?
Plant Cell Physiol 2003, 44:676-686.
The authors go on from the observation that microtubule arrays are
transiently disrupted in freeze-tolerant wheat cultivars during chilling
treatments to demonstrate that a transient depolymerization of cortical
microtubules in freeze-sensitive cultivars will stimulate freeze-tolerance.
56. Komis G, Apostolakos P, Galatis B: Hyperosmotic stress
induces formation of tubulin macrotubules in root-tip cells of
Triticum turgidum: their probable involvement in protoplast
volume control. Plant Cell Physiol 2002, 43:911-922.
57. Eun SO, Youn HS, Lee Y: Lead disturbs microtubule
organization in the root meristem of Zea mays. Physiol Plant
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58. Schwarzerova K, Zelenkova S, Nick P, Opatrny Z:
Aluminum-induced rapid changes in the microtubular
cytoskeleton of tobacco cell lines. Plant Cell Physiol 2002,
43:207-216.
59. Dovgalyuk A, Kalynyak T, Blume YB: Heavy metals have a
different action from aluminium in disrupting microtubules
in Allium cepa meristematic cells. Cell Biol Int 2003,
27:193-195.
www.sciencedirect.com
60. Sivaguru M, Pike S, Gassmann W, Baskin TI: Aluminum rapidly
depolymerizes cortical microtubules and depolarizes the
plasma membrane: evidence that these responses are
mediated by a glutamate receptor. Plant Cell Physiol 2003,
44:667-675.
The authors of this study make an intriguing connection between glutamate receptor activity and aluminum exposure. They provide evidence
that glutamate efflux is a primary response to aluminum treatment, a
treatment that also results in membrane depolarization and microtubule
disorganization.
61. Blancaflor EB, Jones DL, Gilroy S: Alterations in the
cytoskeleton accompany aluminum-induced growth inhibition
and morphological changes in primary roots of maize.
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62. Binet MN, Humbert C, Lecourieux D, Vantard M, Pugin A:
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63. Sedlarova M, Binarova P, Lebeda A: Changes in microtubular
alignment in Lactuca spp. (Asteraceae) epidermal cells
during early stages of infection by Bremia lactucae
(Peronosporaceae). Phyton Annales Rei Botanicae 2001,
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64. Mims CW, Rodriguez-Lother C, Richardson EA: Ultrastructure of
the host–pathogen interface in daylily leaves infected by the
rust fungus Puccinia hemerocallidis. Protoplasma 2002,
219:221-226.
65. Takemoto D, Jones DA, Hardham AR: GFP-tagging of cell
components reveals the dynamics of subcellular reorganization in response to infection of Arabidopsis by
oomycete pathogens. Plant J 2003, 33:775-792.
66. de Almeida Engler J, Van Poucke K, Karimi M, De Groodt R,
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67. Foissner I, Grolig F, Obermeyer G: Reversible protein
phosphorylation regulates the dynamic organization of the
pollen tube cytoskeleton: effects of calyculin A and okadaic
acid. Protoplasma 2002, 220:1-15.
68. Tian GW, Smith D, Gluck S, Baskin TI: Higher plant cortical
microtubule array analyzed in vitro in the presence of the cell
wall. Cell Motil Cytoskeleton 2004, 57:26-36.
This is perhaps the first study to succeed in using the tubulin protofilament hook decoration technique to measure microtubule polarity in the
cortical microtubule arrays of plant cells. It confirms that the cortical array
has microtubules of mixed polarity and shows that microtubules that have
discordant orientation are less stable in these semi-in-vitro assay conditions. This study also shows the involvement of protein phosphorylation
in the regulation of microtubule stability.
69. Gardiner JC, Harper JD, Weerakoon ND, Collings DA, Ritchie S,
Gilroy S, Cyr RJ, Marc J: A 90-kD phospholipase D from tobacco
binds to microtubules and the plasma membrane.
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70. Dhonukshe P, Laxalt AM, Goedhart J, Gadella TW, Munnik T:
Phospholipase D activation correlates with microtubule
reorganization in living plant cells. Plant Cell 2003,
15:2666-2679.
A compelling model for microtubule linkage to the plasma membrane is
developed from studies on the effects of phospholipase D activation.
71. Gardiner J, Collings DA, Harper JD, Marc J: The effects of
the phospholipase D-antagonist 1-butanol on seedling
development and microtubule organisation in Arabidopsis.
Plant Cell Physiol 2003, 44:687-696.
72. Ahad A, Wolf J, Nick P: Activation-tagged tobacco mutants that
are tolerant to antimicrotubular herbicides are cross-resistant
to chilling stress. Transgenic Res 2003, 12:615-629.
73. Reddy VS, Day IS, Thomas T, Reddy AS: KIC, a novel Ca2+
binding protein with one EF-hand motif, interacts with a
microtubule motor protein and regulates trichome
morphogenesis. Plant Cell 2004, 16:185-200.
KIC is identified as an additional contributor to the regulation of KCBP
activity.
Current Opinion in Plant Biology 2004, 7:651–660
660 Cell biology
74. Folkers U, Kirik V, Schobinger U, Falk S, Krishnakumar S,
Pollock MA, Oppenheimer DG, Day I, Reddy AS, Jürgens G
et al.: The cell morphogenesis gene ANGUSTIFOLIA
encodes a CtBP/BARS-like protein and is involved in the
control of the microtubule cytoskeleton. EMBO J 2002,
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76. Wasteneys GO, Galway ME: Remodeling the cytoskeleton
for growth and form: an overview with some new views.
Annu Rev Plant Biol 2003, 54:691-722.
75. Kim GT, Shoda K, Tsuge T, Cho KH, Uchimiya H, Yokoyama R,
Nishitani K, Tsukaya H: The ANGUSTIFOLIA gene of
Arabidopsis, a plant CtBP gene, regulates leaf-cell expansion,
the arrangement of cortical microtubules in leaf cells
77. Fu Y, Li H, Yang ZB: The ROP2 GTPase controls the formation
of cortical fine F-actin and the early phase of directional
cell expansion during Arabidopsis organogenesis. Plant Cell
2002, 14:777-794.
Current Opinion in Plant Biology 2004, 7:651–660
and expression of a gene involved in cell-wall formation.
EMBO J 2002, 21:1267-1279.
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