Journal of Cell Science 115, 1345-1354 (2002)
© 2002 The Company of Biologists Limited
Microtubule organization in the green kingdom: chaos or self-order?
Geoffrey O. Wasteneys
Plant Cell Biology Group, Research School of Biological Sciences, The
Australian National University, GPO Box 475, Canberra ACT 2601,
Australia
(e-mail:
geoffw{at}rsbs.anu.edu.au
)
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Summary
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Plant microtubule arrays differ fundamentally from their animal, fungal and
protistan counterparts. These differences largely reflect the requirements of
plant composite polymer cell walls and probably also relate to the acquisition
of chloroplasts. Plant microtubules are usually dispersed and lack conspicuous
organizing centres. The key to understanding this dispersed nature is the
identification of proteins that interact with and regulate the spatial and
dynamic properties of microtubules. Over the past decade, a number of these
proteins have been uncovered, including numerous kinesin-related proteins and
a 65 kDa class of structural microtubule-associated proteins that appear to be
unique to plants. Mutational analysis has identified MOR1, a probable
stabilizer of microtubules that is a homologue of the TOGp-XMAP215 class of
high-molecular-weight microtubule-associated proteins, and a katanin p60
subunit homologue implicated in the severing of microtubules. The
identification of these two proteins provides new insights into the mechanisms
controlling microtubule assembly and dynamics, particularly in the dispersed
cortical array found in highly polarized plant cells.
Key words: Microtubule-associated protein, Microtubule-organizing centre, Katanin, MOR1, Centrosome, 
-Tubulin, Plant cell
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Introduction
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For over a billion years, plants, animals, fungi and protists have been
acquiring unique characteristics. Microtubules are common to all eukaryotic
kingdoms and are excellent markers of both conservation and specialization. On
the one hand, microtubule fine structure and the tubulin subunits that make up
microtubules are remarkably conserved between kingdoms, as are many
microtubule-associated proteins (MAPs). On the other hand, microtubules are
arranged and organized in highly diverse patterns, relying on a variety of
mechanisms for assembly, orientation and function. We tend to think first
about microtubules as the major structural components of mitotic spindles and
flagellae. In these cases, microtubules emanate from centrosomes,
microtubule-nucleating complexes that are focused around a microtubule-derived
apparatus known as a centriole. Centrosomes can act interchangeably as spindle
poles, anchors for the radial interphase array or basal bodies, in which case
they are referred to as kinetosomes
(Chapman et al., 2000
).
Textbooks tell us that minus ends of microtubules associate with centrosomes
and that their growth by dynamic instability is coordinated by GTP caps at the
fast-growing end. Without centrosomes, the concentration of tubulin subunits
in cells would be far too low to allow microtubule nucleation.
It turns out, however, that centrosomes are a disadvantage for highly
polarized cells. Many microtubules in neurons and epithelial cells are
disconnected from the centrosome, metazoan oocytes have no centrosomes at all
(Megraw and Kaufman, 2000)
and, although yeast spindle pole bodies behave like centrosomes, they lack
centrioles (Balczon, 1996). In
fact, centrosomes are absent from up to half of known eukaryotic species
including most fungi, protists and vascular plants and from the spindles and
interphase arrays of many algae. In vascular and many nonvascular plants,
somatic cells have dispensed with centrosomes altogether
(Vaughn and Harper, 1998).
Among the `higher' seed-producing plants, only two orders of gymnosperm, the
cycads and ginkgoes, retain flagellated sperm
(Southworth and Cresti, 1997).
Dispersed plant microtubule arrays lack tightly focused organizing centres. In
this context, the freedom from centrosomes may be a defining characteristic
that has helped plants to evolve into organisms that are autotrophic and
sessile but highly responsive to their environment.
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Microtubule organization in plants is closely connected to the
special features of cell walls
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To understand microtubule organization in plant cells, it is necessary to
take stock of the full range of arrays. Those arrays found in typical somatic
cells are illustrated in Fig.
1. With the exception of mitotic spindles, whose function in
separating chromosomes and chromatids is conserved, plant microtubule arrays
do things that relate to the development of the plant cell wall and, hence, to
plant cell shape and growth polarity. In the G2 phase of the cell cycle, the
first sign of the approaching mitosis is the preprophase
band*, comprising
prominent bundles of microtubules that generally, but not always, encircle the
midplane (Fig. 1A). Preprophase
bands prepare the eventual site of cell plate attachment, probably by locally
altering the wall properties (Mineyuki,
1999), but disappear long before this event. The recent sighting
of Golgi stacks at the division site during metaphase
(Nebenfuhr et al., 2000)
supports this idea. The signalling mechanisms that determine preprophase band
location, particularly in asymmetrically dividing cells, which are critical
for cell fate determination and tissue differentiation, remain a mystery. Once
determined, however, a tight connection between the preprophase band and
nucleus, via the microtubule- and actin filament-containing
phragmosome*, seems
essential for consolidating the preprophase band
(Gunning and Wick, 1985).
Using a microtubule-binding fluorescent reporter in living cells, Granger and
Cyr recently confirmed that nuclear positioning is critical to downstream
mitotic events (Granger and Cyr,
2001). They propose that the phragmosome-PPB complex plays a key
role in the proper positioning of the preprophase nucleus, which in turn
influences orientation of the mitotic and cytokinetic structures.
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Fig. 1. These schematic illustrations, rendered in 3D at two aspects, show
microtubule arrays through the plant cell cycle. (A) A preprophase band,
linked to the nucleus by phragmosome microtubules, marks the future division
site. (B) Metaphase spindle with a dispersed polar region. (C) In telophase,
the phragmoplast forms as a concentrated cylinder of microtubules between
daughter nuclei. (D) The cytokinetic phragmoplast expands centrifugally,
leading the cell plate towards attachment sites previously established by the
preprophase band. Microtubule plus ends meet at midplane. (E) Once cytokinesis
is complete, microtubules extend from the nucleus toward the cell cortex and
plasma membrane-associated microtubules appear. (F) Plant cells in interphase
and those entering terminal differentiation often expand predominantly in one
direction. During cell elongation, cortical microtubules are usually arranged
in parallel arrays whose predominant orientation is at right angles to the
axis of expansion.
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Dissolution of the preprophase band and nuclear envelope coincides with
formation of the mitotic spindle (Fig.
1B). Spindle poles are typically broad, not tightly focused as in
centrosome-containing cells. At the anaphase-telophase transition, the
phragmoplast
forms (Fig. 1C,D). Like
spindles, phragmoplasts are bipolar complexes with their plus ends meeting at
the midplane. They direct the transport of Golgiderived vesicles towards the
centrifugally expanding cell plate, which matures to become the cross-wall
separating daughter cells (Otegui and
Staehelin, 2000a). Phragmoplast microtubules originate as a
compact cylindrical bundle between the condensing chromatin of daughter nuclei
(Fig. 1C), but gradually form a
self-organizing double ring that increases in circumference in pace with the
cell plate as it expands towards the parent wall
(Fig. 1D). Phragmoplasts are
also part of the cytokinetic apparatus during cellularization of syncytial
cells, which include
endosperm
,
microspores
and
the female gametophyte. In these cells, so-called adventitious phragmoplasts
form at the nuclear-cytoplasmic domains, which are defined by the microtubules
radiating from adjacent nuclei (Brown and
Lemmon, 2001a; Otegui and
Staehelin, 2000b). By observing the incorporation of fluorescent
tubulin, it has been determined that phragmoplast microtubules continually add
subunits at the plus ends, while units are lost at the minus ends
(Asada and Shibaoka, 1991).
This treadmilling maintains the GTP cap, ensuring long-term microtubule
survival.
As cells enter interphase or commit to terminal differentiation,
microtubules are abundant at the periphery of the nucleus and appear to
radiate towards the cell periphery (Fig.
1E). This perinuclear microtubule array is transient but real, as
confirmed in living cells expressing a GFP-tubulin fusion protein
(Hasezawa et al., 2000). Soon
after this stage, microtubules are found throughout the cell periphery, often
in parallel order, in close association with the plasma membrane
(Fig. 1F). These cortical
microtubules play a critical role in controlling growth direction, both in
cells that enlarge by diffuse
growth* and in those
that enlarge by tip
growth
(Bibikova et al., 1999;
Geitmann and Emons, 2000).
They also play important roles in generating localized wall ingrowths after
cells stop expanding in, for example, the formation of vascular
tissues
(Chaffey et al., 1997;
Chaffey et al., 2000;) and
transfer cells
(Bulbert et al., 1998;
Singh et al., 1999).
Microtubule involvement in wall formation and cell expansion is too complex a
subject for this Commentary but is discussed in recent review articles
(Baskin, 2001;
Wasteneys, 2000).
The cortical array exemplifies the enigmatic nature of plant microtubules.
Given no clear organizing centres, where do the cortical microtubules assemble
and what determines their orientation? Despite nearly four decades of study,
we know very little about microtubule assembly and orientation in the various
arrays that coordinate the plant cell through division, polar expansion and
terminal differentiation. In the next section, I speculate on how the
acquisition and evolution of plant-specific features have, by necessity,
influenced microtubule organization, and I outline two models for the
self-organization of plant microtubule arrays.
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Chloroplasts, cell walls and a different sort of motility
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It can be argued that the combination of two defining features of plants,
the cell wall and chloroplasts, was the impetus for the evolution of dispersed
microtubule organization. For chloroplasts to be retained in cell lineages,
primordial chloroplast division needed to be coordinated with that of the host
cell. Many primitive plants including some algae, all bryophytes, certain
vascular cryptogams and at least one group of ferns, have only one large
chloroplast per cell. During division, the plastid surface serves as a
microtubule-organizing centre for the formation of midzone and spindle
microtubules (Brown and Lemmon,
2001b). The need to couple monoplastidic division with cytokinesis
in ancestors of higher plants may have necessitated the abolition of the
nucleus-associated centrosome. Plant cells in more advanced, multiplastidic
vascular plants lack plastid-associated microtubule-organizing centres but
retain the legacy of microtubule-organizing centres that extend beyond the
confines of the nucleus.
The other feature distinguishing plants is of course the cell wall.
Coordinated synthesis and loosening of the largely polysaccharide wall
material to turgor pressure drives plant cell expansion. But this type of
growth requires the bulk of cell volume, which can be considerable, to be
occupied by turgor-regulating vacuoles. As posited by Gunning,
microtubule-based intracellular motility might be inadequate for the metabolic
requirement for cytoplasmic mixing in highly vacuolated plant cells
(Gunning, 1999). This, along
with the need to position chloroplasts optimally in relation to light sources
(Liebe and Menzel, 1995;
Kandasamy and Meagher, 1999),
is likely to have led to actomyosin becoming the dominant motile system in
plant cells. Actin cables provide tracks for movement of myosin-coated
vesicles, endoplasmic reticulum and other organelles, which in animal cells
are largely moved about by microtubule-dependent motors
(Boevink et al., 1998). At
speeds up to 100 µm per second, myosin-driven movement in plant cells is in
a class of its own. There is surprisingly little evidence that plant
microtubules participate in active transport and cytoplasmic streaming but,
unlike in animal cells, microtubules, not actin filaments, are the dominant
element at the plasma membrane. The concept of a
cytoskeleton—plasma-membrane—cell-wall continuum in plant cells
(Wyatt and Carpita, 1993)
necessarily emphasizes the microtubule component.
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Motor proteins organize microtubule converging centres
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If actomyosin-based activity drives much of the intracellular motility,
what are all the kinesin-related proteins that have been identified doing? One
emerging concept is that the ability to organize decentralized microtubule
arrays depends to a large extent on microtubule self-organization. This may be
coordinated largely by the activities of motor proteins, which in plants
include a remarkable variety of minus- and plus-end-directed kinesins
(Liu and Lee, 2001). In
phragmoplasts, the relative activities of several kinesin-related proteins
coordinate the extent of overlapping between antiparallel microtubules at the
midplane (Liu and Lee, 2001;
Lloyd and Hussey, 2001).
Analysis of microtubule patterns in the multinucleate endosperm tissue led to
the concept that microtubule-converging centres link microtubule minus ends to
control the predominant direction of elongation and shortening of microtubule
arrays (Smirnova and Bajer,
1994; Smirnova and Bajer,
1998). Converging centres no doubt also operate at the minus ends
of phragmoplast microtubules to maintain the integrity of the array as it
expands centrifugally.
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Do cortical microtubules grow on fractal trees?
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A variation on the convergence mechanism seems to operate in the formation
of plasma-membrane-associated arrays in walled cells during interphase and
terminal differentiation. Analyzing cortical microtubule patterns after
drug-induced disassembly (Fig.
2) led to the `branching cluster' model for cortical microtubule
assembly in plant cells (Wasteneys and
Williamson, 1989a; Wasteneys,
1992). According to this model, microtubule-initiating factors
move along existing microtubule `tracks' and nucleate the assembly of new
microtubules, which diverge from pre-existing microtubules at acute angles.
The model predicts that plus-end-directed motors disperse initiating factors
along microtubule tracks. The extent of motor activity is likely to be coupled
to the rate of cell expansion; non-growing cells form tightly focused
clusters, whereas rapidly expanding cells re-form cortical microtubules in
more open, fractal tree patterns
(Wasteneys and Williamson,
1989a). This form of self-organized assembly provides a plausible
mechanism for generating even dispersal of microtubule-initiating factors in
expanding cells.
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Fig. 2. Cortical microtubule recovery patterns after drug-induced microtubule
disassembly. (A) Microtubules appear to diverge from the initial assembly
site, forming fractal tree-shaped clusters, with microtubules diverging from
each other at acute angles (figure adapted from
Wasteneys and Williamson,
1989b). (B) Clusters eventually break up. (C) Later in recovery,
parallel microtubule order begins to consolidate but some branching
configurations and discordant microtubules persist. Bar, 10 µm.
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Very recent discoveries of microtubule-associated proteins (MAPs) in plants
provide the first opportunity to test these self-organization models. In the
remainder of this article, the properties of two of these newly identified
MAPs are detailed and I discuss their potential roles in microtubule
organization and function, emphasizing the establishment of the cortical
microtubule array at the onset of cell expansion.
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Molecular approaches have identified several plant
microtubule-organizing and accessory proteins
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Microtubule-associated proteins (MAPs) that nucleate, stabilize and
destabilize, and crosslink and anchor microtubules are all required to
organize plant microtubules. The identity of these so-called structural MAPs
remained elusive for some time in plant cells despite evidence for their
presence in early electron micrographs
(Hardham and Gunning, 1978).
Extracting and identifying MAPs is hindered by the recalcitrance of plant
material to biochemical approaches. Whereas MAPs are relatively easy to purify
from brain tissue, which is teeming with microtubules, plant tissues have
modest concentrations of microtubule proteins. Plant cells are usually
cytoplasm-poor, most cell volume being occupied by vacuoles, whose rupture
releases proteolytic enzymes that hamper purification strategies.
In contrast, genomic and proteomic approaches have proved to be very
effective. Plant tubulins, the building blocks of microtubules, were quickly
identified on the basis of their conserved amino acid sequences
(Hussey et al., 1990;
Kopczak et al., 1992), as were
-tubulin (Liu et al.,
1994) and numerous kinesin-like motor proteins
(Asada and Collings, 1997).
Homology searches for plant structural MAPs were at first largely
unsuccessful, although some candidates, isolated by microtubule affinity,
crossreact with antibodies to the animal MAPs tau
(Vantard et al., 1991) and
MAP4 (Maekawa et al., 1990;
Higashiyama et al., 1996). As
noted by Lloyd and Hussey, homology-based approaches are useful but do not
uncover weakly related proteins, or novel proteins, which may not be
recognized as having MAP function (Lloyd
and Hussey, 2001).
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A plant-specific MAP?
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MAP65 was first isolated from cytoplasmic extracts of evacuolated
protoplasts (Chang-Jie and Sonobe,
1993). Patrick Hussey and colleagues recently isolated three
clones from a tobacco BY-2 cDNA library using antibodies raised to
biochemically isolated protein (Smertenko
et al., 2000). Detailed analysis of one clone, NtMAP65-1,
suggests that its product may be involved in the overlapping of anti-parallel
microtubules in the spindle and phragmoplast. This possibility suggests that
the MAP65 protein works with several kinesin-related proteins to establish and
regulate contact between antiparallel phragmoplast microtubules at the
midplane (Lloyd and Hussey,
2001). NtMAP65-1-specific antibodies also label the preprophase
band and a subset of cortical microtubules
(Smertenko et al., 2000), and
MAP65 isolated from carrot cultures forms regular crossbridges between
adjacent microtubules in vitro (Chan et
al., 1999). These results indicate that MAP65 also maintains
inter-microtubule spacing in preprophase bands and interphase microtubule
arrays. According to its sequence, the 65 kDa family of structural MAPs
appears to be unique to plants.
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Mutational analysis
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In recent years, several groups have adopted mutational approaches to
understand microtubule assembly and function. A collection of
cytoskeleton-defective mutants, identified mainly in thale cress
(Arabidopsis thaliana) and maize (Zea mays), is now
available (Kost et al., 1999;
Wasteneys, 2000;
Azimzadeh et al., 2001). Two
new mutant loci recently joined the collection. One locus encodes a homologue
of the microtubule-severing protein, katanin p60 subunit
(Bichet et al., 2001;
Burk et al., 2001;
McClinton et al., 2001). The
other encodes a member of the TOGp-XMAP215-Dis1 family of high molecular
weight MAPs (Whittington et al.,
2001). The discovery of these mutants, whose phenotypes provide
strong clues about the function of katanin p60 and MOR1 proteins, provides an
opportunity to update our concepts of microtubule organization in plant
cells.
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Katanin p60 adds severing to the repertoire of mechanisms controlling
cortical microtubule organization
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The Arabidopsis homologue of katanin
P60*, AtKSS
McClinton et al., 2001), was
independently identified on the basis of its sequence similarity to the
catalytic subunit of a microtubule-severing protein complex
(McNally, 2000;
Quarmby, 2000). Mutant alleles
including botero1 (bot1)
(Bichet et al., 2001) and
fragile fibre 2 (fra2)
(Burk et al., 2001) were
identified as radially swollen semi-dwarfs with brittle stems. Cortical
microtubule arrays were later found to be disordered, especially during the
onset of post-cytokinetic cell expansion. The katanin P60 subunit is believed
to sever microtubules near their minus ends in neurons
(Ahmad et al., 1999), flagellae
(Lohret et al., 1999) and
mitotic cells (McNally and Thomas,
1998; McNally et al.,
2000). At animal cell spindle poles, severing may increase the
efficiency of the centrosomal release reaction to allow depolymerization of
microtubule minus ends, a force-producing process that may drive anaphase
chromosome movements (Desai et al.,
1998). In plants, a similar role in spindle and phragmoplast
microtubule treadmilling would be anticipated but mitosis and cytokinesis are
apparently unaffected in the bot1 and fra2 mutants, which
include null alleles (Bichet et al.,
2001; Burk et al.,
2001).
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Severing microtubule minus ends to create microtubule-nucleating
templates — why transplant an old tree when you can disperse a
seed?
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Studies of the function of the katanin P60 subunit in neurons suggest that
it is required for severing microtubules from centrosomes
(Ahmad et al., 1999) so that
they can be translocated into the axons. There is some skepticism about this
mechanism (Hollenback and Bamburg,
1999), a study by Chang et al. finding no evidence for microtubule
translocation in axons (Chang et al.,
1999). A similar debate continues in the plant microtubule
community (Stoppin et al.,
1994; Canaday et al.,
2000). Clearly the nuclear surface is one of the preferred sites
for microtubule assembly but cortical assembly has also been verified in
stringent, semi-in-vitro polymerization assays
(Wasteneys et al., 1989;
Kumagai et al., 1999) and in
studies of microtubule recovery after disassembly
(Cleary and Hardham, 1987;
Falconer et al., 1988;
Wacker et al., 1988;
Galway and Hardham, 1989;
Wasteneys and Williamson,
1989a; Wasteneys et al.,
1993). If perinuclear microtubules were the exclusive source of
cortical microtubules, translocation activity should be obvious in
observations of fluorescent microtubules in living cells. In the absence of
such evidence (Wasteneys et al.,
1993; Hush et al.,
1994; Yuan et al.,
1995; Kropf et al.,
1997; Marc et al.,
1998; Himmelspach et al.,
1999; Ueda et al.,
1999; Kumagai and Hasezawa,
2001), I propose that katanin p60 works directly within the
cortical array (Fig. 3).
Periodic severing of the minus ends of microtubules could generate
microtubule-nucleating templates. Dispersing these templates by tracking along
existing microtubules is consistent with the branching assembly model
(Wasteneys, 1992;
Wasteneys and Williamson,
1989a). Templates could be in the form of -tubulin
lock-washer complexes (Moritz et al.,
1995), and this prediction is consistent with the apparent
dispersal of -tubulin along cortical microtubules in plant cells
(Liu et al., 1993;
McDonald et al., 1993;
Liu et al., 1994;
Stoppin-Mellet et al., 2000).
Producing microtubule-nucleating templates by severing microtubule minus ends
would generate a continual supply of microtubules with consistent
protofilament number for the growing cell. Dispersing a seed is easier than
moving a mature tree. Polarized animal cells could have adopted an analogous
strategy for dispersing microtubules.
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Fig. 3. Model for microtubule assembly by severing and transport of nucleating
templates. In this model, a -tubulin ring complex associates with the
minus end of a microtubule, while the microtubule extends by the addition of
tubulin subunits at the fast-growing, GTP-tubulin-containing plus end (dark
green). Severing of the minus end is achieved by the formation of a hexamer of
katanin p60 subunits, whose association with the microtubule wall is
coordinated by the larger p80 subunit, which may transiently dimerize with the
p60 subunits. Microtubule-mediated ATPase activity results in inward movement
of the p60 subunits, an action that cleaves the ring complex from the
microtubule minus end. Katanin subunits dissociate but the lock-washer-shaped
ring complex is transported along the microtubule by a plus-end-directed
kinesin. The extent of transport along the microtubule may be regulated by the
relative activities of plus- and minus-end-directed kinesins. The ring complex
serves as a template for the assembly of additional microtubules. Repeated
generation, severing and transport of nucleating templates at the minus end of
the original microtubule may explain how the fractal tree complexes shown in
Fig. 2A develop.
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MOR1 is a homologue of the TOGp-Dis1 class of high molecular weight
MAPs, and is essential for cortical microtubule organization
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Templated nucleation may explain microtubule assembly in remote parts of
the cell but it does not explain how microtubules are stabilized at these
sites. How, for example, are microtubules organized into the transversely
oriented arrays that typify elongating cells? Reorientation of recently
assembled microtubules by motor MAPs is one feasible mechanism but various
observations of microtubules in living cells indicate that microtubules
assemble at new orientations rather than being actively reoriented
(Yuan et al., 1994;
Yuan et al., 1995;
Wymer et al., 1996;
Himmelspach et al., 1999;
Ueda and Matsuyama, 2000). The
alternative selective stabilization (or biased turnover) model proposes that
only those microtubules assembled in appropriate orientations are stabilized
(Wasteneys and Williamson,
1989b).
The recent discovery that a high molecular weight MAP, called microtubule
organization 1 (MOR1), has an essential function in organizing cortical
microtubules provides a first glimpse at the mechanisms involved in
stabilizing cortical microtubules
(Whittington et al., 2001).
MOR1 was identified by immunofluorescence microscopy screens for
temperature-sensitive microtubule disruption in chemically mutagenized
Arabidopsis seedlings. Two mor1 alleles, whose phenotypes
are similar, have normal cortical microtubule organization and growth at the
permissive temperature of 21°C. At 29°C, microtubules rapidly shorten
and lose their usual parallel alignment in expanding cells
(Fig. 4). Consequently, cells
lose control over their expansion direction. We isolated no null alleles or
mutants with constitutive phenotypes, which suggested that the MOR1
gene, which encodes a calculated 217 kDa protein that has significant sequence
similarity to the XMAP215-TOGp-Dis1 class of structural MAPs
(Whittington et al., 2001),
is vital and non-redundant.
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Fig. 4. Microtubule patterns in the epidermis of Arabidopsis thaliana
cotyledons after 4 hours at 29°C. (A) Cortical microtubules are abundant
and transversely oriented in wildtype. (B) In the mor1 mutant,
microtubules appear short and disoriented. Bar, 10 µm.
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A mor1 allele affects polarization and cell division in male
and female gametophytes
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Although mor1 mutants have normal mitotic and cytokinetic
microtubule arrays, and cell division is unaffected at restrictive
temperatures, this does not rule out a function for MOR1 in cell division.
Twell and Park recently reported that the gemini pollen mutant
phenotype (Park et al., 1998;
Park and Twell, 2001b) is
complemented by the wild-type MOR1 gene
(Eckardt et al., 2001;
Park and Twell, 2001a).
Homozygous gem1 mutants are lethal, supporting a non-redundant
function for MOR1 in Arabidopsis thaliana. In gem1 mutant
lines, aberrant cell division planes in haploid microspores result in a
proportion of sperm-cell-free pollen. Polarization in the female gametophyte
is also altered. These findings suggest that MOR1 organizes microtubules
throughout the cell cycle, in common with all its known homologues
(Wang and Huffaker, 1997;
Chen et al., 1998;
Matthews et al., 1998;
Cullen et al., 1999;
Charrasse et al., 2000;
Dionne et al., 2000;
Graf et al., 2000).
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A temperature-sensitive HEAT repeat may be a key to MOR1 function in
the cortical array
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What is so peculiar about the proteins encoded by the two mor1
mutant alleles that they cause temperature-dependent disruption of interphase
microtubules but not preprophase bands, spindles or phragmoplasts? We
determined that both mutant alleles introduce single amino acid substitutions
in an N-terminal HEAT repeat, one of at least ten HEAT repeats found in MOR1
(Whittington et al., 2001).
Several lines of evidence suggest that conformational changes at the higher
temperature cause the altered microtubule behaviour
(Whittington et al., 2001).
Both disorganization and recovery of cortical microtubules in the
mor1 mutants occur within minutes of the temperature shift, a time
scale too rapid for major transcriptional responses to take effect. In support
of this, RT-PCR analysis demonstrates that mutant transcript is produced at
the restrictive temperature. Finally, the specific amino acid substitutions
found in the mor1 mutants are the sort likely to compromise the
function of HEAT repeats. In mor1-1, phenylalanine replaces leucine
at position 174 and, in mor1-2, lysine replaces glutamic acid at
position 195. Substituting a bulkier, aromatic residue or a charge change may
well alter the structure of HEAT repeats, which are typified by a pattern of
hydrophobic and hydrophilic residues, giving rise to two antiparallel
-helices connected by a short loop
(Kobe et al., 1999). HEAT
repeats have now been found in many different proteins, which have a variety
of roles, including nuclear protein import/export, vesicle trafficking,
regulation of phosphorylation and plasma membrane associations
(Kunz et al., 2000). Thus, a
conformational change in a HEAT repeat, which may be exacerbated at higher
temperatures, is likely to reduce or eliminate a specific protein-protein
interaction. Future studies will concentrate on characterizing the specific
changes that occur at this functional motif at the restrictive
temperature.
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What is the function of this HEAT repeat in the MOR1 protein?
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The N-terminal HEAT repeat targeted by the two mor1 mutations is
also found in homologues of MOR1 (Lemos et
al., 2000; Cassimeris et al.,
2001). This presents a conundrum. Does this motif mediate a
function that is specific to cortical arrays, a specialization of plant cells,
or does it have a more general function? Cortical microtubules are exclusively
affected by the mor1 mutations; so the N-terminal HEAT repeat could
connect microtubules to sites at the plasma membrane, as shown in
Fig. 5A. This is consistent
with evidence from Xenopus (Popov
et al., 2001) and Dictyostelium
(Graf et al., 2000) homologues
that the C-terminus binds the MAP to microtubules whereas the N-terminal
domain has an alternative function. However, one difficulty with this idea is
that the HEAT repeat is highly conserved in all homologues, many of which have
no obvious association with the plasma membrane.
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Fig. 5. Possible functions of the MOR1 HEAT repeat-1 (HR1) in microtubule
stabilization. (A) HR1 links microtubules to the plasma membrane via a
plasma-membrane-associated protein. At restrictive temperature, this loss of
binding dissociates microtubules from the plasma membrane, promoting their
destabilization. (B) HR1 competes with a destabilizing protein (probably a
kin1-like kinesin) for binding. At permissive temperature, the high affinity
of MOR1 for this site prevents destabilization. At 29°C, this affinity is
lost, leading to kin1-dependent destabilization and microtubule
shortening.
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Alternatively, the N-terminal HEAT repeat could be a critical part of a
general microtubule-stabilizing mechanism
(Fig. 5B). The Xenopus
homologue, XMAP215, has N-terminal-specific microtubule-stabilizing activity
(Popov et al., 2001) and works
in balance with XKCM1, a kin-1-class kinesin-related protein that opposes
microtubule stabilization by XMAP215
(Tournebize et al., 2000).
Could the N-terminal HEAT repeat identified in the mor1 mutants
mediate access of a kin-1 kinesin to microtubules? As outlined by Hussey and
Hawkins, the N-terminal HEAT repeat of MOR1 could either interact directly
with the destabilizing kinesin to modulate its microtubule binding or work
less directly by competing for a common microtubule-binding site
(Hussey and Hawkins,
2001).
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Is there more than one MOR1?
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How is MOR1 regulated during the cell cycle? Phosphorylation by increased
cyclin-dependent kinase (CDK) activity is anticipated to reduce the affinity
of MOR1 for microtubules, producing shorter, more dynamic microtubules
(Vasquez et al., 1999).
According to Hussey and Hawkins, candidate CDK targets are found in the MOR1
sequence but experimental confirmation awaits
(Hussey and Hawkins, 2001).
Another way to achieve differential regulation is through alternative splicing
to generate a cell-cycle-specific isoform that is unaffected by the mutations.
The MOR1 gene has 52 introns; thus, it is a good candidate for
transcriptional modification. At least two isoforms of XMAP215, expressed in
different developmental stages, have been identified
(Becker and Gard, 2000).
Southern blotting confirms that the Arabidopsis thaliana genome has
only one copy of MOR1
(Whittington et al., 2001).
However, a protein related to the XMAP215-TOGp family that conserves the HEAT
repeat affected by the mor1 mutations has been discovered in
Drosophila (Lemos et al.,
2000). A putative MAST homologue exists in Arabidopsis
thaliana, and my group is currently investigating the possibility that it
overlaps functionally with MOR1 during the cell cycle.
Determining expression, intracellular distribution and regulation of MOR1
through the cell cycle is an important part of its characterization and likely
to provide significant new information on how microtubules are organized
through this crucial morphogenetic process.
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Conclusion/perspectives
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Without centrosomes, plant microtubule arrays are largely self-organized by
the relative activities of microtubule-associated proteins. The recent
discoveries of katanin p60 and MOR1 provide important clues about how
microtubule assembly and stabilization are achieved, particularly in the
cortical arrays that regulate growth polarity and cell wall deposition in
interphase and terminally differentiating cells. I have put forward the
hypothesis that dispersed microtubule arrays are generated by the severing of
nucleating complexes from microtubule minus ends, and the transport of these
complexes along existing microtubules. The N-terminal HEAT repeat altered in
the two temperature-sensitive mor1 mutants, and conserved in all
eukaryotic homologues of MOR1, is likely to have a key role in the general
mechanism of microtubule stabilization. One goal of future research will be to
elucidate the exact function of this motif. The discovery of MAPs and
microtubule regulatory proteins in plants is still at an early stage. An
essential first step towards understanding how microtubule arrays are
constructed is the isolation of mutants such as bot1/fra2
(Bichet et al., 2001;
Burk et al., 2001),
mor1/gem1 (Park and Twell,
2001b; Whittington et al.,
2001), zwichel
(Oppenheimer et al., 1997),
spiral 1 and 2 (Furutani
et al., 2000), ton
(Traas et al., 1995),
fass (Torresruiz and Jurgens,
1994; McClinton and Sung,
1997) and tan (Smith
et al., 2001). It is now possible to start finding out how plant
MAPs interact and how kinases and phosphatases regulate their activity. This
work will involve classical genetics and more contemporary functional
genomics. Phenotyping double mutants will provide the first clues, and
suspected interactions will be tested by double immunolocalizations,
co-sedimentation assays, in vitro assays, or more elegantly by fluorescence
resonance energy transfer (FRET) in living cells. After four decades of study,
we may finally start to grapple with the real mysteries of plant microtubules.
How are they assembled with such precision, what determines their orientation,
and just how does this orientation control the direction of cell
expansion?
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Acknowledgments
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I am especially grateful to Moira Galway (St Francis Xavier University) for
many helpful discussions and critical reading throughout the preparation of
this manuscript. I thank Brian Gunning (ANU), Ilse Foissner (University of
Salzburg) and Dave Twell (University of Leicester) for their helpful insights
and Angela Whittington, Ke Jun Wei and Madeleine Rashbrooke from my lab, for
their analysis of MOR1's sequence. Finally, I acknowledge Thomas Magill and
Catherine Eadie for expert production of
Fig. 1 and
Fig. 3, respectively.
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Footnotes
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Summary
Introduction
