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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Key words: Microtubule-associated protein, Microtubule-organizing centre, Katanin, MOR1, Centrosome, -Tubulin, Plant cell
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Introduction |
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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 |
---|
|
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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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
cytoskeletonplasma-membranecell-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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Do cortical microtubules grow on fractal trees? |
---|
|
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 |
---|
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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Mutational analysis |
---|
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Katanin p60 adds severing to the repertoire of mechanisms controlling cortical microtubule organization |
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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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|
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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 |
---|
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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A mor1 allele affects polarization and cell division in male and female gametophytes |
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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 the function of this HEAT repeat in the MOR1 protein? |
---|
|
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? |
---|
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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Acknowledgments |
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Footnotes |
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* Phragmosomes are bands of microtubules and actin filaments that span the
distance between the nucleus and the preprophase band, often through
transvacuolar stands.
Phragmoplasts comprise centrifugally expanding bipolar arrays of
microtubules that initiate as a concentrated bundle in the midzone between the
daughter nuclei at telophase.
A triploid tissue derived from one sperm and two polar nuclei of the female
gametophyte; nourishes the zygotic embryo.
Microspores mature into pollen grains, the male gametophyte, by highly
polarized cell divisions, giving rise in angiosperms to a diffuse vegetative
nucleus and two sperm cells, carried within the pollen tube.
* Diffuse growth involves incorporation of new wall material and
turgor-driven stretching of that material over the entire surface of the cell.
Cells typically expand along one axis that is at right angles to the
predominant microtubule orientation.
In tip-growth, wall loosening and synthesis is localized to one part of the
cell.
Vascular tissues provide conduits for water, nutrients and metabolites and
provide mechanical support to enable land plants to become free-standing
structures of impressive stature.
Transfer cells have wall ingrowths to increase the surface area of plasma
membrane for efficient exchange of nutrients.
* Arabidopsis thaliana katanin-like protein small subunit.
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