Fundamental cytoskeletal activities
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3 Expanding beyond the great divide:
the cytoskeleton and axial growth
Geoffrey O. Wasteneys and David A. Collings
Cell expansion is the single most important process in plant morphogenesis. After
all, cell division makes no direct contribution to organ size and relatively little
contribution to organ shape. Cell division in the absence of cell expansion, as in
endosperm tissue (Olsen, 2001), results simply in cells becoming more numerous
with no net gain in organ size. Conversely, there are examples of remarkably
normal organogenesis when the division process is perturbed or absent altogether.
The alga Acetabularia acetabulum can form exquisitely complex shapes as single
cells (Mandoli, 1998). Even in higher plants there is evidence that organogenesis
can proceed despite serious cell division defects. Mutants defective in cell division
plane alignment, including fass and ton of Arabidopsis (Traas et al., 1995) and
tangled of maize (Smith et al., 2001), still develop organs in the right places even
if they are somewhat deformed. These examples highlight the importance of direc-
tional cell expansion for plant morphogenesis.
One success story of higher plant evolution is the production of inconspicuous
meristems and organ primordia. Meristems harbour many small and cytoplasmic-
ally dense cells that retain the potential to give rise to a wide variety of cell types.
The progeny of these stem cells produce tissue precursors, which then follow a
precise series of divisions to give rise to distinct tissue patterns that identify organ
primordia. As the distance and number of partitions from the meristem centre
increases, the basic patterns of organ primordia become established. Cells eventu-
ally undergo a transition, showing a reduced tendency to divide, and sometimes
pass through a post-mitotic isotropic growth phase. But at a critical threshold,
usually triggered by an abiotic signal, cells undergo massive increases in volume
accompanied by vacuole formation. This inflation generally leads to terminal
differentiation, and often precludes any capacity for further partitioning into new
cells. Moreover, most cells grow predominantly or even exclusively in one direction,
an anisotropic process we refer to as axial growth.
In this chapter, we investigate the cytoskeleton-mediated processes that control
axial growth. Axial growth is distinct from polar growth, which strictly involves
highly localized exocytosis. Tip-growing root hairs and pollen tubes expand by
polar growth (see Chapters 7 and 8). While the normal cytokinetic process is also
highly polarized, it plays an important role in setting up cells for axial growth.
In this chapter, we therefore explore the polar secretion that occurs during
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84 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
cytokinesis, and discuss how this process helps to define the properties of cells that
will undergo anisotropic expansion. We then explore the relationship between the
cytoskeleton and axial cell expansion during interphase and post-mitotic expansion.
These two phases are commonly lumped together as interphase but we stress the
importance, in a developmental context, to distinguish these two events.
3.2 Division planes and the establishment of axiality
We begin by considering the cytoskeleton’s function during cytokinesis, the subject
of several recent reviews (microtubules: Otegui & Staehelin, 2000; Hepler et al.,
2002; microfilaments: Schmit, 2000; Hepler et al., 2002). Cell plate construction
during cytokinesis partitions the contents of post-mitotic cells through the activity
of the substantially cytoskeletal phragmoplast. Unlike the primary cell wall, cell
plates are not subjected to turgor pressure-driven strain deformation. Studying cell
plate and cross wall construction thus provides a unique insight into building a
wall from scratch, a process that does not have an existing template of wall material
to build on and one that does not face immediate rearrangement of these materials
by mechanical stresses. Cell plate formation is also of considerable relevance to
the axial expansion that follows because the division plane is usually perpendicular
to the subsequent axis of expansion, even when cells are partitioned along their
long axis (Green, 1984).
3.2.1 Cell plate formation and expansion
Constructing the typical central-forming cell plate begins with fusion of secretory
vesicles between the separated nuclei and is followed by centrifugal plate expansion
towards the parent cell wall. Preservation by high pressure freezing for transmission
electron microscopy shows plate formation progressing through several distinct
stages, including a tubulo-vesicular network, a smooth tubular network, and a
fenestrated cell plate (Samuels et al., 1995; Bednarek & Falbel, 2002). During
plate expansion, these stages can be seen in a radial gradient in individual cells.
In the tubulo-vesicular stage of cell plate formation, phragmoplast microtubules
associate with the plate’s leading edge. This is where the majority of vesicle fusion
as well as the majority of clathrin-coated vesicles are found (Samuels et al., 1995).
Otegui et al. (2001) calculated that endocytosis removes 75% of the membrane
added to the cell plate during Arabidopsis endosperm syncytial divisions. Both
vesicle delivery and clathrin-coated vesicle recovery continue at low levels until
the fenestrated cell plate stage (Samuels et al., 1995), but microtubules do not
associate with this region of the cell plate. Whether the trafficking mechanism and
vesicle content differ from that during formation of the tubulo-vesicular network is
Wall formation is first detected at the smooth tubular network stage with the
deposition of callose, which remains abundant in the new cross wall until some
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THE CYTOSKELETON AND AXIAL GROWTH 85
time after cytokinesis (Samuels et al., 1995; Hong et al., 2001a). A gene for the
catalytic subunit of callose synthase, CALS1, has been cloned from Arabidopsis, and
the encoded protein distributes in the cell plate when expressed as a GFP fusion
protein in tobacco (Hong et al., 2001a). CALS1 interacts with the cell plate-specific
dynamin, phragmoplastin and a UDP-glucose transferase. This entire complex
might be regulated by the small GTPase ROP1 (Hong et al., 2001b). Significant
amounts of cellulose are synthesized at the fenestrated cell plate stage, before the
establishment of any cortical microtubule array. Cellulose synthesis is required for
the completion of cell plate formation, as its inhibition by the cellulose synthase
inhibitor dichlorobenzonitrile results in wavy or incomplete cell plates (Vaughn
et al., 1996). The korrigan mutant of Arabidopsis shows a similar incomplete
cytokinetic phenotype. The KOR gene encodes an endo-1,4-β-glucanase that is
located at the cell plate (Zuo et al., 2000) and is required for normal cellulose
synthesis. A temperature-sensitive KOR allele, rsw2, has decreased amounts of
cellulose at its restrictive temperature (Lane et al., 2001). These observations sug-
gest that the cellulose in the cell plate contributes to the stiffening process required
for completion of partitioning. However, regulatory factors controlling sequential
callose and cellulose deposition have yet to be identified.
3.2.2 Phragmoplast microtubule and microfilament organization
Hepler etal. (2002) define formation and expansion as two distinct stages of
phragmoplast development. The phragmoplast forms late in anaphase, initially
as a poorly defined mass of microfilaments and microtubules between the daughter
nuclei, but soon develops into a barrel-like structure with well-defined polar-
ity. Microtubules develop into two arrays of opposite polarity that interdigitate
at the division plane, with their fast-growing or plus ends towards the centre
(Asada et al., 1997; Wasteneys, 2002). Microfilaments also develop into two
parallel arrays, interdigitated at the division plane and with the barbed-ends central
(Kakimoto & Shibaoka, 1988). The site of intersection of both the microtubules
and microfilaments is the site of active fusion of Golgi-derived secretory vesicles.
The phragmoplast expands outward until it reaches the approximate diameter of
the two adjacent nuclei (Hepler et al., 2002). The second stage of phragmoplast
growth is its outward expansion from the perinuclear region towards the parent
cell wall. During this expansion, microtubules and microfilaments at the centre of
the phragmoplast depolymerize, so that the phragmoplast assumes a ring-like or
toroidal appearance, with the developing and expanding cell plate at its centre
Subtle differences in the organization of microtubules and microfilaments in the
phragmoplast suggest that mechanisms similar to those in force in other highly
polarized exocytotic events may also operate during cell plate formation. In rapidly
growing pollen tubes, root hairs, and lobe-forming epidermal cells, the cytoplasm
and growth are highly polarized. Microtubules are typically excluded whereas fine
networks of microfilaments are closer to but not in direct contact with the sites of
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86 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
Fig. 3. 1 Microfilament and microtubule organization in the Arabidopsis root meristem. Arabidopsis
roots immunolabelled for tubulin and actin (Collings & Wasteneys, unpublished data) were optically sec-
tioned by confocal microsco py. A–D Optical sections separated by 2.10 µm show extensive cytoskeletal
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THE CYTOSKELETON AND AXIAL GROWTH 87
active secretion (reviewed by Wasteneys & Galway, 2003). Hepler et al. (2002)
state that microtubules and microfilaments ‘show a similar dynamic rearrangement
during cytokinesis’. Just how similar is a difficult but important question to
address. Some studies demonstrate that rearrangements of microtubules and
microfilaments, albeit similar, are not identical. Electron micrographs show that
phragmoplast microtubules are more highly ordered than are phragmoplast micro-
filaments (Kakimoto & Shibaoka, 1988). Various studies also indicate that early
phragmoplast microfilaments do not extend as far from the equatorial plane as
microtubules (Kakimoto & Shibaoka, 1988; Liu & Palevitz, 1992; Zhang et al.,
1993; Collings et al., in press) (Fig. 3.1A–D). Moreover, as the phragmoplast
matures, depolymerization of microtubules may precede the loss of microfilaments
from the centre of the phragmoplast. This generates a microtubule annulus that
surrounds either a ring of microfilaments (Zhang et al., 1993) or a microtubule
annulus that is not as broad as the microfilament annulus (Collings et al., in press).
3.2.3 Motor proteins during phragmoplast formation and expansion
Phragmoplast microtubules and microfilaments are both oriented appropriately to
act as tracks for vesicle transport to the cell plate. Identifying phragmoplast motor
proteins is therefore a pursuit of several research groups. There are many motors to
choose from. Higher plants have a wide variety of motor proteins including micro-
filament-associated myosins and microtubule-associated kinesins. In the Arabidopsis
genome alone, there are some 17 different myosins belonging to two different
plant-specific myosin subfamilies (4 class VIII myosins and 13 class XI myosins;
Berg etal., 2000). Arabidopsis also has at least 61 different kinesins, which,
according to biochemical studies and sequence analysis, represent examples from
most of the numerous subfamilies, including both minus and plus end-directed
kinesins (Lawrence et al., 2002; Reddy & Day, 2002). Motor proteins are likely to
serve two major functions during cytokinesis. By converting energy into directed
movement, they are probably active in transporting vesicles to and from the cell plate.
They can also act as structural proteins by binding to the cytoskeleton in either
Fig. 3.1 (continued)
preservation. Interphase cells in the epidermis and cortex contain random endoplasmic microfilaments
and microtubules (en), and transverse cortical microfilaments and microtubules (t). Dividing cells
contain microfilaments and microtubules in phragmoplasts, where microfilaments form a narrower
band than the microtubules (p), but only microtubules in the spindle (s). Dividing cells also have an
increased density of microfilaments adjacent to the entire plasma membrane, except in the zone of actin
depletion at the division plane. (E), (F) Computer-generated vertical sections, showing radial organ-
ization of the cytoskeleton and its pattern on end walls, were calculated for the locations in the optical
sections marked with short bars (indicated as e and f). The planes of the four optical sections are also
marked in the vertical sections with bars (indicated as a and d). An epidermal cell undergoing mitosis
(between asterisks) shows an extensive microfilament array but lacks microtubules on the end wall,
while an interphase epidermal cell has micro filaments and randomly oriented microtubules at the ends
of the cell (between arrows). The scale bar in (A) represents 20µm in all images.
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88 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
an inactivated state or through cytoskeletal binding domains outside of the motor
regions (see Wasteneys, 2002). For example, the tail regions of six Arabidopsis
kinesins contain the calponin-homology domain that can mediate actin binding
(Reddy & Day, 2002) while the tail of the kinesin-like calmodulin-binding protein,
KCBP, contains a further microtubule-binding domain (Narasimhulu & Reddy,
126.96.36.199 Vesicle transport in the phragmoplast could be kinesin-based
In expanding and differentiating cells of higher plants, the transport of organelles
and vesicles seems to depend solely on actomyosin. With the exception of as yet
unproven roles in vesicle transport in growing pollen tubes (Cai et al., 2000) and in
the movement or positioning of nuclei and chloroplasts (Sato et al., 2001), micro-
tubules apparently do not contribute to vesicle and organelle movements. In the
phragmoplast, the opposite situation may occur, though unequivocal evidence is
Phragmoplast microfilaments are oriented in the correct configuration for
myosin-based vesicle delivery, with their barbed-ends towards the division plane.
Several observations, however, suggest that myosin does not participate in cell
plate construction. Treatments that disrupt actin do not prevent cell plate formation
although plates may become misaligned (see Section 188.8.131.52). Immunolabelling
studies with antibodies specific for either myosin VIII (Reichelt et al., 1999) or
myosin XI (Liu etal., 2001) do not label the developing phragmoplast. This is,
however, in contrast to earlier studies using antibodies raised against a 170 kDa
plant myosin (presumably a myosin XI), which labelled punctate vesicle-like
structures within the phragmoplast (Lin et al., 1994). It remains possible that the
configuration of myosins associated with phragmoplast vesicles obscures epitopes
such that access of antibodies raised to be specific to myosins VIII and/or XI is
prevented. Myosin does, however, play at least one role in organelle transport
during cell division. In certain monocots, notably onion, peroxisomes aggregate at
the division plane in late anaphase, and remain associated with the inner edge of
the microtubule phragmoplast through the later stages of cytokinesis. Aggregation
is actomyosin-dependent, as cytochalasin, latrunculin and BDM all generate
cytokinetic cells with randomized peroxisomes (Collings et al., in press). Peroxi-
some aggregation has not, however, been observed at Arabidopsis cell plates so it
may reflect subtle differences between cell wall formation in different plant taxa.
Evidence for microtubule-specific motor activity in cell plate formation is
stronger. Orientation of phragmoplast microtubules is consistent with vesicle
delivery by plus end-directed motors, and an Arabidopsis kinesin, AtPAKRP2, that
was isolated from phragmoplasts fractionates along with endomembranes and
vesicles. Furthermore, immunolabelling for this kinesin gives a punctate pattern
that concentrates at the division plane, but whose localization is disrupted by
brefeldin treatments, consistent with localization on secretory vesicles (Lee et al.,
2001). Unfortunately, this is the only direct evidence for a role of molecular
motors in the delivery of vesicles to the phragmoplast. The association of vesicles
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THE CYTOSKELETON AND AXIAL GROWTH 89
with microtubules in syncytial cell phragmoplasts of Arabidopsis endosperm,
mediated by kinked, rod-like structures that resemble kinesins when viewed in
high resolution electron micrographs (Otegui et al., 2001), has yet to be corroborated
by either biochemical or immunological data.
184.108.40.206 Structural MAPs and kinesins function in phragmoplast
formation and expansion
Two structural microtubule-associated proteins (MAPs) associate with and probably
have critical roles in organizing the phragmoplast. Antibodies to tobacco MAP65
proteins (NtMAP65-1) label the zone of overlapping phragmoplast microtubules
(Smertenko et al., 2000), suggesting that these proteins act as cross-linkers of
anti-parallel microtubules. Recent work also demonstrates that the 217kDa
MOR1/GEM1 protein associates with phragmoplast microtubules (Whittington
et al., 2001; Twell et al., 2002; see Chapter 1). Whereas cell division continues in
the mor1-1 and mor1-2 temperature-sensitive alleles at restrictive temperature,
the stronger gemini alleles of MOR1 in Arabidopsis are homozygous-lethal and
generate gametophytic defects (Twell et al., 2002). Cell plates do not form
properly in gem1 microspores, suggesting MOR1/GEM1 is essential for normal
phragmoplast formation and function (Twell et al., 2002). A tobacco homologue of
MOR1, named TBMP200, was purified from phragmoplast-enriched extracts and
has been shown to cross-link microtubules in vitro (Yasuhara et al., 2002). MOR1/
GEM1’s distribution in the phragmoplast is still not clear. Antibodies generated to
a C-terminal expression fragment localize to the cell plate zone (Twell et al., 2002)
whereas antibodies generated to an N-terminal peptide localize along the entire
length of phragmoplast microtubules (Wasteneys, unpublished data).
The combination of opposing forces created by plus end- and minus end-
directed motors allows the formation and stabilization of the anti-parallel
microtubule arrays of the phragmoplast (Liu & Lee, 2001). The 125 kDa tobacco
kinesin-related protein, TKRP125, is a bimC kinesin. It was isolated from
phragmoplasts of synchronized tobacco BY-2 cells, and localises to the central
overlapping region. Antibodies against TKRP125 prevent the sliding expansion
of the interdigitated microtubule arrays in in vitro assays (Asada et al., 1997).
TKRP125 homologues occur in all plants examined so far including Arabidopsis
(Liu & Lee, 2001; Lawrence et al., 2002; Reddy & Day, 2002). Spatial and
functional data for TKRP125 are consistent with the activities of other bimC
family members. These kinesins generate an outward force between opposing
arrays of microtubules in the overlapping region of animal spindles and form
homotetramers with motor domains at the opposing ends (Lawrence et al ., 2002).
Another plus end-directed motor found at the phragmoplast midplane in Arabidopsis,
AtPAKRP1, may also contribute to phragmoplast stability (Lee & Liu, 2000;
Lee et al., 2001).
Minus end-directed kinesins from the C-terminal motor subfamily act in the
opposite direction to bimC kinesins to help stabilize the antiparallel array. Such
kinesins, including katA, katB and katC in Arabidopsis, localize to the phragmoplast
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90 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
but have not been characterised biochemically (Mitsui et al., 1993; Liu et al., 1996).
The kinesin-like calmodulin-binding protein (KCBP) also associates with the
phragmoplast (Bowser & Reddy, 1997). This minus end-directed, C-terminal motor
domain kinesin was isolated as a calmodulin-binding protein (Reddy et al., 1996),
and is found in all higher plants examined so far but not in other kingdoms. Motor
activity in KCBP is regulated by calcium/calmodulin, so that in the presence of
calcium, the motor is inactive. Antibodies raised against the calmodulin-binding
domain of KCBP prevent calmodulin-binding, giving a constitutively active motor
even in the presence of calcium. Significantly, microinjection of these antibodies
into Tradescantia virginiana stamen hair cells results in aberrant cell division,
suggesting that KCBP activity is downregulated during telophase (Vos et al., 2000).
KCBP was also identified in screens of Arabidopsis trichome branching mutants,
and plays a role in the control of branching (Oppenheimer etal., 1997), but this
role may be unrelated to its functioning in cell division.
220.127.116.11 Expansion of the phragmoplast and cell plate requires
both kinesins and myosins
Recent analysis has demonstrated that kinesins are also required for phragmoplast
expansion. The seedling-lethal hinkel mutant of Arabidopsis has defective cytokin-
esis that often results in multinucleate cells. HIK encodes a member of a plant-
specific subfamily of kinesins with an N-terminal motor, and is presumed to be
plus end-directed. The HIK gene’s expression is cell-cycle dependent, and HIK is
found only at the phragmoplast midplane. In the hik mutant, the central micro-
tubules of the second stage phragmoplast fail to depolymerize and secretory vesi-
cle localization occurs normally. It is therefore assumed that the HIK kinesin is
required for microtubule turnover rather than vesicle transport (Strompen et al.,
2002). The NACK1 protein of tobacco is a HIK homologue and also associates
with the cell plate. Binding of NACK1 to NPK1, a MAP kinase kinase kinase, is
required for the correct expansion of the phragmoplast and cell plate. Over-expression
of a truncated NACK1 that contains the NPK1-binding domain but lacks the motor
domain, prevents microtubule depolymerization and gives a similar cytokinesis-
defective phenotype (Nishihama et al., 2001, 2002) as does virus-induced silencing
of NACK1 in tobacco, or T-DNA knockouts of the NACK1 homologue (HIK) in
Arabidopsis (Nishihama etal., 2002).
Actomyosin activity is essential for cell plate expansion. Treatments that impair
microfilament organization, such as cytochalasins (Palevitz, 1987), or injections of
the actin monomer-binding protein profilin (Valster et al., 1997), disrupt the
formation and correct alignment of the cell plate, as do treatments with the myosin
antagonists BDM and ML-7 (Molchan et al., 2002). Microfilaments extend
between the phragmoplast and plasma membrane (Endlé et al., 1998; Molchan
et al., 2002; Collings et al., in press). Thus, one of the functions of microfilaments
during cytokinesis seems to be to direct the expansion of the phragmoplast, and
hence the cell plate, to the sites on the parent plasma membrane that are marked by
the zone of microfilament depletion (Schmit, 2000).
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THE CYTOSKELETON AND AXIAL GROWTH 91
3.2.4 Cytoskeletal mutants defective in cytokinesis
Several cytokinesis-defective mutants have been isolated. The four PILZ genes of
Arabidopsis, along with KIESEL, are required for the folding of α- and β-tubulin
and the formation of tubulin dimers. Loss of function alleles of these genes cannot
produce microtubules and, not surprisingly, are blocked in cell division and are
embryo-lethal (Steinborn et al., 2002). Laurie Smith’s group has also characterized
plant cytokinesis mutants from maize, including tangled, discordia, pangloss and
brick, that all show similar phenotypes to those induced by actin disruption
(Gallagher & Smith, 2000). Tangled has defective longitudinal divisions, suggest-
ing that the normal mechanism for specifying specialized division planes does not
operate properly. The TAN protein has microtubule-binding properties and is a
homologue of the adenomatous polyposis coli (APC) tumour-suppressing protein
(Smith et al., 2001). APC proteins are activators of a guanine nucleotide exchange
factor that in turn activates a small GTPase. The discovery of TAN shows that
small GTPases may play roles in specifying division planes in plant cells.
3.3 Setting up for axial growth: distinguishing lateral and end walls
For axial growth, end walls and lateral walls must have distinct physiological and
mechanical properties. End walls expand relatively little or sometimes not at all,
while lateral walls generally extend to become many times their original length.
Arabidopsis root cells have perfect anisotropy, with expansion limited to the lat-
eral walls, and no appreciable expansion of the end walls. In this sense, the last
cross walls to form by transverse divisions never become primary walls and
default immediately to secondary wall status (by definition, secondary walls are
not extensible). End walls are therefore fundamentally different to the lateral walls
of the same cells, and are likely to differ considerably in composition, both of
polysaccharide material and wall proteins that include loosening enzymes. Differ-
ences in the organization and dynamic properties of the cytoskeleton reflect these
3.3.1 The cytoskeleton at end walls of elongating cells
Most studies of cytoskeletal organization during cell elongation have focused on
lateral walls, where the majority of growth occurs. End walls have been largely
ignored, partly because imaging these is problematic. One extensive analysis by
electron microscopy compared microtubule patterns in radially expanding and
non-expanding end walls of Azolla pinnata root primordia. Microtubules were
oriented transverse to the major axis of expansion at end walls but were radially
oriented at non-expanding end walls (Busby & Gunning, 1983). We examined
microtubules and microfilament distribution in elongating characean algal internodal
cells by confocal microscopy and observed a sharp shift from abundant transverse
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92 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
cortical microtubules at the lateral walls to sparse and randomly oriented micro-
tubules at the end walls (Wasteneys et al., 1996). We also found that subcortical
actin bundles extended around the ends of these giant cells but were unable to
detect cortical microfilaments as observed along the lateral walls. In higher plants,
however, microfilaments appear to be abundant at the end walls of expanding
cells. Using sectioning analysis of Arabidopsis and maize roots, Balu ka and
coworkers reported that the apical and basal ends of root cells accumulate micro-
filaments, and that in dwarf mutants and latrunculin-treated plants, this actin array
is not found (Balu ka et al., 2001a,b). Moreover, antibodies specific to myosin
VIII were reported to concentrate at the plasma membrane of newly formed trans-
verse cell walls of maize and Arabidopsis roots (Reichelt et al., 1999; amaj et al.,
2000), suggesting a role in anchoring microfilaments rather than a motile function.
Tissue sectioning, however, is not optimal for determining the overall pattern of
cytoskeletal organization in such tissues and can only reveal specific patterns on
cell faces that are tangential to the section plane.
We have recently carried out similar analysis of cytoskeletal distribution in
intact Arabidopsis roots using double immunolocalization and confocal recon-
structions (Collings et al., in press). In both the meristem (Fig. 3.1) and the
distal elongation zone (Fig. 3.2), the cortical arrays of both microtubules and
microfilaments are transversely aligned adjacent to the side walls. This is clearly
visible in epidermal (Fig. 3.1A; t) as well as cortex cells (Fig. 3.1D; t). Interphase
cells contain extensive endoplasmic microfilaments and microtubules, but once
cells reach the elongation zone, endoplasmic microtubules (but not microfilament
bundles) are no longer detected (Fig. 3.2). Using confocal microscopy, we can
reconstruct the organization of the cytoskeleton along the end walls and there,
microtubules appear to have a random orientation pattern (Fig. 3.1E; arrows).
3.4 Establishing axial growth
It is too simplistic to state that the cell cycle stops when terminal elongation
growth begins. Not all tissues stop dividing a set distance from the meristem,
though all must maintain the same rate of elongation (Section 3.6). In roots, the
pericycle tissue layer continues to divide throughout the elongation zone (Tobias
Baskin & Brian Gunning, personal communications), perhaps reflecting how these
cells, which later give rise to lateral roots, remain small and cytoplasmically dense.
In many expanding cells, ploidy levels increase by DNA endoreduplication, a
mechanism that may enable cell enlargement. Presumably, division could also
achieve this but endoreduplication may be optimal at sites distal to the meristem,
where diploidy does not need to be maintained. Ethylene and gibberellins control
endoreduplication in hypocotyls (Gendreau et al., 1999) but recent evidence also
links brassinosteroids and possibly abscisic acid to this process via the topoisomer-
ase VI complex (Sugimoto-Shirazu et al., 2002; Yin et al., 2002). The formation of
large central vacuoles is also a process that enables cells to achieve massive size,
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THE CYTOSKELETON AND AXIAL GROWTH 93
Fig. 3.2 Microfilament and microtubule organization in the elongation zone. Arabidopsis roots
immunolabelled for tubulin and actin were optically sectioned by confocal microscopy. (A) An optical
section shows that changes occur in the cytoskeleton as cells begin to elongate. Although cortical
microtubules remain transverse (t), endoplasmic microtubules (en) disappear. Endoplasmic microfila-
ments are retained, but trichoblast cell files (tr) consistently contain fewer mi crofilaments than
atrichoblast files (atr) especially in the distal parts of the elongation zone. Rapid cell elongation also
coincides with the development of vacuoles (v). (B), (C) Computer-generated transverse sections show
the radial organization of the cytoskeleton. They confirm the differences in microfilament organization
in trichoblasts and atrichoblasts, and confirm the loss of endoplasmic microtubules between the distal
elongation zone (B) and elong ation zone (C). They also show the marked increase in antibody penetration
in the elongation zone including in to the central stele of the root (st). The scale bar in (A) represents
20 µm in all images.
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94 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
since these compartments, which are essential for regulating turgor, occupy most
of the volume. The efficient mixing of the thin layer of cytoplasm at the cell
periphery depends on the myosin motor activity along microfilament networks
3.4.1 A transverse cortical microtubule array is essential
for axial growth
Establishing a transverse cortical microtubule array during the onset of rapid
expansion is a critical feature of axial growth. This is a process of both dispersal
and alignment that is probably mediated by a combination of mechanisms includ-
ing microtubule nucleation, assembly, severing, cross-linking, stabilization and
selective disassembly. Candidate proteins and protein complexes that fulfil these
activities are rapidly being discovered. One recent model proposes that severing of
nucleating templates from the slow-growing ends of microtubules, and motor-
dependent movement of these templates along microtubules is one way to generate
a dispersed microtubule array throughout the cortex (Wasteneys, 2002). This
dispersal process begins in early G1-phase of the cell cycle as a perinuclear array.
The requirement for severing in the process is inferred from mutational analysis of
the katanin p60 subunit homologue AtKSS (McClinton et al., 2001), whose ATP-
dependent severing activity has recently been demonstrated (Stoppin-Mellet et al.,
2002). Mutants with defective katanin p60 fail to produce a transverse cortical
microtubule array during the onset of elongation, resulting in loss of growth
anisotropy (Bichet et al., 2001; Burk et al., 2001), problems with cell fate specifi-
cation (Webb et al., 2002) and wall defects (Burk et al., 2001; Burk & Ye, 2002).
Just how these mutations affect the dynamic properties and ability of microtubules
to function properly remains to be determined. Nucleating complexes are likely
to include the recently identified Spc98p homologue (Erhardt et al., 2002) and
γ-tubulins, whose mutational analysis in plant cells is anticipated.
Selective stabilization of microtubules at the plasma membrane requires the
activity of MAPs. The high molecular mass MOR1 protein, a DIS1-TOGp-XMAP215
homologue, is the strongest contender for this role. Two mutant alleles with single
point mutations, mor1-1 and mor1-2, generate temperature-dependent disorganiza-
tion of the cortical arrays, left-handed twisting of organs (Section 3.6.2) and loss of
growth anisotropy (Whittington et al ., 2001). MOR1 may act as a key component of
a protein complex, with multiple binding partners. Mechanisms by which MOR1
stabilizes cortical microtubules have been proposed (Wasteneys, 2002), including
the obvious cross-linking of microtubules to the plasma membrane. In Xenopus
egg extracts it has been shown that MOR1’s homologue, XMAP215, stabilizes
microtubules by opposing the activity of the kinesin XKCM1 (Tournebize et al.,
2000; Hussey & Hawkins, 2001). Whether MOR1 interacts with homologues
of XKCM1 in plant cells is one avenue to explore in understanding the balanced
turnover of microtubules that is required to maintain a fully functional cortical
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THE CYTOSKELETON AND AXIAL GROWTH 95
3.4.2 Microtubules and their relationship with cellulose microfibrils
Over the years there has been considerable acceptance, despite only limited evi-
dence, for the cellulose synthase constraint model. The model proposes that cortical
microtubules, through their interaction with the plasma membrane, constrain the
movement of cellulose synthase complexes so as to control the orientation of cel-
lulose microfibrils. Some recent studies have raised doubts about this model. Fisher
and Cyr (1998) perturbed microtubule organization in tobacco BY-2 culture cells using
drugs that reduced cellulose synthesis and Sugimoto et al. (2000) showed that
cellulose microfibrils in Arabidopsis roots are more uniformly aligned than micro-
tubules, as cells enter elongation. The Arabidopsis fra2 mutant has disordered
microtubules and also has somewhat altered cellulose microfibril patterns. Nevertheless,
the microfibril texture is not as disturbed as would be predicted from the grossly
disturbed microtubule organization. Furthermore, the fra2 mutants have significantly
depleted cellulose levels, a condition that alone generates disordered microfibrils
in dichlorobenzonitrile-treated seedlings (Sugimoto et al., 2001), and the rsw1-1
(Sugimoto et al., 2001) and kobito (Pagant et al., 2002) mutants.
New data demonstrate that disturbance or removal of microtubules causes
radial swelling without altering the parallel orientation of cellulose microfibrils
(Sugimoto et a l., 2003). Himmelspach et al. (2003) have extended these studies,
showing that cellulose microfibrils can re-establish parallel transverse order during
microtubule disruption in the mor1 mutant, even when a template of transverse
microfibrils is destroyed by dichlorobenzonitrile treatment. Together, these studies
in Arabidopsis roots suggest that: (1) cellulose microfibril orientation is largely
self-ordered; (2) transverse orientation of both cortical microtubules and cellulose
microfibrils is essential for axial growth; and (3) cortical microtubules do not
directly regulate the movement of cellulose synthase complexes.
In exploring the cytoskeleton’s role during axial growth, most emphasis has
been placed on the relationship between cortical microtubules and cellulose micro-
fibril alignment in lateral walls (Baskin, 2001). Yet it is now clear that this is not
the only process responsible for maintaining anisotropy. The rsw4 and rsw7 mutants
have substantially normal microtubule and cellulose microfibril organization and
yet undergo radial swelling under restrictive conditions (Wiedemeier et al., 2002).
Relatively little focus has been given to how the cytoskeleton might modulate
secretion of other wall components or the activity of wall loosening enzymes.
Recent work, however, links microtubule organization and xyloglucan metab-
olism, though microtubule activity seems to be more responsive than regulatory.
Takeda et al. (2002) demonstrated that xyloglucan application to pea stems can
suppress cell elongation and stimulate a shift in microtubule orientation from
transverse to longitudinal, while application of an oligosaccharide derivative of
xyloglucan stimulates elongation. Some results of this study are notable from the
perspective of microtubule organization. Like the findings of gravitropic bending
studies (Section 3.6.1), neither microtubule reorientation from transverse to
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96 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
longitudinal nor its maintenance in a transverse direction controlled the growth
response. Taxol stabilization of transverse microtubule orientation did not affect
the ability of the xyloglucan application to reduce growth, while longitudinal
microtubule reorientation generated by the kinase inhibitor 6-dimethylaminopurine
did not prevent growth stimulation by the oligosaccharide derivative. Nevertheless,
taxol treatments enhanced the elongation stimulated by oligosaccharide application,
and 6-dimethylaminopurine-treatments enhanced oligosaccharide-induced growth
inhibition. These results support the concept that cellulose orientation is largely
self-ordered and dependent on the correct levels of polymer synthesis, while
microtubule coalignment with microfibrils contributes to, but is not sufficient to
generate axial growth. The results also support the concept that microtubule ori-
entation depends on cues from the cell wall (Fisher & Cyr, 1998; Wasteneys, 2000;
3.4.3 Does the cytoskeleton regulate wall polysaccharide and protein
Axial growth requires that different processes act simultaneously on end and
lateral walls. At the end walls, loosening needs to be greatly restricted or prevented
altogether. At lateral walls, loosening activity must be upregulated relative to that
at the end walls and yet restricted by the mechanical properties of the wall to
expand in only one direction. Constraining expansion of end walls while maximizing
loosening of lateral walls requires differential synthesis and secretion of matrix
polysaccharides as well as wall-specific enzymes and other proteins. Current
models of wall extensibility consider the interactions between xyloglucans and
cellulose microfibrils to be the major load-bearing mechanism, though the import-
ance of pectic polysaccharides is also recognized (Cosgrove, 2001). The ratio of
xyloglucan to cellulose, and perhaps also the amount and activity of xyloglucan
endotransglycosylase, are critical determinants of differential growth on end and
lateral walls, and between growing and non-growing cells. We envisage that wall
enzymes, including xyloglucan endotransglycosylases and endoglucanases, but
also proteins like expansins are likely to differ in abundance and/or activity at the
lateral and end walls. This can now be tested by immunocytochemistry. Arabi-
nogalactan proteins (AGPs) are also likely to play important roles in elongation
growth (van Hengel & Roberts, 2002). Interestingly, some AGPs influence micro-
tubule organization. This has been shown for AGPs in the case of the reb1 mutant,
in which reduced AGP levels in trichoblasts lead to bulging cells with disturbed
microtubule organization (Andème-Onzighi etal., 2002).
3.4.4 Hormones, cytoskeleton and wall extensibility
As demonstrated by Baskin et al. (1999), environmental conditions dictate the
degree of anisotropy. In response to abiotic cues, hormones reinforce or reduce
anisotropy. Ethylene is frequently associated with altering the expansion axis of
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THE CYTOSKELETON AND AXIAL GROWTH 97
diffusely expanding cells. It may act to alter microtubule and microfibril orienta-
tion and change the growth direction from longitudinal to radial (Lang et al., 1982)
but it can also stimulate elongation, as has been shown in light-grown Arabidopsis
hypocotyls (Smalle et al., 1997) or stems of submerged rice (van der Straeten et al.,
2001). Gibberellins promote both anisotropic expansion and transverse cortical
microtubules in aerial tissues. In barley, Wenzel et al. (2000) demonstrated that
gibberellin-deficient dwarfs undergo considerable radial swelling during the early
phase of cell expansion when microtubules are in fact still transversely aligned,
supporting the idea that gibberellins act independently on wall loosening and micro-
tubule orientation. Auxin has a similar major role on wall loosening by inducing
matrix polysaccharide hydrolysis. As summarized by Cosgrove (2001) it also stimul-
ates wall synthesis, increases wall plasticity and activates genes for wall enzymes and
proton ATPases. The involvement of the cytoskeleton in polar auxin transport is
considered in detail in Section 3.5.
3.4.5 How does the actin cytoskeleton contribute to cell elongation?
How are microfilaments organized during the crucial phase of cell development
when rapid cell elongation begins? Bundles of microfilaments are located in the
subcortex and in the transvacuolar strands where they generate cytoplasmic stream-
ing, but a fine network of microfilaments is also found at the plasma membrane,
parallel to the cortical microtubules (for example, Collings et al., 1998; Blancaflor,
2000; Collings & Allen, 2000). The presence of this network in diffusely expanding
cells is consistent with the actin cytoskeleton’s presence near sites of rapid exocytosis
during tip growth and wound wall formation (Wasteneys & Galway, 2003). The actin
cytoskeleton does not undergo significant modification as elongation commences
or as cells undergo rapid vacuolation. By comparison, the endoplasmic microtubules
rapidly disappear (Fig. 3.2). Microfilaments are consistently more abundant in
atrichoblasts than in trichoblasts (Fig. 3.2A), a result confirmed in computer-
generated reconstructions (Fig. 3.2B) but opposite to that reported previously
(Balu ka et al., 2001b).
Disruption of the actin cytoskeleton in Arabidopsis and maize with low concen-
trations of cytochalasin or latrunculin reduces cell elongation (Baskin & Bivens,
1995; Balu ka et al., 2001b). Actin disruption also generates a swollen root
phenotype (Baskin & Bivens, 1995; Blancaflor, 2000) as does the myosin inhibitor
2,3-butanedione 2-monoxime (BDM) (Baskin & Bivens, 1995). This phenotype is
similar to the effects of drug-induced microtubule depolymerization, the inhibition
of exocytosis with brefeldin or monensin, or the inhibition of cellulose synthesis.
The phenotype is not, however, induced by a large range of other metabolic or
growth inhibitors (Baskin & Bivens, 1995). Furthermore, the transgenic over- and
under-expression of certain actin-binding proteins in Arabidopsis, including profi-
lin, actin depolymerization factor (ADF) and cyclase-associated protein, generate
morphological changes that sometimes extend beyond simply altered growth rates
and suggest effects on microtubule organization and anisotropic cell elongation
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98 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
(Ramachandran et al., 2000; Dong et al ., 2001; Barrero et al., 2002). Over-expressing
profilin also generated other morphological phenotypes including apparent left-
handed stem twisting, and a reduction in the lobing of epidermal pavement cells
(Ramachandran et al., 2000), both features of mutants that show microtubule dis-
ruption (Whittington et al., 2001; Thitamadee et al., 2002).
How then do microfilaments contribute to cell elongation? One possible mechanism
is through interactions with cortical microtubules. Numerous drug studies indicate
that such interactions occur (Collings & Allen, 2000), although it is uncertain
whether these interactions are direct or whether the coordination between the two
systems is via an indirect pathway. Actomyosin-dependent streaming is needed
to efficiently mix the thin layer of cytoplasm of vacuolated cells and to transport
transcription and translation products to all parts of the expanding periphery.
Furthermore, auxin transport depends on the activity of microfilament networks
(Section 3.5.1) and not microtubules (Hasenstein et al., 1999). But as microtubule
organization is also one of the targets of auxin signalling (Takasue & Shibaoka,
1999), and as auxin promotes cell wall loosening (Taguchi et al. 1999), any changes
in auxin levels by disruption to the actin cytoskeleton will have downstream
consequences on microtubules and microtubule-dependent anisotropy.
3.5 Polar auxin transport and its regulation by the actin
What factors define asymmetry in organs and individual cells, and how is this
determined? Plant hormones help to define, reinforce and/or perturb cell polarity
and axiality. Their effects on growth through the cytoskeleton were extensively
reviewed several years ago (Shibaoka, 1994), but recent years have seen considerable
advances in understanding how the cytoskeleton and plant hormones interact at a
molecular level. Auxin is the best example of this, and in this section, we discuss how
the cytoskeleton is involved in auxin transport and how it responds to auxin gradients.
3.5.1 Auxin transport and the chemiosmotic theory
The chemiosmotic theory of polar auxin transport (Raven, 1975) established a
theoretical framework that accounts for the apical to basal movement through
plants of indoleacetic acid (IAA), the most common biologically occurring auxin
(Fig. 3.3A). The relative acidity of the cell wall (pH 5) causes weak acids such as
auxin to protonate, forming a non-charged species that can diffuse across the
plasma membrane into the cytoplasm where it then deprotonates to give a charged
species that cannot re-cross the plasma membrane. Movement of auxin out of the
cell is limited to an efflux carrier that is restricted solely to the basal plasma
membrane, resulting in the polarised auxin transport downwards through the stem
and roots (Fig. 3.3B). This theory has been validated experimentally in recent
years. Eight auxin efflux carriers, encoded by members of the PIN gene family,
have been identified in Arabidopsis, initially through the cloning of genes from
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THE CYTOSKELETON AND AXIAL GROWTH 99
mutants whose auxin-dependent development and/or gravitropic responses are
compromised (Chen etal., 1998; Gälweiler etal., 1998; Luschnig et al., 1998;
Müller et al., 1998; Friml & Palme, 2002). Localizations of these efflux carriers
match the chemiosmotic theory’s predictions (Fig. 3.3B). AtPIN1 is found at the
lower or basal plasma membrane in shoots (relative to the shoot apical meristem),
and at the lower or apical plasma membrane (towards the root apex) within the
central stele of the root (Gälweiler et al., 1998). AtPIN2 distributes to the basal
plasma membrane of epidermal and cortical cells of the root tip (Müller et al.,
1998). Within the root tip, AtPIN4 may contribute to the increased auxin levels found
in the quiescent zone cells (Friml et al., 2002a), while AtPIN3 distributes evenly
throughout the plasma membrane of root cap columella cells but redistributes to
the lower side of the root after gravistimulation (Friml et al., 2002b). These patterns
are consistent with the paths that auxin takes through the plant. Interestingly, the
auxin influx carrier AUX1 has an asymmetric distribution to the basal plasma
membrane of cells within the protophloem of the root tip, again consistent with
auxin movements (Swarup et al., 2001) (Fig. 3.3B).
How do plants maintain these asymmetries? Separate streams of evidence
indicate that the actin cytoskeleton, through its function in vesicle trafficking,
helps to regulate the polarized location of auxin efflux carriers at the plasma mem-
brane. Auxin efflux carriers undergo rapid cycling between the plasma membrane
and a perinuclear endomembrane compartment (Fig. 3.3C,D). Brefeldin A, by
inhibiting vesicle trafficking and endocytosis, causes a reversible build-up of PIN
proteins and AUX1 within the perinuclear compartment (Steinmann et al., 1999;
Geldner et al., 2001; Grebe etal., 2002). Cycloheximide treatments that block
protein synthesis do not prevent the build-up of AtPIN1, demonstrating that it
cycles between the perinuclear compartment and the plasma membrane (Geldner
et al., 2001). Disruption of microfilaments with either cytochalasin or latrunculin
has little effect on AtPIN1 localization, but cytochalasin pre-treatment prior to
brefeldin A prevents AtPIN1 build-up in the perinuclear compartment, while
cytochalasin addition after brefeldin A’s removal prevents redirection of AtPIN1
to the plasma membrane (Geldner et al., 2001). Similar results were also obtained
for AtPIN3 (Friml etal., 2002b). These results show that an intact and func-
tional microfilament network is essential for both delivery to and recovery of PIN
proteins from the plasma membrane (Fig. 3.3D). This functioning likely includes
both myosin-dependent movement of vesicles along microfilament bundles, but
may also indicate further roles for microfilaments in exocytosis and/or endocytosis.
Gloria Muday and colleagues have theorised that proteins regulating the auxin
efflux carrier bind to the actin cytoskeleton (Fig. 3.3C; reviewed in Muday, 2000;
Muday & DeLong, 2001). Naphthylphthalamic acid (NPA), an auxin transport
inhibitor that inhibits auxin-dependent growth and gravitropism (Rashotte et al.,
2000), functions through high-affinity coupling to the NPA-binding protein that
regulates the activity of auxin efflux carriers at the plasma membrane. If micro-
filaments are disrupted, NPA-binding activity recovered from plasma membrane
preparations is reduced (Butler etal., 1998). NPA-binding activity can also be
recovered from F-actin affinity columns, and although the protein(s) responsible
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100 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
could not be identified, the results are consistent with the NPA-binding protein
being an actin-binding protein (Hu et al., 2000). Recent developments have,
however, complicated the interpretation of the NPA-binding protein being a link
between auxin efflux carriers and the actin cytoskeleton. NPA and other auxin
transport inhibitors have a complicated mode of action that includes the brefeldin
A-like suppression of efflux carrier cycling between the plasma membrane and
perinuclear compartment (Geldner et al., 2001), although these effects occur at
significantly higher concentrations (about 200 µm) than required to block gravi-
Fig. 3.3 Microfilaments contribute to cell elongation by maintaining polar auxin transport. (A) The
chemiosmotic theory explains polar auxin transport through the asymmetric distribution of an auxin
efflux carrier. Auxins are weak acids that occur in the membrane-permeant protonated form (IAAH) in the
acidic cell wall, and which accumulate in the cytoplasm because they deprotonate to give a membrane-
impermeant ion (IAA–). The asymmetric distribution of auxin efflux carriers (circles) to the basal
plasma membrane generates basipetal movement of auxin. This process is aided by the localization to
the apical plasma membrane of auxin influx carriers (squares), which use the trans-membrane proton
gradient to speed aux in uptake. (B) In Arabidopsis seedlings, basipetal auxin transport occurs through the
central stele of the hypocotyl to the root tip, from where auxin circulates back through the cortical and
epidermal cells. Auxin efflux carriers from the PIN family of proteins and the auxin influx carrier
AUX1, whose locations match the predictions of the chemiosmotic theory, generate this pattern. In
roots, AtPIN1 localises to the lower (tipward or apical) plasma membrane of vascular cells, while
AtPIN2 localises to the upper (basal) plasma membrane of epidermal and cortex cells. AtPIN3 occurs
in root cap cells, while AtPIN4 localises asymmetrically to the apical plasma membrane of cells adja-
cent to the quiescent centre. AUX1 also distributes asymmetrically to the basal ends of protophloem
cells, the opposite end to AtPIN1. Cell types are: 1 =epidermis; 2 =cortex; 3 =endodermis; 4 =pericycle;
5=vascular tissue, which includes protophloem (6); 7 =quiescent zone and cell file initials; 8=columella;
and 9 =lateral root cap.
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THE CYTOSKELETON AND AXIAL GROWTH 101
tropic bending (about 1µm; Rashotte etal., 2000). Moreover, several Arabidopsis
proteins involved in auxin transport, including aminopeptidases (Muday & Murphy,
2002; Murphy etal., 2002), bind NPA with low affinity. These proteins have not
been shown to bind to the actin cytoskeleton, and high-affinity NPA-binding
proteins that interact with the actin cytoskeleton have not been identified (Muday
& Murphy, 2002).
3.5.2 Important questions concerning auxin transport and the actin
Why do the PIN proteins undergo rapid cycling? In their recent review of polar
auxin transport, Friml and Palme (2002) suggested possible reasons for rapid
cycling. Perhaps cycling allows for rapid changes in auxin efflux carrier location
Fig. 3.3 (continued) (C), (D) Auxin efflux carriers such as AtPIN1 (circles) undergo rapid actin-dependent
cycling between the plasma membrane and an endocytotic compartment, here labelled as an endosome,
and may be regulated at the plasma membrane by the NPA-binding protein (triangles) and microfilaments
(C). In control root cells (D), AtPIN1 (shading) distributes to the apical plasma membrane. Brefeldin A
inhibits exocytosis and causes a rapid accumulation of AtPIN1 into a perinuclear endocytotic compart-
ment, but microfilament disruption with cytochalasin (CD) has little effect on AtPIN1 localization.
Serial brefeldin and cytochalasin treatments, however, demonstrate that microfilaments are required
for cycling, and show that brefeldin blocks exocytosis, but microfilament disruption with cytochalasin
D inhibits both exocytosis and endocytosis. This inhibition may involve both the prevention of vesicle
movement along actin bundles, and/or a direct involvement in exocytosis and endocytosis.
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102 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
to be achieved or perhaps there is some sensing role to the cycling, with efflux
carriers acting as membrane receptors. Alternatively, when vesicles containing
PIN efflux carriers move along the actin cytoskeleton, they are also actively trans-
porting auxin through the cell as a specific cargo. We view this third possibility as
unlikely because the majority of auxin efflux carrier proteins such as AtPIN1 are
found in polarized locations at the plasma membrane rather than associated with
vesicles (Gälweiler et al., 1998). Moreover, the acidification of secretory vesicles,
which occurs before secretion in animal cells (Miesenböck et al., 1998), and pre-
sumably also in plants cells, would limit auxin build-up because protonated auxin
could diffuse back into the cell. Instead, we suggest that the rapid cycling of auxin
efflux carriers may also contribute to their asymmetric distribution. Were targeted
secretion to correctly deliver efflux carriers to one part of the plasma membrane,
the rapid recovery of proteins from throughout the plasma membrane would limit
diffusion of the efflux carriers through the cell. This would also eliminate the need
for the efflux carriers to be tethered into the correct location through a cytoskeleton-
based fixing mechanism such as the NPA-binding protein.
How are the PIN proteins targeted asymmetrically to the plasma membrane?
A comparison of AtPIN1 and AUX1 location in the protophloem of roots shows
that they are found at opposite ends of the cell (Gälweiler et al., 1998; Swarup
et al., 2001) (Fig. 3.3B), confirming that cells must have multiple methods of
ensuring different types of asymmetric exocytosis. How differences between the
contents of exocytotic vesicles at end- and side-walls are generated and how the
polar distribution of auxin efflux carriers re-establish after cell division are also
unresolved questions. When a cell that has asymmetrically localized auxin efflux
carriers divides, the daughter cells need to re-establish this polarization to ensure
continued polar auxin transport. Initial evidence shows that AtPIN1 colocalizes
with the KNOLLE protein in the cell plate, presumably without there being any
distribution asymmetry (Geldner et al., 2001). How and how quickly the polarized
pattern re-establishes itself has not been determined.
3.5.3 Small GTPases may be a key to the shuttling of auxin
ROP (Rho of Plants) proteins are small GTPases that act as molecular switches.
In tobacco suspension cells, the Arabidopsis ROP4-GFP fusion protein localises to
the cell plate and cross wall. Immunolocalization suggests that ROP4 (and prob-
ably other closely related ROPs such as ROP6) is preferentially distributed at the
apical and basal ends of elongating cells in Arabidopsis root meristems (Molendijk
et al., 2001). In the same study, expression of a constitutively active GFP-AtROP4
construct under an inducible promoter caused isotropic swelling of epidermal cells
of hypocotyls, cotyledons and roots (Molendijk etal., 2001). The identification of
an end-wall associated ROP is consistent with a general ROP association with sites
of active vesicle secretion, as well as the formation of fine actin networks, such as
those reported at the ends of cells (Balu ka et al., 2001b). These ROPs could
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THE CYTOSKELETON AND AXIAL GROWTH 103
regulate the shuttling of auxin efflux carriers between their endomembrane
compartment and the plasma membrane at the ends of cells (Section 3.5.1), either
directly or by remodelling the actin cytoskeleton.
3.5.4 Auxin and gene expression
Auxin elicits many and diverse effects in plants. As discussed below (Section 3.6.1),
auxin’s involvement in gravitropism includes separate but tightly integrated
effects on microtubule orientation and cell elongation. However, auxin also causes
the rapid induction and repression of many different genes through multiple
pathways (Guilfoyle etal., 1998; Leyser, 2002) that may be modulated, at least in
part, through ROP GTPases (Li et al., 2001; Tao et al., 2002). Some of the auxin-
induced changes in gene expression directly affect the cytoskeleton. The Arabidopsis
ACT7 gene encodes an actin that is preferentially expressed in rapidly elongating
tissue. ACT7 expression responds to hormones, including auxin, abscisic acid and
cytokinins. Exogenous auxin reduces ACT7 expression along with reducing
root elongation (McDowell et al., 1996) but increases ACT7 expression during
auxin-dependent callus formation (Kandasamy et al., 2001). Significantly, act7-1,
a T-DNA insertional mutant, was slow to form callus in response to external auxin,
suggesting a role for this actin isoform in callus formation (Kandasamy et al.,
3.6 Bending and twisting – the consequences of differential growth
For plant organs to grow in one direction, the different tissue layers must all
elongate at the same rate. Any major differences in elongation rates between tissue
layers result in buckling and rupture, with devastating consequences. Subtle
differences in growth rates between cells in adjacent tissue layers can, however,
alter the direction of organ growth. Gravitropic and phototropic bending responses
all involve coordinated differential flank growth, in which there is evidence for
cytoskeletal involvement. Uncoordinated differences in growth can lead to the
helical arrangement of cell files or the twisting of whole organs. Microtubules may
have a regulatory role in twisting. Mutational, transgenic and drug-dependent
perturbation of microtubule organization can generate either left- or right-handed
twisting of both axial and lateral organs (Table 3.1). We present, in turn, evidence
for cytoskeletal involvement in bending and twisting.
3.6.1 Tropic bending responses
Tropic responses to light (phototropism), gravity (gravitropism) and other stimuli
are adaptive growth responses in which the differential expansion of cells on oppos-
ite sides of organs, and not cell division, generate bending. The cytoskeleton’s
involvement in the perception of a gravity stimulus through amyloplast sedimentation
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104 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
and the generation of the auxin asymmetry that results in asymmetric growth have
been extensively debated, and recently reviewed (Blancaflor, 2002). The con-
tradictory nature of many of the observations derives, in part, from the variety of
tissues studied (e.g. roots, hypocotyls, coleoptiles and various types of stems).
These may show different adaptations of an underlying gravitropic process. Using
the bending response as a measure of signal perception also confounds analysis of
the events in perception.
In this section, we consider the possible roles of the cytoskeleton in gravitropic
bending responses. These responses require cell elongation, so microtubules
should play a critical role. The relationship between microtubules and gravitropic
bending has been extensively studied in maize coleoptiles, roots and stems. In
Table 3.1 Genotypes and treatments generating organ-twisting phenotypes in Arabidopsis thaliana
References : 1, Migliaccio et al. (2000); 2, Furutani et al. (2000); 3, Whittington et al. (2001);
4, Wasteneys (2002); 5, Thitamadee et al. (2002); 6, Hashimoto (2002).
Genoty pe or
treatment Cortical microtubules Protein targeted
wild type transverse ecotype-
tubulin left 2, 6
taxol (low concs) stabilized tubulin left 2, 6
spr1 diso rder ed in root
endodermis and cortex;
left-handed helical in
unknown axial organs;
spr2 not determined unknown right
short and disordered 217 kDa MAP all organs left (loss of
after 24 h)
lefty1 right-handed helix in
α-tubulin 4 all organs left 5, 6
lefty2 right-handed helix in
α-tubulin 6 all organs left 5, 6
35S::GFP-TUA6 transverse overexpression
35S::MBD-GFP transverse overexpression of
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THE CYTOSKELETON AND AXIAL GROWTH 105
gravistimulated coleoptiles and roots, there is a consistent change in microtubule
orientation from transverse to longitudinal in the epidermal cells whose growth is
inhibited. In upward bending coleoptiles these are the cells on the upper flank
where auxin levels are reduced (Nick et al., 1990; Himmelspach et al., 1999)
whereas in downward bending roots this occurs on the lower flank where auxin
accumulates (Blancaflor & Hasenstein, 1995). This microtubule reorientation in
coleoptiles is not coupled to growth and even occurs when the coleoptiles are pre-
vented from growing and bending by gluing them to glass slides (Himmelspach &
Nick, 2001). Similarly, Takesue and Shibaoka (1999) showed that auxin-induced
longitudinal to transverse microtubule reorientation can occur when growth is
suppressed by anaerobic conditions. In downward bending maize roots, micro-
tubule reorientation does not begin until after bending has commenced and can be
mimicked by applying auxin (Blancaflor & Hasenstein, 1995). These observations
demonstrate that the reorientation of microtubules to the longitudinal direction has
no intrinsic function in the bending response. Indeed, transverse to longitudinal
reorientation of microtubules is a normal consequence of growth cessation (Liang
et al., 1996; Sugimoto et al., 2000).
Reinforcement of transverse microtubules in the growth-stimulated cells on the
outer side of the bend is potentially of greater significance to gravitropic bending
but is likely to be a requirement rather than a regulatory mechanism. Indeed,
microtubule depolymerization does not inhibit gravitropic bending of roots (Balu ka
et al., 1996; Hasenstein et al., 1999), though the time required for bending is
unlikely to alter the mechanical properties of walls of the expanding cells. These
experiments serve to demonstrate that the key to a bending response is the stimula-
tion of growth on one flank and suppression on the other regardless of whether the
stimulated growth is isotropic or anisotropic. The opposite responses of coleoptiles
and roots to auxin (growth stimulation and maintenance of transverse microtubules
in coleoptiles; growth inhibition and loss of transverse microtubules in roots) are
The maize stem pulvinus, like the coleoptile, generates upward bending
responses, with stimulation of growth on the lower flank by auxin accumulation
(Long et al., 2002). This system, however, is fundamentally different from gravi-
responding roots and coleoptiles in that the pulvinal cells, which occur at nodes
along the stem, remain in a quiescent state until gravistimulated. Once the stem is
induced to bend upward by horizontal placement, the microtubules remain
transverse in pulvinal cells on both the upper, non-elongating and lower, rapidly
elongating flanks (Collings et al., 1998). Here, the key to microtubules remaining
transverse is the need to retain versatility. The pulvinus enables maize stems to
recover vertical growth when flattened by wind, crop circle devotees (or alien
spaceships) (Levengood & Talbott, 1999) but also allows for more moderate
adjustments to stem attitude requiring sequential bending in more than one direction
from a single node.
Several genes involved in gravity perception, signal transmission and response
have also been identified through mutational strategies, and the cytoskeleton’s
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106 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
roles in these processes dissected. As discussed in Section 3.5.1, the PIN family of
auxin efflux carriers was first identified in agravitropic mutants in Arabidopsis
hypocotyls and roots (Chen et al., 1998; Gälweiler et al., 1998; Luschnig et al.,
1998; Müller et al., 1998), as was the auxin influx carrier AUX1 (Bennett et al.,
1996), and actin plays a fundamental role in the distribution of these proteins. The
gene affected in the altered response to gravity (arg1) mutant of Arabidopsis
encodes a DNA-J-like protein, with a coiled-coil region that may interact with the
cytoskeleton (Sedbrook et al., 1999) although this has yet to be shown conclusively.
Another gravity-response mutant with links to the cytoskeleton is the Yin-Yang
mutant of rice. The coleoptiles of Yin-Yang are ten times more sensitive to cyto-
chalasin D than wild-type coleoptiles but this sensitivity is auxin-dependent. When
exposed to auxin, the actin cytoskeleton of the mutant and its gravitropic response
resemble those of the cytochalasin-D-treated wild type. Despite its clear actin-based
phenotype, Yin-Yang was isolated on the basis of its cell elongation being resistant
to microtubule inhibitors, suggesting that its altered actin organization influences
microtubule-dependent processes (Wang & Nick, 1998). The gene affected in
Yin-Yang has not yet been identified.
Twisting of organs is a feature of circumnutation and thigmotropic responses such
as twining. Microtubule disruption can generate twisting in axial and lateral
organs. That different perturbations to the microtubule cytoskeleton cause twisting
that is consistently either left- or right-handed (Table 3.1), indicates the presence
of some basic underlying mechanism. Can we understand the involvement of
microtubules in cell expansion by studying how microtubule disruption influences
The most likely cause of twisting is in fact the differences in expansion between
the outer and inner tissue layers. In the spr1 mutant, Furutani etal. (2000) demon-
strated that while the epidermal cells maintain normal anisotropic expansion, the
cortex and endodermal cells grow isotropically. This more rapid elongation in the
epidermis introduces physical stresses to which the epidermis can respond in two
1. It can buckle, leading to tissue damage and probably seedling death.
2. If the difference in growth rates is small, the epidermal files can change their
growth direction, resulting in twisting.
Although a detailed assessment has not been carried out in the spr1 mutant,
it seems likely that the twisting results from the third type of response.
All mutations and treatments that generate twisting also reduce anisotropic
expansion to some extent, although this in itself is not enough to cause twisting.
For example, taxol and propyzamide cause twisting but at higher concentrations
produce radial swelling (Furutani et al., 2000), while at its restrictive temperature,
Pccdc03.fm Page 106 Wednesday, October 8, 2003 6:14 PM
THE CYTOSKELETON AND AXIAL GROWTH 107
the temperature-sensitive mor1-1 mutant gradually loses anisotropic expansion
(Whittington et al., 2001). The lefty1 and lefty2 mutants have thicker roots than
wild-type, and lefty1lefty2 (Thitamadee et al., 2002) and spr1spr2 (Furutani et al.,
2000) double mutants have strong radial swelling phenotypes. Many radial swelling
mutants do not, however, show any twisting growth. Mutants with reduced cellu-
lose levels, or plants in which microfilaments have been disrupted, show radial
swelling but not twisting (although Arabidopsis plants over-expressing profilin
show radially swollen hypocotyls that may show twisting – see Fig. 3.3 in
Ramachandran et al., 2000). Mutations affecting the microtubule severing protein
katanin p60 also do not generate twisting. While this seems at odds with the strong
correlation between twisting and microtubule defects, cellulose levels are signifi-
cantly depressed in the fra2 mutant (Burk et al., 2001), unlike the left-twisting mor1-1
mutant (Sugimoto et al., 2003). Thus, mutations and treatments affecting micro-
tubules, but not cellulose synthesis, can generate twisting.
What determines the direction of twisting? Hashimoto and coworkers observed
that microtubule orientation is right handed in the left-twisting lefty mutants, and
left-handed in the right-twisting spr1 mutant, and have hypothesized that it is the
handedness of cortical microtubules in the epidermis that determines the handed-
ness of twisting (Furutani et al., 2000; Hashimoto, 2002; Thitamadee et al., 2002).
We note, however, that if the orientation of the root is taken into account, the
cortical microtubules are almost perpendicular to the gravity vector in these mutants,
and although there is no evidence to suggest that microtubules sense gravity, this
possibility should not be discounted. Furthermore, the model of Hashimoto and
coworkers fails to explain the strong left-twisting of mor1 mutants in which micro-
tubules become disordered with no preferred handedness, and it also cannot
explain the strong right-handed twisting caused by the transgenic expression of the
MBD-GFP and GFP-TUA6 fusion proteins.
We suggest that various radial gradients generate at least some of the twisting
phenotypes, acting in concert with an inherent torsional handedness of the whole
organ. This torsion could be generated by the staggered helical division patterns
that follow the periclinal divisions of the lateral root cap/epidermis initial cells.
Defects in these divisions generate strong twisting in the tornado mutants (Cnops
et al., 2000). This inherent torsion might also explain why microtubules consist-
ently reorient from transverse to longitudinal via a right-handed helix as root cells
of maize and Arabidopsis (Liang et al., 1996; Sugimoto et al., 2000) cease elonga-
tion. Inward gradients, or treatments that first reduce anisotropic growth in the
outer tissues, will generate left-handed twisting of cell files. This is consistent
with mor1’s temperature-dependent phenotype (a temperature gradient) and drug
treatments such as propyzamide and taxol that diffuse inward from the epidermal
layer (Furutani et al., 2000). We would also predict that in the lefty mutants,
expression of the mutated α-tubulin 4 and α-tubulin 6 would be greater in the
epidermis than in the inner tissues, although this has not yet been determined.
Conversely, right-handed twisting phenotypes may be generated by outward flow-
ing gradients. This is clearly the case for the spr1 mutant, with endodermis and
Pccdc03.fm Page 107 Wednesday, October 8, 2003 6:14 PM
108 FUNDAMENTAL CYTOSKELETAL ACTIVITIES
cortex cells broader and shorter than the epidermal cells (Furutani et al., 2000).
Clearly, however, more detailed analyses of microtubule handedness in twisting
organs other than roots are also required, because agar-based seedling culture
introduces considerable artefacts. Growth on hard agar surfaces means that root
twisting will cause skewing of roots in one direction across the plate, whereas sim-
ilarly affected soil-grown roots would grow in a helical direction like a corkscrew
(Buer et al., 2000).
3.7 Conclusions and future perspectives
Axial growth is an inherent feature of plant function and essential at all stages of
development, beginning with the zygotic embryo and continuing through until all
reproductive organs are complete. Anisotropic expansion enables plants to produce
organs of unlimited complexity, with features that help them compete for light,
bend against gravity or towards light, minimize wind damage, twine up other plants,
penetrate seemingly impenetrable soils and even optimize pollination strategies.
We have examined the role microtubules and microfilaments play in axial growth,
from the onset of cell plate formation in telophase, through the establishment of
axial growth and on to more complex processes of differential expansion. Recent
investigations assisted by the genome projects have uncovered a plethora of new
factors, whose functions will begin to become clear as thoughtful investigations
are carried out. Unfortunately, picking through the morass of confusing results will
undoubtedly continue for some time.
Geoffrey Wasteneys and David Collings are supported by ARC Discovery Project
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