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Plant Science 175 (2008) 747–755
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Review
The roles of microtubules in tropisms
Sherryl R. Bisgrove *
Department of Biological Sciences, Simon Fraser University, 8888 University Drive, Burnaby, B.C., Canada V5A 1 s6
A R T I C L E I N F O
A B S T R A C T
Article history:
Received 11 March 2008
Received in revised form 19 August 2008
Accepted 19 August 2008
Available online 7 September 2008
Plant tropisms, or growth towards or away from a stimulus, usually involve the bending of shoots or roots
which reorient growth in a new direction. Plant responses to tropic cues, especially gravity and light, have
been active areas of investigation for many years. Despite all of this attention we still do not understand
how these responses are regulated. In this review possible roles for microtubules in tropisms are
discussed. Tropisms occur in a series of steps; directional cues are perceived and converted into
biochemical signals that induce bending in roots and shoots. One model suggests that microtubules
function late in the response pathway, during organ bending. Microtubules have been linked to organ
bending by virtue of their role in regulating the direction of cell elongation. In elongating cells
microtubules appear to function as guides for the deposition of cellulose microfibrils into the cell wall and
the placement of the microfibrils in the wall is thought to constrain the direction of cell elongation.
According to the model bending occurs when different microtubule/microfibril alignments across the
organ cause cells on the outer flank to elongate more than cells on the inner flank. In support of this idea is
the observation that tropic signals can induce the appropriate changes in microtubule orientations across
a bending organ. However, attempts to validate the hypothesis have produced conflicting results and the
idea that microtubule alignment regulates cell expansion during organ bending is controversial.
Microtubules have also been linked to organizational events associated with the plasma membrane.
Although speculative, one possibility is that microtubules influence tropisms by positioning regulatory
proteins and/or complexes in the plasma membrane. Two possible mechanisms by which microtubules
could contribute to organizational events associated with the plasma membrane are outlined. In addition
to cell expansion, microtubules are postulated to have roles in the perception of touch and gravity signals.
Although microtubules are associated with touch sensing in animals, we know very little about the
relevant receptors in plants. One way to assess how microtubules function during tropisms is to identify
and study proteins that function in concert with microtubules. In particular, the analysis of microtubuleassociated proteins whose mutant forms confer defects in tropic responses promises to provide
additional insights into the roles of microtubules in tropisms.
ß 2008 Elsevier Ireland Ltd. All rights reserved.
Keywords:
Microtubule
Cytoskeleton
Cell expansion
Cell wall
Tropism
Contents
1.
2.
3.
4.
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1.1.
Signal perception . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1.2.
Auxin redistribution and differential growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
1.3.
Possible roles for microtubules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Cortical microtubule organization and cell elongation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.1.
Cellulose deposition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
2.2.
Microtubule reorientations and tropisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Beyond cellulose deposition: new roles for microtubules? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Summary and future directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Acknowledgements. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
* Tel.: +1 778 782 5269; fax: +1 778 782 3496.
E-mail address: [email protected].
0168-9452/$ – see front matter ß 2008 Elsevier Ireland Ltd. All rights reserved.
doi:10.1016/j.plantsci.2008.08.009
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S.R. Bisgrove / Plant Science 175 (2008) 747–755
1. Introduction
Plants are sessile and cannot move when conditions become
unfavorable. They can, however, redirect their growth to
position stems, roots, leaves, and flowers towards the best
possible locations. Shoots grow towards light, maximizing
photosynthetic activity. Roots point down along the gravity
vector and can be attracted to areas of higher moisture or
nutrient content in the soil. When conditions change, plants
respond by redirecting their growth in accordance with the new
signals. Tropisms, these changes in the direction of growth in
response to stimuli, involve the bending of stems or roots that
reorient growth in the most favorable direction. Tropisms
include responses to a number of cues including gravity
(gravitropism), light (phototropism), and touch (thigmotropism), as well as gradients of moisture (hydrotropism), chemicals
(chemotropism), and temperature (thermotropism). Although
these tropisms have all been documented many have not been
investigated in any further detail [1]. Gravitropism and
phototropism have received the most attention, although some
studies addressing hydrotropism and thigmotropism have been
published in recent years [2–10].
Tropisms can be either positive or negative depending on
whether growth occurs towards or away from the stimulus. The
response occurs through a series of steps. Directional cues are
sensed and converted into biochemical signals. Under normal
conditions, plants receive many cues at once often from different
directions. To cope with all of this information incoming signals
must be integrated and transmitted to the responding cells where
the changes in growth occur (see ref. [11] for a review). Gravity is a
constant stimulus and changes in growth occur only when
opposing cues out compete or over-ride gravity [11]. Because of
the large amount of incoming information and the requirement for
an integrated growth response, the signaling pathways that
underlie tropisms are complex and they are not well understood.
However, this is an active area of investigation and it is discussed in
several recent reviews [1,6,11–19].
1.1. Signal perception
The receptors responsible for initiating phototropism, the
phototropins, were the first to be described at the molecular level
(see ref. [12] for a review). They are autophosphorylating protein
kinases that are activated by blue light [12,17,20]. In addition to
phototropins cryptochromes, another class of blue light receptors,
and phytochromes, red/far-red reversible photoreceptors, also
modulate phototropism [17,19,21]. Once activated, the phototropins transfer the light signal to proteins in downstream
signaling pathways, but as of yet only a few of these intermediates
have been identified [12,17,22].
Little is known about mechanoperception in plants, although
studies in bacteria and animals are providing information about
how mechanical stimuli are sensed in these organisms. The
relevant receptors appear to be mechanosensitive ion channels
[16]. These channels are transmembrane complexes that open in
response to mechanical forces exerted on the plasma membrane.
In some cases mechanical forces are thought to induce a
conformational change in the channel that opens it while in other
instances channels are opened indirectly via interactions with
signaling molecules or cytoskeletal proteins (see ref. [16] for
review). The first molecular characterization of mechanosensitive
ion channel proteins in plant membranes has recently been
published [7]. These MscS (mechanosensitive channel of small
conductance)-like proteins form pressure-sensitive channels in the
plasma membrane of protoplasts suggesting that they could be
mechanosensors [7]. However, the physiological relevance of these
channels in intact plants is unknown.
The receptors that mediate gravity perception have not been
identified, although two models that describe how gravity is
sensed have been put forward in the literature. The starch-statolith
hypothesis postulates that gravity-sensitive cells detect the falling
of intracellular masses called statoliths (see refs. [18,23] for
reviews). The gravitational pressure model proposes that the
settling of the protoplast within the cell wall is sensed by receptors
at the plasma membrane–extracellular matrix junction [18,24].
These receptors would be capable of detecting differences in
tension and compression between the plasma membrane and the
extracellular matrix at the top and bottom of the cell. Higher plant
cells that can sense gravity commonly contain starch-filled
amyloplasts that sediment and it is thought that this is the
primary mechanism by which gravity is perceived, although it is
possible that protoplast settling is also detected [18,23]. How
amyloplast sedimentation is converted into a biochemical signal is
not understood, although it has been proposed that mechanosensitive ion channels are involved [11,14,16,25,26].
With the exception of hydrotropism, plant responses to most
other directional cues have been described mainly from a
phenomenological perspective [1]. Recently genetic approaches
have been used in efforts to understand the mechanisms that
regulate hydrotropism [3–6,27,28]. Screens for mutants that are
defective in hydrotropic responses have been done [29] and the
characterization of the corresponding genes promises to yield
insight into the molecular mechanisms that regulate hydrotropism
[1].
1.2. Auxin redistribution and differential growth
Signals controlling tropisms often result in a concentration
gradient of auxin across a responding organ and this auxin gradient
is responsible for redirecting growth [17,19,30]. Consider, for
example, a seedling that has been placed on its side. Auxin
accumulates to a higher level on the lower flanks of the hypocotyl
and root. According to the Cholodony–Went hypothesis [31], an
auxin gradient triggers a differential growth response in which
cells on one side of the organ elongate more than cells on the other
side. Because the cells are held together and cannot move apart
from one another, differential growth results in the formation of a
Fig. 1. Bend formation in the hypocotyl and root of a seedling responding to light
and gravity. In the seedling on the left, the shoot (shaded in green) grows up
towards light; it is negatively gravitropic and positively phototropic. The root, on
the other hand, has the opposite response; it grows down and away from the light.
Cells in both the hypocotyl and the root expand mainly in the longitudinal direction
(indicated by double-headed arrows). The seedling in the middle has been placed on
its side and it now perceives a change in the direction of the light and gravity
vectors. Cells on the lower flank of the hypocotyl elongate more than the cells on the
upper flank (designated by large and small double-headed arrows respectively).
This differential growth response produces a bend that reorients the shoot into an
upright position (shown in the seedling on the right). A similar differential growth
response occurs in the root, but in this case cells on the upper flank elongate more
than the cells on the lower flank and the root bends down, towards the gravity
vector.
S.R. Bisgrove / Plant Science 175 (2008) 747–755
bend (Fig. 1). The Cholodony–Went hypothesis and auxin transport
have received a considerable amount of attention in the literature
and the reader is referred to several other articles for more detailed
discussions [25,30,32–37]. The mechanism by which auxin induces
differential cell elongation during organ bending is not understood,
but it is thought that auxin alters the rate at which cells elongate by
inducing changes in cell wall extensibility [38,39]. Ethylene also
influences organ bending by a mechanism that is not well
understood but appears to involve the modulation of auxin
transport [36,40].
1.3. Possible roles for microtubules
How might microtubules contribute to these growth
responses? Microtubules are thought to be involved in the
differential growth response that leads to organ bending (see
ref. [41] for review). Suggestions have also been made that
microtubules function earlier in the gravity and touch pathways,
possibly during signal perception or transduction [16,26,41]. These
ideas and the evidence that supports them are discussed in this
review. In addition, recent analyses that hint at new possibilities
for how microtubules might function during tropic responses are
presented.
2. Cortical microtubule organization and cell elongation
Microtubules are long, tubule-shaped polymers of a- and btubulin. Microtubules are actually highly dynamic, although they
appear stable in still photographs. They are almost always growing
or shrinking, providing them the flexibility to rearrange into
different arrays within the cell. Microtubules function in concert
with a large group of microtubule-associated proteins or MAPs.
MAPs bind to microtubules and while bound they modify
microtubule dynamics or mediate microtubule interactions with
other proteins or structures in the cell. The microtubule arrays that
appear in higher plants include preprophase bands, spindles, and
phragmoplasts in dividing cells, as well as the cortical arrays seen
in elongating cells. Plant MAPs are discussed in detail in several
recent reviews [42–46].
Microtubules are found in the periphery (or cortex) of
elongating cells just below the plasma membrane where they
are arranged in parallel hoops that encircle the cell (Fig. 2).
Elongation occurs in a direction that is perpendicular to the cortical
microtubules and the arrangement of the cortical microtubules is
important for controlling the direction in which cells elongate (as
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discussed below). When microtubules are disrupted by pharmacological or herbicide treatments or by mutation, cells stop
elongating and swell by expanding radially (see refs. [47–53] for
examples). In addition, there are several mutations that alter
cortical microtubule organization and plants with these mutations
have defects in the control of directional cell elongation. For
example, microtubules become very short and cells expand
radially when the temperature-sensitive mor1-1 mutants are
grown at the restrictive temperature [51]. The lefty and spiral
mutants comprise another class of microtubule mutants that have
a twisted pattern of growth [54–57]. These mutant plants can
organize their microtubules into parallel arrays, but instead of a
transverse alignment, the arrays form shallow helices with either a
right- or a left-handed orientation. Theoretically, twisted growth
occurs because cells are expanding perpendicularly to microtubules that have a helical rather than a transverse orientation
[58,59]. The radially swollen 6 (rsw6) mutants represent a third
class of temperature-sensitive microtubule mutants that have
defects in cell expansion. At the restrictive temperature rsw6
mutants have microtubules that are well organized into parallel
arrays within each cell but the orientations of microtubules among
neighboring cells are highly variable [60]. Apparently rsw6
mutants are defective in a mechanism that controls global
microtubule array changes across cells.
The examples described above all illustrate the importance of
cortical microtubules in cell elongation. Although the mechanisms
by which microtubules affect cell elongation are not completely
understood, it is thought they exert their influence by modifying
the cell wall and the wall then acts as a constraint for expansion
[61,62,52]. Expansion is driven by water uptake into the vacuole.
Water entering the vacuole causes an increase in volume that
exerts turgor pressure on the cell wall. When turgor exceeds the
resistance imposed by the wall the cell expands (see refs. [62–66]
for reviews).
Cellulose microfibrils and the linkages that hold them together
are thought to be the major load-bearing elements in the wall.
Because long and intact microfibrils are resistant to stretching
along their lengths, the wall yields to turgor by moving adjacent
microfibrils apart from one another (Fig. 2). Cellulose arrays in the
wall, therefore, constrain expansion to a direction that is
perpendicular to the microfibril orientation. The microfibrils are
embedded in a matrix of hemicelluloses, pectins and proteins, and
the extensibility of the wall also depends on the level of crosslinking between these molecules and the cellulose microfibrils.
When some of the connections that hold adjacent microfibrils
Fig. 2. Cells expand in a direction that is perpendicular to cortical microtubules and cellulose microfibrils in the cell wall. (A) A confocal section taken through expanding root
cells reveals cortical microtubules (green) in the upper cell that are aligned transverse to the long axis of the cell. The section has passed through more internal regions of the
lower cells. (B) Cartoon showing cellulose synthases moving through the plasma membrane. Microtubules below the plasma membrane guide the synthases and the wall
microfibrils are positioned parallel with the underlying microtubules. When the cell expands adjacent cellulose microfibrils move apart from one another and elongation
occurs in a direction that is perpendicular to the microtubule/microfibril alignment (indicated by double-headed arrows in A and B). Microtubules in (A) were visualized in a
transgenic Arabidopsis plant expressing a microtubule binding domain – green fluorescent protein fusion [144].
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S.R. Bisgrove / Plant Science 175 (2008) 747–755
together are disrupted, the wall becomes more extensible and cell
expansion increases. Thus, the orientation of cellulose microfibrils
in the wall and the degree of cross-linking largely determine the
direction and the rates at which cells expand [61–68].
2.1. Cellulose deposition
Cellulose microfibrils are produced by cellulose synthase
complexes embedded in the plasma membrane. The synthases
travel laterally in the membrane, extruding microfibrils to the
outside as they move [64–66]. This means that the arrangement of
microfibrils in the wall is determined by the paths that are taken by
the synthase complexes during cellulose synthesis. Microtubules
appear to be involved in guiding synthase movement, but the
mechanism is not completely understood. Cortical microtubules in
elongating cells are closely associated with the plasma membrane
and co-aligned with the microfibrils in the wall (Fig. 2). Based on
this observation, the microtubule/microfibril co-alignment model
postulates that cortical microtubules act as guides for the cellulose
synthase complexes as they move through the plasma membrane
(as reviewed in refs. [53,69,70]). This co-alignment model was
accepted for many years as the textbook version [71] for how
ordered arrays of microfibrils are deposited into the wall, even
though direct evidence linking microtubules with cellulose
synthase movement was lacking [69].
A direct connection between synthase movements and microtubules in elongating cells was finally established in 2006.
Synthase complexes moving in paths that coincided with cortical
microtubules were visualized in transgenic Arabidopsis plants
expressing cellulose synthase – yellow fluorescent protein fusions
[72]. Synthases were seen tracking along curved microtubules and
when microtubules were reoriented the synthases altered their
movement according to new microtubule positions.
Although these observations clearly link synthase movement to
microtubules, there is also evidence that suggests this is not the
whole story. When microtubules are disassembled either by
treatment with pharmacological or herbicidal agents, or by
mutation, synthase complexes continue to move in tracks that
are roughly parallel with each other and aligned arrays of
microfibrils continue to be deposited into the wall [50,72]. How
synthase movements might be guided in the absence of microtubules is not clear, but the findings do suggest that there is an
alternative or additional guidance mechanism [53,69,73].
Cells with compromised microtubules lose the ability to
elongate and they swell by expanding radially even though
ordered arrays of cellulose are deposited into their walls [50]. This
suggests that organized cellulose arrays alone are not sufficient for
directional control of cell expansion and that microtubules may be
influencing the wall in additional ways [50,53,73]. One explanation
is that microtubule disruption leads to the deposition of short
cellulose microfibrils [53]. According to the microfibril length
regulation model, walls with shorter microfibrils will be less able
to restrict cell expansion to one direction [53]. One could also
speculate that microtubules control the spatial arrangement of
additional molecules involved in cell wall modification and/or
synthesis. If so, microtubule disruption could result in the loss or
aberrant localization of molecules that stabilize or loosen the wall
leading to altered cell expansion. Microtubules have been
implicated in the localization of two other proteins involved in
cellulose synthesis, KORRIGAN (an endo-1,4-b-D-glucanase
involved in cellulose synthesis [74]) and COBRA (a glycosylphosphatidyl inositol (GPI) containing protein [75]). Both KORRIGAN and COBRA are distributed in linear arrays along the plasma
membrane and treatment with the microtubule depolymerizing
herbicide oryzalin disrupts this localization pattern [74,75].
2.2. Microtubule reorientations and tropisms
The concept that microtubules determine the orientation of
cellulose microfibrils in the wall has been put forth as an
explanation for how bend formation might be regulated [76,77].
Microtubule reorientations have been observed in the cells that are
involved in bend formation in plants that are responding to tropic
cues. Microtubules on the inner flanks of bending organs become
oriented parallel with the long axis of the organ while microtubules on the outer sides are transversely oriented [76–81]. If
cellulose synthase complexes follow microtubule tracks, then
microfibrils will be deposited in the same direction as the
reoriented microtubules. Since cells with transverse microfibrils
elongate more than cells with longitudinal microfibrils, the end
result will be organ bending [41,76,77,79,80]. In line with this idea
is the observation that microtubules reorient in response to auxin
[82–88]. Hence, the formation of an auxin gradient across a
tropically stimulated root or stem could trigger microtubule/
microfibril reorientations that lead to bending [41,77,80]. These
microtubule reorientations must also be coordinated across all of
the cells in the bend.
Although a model in which auxin-induced microtubule
reorientations are responsible for organ bending has been
suggested (see ref. [41] for a review) attempts to validate this
hypothesis have produced conflicting results [41]. One set of
experiments involves treating seedlings with agents that disrupt
microtubule organization or function. In some cases tropic bending
occurred while in other instances it was inhibited [84,89,90]. For
example, in some experiments maize roots pretreated with
microtubule depolymerizing agents were able to bend in response
to a gravity stimulus [89,84], although the total amount of
curvature was less than in untreated controls [84]. However, in
other experiments treating maize coleoptiles with herbicides that
depolymerized microtubules inhibited gravitropic bending even at
low concentrations that only partially eliminated microtubules
[90]. In contrast, phototropism proceeded even after complete
removal of cortical microtubules [90].
Imposing mechanical strain on an organ can also cause
microtubules to reorient [91]. This raises the possibility that the
reorientations during tropic responses result from the mechanical
strain produced when the organ bends. When mechanical
counterforces were used to prevent or reverse tropic curvatures
in maize coleoptiles and bean epicotyls, microtubules reoriented in
response to the mechanical strain rather than the tropic stimuli
[91–93]. On the other hand, microtubules in maize coleoptiles did
reorient in response to gravity when bending was physically
prevented, suggesting that tropic stimuli can induce microtubule
reorientations in the absence of a bend [80].
Issues have also been raised regarding the timing of the
microtubule reorientations that are proposed to result in
differential growth and organ bending. In some cases microtubule
reorientations that preceded organ bending were observed. For
example, in gravity stimulated maize coleoptiles [80] and cut
snapdragon spikes [94] microtubule reorientations occurred
before the organs bent. However, microtubule reorientations do
not always precede organ bending. For example, in gravity
stimulated maize roots organ bending was initiated before
microtubule reorientations occurred [78]. In addition, microtubule
reorientations that preceded organ bending were not consistently
observed when maize coleoptiles responding to light and gravity
stimuli were analyzed [95]. Another issue that has been raised is
based on the idea that changes in cell elongation rates due to
microtubule reorientations are likely to be time-consuming as they
involve reinforcing the cell wall through cellulose synthesis after
the microtubule reorientations occur [96]. In support of this idea,
S.R. Bisgrove / Plant Science 175 (2008) 747–755
correlations of cell elongation rates and microtubule/microfibril
alignments in elongating Arabidopsis roots indicate that microfibrils remain transversely aligned until well after microtubules
have reoriented [97]. Since the changes in cell elongation rates that
occur during plant tropisms are initiated quickly, it has been
argued that microtubule reorientation is unlikely to play a role in
the early stages of bend formation even if the reorientations occur
before the bend forms [96]. However, another suggestion is that
microtubule reorientations play a role later in bend formation,
perhaps as a mechanism to reinforce bending once it is initiated
[78].
Given all of the results discussed above, a simple model in
which auxin-induced microtubule reorientations lead to differential growth and organ bending is unlikely, although microtubules could influence cell expansion and organ bending in other
ways. In addition to affecting microtubule organization, auxin also
alters gene expression and ion homeostasis in cells [33–36,38,98–
102]. According to the ‘‘Acid Growth Theory’’ [38,98] auxin
stimulates cell expansion by triggering the extrusion of protons
into the wall. Acidification activates pH-sensitive enzymes and
these proteins increase cell expansion by cleaving the linkages
between polysaccharide components in the wall [38,39,98].
Another well-known effect of auxin is its ability to alter gene
transcription and it follows that a gradient of auxin across a stem or
root would result in the expression of different sets of genes on
each side of the organ. Proteins that enhance cell expansion are
synthesized preferentially on the faster growing flank where they
can stimulate growth. The cell wall loosening proteins known as
expansins localize preferentially to the more rapidly expanding
outer flank of gravistimulated maize roots [103]. Expression
analyses in Brassica oleracea seedlings have identified a set of
tropically stimulated genes that are preferentially expressed in the
cells where auxin levels are the highest. Among the up-regulated
genes were enzymes that have been associated with roles in cell
wall expansion [33,68]. These newly synthesized proteins must be
transported to their sites of action in the plasma membrane or cell
wall. Although speculative, it is possible that microtubules are
involved in positioning these molecules in the proper places at the
appropriate times (see Fig. 3 and discussion below). Note, in this
scenario microtubule disruption would not necessarily prevent
localization to the plasma membrane or cell wall, although it could
affect placements with respect to space and/or time.
3. Beyond cellulose deposition: new roles for microtubules?
Several lines of evidence suggest that microtubules could be
involved in organizational events associated with the plasma
membrane [104]. As mentioned above, the localization of both
COBRA and KORRIGAN, two proteins with roles in cell wall
biosynthesis, are microtubule-dependent [74,75]. Although these
proteins could be part of cellulose synthase complexes, their
localization in the plasma membrane is microtubule-dependent. A
link between microtubules and the localization of an arabinogalactan cell surface GPI-linked protein from tomato has also been
reported [105]. Microtubule disruption caused a relocalization of
this protein [105]. GPI-anchored proteins have been associated
with protein-containing membrane microdomains or lipid rafts in
the plasma membrane [106–108]. Evidence from the animal
literature suggests that lipid rafts may have roles in signaling
processes and it is thought that they could serve as centers for
signaling cascades [109,110]. It has been proposed that similar
membrane microdomains may also be present in plant cells
[111,112]. Although speculative, one possibility is that microtubules play a role in positioning signaling proteins and/or
complexes in the plasma membrane in a manner similar to the
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mechanism by which cellulose synthase complexes are guided
during cellulose deposition (Fig. 3A and B). In such a scenario,
microtubules would influence cell elongation and organ bending
by coordinating the positions of signaling molecules in the plasma
membrane with the structural components of the wall that are
modified during the response (Fig. 3B).
In animal and fungal cells the rapidly growing or plus-ends of
microtubules also interact with proteins localized in the cortex of
the cell next to the plasma membrane (reviewed in refs. [113–
115]) and there is evidence that suggests microtubule plus-ends
have roles in plant tropisms. Arabidopsis plants carrying mutations
in the microtubule plus-end binding protein END BINDING 1 (EB1)
are slower to initiate bends after touch and/or gravity stimuli
[116]. The delay appears to be specific for the initiation of
differential growth as mutants grow at the same rate as wild type
plants. Mutant roots have a tendency to grow in loops on tilted agar
surfaces, indicating that they also have difficulty turning off
differential growth [116].
How might EB1 influence differential growth during tropic
responses? Although the mechanism is unknown, EB1 proteins have
been the object of intense scrutiny in animal and fungal cells and
results from these studies could provide insight into the roles of EB1
in plants. EB1 accumulates preferentially on the rapidly growing (or
plus) ends of microtubules in all of the organisms in which it has
been studied [113,115,117–120]. While bound to the microtubule
end EB1 proteins usually make microtubules more dynamic [113].
The increase in dynamics is thought to help microtubules search
the cytoplasm for ‘‘capture’’ sites. When an appropriate site is
encountered EB1 mediates interactions between the microtubule
end and other proteins localized at the site [113,121]. Capture sites
often contain actin and EB1 is known to mediate interactions
between microtubules and actin in some cell types (reviewed in ref.
[113]). The interactions of microtubule ends with capture sites
serves several functions including the targeted delivery of vesicles,
signaling molecules, and ion channels to specific places in the cell
[113,114,122–131]. Whether a similar system operates in plant cells
is not known. However, one could speculate that the interaction of
microtubule ends with specific sites in the cortex of the cell plays a
role in tropic responses (see Fig. 3C). A captured microtubule could
direct the insertion, removal, or position of proteins and/or
complexes at specific sites in the cortex of the cell by serving as a
track along which cargo-bearing microtubule-based motors like
kinesins would travel. Alternatively, interactions between microtubule ends and capture sites could also be transient in nature.
Microtubule plus-end binding proteins like EB1 appear to serve as
platforms for recruiting signaling molecules to the active ends of
microtubules (reviewed in ref. [113]). In this case a brief encounter at
an appropriate capture site would be sufficient to deliver a specific
factor.
The idea that microtubules might be involved in the spatial
integration of endo- and exocytotic events has been proposed
[104]. In addition to a role in the localization of the secreted
proteins COBRA, KORRIGAN, and the arabinogalactan protein
discussed above, microtubules also appear to have roles in
endocytosis in plant cells [132,133]. During preprophase band
formation in cultured tobacco BY-2 cells endocytotic vesicles are
internalized in trans-vacuolar cytoplasmic strands that form
connections between the nucleus and the cortex of the cell. These
cytoplasmic strands contain microtubules and treatment of cells
with an herbicide that depolymerizes microtubules disrupts
vesicle internalization [132]. Ligand-mediated endocytosis of a
receptor involved in plant defense responses (the flagellin receptor
FLS2) also utilizes microtubules [133]. Endocytosis of FLS2 is
thought to be a regulatory step in the signaling pathway [133]. One
could speculate that these microtubule-dependant endocytotic
752
S.R. Bisgrove / Plant Science 175 (2008) 747–755
Fig. 3. Possible roles for microtubules in plant tropisms. (A) According to the microtubule/microfibril co-alignment model microtubules serve as guides that influence the
movement of cellulose synthase complexes during cellulose deposition. In response to tropic cues, there is a microtubule reorientation that has been proposed to change the
direction of cellulose deposition and reduce the amount of cell elongation on the inner flank of a bending organ. Although the change in cell elongation rate may be too slow to
play a role in bend initiation, the reorientations could be involved at later stages, perhaps to reinforce the bend once it is initiated. (B) Although speculative, it is possible that
microtubules could constrain the positions of signaling proteins and/or complexes in the plasma membrane in a way that is similar to the mechanism postulated in the
microtubule/microfibril co-alignment model. This would enable the plant to spatially coordinate structural components of the wall with the signaling molecules that trigger
changes in wall extensibility during tropic responses. (C) Another way that microtubules could influence tropism is through interactions of their ends with specific sites in the
cortex of the cell. Proteins that preferentially bind the more rapidly growing or plus-ends of microtubules, such as EB1, can mediate both microtubule dynamics and
interactions of the microtubule end with other proteins or structures in the cell. EB1 tends to make microtubules more dynamic and this increase in dynamics is thought to
facilitate microtubule searching of the cytoplasm for specific ‘‘capture’’ sites. In animal and fungal cells EB1 is also known to interact with several other proteins including
signaling molecules. When the microtubule end encounters a suitable site proteins associated with the microtubule end (yellow balls) and proteins localized at the capture
site (blue balls) can interact with one another. Capture sites often contain actin and EB1 is sometimes involved in crosstalk between the two cytoskeletal structures (for
simplicity actin is not shown in A or B). Transient encounters would be sufficient for the release of factors associated with the microtubule end while stably connected
microtubules could serve as tracks that direct the movement of cargo-bearing motor proteins (not shown) to and from the capture site. These interactions could play a role in
tropisms by delivering signals or materials that are needed to regulate organ bending to the appropriate sites in the cortex of the cell. They could also be involved in signal
perception either via connections to mechanosensory ion channels in the plasma membrane or by mediating the removal or insertion of signaling molecules such as receptors
into or out of the plasma membrane.
events involve the interactions of microtubule ends with cortical
capture sites. Other plasma membrane proteins, including
hormone receptors and ion channels, are also internalized in
plant cells, although the cytoskeletal requirements for many of
these endocytotic events have not been determined [134–136].
Microtubules have been implicated with roles in the perception
of touch and gravity signals [16,26,41,90,95,137–139]. There are
reports that microtubule inhibitors modify the rate of amyloplast
sedimentation and inhibit auxin transport during gravitropism
(reviewed in ref. [41]). The mechanism by which gravity is converted
S.R. Bisgrove / Plant Science 175 (2008) 747–755
into a biochemical signal is unknown, although one possibility is that
falling amyloplasts could be sensed by mechanosensitive ion
channels [26]. Touch sensing is also thought to be mediated by
mechanosensitive ion channels, a hypothesis that is based in part on
analyses conducted on touch receptor neurons in the nematode
worm Caenorhabditis elegans [16,26]. Genetic analyses indicate that
mutations in tubulins associated with these neurons cause loss of
mechanoresponse [140]. These microtubules are thought to form
tethers that are linked, either directly or indirectly, to ion channels in
the plasma membrane [141,142]. Force from a touch stimulus is
thought to displace the microtubule tethers and this induces a
conformational change in the channel that activates it [26,140,141].
Ca2+ channels have been implicated in touch responses in plants
[9,16,26] and patch-clamping analyses have detected mechanosensitive calcium channels that are affected by microtubule
inhibitors [143]. However, the molecular identity of these channels
is unknown. Proteins that function as mechanosensitive ion
channels in plants have recently been identified [7,10]. Whether
these channels are involved in touch or gravity sensing and how
their activities are regulated is unknown.
4. Summary and future directions
Although microtubules have been associated with plant
tropisms for many years our understanding of how they participate
in these growth responses is far from complete. One proposal is
that microtubules function during the differential growth response
that leads to organ bending [41,76,77,79,80]. This idea is based on
the concept that microtubules determine the direction of cell
elongation by guiding the deposition of cellulose microfibrils into
the cell wall. However, the hypothesis is controversial. In particular
there are questions surrounding the timing of microtubule/
microfibril reorientations with respect to the onset of differential
growth [78,80,94,96,97]. Several recent reports indicate that
microtubules also have roles in organizational events associated
with the plasma membrane [74,75,105,132,133]. In addition,
plants carrying mutations in the microtubule plus-end binding
protein EB1 exhibit delayed responses to touch/gravity signals
[116]. Although speculative, these studies raise the possibility that
microtubules could influence the wall in ways that are independent of a role in guiding cellulose deposition.
Another possibility is that microtubules function in the
perception of touch and gravity signals [16,26,41,90,95,137–
139]. There is evidence that indicates microtubules are involved
in the activation of mechanosensory ion channels in animal cells
[140,141,142] and microtubules appear to affect the activity of
mechanosensitive Ca2+ channels in plant membranes [143].
However, mechanistic links between microtubules and the
mechanoreceptors that might be involved in touch/gravity sensing
are speculative.
One way to assess how microtubules function during tropisms
is to analyze proteins that interact with microtubules. Several
microtubule-associated proteins have been identified in plants
(see refs. [43,46] for reviews). Many of these proteins affect the
stability or organization of microtubule arrays [46] and the
corresponding mutants often have cell expansion defects that
make it difficult to measure tropic responses. Because plants
carrying mutations in EB1 genes have defects in touch/gravity
responses they hold promise for providing further insights into the
role of microtubules in tropisms. Biochemical screens for proteins
that interact directly with EB1 could identify proteins that are
important for either sensing touch/gravity cues or initiating
differential growth. Genetic screens for enhancers or suppressors
of the eb1 mutant phenotype could identify genes that function at
different steps in the same or parallel pathways. These analyses
753
hold the potential to yield new insights into how microtubules
function in plant cells.
Acknowledgement
S.R.B. is funded by a Natural Sciences and Engineering Research
Council of Canada Discovery Grant (Application 331017).
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