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Research Article
Microtubules and CESA tracks at the inner epidermal
wall align independently of those on the outer wall of
light-grown Arabidopsis hypocotyls
Jordi Chan1,*, Magdalena Eder1,‡, Elizabeth Faris Crowell2,§, Janet Hampson1, Grant Calder1 and Clive Lloyd1
1
Department of Cell and Developmental Biology, John Innes Centre, Colney, Norwich NR4 7UH, UK
Institut Jean-Pierre Bourgin, UMR1318 INRA-AgroParisTech, INRA Centre de Versailles-Grignon, Route de St-Cyr (RD10), 78026 Versailles
Cedex France
2
*Author for correspondence ([email protected])
‡
Present address: INM–Leibniz Institute for New Materials, Campus D2-2, 66123 Saarbrücken, Germany
§
Present address: Membrane Traffic and Cell Division Research Group, Institut Pasteur, 28 rue du Dr Roux, 75015 Paris, France
Journal of Cell Science
Accepted 20 February 2011
Journal of Cell Science 124, 1088-1094
© 2011. Published by The Company of Biologists Ltd
doi:10.1242/jcs.086702
Summary
Microtubules are classically described as being transverse, which is perpendicular to the direction of cell elongation. However, fixation
studies have indicated that microtubules can be variably aligned across the epidermis of elongating shoots. In addition, microtubules
are reported to have different orientations on inner and outer epidermal surfaces, undermining the idea of hoop-reinforcement. Here,
long-term movies of Arabidopsis seedlings expressing GFP–TUA6 allowed microtubule alignment to be directly correlated with the
rate of elongation within individual growing cells. We also investigated whether microtubule alignment at the inner or the outer
epidermal wall better reflected the growth rate. Movies confirmed that transverse microtubules form on the inner wall throughout
elongation, but orientation of microtubules is variable at the outer wall, where they tend to become transverse only during episodes of
accelerated growth. Because this appears to contradict the concept that circumferential arrays of transverse microtubules or microfibrils
are essential for cell elongation, we checked the organisation of cellulose synthase tracks using GFP–CESA3 and found a similar
mismatch between trajectories on inner and outer epidermal surfaces. We conclude that microtubule alignment on the inner wall
appears to be a more stable predictor of growth anisotropy, whereas outer-wall alignment is more sensitive to the elongation rate.
Key words: Microtubules, Cell wall, Cellulose microfibrils, Cellulose synthases, Cell elongation
Introduction
When they were first seen in the electron microscope, cortical
microtubules were described as transverse ‘hoops’, matching the
alignment of overlying cellulose microfibrils in the cell wall
(Ledbetter and Porter, 1963). This agreed with the biophysical
concept of hoop-reinforcement (Green, 1962), in which transversely
aligned, load-bearing cellulose microfibrils maintained the girth of
the expanding cell, directing growth into elongation orthogonal to
the net alignment. This picture seems to apply to the root where
microtubules are transverse in rapidly elongating cells, reorienting
to oblique or longitudinal as growth declines (Baskin, 2001;
Granger and Cyr, 2001; Sugimoto et al., 2000). However, reports
suggest that the situation is not so straightforward in shoots, where
cortical microtubules beneath the outermost epidermal wall can be
oblique or longitudinal or mixed, as well as transverse, apparently
during cell elongation (Duckett and Lloyd, 1994; Le et al., 2005;
Sawano et al., 2000; Takeda and Shibaoka, 1981; Yuan et al., 1995;
Zandomeni and Schopfer, 1993).
Various studies have indicated that rather than representing
stable configurations, microtubule arrays in shoots might undergo
a regular programme of reorientation (e.g. Mayumi and Shibaoka,
1996). Microtubules in pea shoot epidermal cells, which had been
microinjected with fluorescent tubulin, have been seen reorienting
from transverse to longitudinal (Yuan et al., 1995) and in the
reverse direction upon addition of gibberellic acid (Lloyd et al.,
1996). In their immunofluorescence studies on mung bean shoots,
the Shibaoka laboratory observed longitudinal or oblique, as well
as transverse microtubules in elongating tissue and suggested this
variability to be an expression of a cyclic reorientation
physiologically regulated by a range of hormonal factors (Mayumi
and Shibaoka, 1996; Takesue and Shibaoka, 1998). In his
immunofluorescence study of rapidly elongating epidermal cells in
the sunflower hypocotyl, Hejnowicz (Hejnowicz, 2005) also saw
microtubule arrays in a variety of alignments. He hypothesised that
the variability of alignment reflected stages sampled from a
rotational clock-like cycle, rather than the oscillatory cycle proposed
by the Shibaoka laboratory. Chan and colleagues (Chan et al.,
2007) then made long-term movies of Arabidopsis hypocotyl
epidermal cells, confirming that microtubules beneath the outer
epidermal wall do indeed undergo both discontinuous shifts
(‘jumps’) in alignment and continuous rotary movements in a
clockwise or anticlockwise direction (which are collectively termed
‘persistent reorientation’). However, neither rotations like this, nor
Shibaoka-like oscillatory cycles have been seen in living
Arabidopsis root epidermal cells imaged with GFP–MBD (Granger
and Cyr, 2001), underlining the differences between roots and
shoots.
Similarly to microtubules, cellulose microfibrils are classically
considered to be transverse in rapidly elongating tissue, providing
hoop-reinforcement during expansion. Baskin (Baskin, 2005),
however, concludes that the behaviour of the stem epidermis is
‘paradoxical’ because the epidermis of aerial tissue has been
Microtubule alignment in Arabidopsis epidermis
Journal of Cell Science
reported to contain longitudinal microfibrils, even in rapidly
elongating areas. Non-hoop-like behaviour is also suggested by
reports that microtubules on the outer epidermal surface are not
necessarily co-aligned with microtubules on the inner epidermal
surface (Busby and Gunning, 1983; Flanders et al., 1989).
This major difference in apparent behaviour of cortical
microtubules in shoot and root tissues prompted us to look at the
relationship between the alignment of microtubules and cell
elongation in light-grown hypocotyls, paying attention to both
outer and inner tangential walls of epidermal cells. Arabidopsis
seedlings expressing GFP–TUA6 were grown in special growth
chambers (Chan et al., 2007) and live changes in microtubule
alignment were monitored over several hours. In this way, instead
of basing conclusions on populations of cells with potentially
unsynchronised microtubule orientations, it was possible to
dynamically correlate the behaviour of microtubules with the
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variable growth rate for up to 24 hours at a time, within individual
cells. This showed that microtubules on the outer epidermal wall
behaved differently to microtubules on the inner wall of the same
cell; a finding that we confirmed with a cellulose synthase marker.
This allows us to discuss the role of microtubules in growth
anisotropy.
Results
Relationship between growth and microtubule alignment
at the outer epidermal face
In a previous study (Chan et al., 2007), persistent microtubule
reorientation was observed in Arabidopsis seedlings expressing
GFP–EB1a. Here, we used the same experimental procedure except
that microtubules in 2- to 5-day-old seedlings were labelled with
GFP–TUA6 (Ueda et al., 1999). This probe maintains bright
fluorescence, enabling confocal z-stacks of epidermal cells in the
Fig. 1. Changes in microtubule alignment
during elongation of epidermal cells of
Arabidopsis hypocotyls expressing TUA6–GFP.
(A)Low-magnification image of cells from slowgrowing hypocotyl displaying non-coordinated
orientations of microtubules. (B)Lowmagnification image illustrating coordinated
transverse arrays that develop across neighbouring
cells during phases of rapid cell elongation.
(C)Montage of movie frames of the outer surface
of an epidermal cell displaying a fixed transverse
alignment of microtubules during rapid
elongation. (D)Microtubules displaying oblique
or longitudinal arrays when growth has ceased.
(E)Kymograph along the long axis of an
elongating cell showing transitions between slow
and rapid elongation (single arrowhead) and rapid
to no elongation (double arrowhead). Rate of
elongation is indicated inm per hour.
(F)Montage of microtubule alignments that
occurred during the changes in elongation rate
shown in E. Note that microtubules tend to net
transverse orientation when elongation rate
increases and then change to oblique when
elongation ceases. Orientation of microtubules is
indicated by white lines. (G)Low-magnification
images of neighbouring cells during slow
elongation (t0), variable alignment; rapid
elongation (t210), transverse alignment; and no
elongation (t645), longitudinal or oblique
alignment, respectively. Scale bars: 12m (A,B),
18m (E, y axis), 119 minutes (E, x axis), 20m
(G). Strip width: 4m, taken every 20 minutes;
numerals show hours (C,D).
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middle portions of hypocotyls to be captured every 15–30 minutes
for up to 24 hours.
Three different general patterns of microtubules were seen at the
outer epidermal surface of the hypocotyl, which appeared to change
in a growth-related fashion. Most commonly, microtubule alignment
differed between neighbouring cells (Fig. 1A). This particular
pattern was associated with relatively slow growth rates (average
cellular elongation rate of 1.27 m/hour; s.d.1.4, s.e.0.14 or an
average relative elongation rate of 0.02/hour; s.d.0.02, s.e.0.002,
n99 cells; 12 hypocotyls). Movies confirmed that microtubules
undergo persistent reorientation in such tissues (Chan et al., 2007).
Importantly, although microtubules could be transiently transverse
in these persistently reorienting arrays, in such slower-growing
cells the phase of reorientation was not synchronised across the
epidermis (Hejnowicz, 2005; Chan et al., 2007).
A different pattern of microtubule alignment was observed in
relatively fast-growing hypocotyls (with an average cellular
elongation rate of 3.19 m/hour; s.d.3.78; s.e.0.54 or an average
relative elongation rate of 0.06/hour; s.d.0.07, s.e.0.01, n49
cells). In these tissues, transverse alignments of microtubules were
observed that were coordinated across fields of neighbouring cells
(Fig. 1B). Transverse alignment could be maintained over several
hours, illustrating that it is not a fixed stage extracted from the
rotary process (Fig. 1C).
A third pattern was observed between days four and five, when
growth ceased. In such tissues, coordinated alignments were also
sustained between neighbouring cells, except in these cases,
microtubules were arranged in oblique or longitudinal arrays (Fig.
1D). Such oblique alignments – similarly to the transverse
alignment of faster-growing cells (Fig. 1B) – could be maintained
over several hours.
Analysis of movies showed that hypocotyls did not maintain
constant growth rates and transitions between relatively fast and
slow elongation rates (and vice-versa) were captured. Such movies
were used to analyse the dynamics of the development of transverse
alignments with respect to growth (supplementary material Movie
1). Fig. 1F provides an example of microtubule alignment as the
growth rate changes. To obtain these data, the length of the cell
within a movie frame was measured by projecting a one-pixelwide line from one end wall to the other. After extracting a time
series of such lines from the movie, the bottom ends were aligned
horizontally, so that the upper ends traced a curve that shows the
rate of cell elongation (Fig. 1E). A corresponding montage of the
different microtubule alignments that occurred over the same period
of elongation is shown in Fig. 1F (which is best seen by magnifying
the image online). During more rapid growth (Fig. 1E),
microtubules tended towards transverse alignment (supplementary
material Movie 2), which was coordinated across fields of cells
(Fig. 1G; t210, compare with the more variable alignments at
t0). But as the growth rate reached a plateau (Fig. 1E), the
corresponding panels of microtubules (which are aligned in Fig.
1F), switch to a more oblique alignment. This alignment was also
coordinated across the tissue (Fig. 1G; t645).
Observation of those transitions where the rate of elongation
accelerated revealed two features of the evolution of transverse
microtubule alignment. First, transverse arrays are a transient
phenomenon and tend to develop before phases of more rapid
elongation (Fig. 2A,B). Out of 41 cells monitored long-term in five
hypocotyls filmed as they changed from slow to faster elongation,
35 evolved transverse alignment an average of 184 minutes (3.07
hours) (s.d.170 minutes; s.e.27) before a measurable burst of
Fig. 2. Transverse microtubule alignments develop at the outer epidermal
surface of hypocotyl cells during rapid elongation. (A)Kymograph along
the long axis of an elongating cell showing a transition from slow
(0.6m/hour) to more rapid elongation (3.6m/hour). (B)Corresponding
montage of microtubule alignments that occurred during elongation shown in
A. Note that microtubules become transverse before the transition to rapid
elongation. (C)Kymograph along the long axis of an elongating cell showing
a transition from slow (0.4m/hour) to rapid elongation (3.0m/hour).
(D)The corresponding montage of microtubule alignments that occurred
during elongation shown in C. Note that microtubules become transverse at the
transition to rapid elongation. Scale bar: 28m (A, y axis), 16m (C, y axis),
660 minutes (A,C, x axis). Strip width: 3m (B,D). Arrows denote distance (y
axis) and time (x axis).
elongation, whereas the arrays of six cells became transverse at the
transition to more rapid growth (Fig. 2C,D). During more rapid
elongation, microtubules spent on average 370 minutes (6.2 hours)
in transverse alignment (s.d.212 minutes; s.e.42, 25 cells, 4
hypocotyls). Second, transverse alignment is not necessary to
maintain rapid elongation. For instance, supplementary material
Movie 1 shows transverse arrays reorienting towards the
longitudinal axis despite the fact that analysis showed no decrease
from the maximum elongation rate. This demonstrates that although
arrays might have a tendency to become transverse before rapid
elongation, it is not essential for this alignment to be maintained
throughout the entire process of more rapid elongation.
In summary, persistent reorientation is mainly seen at the outer
epidermal surface during hypocotyl growth (between days two and
three and days three and four) but can be interrupted by transient
periods of coordinated transverse alignment before rapid elongation.
When growth diminishes between days four and five, there is a
switch towards an oblique or longitudinal alignment that is also
coordinated between groups of cells. Alignment of microtubules
Microtubule alignment in Arabidopsis epidermis
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beneath the outer epidermal wall, coupled with the extent to which
this is coordinated between neighbouring cells, therefore serves as
a general indicator of the elongation status of the tissue.
Journal of Cell Science
Relationship between growth and microtubule alignment
at the inner epidermal face
Next, we investigated whether microtubules on the inner
tangential wall (in contact with the cortical cells) of the epidermis
display similar behaviour to those on the outer tangential surface
(in contact with the external environment). First we collected zstacks of 2-, 3- and 4-day-old hypocotyls. By separating the outer
and inner epidermal surfaces within z-stacks, it was possible to
compare microtubules on inner as well as outer tangential surfaces
of the same cell. Arrays were categorised on a ‘one-cell–onemicrotubule alignment’ basis (transverse0±22.5°, oblique
45±22.5°, longitudinal90±22.5°), but where an overall alignment
of domains could not be clearly determined they were classified
as ‘mixed’.
Quantification showed that microtubule alignments on the inner
epidermal wall were distributed differently to those on the outer
surface (Fig. 3A). In 2-day-old hypocotyls, which had just emerged
from the seed, microtubules of various orientations (oblique,
transverse, mixed and longitudinal) were present on both the inner
and outer epidermal walls, consistent with uncoordinated
reorientations described above. However, in elongating 3-day-old
seedlings, mixed arrays formed the predominant category on the
outer wall. By contrast, on the inner epidermal surface, the
predominant alignment was transverse, with 62% (s.d.23%, 78
cells in 7 hypocotyls) of cells organised in this way. This was
double the number of transverse arrays sampled at the outer surface
(33%, s.d.18%, 78 cells in 7 hypocotyls). This was consistent
with findings from the dynamic studies, which showed that
microtubules on the inner epidermal wall of 3-day-old hypocotyl
cells had stopped reorienting and became aligned in coordinated
transverse alignments. At this stage, none of the cells displayed
longitudinal arrays. At day four, approximately equal proportions
of oblique and transverse microtubules were seen on both inner
and outer surfaces.
This analysis indicated that microtubules could be differently
aligned on different surfaces of the same cell, with microtubules
having an increased tendency to maintain transverse alignment at
the inner wall of three-day-old hypocotyls. Movies made of the
inner and outer walls of the same cell supported this conclusion.
The key frames from supplementary material Movie 3 (see Fig. 3)
show that microtubules upon the inner epidermal wall remain
transverse (Fig. 3B, 0–270 minutes) throughout a period when
microtubules rotate through 180 degrees upon the outer surface of
the same cell (Fig. 3C). Microtubules associated with the inner
epidermal wall therefore serve as a better indicator of the axis of
cell or tissue elongation.
The sensitivity of inner wall alignment to growth was examined
between days three and four by making kymographs of hypocotyl
cells found to be undergoing fluctuations in the elongation rate.
Only cells whose inner tangential wall was not too facetted by
contact with subjacent cells had a sufficiently flat surface for this
study. In contrast to the upper surface, detailed kymographic
analysis of six cells revealed that microtubule alignment on the
inner surface was less sensitive to decelerations and accelerations
in the rate of growth. The kymograph in Fig. 4A shows the endwall trace of an elongating cell in which growth plateaus for a
while before undergoing a sudden acceleration.
Fig. 3. Microtubule orientation can be uncoupled on different faces of the
same cell. (A)Quantification of microtubule orientation along the outer versus
inner surfaces of hypocotyl cells at different days of development. Note that
microtubule orientation is uncoupled on day two, when inner walls maintain a
higher percentage of transverse alignments; this is more pronounced on day
three. Values are mean ± s.e. (B,C)Montage of movie frames showing the
inner (B) and outer (C) tangential surfaces of a pair of cells. Note that
microtubules rotate over time at the outer surface (C) whilst maintaining
transverse alignment on the inner surface (B). Time is shown in minutes. Scale
bar: 15m (B,C).
Fig. 4A shows an end-wall trace where growth plateaus then
accelerates. In Fig. 4B, each strip shows microtubule alignment at
the outer epidermal wall, corresponding to the end-wall trace
directly above it (in Fig. 4A). This shows that the alignment of
microtubules at the outer epidermal surface varies over time, but
that after a period in which the growth rate plateaus, microtubules
tend towards transverse alignment as the growth rate accelerates
once more. By contrast, the microtubules upon the inner surface of
the same cell (Fig. 4C) were notably less variable, remaining
mainly transverse throughout. Microtubules upon the inner surface
could remain net transverse for prolonged periods of time. Such an
array was tracked for 20 hours (Fig. 4C; supplementary material
Movie 4), in contrast to the shorter episodes of transverseness on
the upper surface of the same cell. This suggests that transverse
alignment of microtubules on the inner epidermal wall is relatively
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Journal of Cell Science
Fig. 4. Mismatch between inner and outer wall microtubule alignment
during changes in growth rate. (A)Kymograph along the long axis of an
elongating cell showing a change in the rate of elongation, from slow
(0.2m/hour) to more rapid elongation (2.0m/hour). (B)Corresponding
montage of microtubule alignments that occurred along the outer surface of
the cell during elongation. (C)Montage of microtubule alignments that
occurred along the epidermal surface of the same cell. Note that microtubules
remained in a net transverse orientation on the inner wall regardless of changes
in growth rate. Scale bar: 298 minutes (A, x axis), 10.5m (A, y axis).
insensitive to fluctuations in elongation rate. By contrast,
microtubules on the outer face rotate in slow-growing cells and
only display an increased tendency towards transverseness during
phases of accelerating growth.
In addition to examining outer and inner tangential or periclinal
epidermal surfaces, we looked at the connecting radial or anticlinal
walls. Similarly to the inner surface, these maintained transverse
microtubule alignment, whereas a more pronounced realignment
was confined to the outer surface (see supplementary material
Movie 4).
Orientation of cellulose microfibrils at the inner epidermal
surface
The preceding analysis indicated that microtubules can adopt
various alignments on the inner epidermal surface at day two, but
at day three, they have a stronger tendency than those in the outer
wall for transverse alignment. To view the most recently deposited
layer of cellulose microfibrils upon this inner epidermal surface,
the outer epidermal wall was removed by sectioning, the cell
contents and plasma membrane were removed with detergent, and
the exposed surface of the inner epidermal wall examined by field
emission scanning electron microscopy (FESEM). In 2-day-old
hypocotyls (77 cells, 5 hypocotyls), the inner wall showed various
alignments of microfibrils that differed from cell to cell, with
‘oblique’ forming the largest category (Fig. 5A–E). In 3-day-old
hypocotyls (51 cells, 6 hypocotyls) there was a doubling in the
percentage of transversely aligned microfibrils, which now formed
the predominant category, followed by oblique, whereas virtually
no longitudinal alignments were seen at this stage (Fig. 5A). This
pattern was reversed at day four (34 cells, 4 hypocotyls) when
transverse alignments were almost absent with oblique or
longitudinal forming the predominant category Fig. 5A.
Comparison of cellulose synthase tracks on inner and
outer cell surfaces
After observing that microtubules can display different alignments
on opposing tangential surfaces of the same cell, we investigated
Fig. 5. Analysis of cellulose microfibrils along the inner walls of hypocotyl
epidermal cells. (A)Quantification of cellulose microfibril alignment along
the inner walls of hypocotyl epidermal cells at different days of development.
Note that cellulose microfibrils along inner walls have a greater tendency
towards transverseness on day 3. Results are mean ± s.e. (B–E) FESEM
images of cellulose microfibril orientations at the inner wall of Arabidopsis
epidermal cells: (B) longitudinal, (C) oblique, (D) transverse and (E) mixed
orientation. Scale bars: 200 nm.
whether this also applies to cellulose synthase tracks on the outer
and inner tangential epidermal walls. This was performed with
seedlings expressing GFP–CESA3 under control of its own
promoter in the cesa3je5 mutant background (Desprez et al., 2007).
The GFP–CESA3 labelling pattern in light-grown hypocotyl cells
was consistent with that seen in dark-grown cells, i.e. cellulosesynthesising particles were observed upon the cortex, whereas
faster-moving Golgi bodies were seen deeper in the cytoplasm
(Desprez et al., 2007).
In particular, the CESA particles moved along the plasma
membrane in linear tracks, as previously described (Crowell et al.,
2009; Paredez et al., 2006). Although this was difficult to discern
upon the inner epidermal wall, as a result of the dimmer
fluorescence and basal accumulation of Golgi bodies (particularly
in the small cells of 2-day-old seedlings), it was possible to identify
cells in which CESA tracks on the inner wall did not mirror the
alignment of tracks on the outer wall of the same cell. This was
observed more clearly in the longer cells of 3-day-old hypocotyls.
In this case, GFP–CESA3 tracks were aligned transversely upon
the inner epidermal surface, whereas a variety of alignments were
observed on the outer surface (Fig. 6A–D).
In summary, just as microtubules do not necessarily form
transverse ‘hoops’ around illuminated epidermal cells, the cellulose
synthases also form tracks that can be aligned in different
orientations on different surfaces of the same cell.
Journal of Cell Science
Microtubule alignment in Arabidopsis epidermis
Fig. 6. Comparison of cellulose synthase (CESA3) tracks along outer and
inner surfaces of hypocotyl epidermal cells. Limited z-stack projections of
3-day-old hypocotyl cells expressing GFP–CESA3. CESA tracks on outer
(A,C) and inner walls (B,D) of the same cell. This shows that cellulose
synthase tracks can be differently oriented on opposite faces of the same cell.
Scale bar: 10m.
Discussion
This study of individual hypocotyl epidermal cells shows that
microtubules display complex three-dimensional reorganisations
in which alignment of microtubules on different faces of the cell
can vary in a growth-sensitive manner. In the early stages of
growth, 2-day-old hypocotyl cells had various alignments
(transverse, oblique, longitudinal or mixed) on both inner and
outer surfaces. But as growth proceeded, microtubules on the inner
walls of 3-day-old hypocotyls tended to display coordinated
transverse alignments. These illuminated Arabidopsis seedlings do
not grow as fast as etiolated plants (Gendreau et al., 1997; Le et
al., 2005) and microtubules at their outer epidermal surface can
undergo rotary movements during slow growth, but during phases
of accelerating growth, hypocotyls show a tendency to switch to
transverse alignment when imaged for long periods. Although this
switch in alignment tended to occur before accelerations in growth
rate, it was not tightly coupled with the actual change in elongation
rate, nor was it essential for this alignment to be maintained
throughout the process of rapid elongation. Then, as growth
diminished, the pattern changed once more, consisting of arrays in
which oblique and steeply oblique or longitudinal microtubules
were present on both outer and inner surfaces. The same trend was
noted for microtubule alignment on the outer surface of lightgrown Arabidopsis seedlings expressing GFP–TUA6 (Le et al.,
2005). This pattern is also very similar to that reported in a live
cell study of Arabidopsis root epidermal cells expressing GFP–
MBD (Granger and Cyr, 2001). There, transverse microtubules
were mainly associated with accelerating growth, whereas a variety
of orientations was found as the growth rate declined. From their
investigation on fixed Arabidopsis root cells, Sugimoto and co-
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workers (Sugimoto et al., 2000) came to similar conclusions:
cortical microtubule alignment can vary between neighbouring
cells and microtubules tend to become transverse during phases of
accelerating growth and reorient to oblique as the growth rate
declines. One difference between Granger and Cyr’s (Granger and
Cyr, 2001) investigation of the living Arabidopsis root and ours on
the living Arabidopsis hypocotyl is that although they found no
evidence for a cyclic, oscillatory reorientation of microtubules of
the kind deduced to be occurring in azuki bean epicotyls (Mayumi
and Shibaoka, 1996), our observations show that microtubules can
rotate or jump. This might be due to inherent differences between
roots and light-grown shoots, such as their different rates of growth.
Our observations indicate that both microtubule tracks and
CESA trajectories can have different alignments in different cells
of growing shoot tissue and, furthermore, that alignment can also
differ between outer and inner tangential surfaces of the same cell.
The classical view of the cortical array as a continuous hoop-like
structure is evidently overly simple, because the present
observations demonstrate that the cortical array behaves as if it is
composed of domains or groups of mobile microtubules that can
move independently of one another (Chan et al., 2007). The idea
that microtubules and cellulose microfibrils form transverse hoops
originated with freely-growing filamentous cells, such as Nitella
internodal cells (Green, 1960; Green, 1962; Green, 1963). However,
as the above observations indicate, shoot epidermal cells do not
necessarily show this strict hoop reinforcement. This mismatch
between inner and outer surfaces is seen for CESA tracks, as well
as for microtubules. Despite their relatively slow rate of growth,
illuminated hypocotyls nonetheless produce a well-defined axis.
This study shows that such anisotropic growth can occur without
the constantly coordinated alignments of microtubule tracks or
CESA trajectories on the outer epidermal surfaces that comprise
the ‘organ wall’.
An interesting aspect of this study is that microtubules and
CESA tracks on the inner epidermal wall tend to show more highly
coordinated transverse alignments than seen with those on the
outer wall. This is relevant to discussions about the role of inner
wall versus the function of the outer epidermal ‘organ’ wall.
Because the epidermis shrinks when the stem is cut whereas inner
tissues swell, it has been concluded that the epidermis constrains
the driving force provided by the expansion of inner tissues (Baskin,
2005; Hejnowicz et al., 2000; Kutschera and Niklas, 2007;
Schopfer, 2006). According to these ideas, the thicker outer
epidermal cell wall resists stress isotropically, whereas the inner
tissues channel expansion anisotropically and show better ‘hoop
reinforcement’ (Schopfer, 2006). This could explain why the
microtubules on the inner epidermal wall display transverse
microtubules during phases of cell elongation, consistent with
organ-level hoop reinforcement, whereas transverse alignment is
not necessarily strictly observed on the more isotropic outer face,
which forms a thicker, strong mechanical barrier without dictating
the direction of organ growth.
Materials and Methods
Plant material
Arabidopsis thaliana plants expressing 35S::GFP:TUA6 and CESA3::CESA3:GFP
have been described previously (Chan et al., 2010). Seeds were sterilised in 5% (v/v)
sodium hypochlorite before transfer onto Petri dishes containing 0.43% (w/v)
Murashige and Skoog medium (Formedium, Hunstanton, UK) containing 1% (w/v)
sucrose (Sigma) and 0.5% (w/v) Phytagel (Sigma). Seeds were placed at 4°C for 2
days and then incubated at 25°C under continuous light. 2- to 4-day-old seedlings
were transferred to microscopy chambers (Chan et al., 2007) and then imaged under
the confocal microscope.
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Confocal imaging and image analysis
The middle regions of hypocotyls were imaged using either a VisiTech (UK) spinning
disc confocal microscope or a Bio-Rad 1024 confocal laser-scanning microscope
using a 40⫻/1.3 NA oil objective lens. GFP was excited using the 488 nm line of
an argon ion laser and emitted light filtered through a 500–550 nm band-pass filter.
For the spinning disk microscopy, fluorescence was detected using a Hamamatsu
Orca ER cooled CCD camera with 1⫻ binning, set at 0.7–1.0 second (to visualise
GFP–tubulin) and 2–3 second (GFP–CESA3) exposure time. Time-lapse images
were acquired with a time delay of 15–30 minutes. Cell measurements and
kymographs were constructed using the re-slice tool of ImageJ
(http://reb.info.nih.gov/ij/). Long-term movies of growing hypocotyls were aligned
using Stackreg (http://bigwww.epfl.ch/thevenaz/stackreg/) plug-ins of ImageJ
(http://reb.info.nih.gov/ij/). Movies were edited using the Brightness/Contrast tool of
imageJ and the Time stamper plug-in (http://rsb.info.nih.gov/ij/plugins/stamper.html).
Journal of Cell Science
Field emission scanning electron microscopy
Hypocotyls were fixed in 4% (v/v) formaldehyde (Sigma) prepared in PCM buffer
(25 mM PIPES, 0.5 mM CaCl2, 0.5 mM MgSO4, pH 7.0; all from Sigma) for 30
minutes and washed three times in PCM buffer. The left-hand cotyledon was removed
for orientation and the root excised. Hypocotyls were cryo-protected in 25% and
50% (v/v) DMSO (in PCM buffer) for 10 minutes each, then placed on a nail head
and immediately frozen in liquid nitrogen. The outer epidermal wall was sliced off
with a glass knife mounted on a Leica Ultracut EM FC6 cryo-ultramicrotome at
–120°C and a knife angle of 6°. The specimen was thawed in 50% DMSO and
transferred to PCM buffer.
The cytoplasm was extracted under mild agitation with sodium hypochlorite
solution (Sigma; diluted 1:10) for 10 minutes followed by three washing steps in
PCM buffer for 10 minutes each. After dehydration in an ethanol series (30%, 50%,
70%, 95% and 100% three times), 15 minutes for each step, hypocotyls were
critical-point dried using CO2, transferred to scanning electron microscopy pin stubs,
and sputtered with platinum (Agar, Essex, UK) for 30 seconds. Samples were
investigated in a Supra 55 VP PEG SEM (Zeiss, Cambridge, UK) using an in-lens
detector and a secondary electron detector at an acceleration voltage of 5 kV and a
working distance of 4 mm. The orientation of cellulose fibrils was measured using
the line tool of ImageJ.
This work was supported by a grant-in-aid by the BBSRC to the
John Innes Centre. We thank the Gatsby Foundation for financing
M.E. E.F.C. was supported by the National Agency for Research
Project ‘IMACEL’ ANR-06-BLAN-0262. We also thank the European
Union Framework Program 6 (FP6) ‘CASPIC’ NSET-CT-2004-028974,
the National Agency for Research Project ‘Wall Integrity’ ANR-08BLAN-0292, and the FP6 program 037704 ‘AGRONOMICS’ for their
support.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/124/7/1088/DC1
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