1088 Research Article

IntroductionWhen they were first seen in the electron microscope, corticalmicrotubules were described as transverse ‘hoops’, matching thealignment of overlying cellulose microfibrils in the cell wall(Ledbetter and Porter, 1963). This agreed with the biophysicalconcept of hoop-reinforcement (Green, 1962), in which transverselyaligned, load-bearing cellulose microfibrils maintained the girth ofthe expanding cell, directing growth into elongation orthogonal tothe net alignment. This picture seems to apply to the root wheremicrotubules are transverse in rapidly elongating cells, reorientingto oblique or longitudinal as growth declines (Baskin, 2001;Granger and Cyr, 2001; Sugimoto et al., 2000). However, reportssuggest that the situation is not so straightforward in shoots, wherecortical microtubules beneath the outermost epidermal wall can beoblique or longitudinal or mixed, as well as transverse, apparentlyduring 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 representingstable configurations, microtubule arrays in shoots might undergoa regular programme of reorientation (e.g. Mayumi and Shibaoka,1996). Microtubules in pea shoot epidermal cells, which had beenmicroinjected with fluorescent tubulin, have been seen reorientingfrom transverse to longitudinal (Yuan et al., 1995) and in thereverse 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 wellas transverse microtubules in elongating tissue and suggested thisvariability to be an expression of a cyclic reorientationphysiologically regulated by a range of hormonal factors (Mayumiand Shibaoka, 1996; Takesue and Shibaoka, 1998). In hisimmunofluorescence study of rapidly elongating epidermal cells inthe sunflower hypocotyl, Hejnowicz (Hejnowicz, 2005) also sawmicrotubule arrays in a variety of alignments. He hypothesised thatthe variability of alignment reflected stages sampled from arotational clock-like cycle, rather than the oscillatory cycle proposedby the Shibaoka laboratory. Chan and colleagues (Chan et al.,2007) then made long-term movies of Arabidopsis hypocotylepidermal cells, confirming that microtubules beneath the outerepidermal wall do indeed undergo both discontinuous shifts(‘jumps’) in alignment and continuous rotary movements in aclockwise or anticlockwise direction (which are collectively termed‘persistent reorientation’). However, neither rotations like this, norShibaoka-like oscillatory cycles have been seen in livingArabidopsis root epidermal cells imaged with GFP–MBD (Grangerand Cyr, 2001), underlining the differences between roots andshoots.

Similarly to microtubules, cellulose microfibrils are classicallyconsidered to be transverse in rapidly elongating tissue, providinghoop-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

SummaryMicrotubules are classically described as being transverse, which is perpendicular to the direction of cell elongation. However, fixationstudies have indicated that microtubules can be variably aligned across the epidermis of elongating shoots. In addition, microtubulesare 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 therate of elongation within individual growing cells. We also investigated whether microtubule alignment at the inner or the outerepidermal wall better reflected the growth rate. Movies confirmed that transverse microtubules form on the inner wall throughoutelongation, but orientation of microtubules is variable at the outer wall, where they tend to become transverse only during episodes ofaccelerated growth. Because this appears to contradict the concept that circumferential arrays of transverse microtubules or microfibrilsare essential for cell elongation, we checked the organisation of cellulose synthase tracks using GFP–CESA3 and found a similarmismatch between trajectories on inner and outer epidermal surfaces. We conclude that microtubule alignment on the inner wallappears 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

Accepted 20 February 2011Journal of Cell Science 124, 1088-1094 © 2011. Published by The Company of Biologists Ltddoi:10.1242/jcs.086702

Microtubules and CESA tracks at the inner epidermalwall align independently of those on the outer wall oflight-grown Arabidopsis hypocotyls Jordi Chan1,*, Magdalena Eder1,‡, Elizabeth Faris Crowell2,§, Janet Hampson1, Grant Calder1 and Clive Lloyd1

1Department of Cell and Developmental Biology, John Innes Centre, Colney, Norwich NR4 7UH, UK2Institut Jean-Pierre Bourgin, UMR1318 INRA-AgroParisTech, INRA Centre de Versailles-Grignon, Route de St-Cyr (RD10), 78026 VersaillesCedex France*Author for correspondence (jordi.chan@bbsrc.ac.uk)‡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

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reported to contain longitudinal microfibrils, even in rapidlyelongating areas. Non-hoop-like behaviour is also suggested byreports that microtubules on the outer epidermal surface are notnecessarily co-aligned with microtubules on the inner epidermalsurface (Busby and Gunning, 1983; Flanders et al., 1989).

This major difference in apparent behaviour of corticalmicrotubules in shoot and root tissues prompted us to look at therelationship between the alignment of microtubules and cellelongation in light-grown hypocotyls, paying attention to bothouter and inner tangential walls of epidermal cells. Arabidopsisseedlings expressing GFP–TUA6 were grown in special growthchambers (Chan et al., 2007) and live changes in microtubulealignment were monitored over several hours. In this way, insteadof basing conclusions on populations of cells with potentiallyunsynchronised microtubule orientations, it was possible todynamically correlate the behaviour of microtubules with the

1089Microtubule alignment in Arabidopsis epidermis

variable growth rate for up to 24 hours at a time, within individualcells. This showed that microtubules on the outer epidermal wallbehaved differently to microtubules on the inner wall of the samecell; a finding that we confirmed with a cellulose synthase marker.This allows us to discuss the role of microtubules in growthanisotropy.

ResultsRelationship between growth and microtubule alignmentat the outer epidermal faceIn a previous study (Chan et al., 2007), persistent microtubulereorientation was observed in Arabidopsis seedlings expressingGFP–EB1a. Here, we used the same experimental procedure exceptthat microtubules in 2- to 5-day-old seedlings were labelled withGFP–TUA6 (Ueda et al., 1999). This probe maintains brightfluorescence, enabling confocal z-stacks of epidermal cells in the

Fig. 1. Changes in microtubule alignmentduring elongation of epidermal cells ofArabidopsis hypocotyls expressing TUA6–GFP.(A)Low-magnification image of cells from slow-growing hypocotyl displaying non-coordinatedorientations of microtubules. (B)Low-magnification image illustrating coordinatedtransverse arrays that develop across neighbouringcells during phases of rapid cell elongation.(C)Montage of movie frames of the outer surfaceof an epidermal cell displaying a fixed transversealignment of microtubules during rapidelongation. (D)Microtubules displaying obliqueor longitudinal arrays when growth has ceased.(E)Kymograph along the long axis of anelongating cell showing transitions between slowand rapid elongation (single arrowhead) and rapidto no elongation (double arrowhead). Rate ofelongation is indicated inm per hour.(F)Montage of microtubule alignments thatoccurred during the changes in elongation rateshown in E. Note that microtubules tend to nettransverse orientation when elongation rateincreases and then change to oblique whenelongation ceases. Orientation of microtubules isindicated by white lines. (G)Low-magnificationimages of neighbouring cells during slowelongation (t0), variable alignment; rapidelongation (t210), transverse alignment; and noelongation (t645), longitudinal or obliquealignment, respectively. Scale bars: 12m (A,B),18m (E, y axis), 119 minutes (E, x axis), 20m(G). Strip width: 4m, taken every 20 minutes;numerals show hours (C,D).

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middle portions of hypocotyls to be captured every 15–30 minutesfor up to 24 hours.

Three different general patterns of microtubules were seen at theouter epidermal surface of the hypocotyl, which appeared to changein a growth-related fashion. Most commonly, microtubule alignmentdiffered between neighbouring cells (Fig. 1A). This particularpattern was associated with relatively slow growth rates (averagecellular elongation rate of 1.27 m/hour; s.d.1.4, s.e.0.14 or anaverage relative elongation rate of 0.02/hour; s.d.0.02, s.e.0.002,n99 cells; 12 hypocotyls). Movies confirmed that microtubulesundergo persistent reorientation in such tissues (Chan et al., 2007).Importantly, although microtubules could be transiently transversein these persistently reorienting arrays, in such slower-growingcells the phase of reorientation was not synchronised across theepidermis (Hejnowicz, 2005; Chan et al., 2007).

A different pattern of microtubule alignment was observed inrelatively fast-growing hypocotyls (with an average cellularelongation rate of 3.19 m/hour; s.d.3.78; s.e.0.54 or an averagerelative elongation rate of 0.06/hour; s.d.0.07, s.e.0.01, n49cells). In these tissues, transverse alignments of microtubules wereobserved that were coordinated across fields of neighbouring cells(Fig. 1B). Transverse alignment could be maintained over severalhours, illustrating that it is not a fixed stage extracted from therotary process (Fig. 1C).

A third pattern was observed between days four and five, whengrowth ceased. In such tissues, coordinated alignments were alsosustained between neighbouring cells, except in these cases,microtubules were arranged in oblique or longitudinal arrays (Fig.1D). Such oblique alignments – similarly to the transversealignment of faster-growing cells (Fig. 1B) – could be maintainedover several hours.

Analysis of movies showed that hypocotyls did not maintainconstant growth rates and transitions between relatively fast andslow elongation rates (and vice-versa) were captured. Such movieswere used to analyse the dynamics of the development of transversealignments with respect to growth (supplementary material Movie1). Fig. 1F provides an example of microtubule alignment as thegrowth rate changes. To obtain these data, the length of the cellwithin a movie frame was measured by projecting a one-pixel-wide line from one end wall to the other. After extracting a timeseries of such lines from the movie, the bottom ends were alignedhorizontally, so that the upper ends traced a curve that shows therate of cell elongation (Fig. 1E). A corresponding montage of thedifferent microtubule alignments that occurred over the same periodof elongation is shown in Fig. 1F (which is best seen by magnifyingthe image online). During more rapid growth (Fig. 1E),microtubules tended towards transverse alignment (supplementarymaterial Movie 2), which was coordinated across fields of cells(Fig. 1G; t210, compare with the more variable alignments att0). But as the growth rate reached a plateau (Fig. 1E), thecorresponding panels of microtubules (which are aligned in Fig.1F), switch to a more oblique alignment. This alignment was alsocoordinated across the tissue (Fig. 1G; t645).

Observation of those transitions where the rate of elongationaccelerated revealed two features of the evolution of transversemicrotubule alignment. First, transverse arrays are a transientphenomenon and tend to develop before phases of more rapidelongation (Fig. 2A,B). Out of 41 cells monitored long-term in fivehypocotyls filmed as they changed from slow to faster elongation,35 evolved transverse alignment an average of 184 minutes (3.07hours) (s.d.170 minutes; s.e.27) before a measurable burst of

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elongation, whereas the arrays of six cells became transverse at thetransition to more rapid growth (Fig. 2C,D). During more rapidelongation, microtubules spent on average 370 minutes (6.2 hours)in transverse alignment (s.d.212 minutes; s.e.42, 25 cells, 4hypocotyls). Second, transverse alignment is not necessary tomaintain rapid elongation. For instance, supplementary materialMovie 1 shows transverse arrays reorienting towards thelongitudinal axis despite the fact that analysis showed no decreasefrom the maximum elongation rate. This demonstrates that althougharrays might have a tendency to become transverse before rapidelongation, it is not essential for this alignment to be maintainedthroughout the entire process of more rapid elongation.

In summary, persistent reorientation is mainly seen at the outerepidermal surface during hypocotyl growth (between days two andthree and days three and four) but can be interrupted by transientperiods of coordinated transverse alignment before rapid elongation.When growth diminishes between days four and five, there is aswitch towards an oblique or longitudinal alignment that is alsocoordinated between groups of cells. Alignment of microtubules

Fig. 2. Transverse microtubule alignments develop at the outer epidermalsurface of hypocotyl cells during rapid elongation. (A)Kymograph alongthe long axis of an elongating cell showing a transition from slow(0.6m/hour) to more rapid elongation (3.6m/hour). (B)Correspondingmontage of microtubule alignments that occurred during elongation shown inA. Note that microtubules become transverse before the transition to rapidelongation. (C)Kymograph along the long axis of an elongating cell showinga transition from slow (0.4m/hour) to rapid elongation (3.0m/hour).(D)The corresponding montage of microtubule alignments that occurredduring elongation shown in C. Note that microtubules become transverse at thetransition to rapid elongation. Scale bar: 28m (A, y axis), 16m (C, y axis),660 minutes (A,C, x axis). Strip width: 3m (B,D). Arrows denote distance (yaxis) and time (x axis).

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beneath the outer epidermal wall, coupled with the extent to whichthis is coordinated between neighbouring cells, therefore serves asa general indicator of the elongation status of the tissue.

Relationship between growth and microtubule alignmentat the inner epidermal faceNext, we investigated whether microtubules on the innertangential wall (in contact with the cortical cells) of the epidermisdisplay similar behaviour to those on the outer tangential surface(in contact with the external environment). First we collected z-stacks of 2-, 3- and 4-day-old hypocotyls. By separating the outerand inner epidermal surfaces within z-stacks, it was possible tocompare microtubules on inner as well as outer tangential surfacesof the same cell. Arrays were categorised on a ‘one-cell–one-microtubule alignment’ basis (transverse0±22.5°, oblique45±22.5°, longitudinal90±22.5°), but where an overall alignmentof domains could not be clearly determined they were classifiedas ‘mixed’.

Quantification showed that microtubule alignments on the innerepidermal wall were distributed differently to those on the outersurface (Fig. 3A). In 2-day-old hypocotyls, which had just emergedfrom the seed, microtubules of various orientations (oblique,transverse, mixed and longitudinal) were present on both the innerand outer epidermal walls, consistent with uncoordinatedreorientations described above. However, in elongating 3-day-oldseedlings, mixed arrays formed the predominant category on theouter wall. By contrast, on the inner epidermal surface, thepredominant alignment was transverse, with 62% (s.d.23%, 78cells in 7 hypocotyls) of cells organised in this way. This wasdouble the number of transverse arrays sampled at the outer surface(33%, s.d.18%, 78 cells in 7 hypocotyls). This was consistentwith findings from the dynamic studies, which showed thatmicrotubules on the inner epidermal wall of 3-day-old hypocotylcells had stopped reorienting and became aligned in coordinatedtransverse alignments. At this stage, none of the cells displayedlongitudinal arrays. At day four, approximately equal proportionsof oblique and transverse microtubules were seen on both innerand outer surfaces.

This analysis indicated that microtubules could be differentlyaligned on different surfaces of the same cell, with microtubuleshaving an increased tendency to maintain transverse alignment atthe inner wall of three-day-old hypocotyls. Movies made of theinner 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 remaintransverse (Fig. 3B, 0–270 minutes) throughout a period whenmicrotubules rotate through 180 degrees upon the outer surface ofthe same cell (Fig. 3C). Microtubules associated with the innerepidermal wall therefore serve as a better indicator of the axis ofcell or tissue elongation.

The sensitivity of inner wall alignment to growth was examinedbetween days three and four by making kymographs of hypocotylcells found to be undergoing fluctuations in the elongation rate.Only cells whose inner tangential wall was not too facetted bycontact with subjacent cells had a sufficiently flat surface for thisstudy. In contrast to the upper surface, detailed kymographicanalysis of six cells revealed that microtubule alignment on theinner surface was less sensitive to decelerations and accelerationsin the rate of growth. The kymograph in Fig. 4A shows the end-wall trace of an elongating cell in which growth plateaus for awhile before undergoing a sudden acceleration.

1091Microtubule alignment in Arabidopsis epidermis

Fig. 4A shows an end-wall trace where growth plateaus thenaccelerates. In Fig. 4B, each strip shows microtubule alignment atthe outer epidermal wall, corresponding to the end-wall tracedirectly above it (in Fig. 4A). This shows that the alignment ofmicrotubules at the outer epidermal surface varies over time, butthat after a period in which the growth rate plateaus, microtubulestend towards transverse alignment as the growth rate acceleratesonce more. By contrast, the microtubules upon the inner surface ofthe same cell (Fig. 4C) were notably less variable, remainingmainly transverse throughout. Microtubules upon the inner surfacecould remain net transverse for prolonged periods of time. Such anarray was tracked for 20 hours (Fig. 4C; supplementary materialMovie 4), in contrast to the shorter episodes of transverseness onthe upper surface of the same cell. This suggests that transversealignment of microtubules on the inner epidermal wall is relatively

Fig. 3. Microtubule orientation can be uncoupled on different faces of thesame cell. (A)Quantification of microtubule orientation along the outer versusinner surfaces of hypocotyl cells at different days of development. Note thatmicrotubule orientation is uncoupled on day two, when inner walls maintain ahigher percentage of transverse alignments; this is more pronounced on daythree. Values are mean ± s.e. (B,C)Montage of movie frames showing theinner (B) and outer (C) tangential surfaces of a pair of cells. Note thatmicrotubules rotate over time at the outer surface (C) whilst maintainingtransverse alignment on the inner surface (B). Time is shown in minutes. Scalebar: 15m (B,C).

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insensitive to fluctuations in elongation rate. By contrast,microtubules on the outer face rotate in slow-growing cells andonly display an increased tendency towards transverseness duringphases of accelerating growth.

In addition to examining outer and inner tangential or periclinalepidermal surfaces, we looked at the connecting radial or anticlinalwalls. Similarly to the inner surface, these maintained transversemicrotubule alignment, whereas a more pronounced realignmentwas confined to the outer surface (see supplementary materialMovie 4).

Orientation of cellulose microfibrils at the inner epidermalsurfaceThe preceding analysis indicated that microtubules can adoptvarious alignments on the inner epidermal surface at day two, butat day three, they have a stronger tendency than those in the outerwall for transverse alignment. To view the most recently depositedlayer of cellulose microfibrils upon this inner epidermal surface,the outer epidermal wall was removed by sectioning, the cellcontents and plasma membrane were removed with detergent, andthe exposed surface of the inner epidermal wall examined by fieldemission scanning electron microscopy (FESEM). In 2-day-oldhypocotyls (77 cells, 5 hypocotyls), the inner wall showed variousalignments of microfibrils that differed from cell to cell, with‘oblique’ forming the largest category (Fig. 5A–E). In 3-day-oldhypocotyls (51 cells, 6 hypocotyls) there was a doubling in thepercentage of transversely aligned microfibrils, which now formedthe predominant category, followed by oblique, whereas virtuallyno longitudinal alignments were seen at this stage (Fig. 5A). Thispattern was reversed at day four (34 cells, 4 hypocotyls) whentransverse alignments were almost absent with oblique orlongitudinal forming the predominant category Fig. 5A.

Comparison of cellulose synthase tracks on inner andouter cell surfacesAfter observing that microtubules can display different alignmentson opposing tangential surfaces of the same cell, we investigated

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whether this also applies to cellulose synthase tracks on the outerand inner tangential epidermal walls. This was performed withseedlings expressing GFP–CESA3 under control of its ownpromoter in the cesa3je5 mutant background (Desprez et al., 2007).The GFP–CESA3 labelling pattern in light-grown hypocotyl cellswas consistent with that seen in dark-grown cells, i.e. cellulose-synthesising particles were observed upon the cortex, whereasfaster-moving Golgi bodies were seen deeper in the cytoplasm(Desprez et al., 2007).

In particular, the CESA particles moved along the plasmamembrane in linear tracks, as previously described (Crowell et al.,2009; Paredez et al., 2006). Although this was difficult to discernupon the inner epidermal wall, as a result of the dimmerfluorescence and basal accumulation of Golgi bodies (particularlyin the small cells of 2-day-old seedlings), it was possible to identifycells in which CESA tracks on the inner wall did not mirror thealignment of tracks on the outer wall of the same cell. This wasobserved more clearly in the longer cells of 3-day-old hypocotyls.In this case, GFP–CESA3 tracks were aligned transversely uponthe inner epidermal surface, whereas a variety of alignments wereobserved on the outer surface (Fig. 6A–D).

In summary, just as microtubules do not necessarily formtransverse ‘hoops’ around illuminated epidermal cells, the cellulosesynthases also form tracks that can be aligned in differentorientations on different surfaces of the same cell.

Fig. 4. Mismatch between inner and outer wall microtubule alignmentduring changes in growth rate. (A)Kymograph along the long axis of anelongating cell showing a change in the rate of elongation, from slow(0.2m/hour) to more rapid elongation (2.0m/hour). (B)Correspondingmontage of microtubule alignments that occurred along the outer surface ofthe cell during elongation. (C)Montage of microtubule alignments thatoccurred along the epidermal surface of the same cell. Note that microtubulesremained in a net transverse orientation on the inner wall regardless of changesin growth rate. Scale bar: 298 minutes (A, x axis), 10.5m (A, y axis).

Fig. 5. Analysis of cellulose microfibrils along the inner walls of hypocotylepidermal cells. (A)Quantification of cellulose microfibril alignment alongthe inner walls of hypocotyl epidermal cells at different days of development.Note that cellulose microfibrils along inner walls have a greater tendencytowards transverseness on day 3. Results are mean ± s.e. (B–E) FESEMimages of cellulose microfibril orientations at the inner wall of Arabidopsisepidermal cells: (B) longitudinal, (C) oblique, (D) transverse and (E) mixedorientation. Scale bars: 200 nm.

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DiscussionThis study of individual hypocotyl epidermal cells shows thatmicrotubules display complex three-dimensional reorganisationsin which alignment of microtubules on different faces of the cellcan vary in a growth-sensitive manner. In the early stages ofgrowth, 2-day-old hypocotyl cells had various alignments(transverse, oblique, longitudinal or mixed) on both inner andouter surfaces. But as growth proceeded, microtubules on the innerwalls of 3-day-old hypocotyls tended to display coordinatedtransverse alignments. These illuminated Arabidopsis seedlings donot grow as fast as etiolated plants (Gendreau et al., 1997; Le etal., 2005) and microtubules at their outer epidermal surface canundergo rotary movements during slow growth, but during phasesof accelerating growth, hypocotyls show a tendency to switch totransverse alignment when imaged for long periods. Although thisswitch in alignment tended to occur before accelerations in growthrate, it was not tightly coupled with the actual change in elongationrate, nor was it essential for this alignment to be maintainedthroughout the process of rapid elongation. Then, as growthdiminished, the pattern changed once more, consisting of arrays inwhich oblique and steeply oblique or longitudinal microtubuleswere present on both outer and inner surfaces. The same trend wasnoted for microtubule alignment on the outer surface of light-grown Arabidopsis seedlings expressing GFP–TUA6 (Le et al.,2005). This pattern is also very similar to that reported in a livecell study of Arabidopsis root epidermal cells expressing GFP–MBD (Granger and Cyr, 2001). There, transverse microtubuleswere mainly associated with accelerating growth, whereas a varietyof orientations was found as the growth rate declined. From theirinvestigation on fixed Arabidopsis root cells, Sugimoto and co-

1093Microtubule alignment in Arabidopsis epidermis

workers (Sugimoto et al., 2000) came to similar conclusions:cortical microtubule alignment can vary between neighbouringcells and microtubules tend to become transverse during phases ofaccelerating growth and reorient to oblique as the growth ratedeclines. One difference between Granger and Cyr’s (Granger andCyr, 2001) investigation of the living Arabidopsis root and ours onthe living Arabidopsis hypocotyl is that although they found noevidence for a cyclic, oscillatory reorientation of microtubules ofthe kind deduced to be occurring in azuki bean epicotyls (Mayumiand Shibaoka, 1996), our observations show that microtubules canrotate or jump. This might be due to inherent differences betweenroots and light-grown shoots, such as their different rates of growth.

Our observations indicate that both microtubule tracks andCESA trajectories can have different alignments in different cellsof growing shoot tissue and, furthermore, that alignment can alsodiffer between outer and inner tangential surfaces of the same cell.The classical view of the cortical array as a continuous hoop-likestructure is evidently overly simple, because the presentobservations demonstrate that the cortical array behaves as if it iscomposed of domains or groups of mobile microtubules that canmove independently of one another (Chan et al., 2007). The ideathat microtubules and cellulose microfibrils form transverse hoopsoriginated with freely-growing filamentous cells, such as Nitellainternodal cells (Green, 1960; Green, 1962; Green, 1963). However,as the above observations indicate, shoot epidermal cells do notnecessarily show this strict hoop reinforcement. This mismatchbetween inner and outer surfaces is seen for CESA tracks, as wellas 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 withoutthe constantly coordinated alignments of microtubule tracks orCESA trajectories on the outer epidermal surfaces that comprisethe ‘organ wall’.

An interesting aspect of this study is that microtubules andCESA tracks on the inner epidermal wall tend to show more highlycoordinated transverse alignments than seen with those on theouter wall. This is relevant to discussions about the role of innerwall versus the function of the outer epidermal ‘organ’ wall.Because the epidermis shrinks when the stem is cut whereas innertissues swell, it has been concluded that the epidermis constrainsthe 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 outerepidermal cell wall resists stress isotropically, whereas the innertissues channel expansion anisotropically and show better ‘hoopreinforcement’ (Schopfer, 2006). This could explain why themicrotubules on the inner epidermal wall display transversemicrotubules during phases of cell elongation, consistent withorgan-level hoop reinforcement, whereas transverse alignment isnot necessarily strictly observed on the more isotropic outer face,which forms a thicker, strong mechanical barrier without dictatingthe direction of organ growth.

Materials and MethodsPlant materialArabidopsis thaliana plants expressing 35S::GFP:TUA6 and CESA3::CESA3:GFPhave 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 2days and then incubated at 25°C under continuous light. 2- to 4-day-old seedlingswere transferred to microscopy chambers (Chan et al., 2007) and then imaged underthe confocal microscope.

Fig. 6. Comparison of cellulose synthase (CESA3) tracks along outer andinner surfaces of hypocotyl epidermal cells. Limited z-stack projections of3-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 cellulosesynthase tracks can be differently oriented on opposite faces of the same cell.Scale bar: 10m.

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Confocal imaging and image analysisThe middle regions of hypocotyls were imaged using either a VisiTech (UK) spinningdisc confocal microscope or a Bio-Rad 1024 confocal laser-scanning microscopeusing a 40�/1.3 NA oil objective lens. GFP was excited using the 488 nm line ofan 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 HamamatsuOrca ER cooled CCD camera with 1� binning, set at 0.7–1.0 second (to visualiseGFP–tubulin) and 2–3 second (GFP–CESA3) exposure time. Time-lapse imageswere acquired with a time delay of 15–30 minutes. Cell measurements andkymographs were constructed using the re-slice tool of ImageJ(http://reb.info.nih.gov/ij/). Long-term movies of growing hypocotyls were alignedusing 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 ofimageJ and the Time stamper plug-in (http://rsb.info.nih.gov/ij/plugins/stamper.html).

Field emission scanning electron microscopyHypocotyls 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 30minutes and washed three times in PCM buffer. The left-hand cotyledon was removedfor orientation and the root excised. Hypocotyls were cryo-protected in 25% and50% (v/v) DMSO (in PCM buffer) for 10 minutes each, then placed on a nail headand immediately frozen in liquid nitrogen. The outer epidermal wall was sliced offwith 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 andtransferred to PCM buffer.

The cytoplasm was extracted under mild agitation with sodium hypochloritesolution (Sigma; diluted 1:10) for 10 minutes followed by three washing steps inPCM 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 werecritical-point dried using CO2, transferred to scanning electron microscopy pin stubs,and sputtered with platinum (Agar, Essex, UK) for 30 seconds. Samples wereinvestigated in a Supra 55 VP PEG SEM (Zeiss, Cambridge, UK) using an in-lensdetector and a secondary electron detector at an acceleration voltage of 5 kV and aworking distance of 4 mm. The orientation of cellulose fibrils was measured usingthe line tool of ImageJ.

This work was supported by a grant-in-aid by the BBSRC to theJohn Innes Centre. We thank the Gatsby Foundation for financingM.E. E.F.C. was supported by the National Agency for ResearchProject ‘IMACEL’ ANR-06-BLAN-0262. We also thank the EuropeanUnion Framework Program 6 (FP6) ‘CASPIC’ NSET-CT-2004-028974,the National Agency for Research Project ‘Wall Integrity’ ANR-08-BLAN-0292, and the FP6 program 037704 ‘AGRONOMICS’ for theirsupport.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/124/7/1088/DC1

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