RESEARCH ARTICLE
Plant Cell Advance Publication. Published on May 26, 2017, doi:10.1105/tpc.16.00782
Cellulose synthesis and cell expansion are regulated by different mechanisms in growing
Arabidopsis hypocotyls
Alexander Ivakov1,2†*$, Anna Flis2†$, Federico Apelt2, Maximillian Fünfgeld2, Ulrike Scherer2‡, Mark Stitt2, Friedrich
Kragler2, Kris Vissenberg3,4, Staffan Persson1,2#, Dmitry Suslov3,5#
1
School of Biosciences, University of Melbourne, Parkville, VIC 3010, Melbourne, Australia
Max-Planck Institute of Molecular Plant Physiology, Am Muehlenberg 1, 14476 Potsdam, Germany
3
Biology Department, Integrated Molecular Plant Physiology Research, University of Antwerp, Groenenborgerlaan 171,
2020 Antwerpen, Belgium
4
UASC-TEI, Plant Biochemistry & Biotechnology Lab, Dept. Agriculture, School of Agriculture, Food & Nutrition,
Stavromenos, Heraklion, Crete, Greece
5
Saint Petersburg State University, Faculty of Biology, Department of Plant Physiology and Biochemistry,
Universitetskaya emb. 7/9, 199034 Saint Petersburg, Russia
†Present address: ARC Centre of Excellence in Translational Photosynthesis, Research School of Biology, Australian
National University, GPO Box 475, Canberra, ACT 2601, Australia
‡Present address: Zoological Institute, University of Hamburg, Martin-Luther-King-Platz 3, 20146, Hamburg, Germany
$
Authors contributed equally to the presented work
#
Co-senior authors
*
Corresponding author: Alexander Ivakov ([email protected])
2
Short title: Regulation of cell wall synthesis and growth
One-sentence summary:
The production of cellulose in the plant cell wall is controlled by metabolic signals while the expansion of cells is regulated
by changes in cell wall properties controlled by the biological clock.
The author responsible for distribution of materials integral to the findings presented in this article in accordance with the
policy described in the Instructions for Authors (www.plantcell.org) is: Alexander Ivakov ([email protected])
Abstract
Plant growth is sustained by two complementary processes – biomass biosynthesis and cell expansion. The cell wall
is crucial to both as it forms the majority of biomass, while its extensibility limits cell expansion. Cellulose is a major
component of the cell wall and cellulose synthesis is pivotal to plant cell growth and its regulation is poorly understood.
Using periodic diurnal variation in Arabidopsis thaliana hypocotyl growth, we found that cellulose synthesis and cell
expansion can be uncoupled and are regulated by different mechanisms. We grew Arabidopsis plants in very short
photoperiods and used a combination of extended nights, continuous light, sucrose feeding experiments and photosynthesis
inhibition to tease apart the influences of light, metabolic and circadian clock signaling on rates of cellulose biosynthesis
and cell wall biomechanics. We demonstrate that cell expansion is regulated by protein-mediated changes in cell wall
extensibility driven by the circadian clock. By contrast, the biosynthesis of cellulose is controlled through intracellular
trafficking of cellulose-synthase enzyme complexes regulated exclusively by metabolic signaling related to the carbon status
of the plant, and independently of the circadian clock or light signaling.
©2017 American Society of Plant Biologists. All Rights Reserved
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Introduction
Plant growth is sustained by two complementary processes – biomass biosynthesis, resulting in dry matter
accumulation, and cell expansion accompanied by water uptake leading to irreversible increases in size. Both
processes are tightly linked to the cell wall, a structure that encases all plant cells. A major proportion of plant
biomass is in the cell wall and large fluxes of photosynthetic products are directed to cell walls in growing cells
(Wang et al., 2013; Boex-Fontvielle et. al., 2014; Haigler et al., 2001). On the other hand, plant cell expansion is
limited by cell wall extensibility (Cosgrove, 1986, 2016a). The cell wall is thus a key player in cell expansion,
morphology and size as well as plant biomass accumulation.
Cell expansion is controlled by changes in cell wall properties, which have been extensively studied under
different conditions (Cosgrove and Li, 1993; Schopfer et al., 2002; Suslov et al., 2009; Peaucelle et al., 2015).
More specifically, cell expansion is linked to cell wall extensibility, which can be defined as the ability of the cell
wall to increase in surface area irreversibly during growth (Cosgrove, 2016a). Cell wall extensibility is regulated
through selective breakage and formation of links between wall polymers driven by loosening and tightening
proteins secreted by the cell (Cosgrove, 2016b). By contrast, how different external and internal cues control cell
wall synthesis is only starting to be unravelled (Bischoff et al., 2011; Boex-Fontvieille et al., 2014). To protect
expanding cell walls from excessive thinning and failure during growth, their synthesis and expansion need to be
coordinated (Voxeur and Höfte, 2016). While several reports have observed that these processes can occur
independently of each other (Edelmann et al., 1989; Edelmann and Fry, 1992; Refrégier et al., 2004; Derbyshire
et al., 2007), the principles behind such uncoupling are not clear.
How cell wall synthesis is controlled and whether it is coordinated with cell expansion can be investigated
in plants growing in conditions of alternating light and darkness during 24 hour cycles, whereby light, temperature
and metabolic resource supply fluctuate. In these conditions dicot plants exhibit diurnal variation in expansion
growth. Studies on diurnal growth dynamics (Wiese et al., 2007; Yazdanbakhsh et al., 2011; Apelt et al., 2015)
have shown that the periodic patterns are generated through the interplay between light signaling and an internal
timing mechanism known as the circadian clock (Nozue et al., 2007; Nusinow et al., 2011), and that the patterns
persist in constant-light, constant-temperature conditions for extended periods. However, the mechanistic basis
for these growth variations at the level of cell wall synthesis and cell expansion remains unclear and it is unknown
to what extent the synthesis of cell wall varies with changing growth rates and what regulatory inputs impact on
it. In addition, how the cell wall extensibility modulates these rapid changes in growth and to what extent cell
wall biosynthesis and cell expansion are coordinated as growth rates vary remain largely unknown.
Any diurnal pattern observed in a plant may be the confluence of separate responses to light, metabolic
and endogenous cues. Untangling these effects requires an experimental system where all three can be
manipulated separately but all remain functional. Cell wall biosynthesis relies on carbon newly fixed through
photosynthesis during the day and remobilized from starch reserves at night (Smith and Stitt, 2007; Stitt and
Zeeman, 2012; Sulpice et al., 2014). When days are very short, the limited starch reserves accumulated during
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the day are only sufficient to maintain respiration at night (Gibon et al., 2009; Sulpice et al., 2014), leaving little
for biosynthesis, while still allowing a circadian rhythm to be maintained in a natural diurnal cycle. These
conditions offer an excellent experimental system to explore the regulatory inputs that control growth rates at the
level of cell wall synthesis and re-modelling.
Cellulose is a major component of plant cell walls. In contrast to other cell wall components, cellulose is
produced by plasma-membrane cellulose synthase (CesA) enzyme complexes (CSCs). Fluorescently-labelled
CSCs can be observed microscopically as moving foci in the plasma membrane of plant cells (Paredez et al.,
2006). Their movement is thought to be driven by the polymerization and extrusion of cellulose chains into the
cell wall, which push the enzyme complex in the plane of the plasma membrane, and this has been used as a proxy
for the rate of cellulose synthesis. CSCs are also observed in intracellular membrane compartments thought to be
involved in their trafficking and the CSCs are thought to be inactive until they reach the plasma membrane
(Paredez et al., 2006; Gutierrez et al., 2009; Crowell et al., 2009). Here, we investigated the regulation of cellulose
synthesis and cell wall extensibility in hypocotyls of Arabidopsis thaliana (L.) Heyn. plants grown in short diurnal
cycles.
Results
CesA abundance at the plasma membrane exhibits diurnal fluctuations in low-carbon conditions
To create a strong diurnal alternation between periods of growth and no growth, and to rely purely on
photosynthesis-derived carbon for cell wall biosynthesis, we grew Arabidopsis seedlings in short photoperiods (4
h day/20 h night) on sucrose-free media for 7 days. It has been shown that seed storage lipids are essentially
depleted as early as three days after germination (Andre & Benning, 2007; Kelly et al., 2011), and the plants must
then rely on photosynthesis for their carbon supply, although it is not known whether the reserves last longer or
shorter under certain conditions. We observed a fluorescently-tagged subunit of the CSC, YFP-CesA6, in
hypocotyl cells using spinning disc confocal microscopy as an indicator of cellulose synthesis. At the end of the
night, there was a low abundance of CesA particles in the plasma membrane, with the overwhelming majority
associated with intracellular membrane compartments (Supplemental Movie 1; Supplemental Figure 1), referred
to as small CesA compartments (smaCCs; Gutierrez et al., 2009), or microtubule-associated CesA compartments
(MASCs; Crowell et al., 2009), which are thought to contain inactive CSCs and to be involved in either exo- or
endocytosis of the CSCs and exhibit erratic, irregular movement. CesA particles began to re-populate the plasma
membrane during the light period, while the number and intensity of SmaCCs decreased. The density of plasma
membrane-based particles varied significantly with time (ANOVA p<0.001) and steadily increased to a peak at
the end of the day (ED; Figure 1A, Supplemental Table 1) and declined thereafter, persisting for several hours
into the night. This diurnal pattern of CesA dynamics suggests that light signalling and/or metabolic status and/or
endogenous signals may regulate the secretion of CesA to the plasma membrane during the diurnal cycle.
Extended darkness prevents the mobilization of CesA to the plasma membrane
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To assess whether endogenous factors or signals regulated the observed CesA behaviour, we performed
an extended night experiment where, on one day, plants were kept in the dark for four hours instead of in the
light, and were imaged at end of the subjective day. Here, plasma membrane CesA particle abundance did not
increase significantly during the extended night (Figure 1A, Supplemental Movie 1, Supplemental Figure 1).
This suggests that the diurnal fluctuations in CesA abundance at the plasma membrane were not the result of
endogenous rhythms and indicated that light signalling (i.e., photoreceptors) or the metabolic/signalling effects
associated with photosynthesis in the light must be required to mobilise the CesAs to the plasma membrane.
Inhibition of photosynthesis in the light prevents CesA mobilization
To separate the roles of light signalling and photosynthesis, we pre-treated short day-grown seedlings with
the photosynthesis inhibitor 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU) overnight and then exposed them
to light in the absence or presence of exogenous sucrose to provide a carbon source independent of photosynthesis.
Treatment with DCMU reduced the quantum efficiency of photosystem II to very low levels both in the presence
and absence of sucrose (Supplemental Figure 2). In the absence of sucrose, CesA abundance at the plasma
membrane did not increase significantly during the day after inhibition of photosynthesis by DCMU (Figure 1B,
Supplemental Movie 2, Supplemental Figure 3. However, in the presence of DCMU and sucrose, CesA complexes
were mobilised to the plasma membrane despite the severe inhibition of photosynthesis. These results suggest
that metabolic signalling associated with the supply of carbon triggers the mobilization of CesA complexes to the
plasma membrane in the light.
A metabolic signal is necessary and sufficient to trigger CesA mobilization
To test whether a favorable metabolic status alone is sufficient to mobilize CesAs to the plasma membrane,
we performed the extended night experiment while transferring seedlings to media containing 2% sucrose. Plasma
membrane CesA particles were abundant after 4 h of light at the subjective dusk (Figure 1C, Supplemental Movie
3, Supplemental Figure 4). This increase was absent in control seedlings (2% mannitol; Supplemental Movie 3,
Supplementary Figure 5). This indicates that a favorable metabolic status is necessary and sufficient to trigger the
mobilization of CesA to the plasma membrane and that a light stimulus is not required.
Cellulose synthase dynamics are constant if carbon resources are constantly available
Finally, we observed plants grown in short-day conditions on media supplemented with sucrose. In these
conditions, CesA abundance at the plasma membrane was high and exhibited no significant variation throughout
the diurnal cycle (Figure 1D, ANOVA P=0.07, Supplemental Movie 4, Supplemental Figure 5, Supplemental
Table 2). The speed of particle movement also did not vary significantly (Figure 1E, Kruskal-Wallis rank ANOVA
P=0.114, Supplemental Table 3). Furthermore, extension of the night did not result in a decrease in CesA particle
abundance at the plasma membrane or their speed of movement. Taken together, the unchanging abundance and
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movement speed of CesA in these conditions implies that cellulose synthesis rates are constant when carbon
resources are constantly available.
Substrate levels do not underlie diurnal variation in cellulose synthase dynamics
The fluxes through many enzymes are limited by the supply of substrate and it may be assumed that severe
metabolic limitation may reduce the pool of substrate available to the CesA, limiting its activity or affecting its
trafficking. To investigate this, we measured the levels of uridine disphosphateglucose (UDP-glucose), the
substrate of CesA, throughout diurnal cycles in seedlings grown in 4-hour days with and without sucrose
supplementation and also in extended night conditions. Surprisingly, UDP-glucose levels did not vary
significantly without sucrose supplementation (Figure 1F) despite altered CesA localization. Extension of the
night by 4 h also did not significantly alter UDP-glucose levels compared to normal short-day conditions. The
presence of sucrose in the growth medium significantly increased UDP-glucose levels (ANOVA, P<0.001,
Supplemental Table 4), with 48–78% higher levels at the end of the day and end of the night in sucrose-grown
plants compared to no sucrose, respectively. However, sucrose-grown plants had similar levels of UDP-glucose
at the end of the day and end of the night (Supplemental Table 4). These results suggest that the CesA dynamics
are not due to substrate limitation as UDP-glucose is present at similar levels during the day, night and extended
night, and implies that a signalling process might be responsible for the observed changes in CesA trafficking.
Extension growth is independent of cellulose synthesis
To assess how the regulation of cellulose synthesis relates to rates of expansion growth, we monitored
hypocotyl elongation in the presence of sucrose under the same short-day conditions as above using an infra-red
camera. Hypocotyl growth exhibited a diurnal rhythm similar to that reported in previous studies (Nozue et al.,
2007; Dowson-Day and Millar, 1999), with a peak in growth at the end of the night and a slowdown during the
day (Figure 2A). Growth rates varied 3.0-fold, despite constant high abundance and speed of plasma membrane
CesA particles and, by implication, cellulose synthesis rates in the same conditions. This indicates that cellulose
synthesis and plant expansion growth are uncoupled on short timescales.
Protein-dependent cell wall modification underlies diurnal variation in extension growth
Next, we investigated whether changes in cell wall extensibility can explain the observed diurnal growth
differences. We harvested hypocotyls at the end of the night (EN; fast growth) and end of the day (ED, slow
growth). The frozen/thawed hypocotyls were extended in vitro using the constant load (creep) method in buffered
conditions at pH5, pH6 and pH5 with heat-inactivation at 90°C (Suslov et al., 2015). At pH5, creep rates were
two- to six-fold higher than at pH6, reflecting the higher activity of expansins under acidic conditions (Figure
3A). Fast-growing plants harvested at EN had 1.7- to 2.7-fold higher creep rates than slow-growing plants
harvested at ED at pH5, the difference being significant at higher loads (1600 and 2000 mg). Up to 2.0-fold
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difference was observed between EN and ED at pH6, but this was significant at lower loads. There was no
significant difference in creep rates in heat-inactivated hypocotyls, indicating that the observed differences at pH5
and pH6 are protein-mediated. Thus, diurnal variation in hypocotyl growth rate is accompanied by variations in
the activity of at least two groups of cell wall-loosening proteins: those active at pH5 (high loads) and at pH6
(low loads).
The circadian clock controls cell wall modification to control plant growth
Next, we sought to investigate to what extent the diurnal variations in creep rates are controlled by the
circadian clock. We did this by first entraining plants in our short-day conditions and then transferring them to
constant-light conditions and recording their growth. Periodic peaks in extension growth rate continued to occur
every 24 h (Figure 2B). The timing of the peak in growth rate was shifted forward by approximately 4 h and
maximum growth now occurred at the subjective ED, while minimum growth was observed at the subjective EN,
similar to what has been reported in previous studies (Nozue et al., 2007). Growth rates were 3.6-fold higher at
the subjective ED than at the start of the subjective day.
We harvested hypocotyls at subjective dawn and subjective dusk, close to the minimum and maximum
elongation rate in continuous light and subjected them to creep measurements. In contrast to diurnal cycles, creep
rates at pH6 did not vary significantly between time-points in continuous light (Figure 3B). However, a large,
significant difference in creep rates was observed at pH5 under 1600 and 2000 mg loads, where fast-elongating
hypocotyls harvested at subjective dusk had 2.2- to 2.7-fold higher creep rates than slow-growing hypocotyls
harvested at subjective dawn. Creep rates in heat-inactivated hypocotyls did not vary significantly between time
points, confirming that the changes observed at pH5 are protein-dependent.
To confirm the involvement of the circadian clock, we measured growth and estimated cell wall
extensibility in the elf3 loss-of-function mutant, deficient in a key component of the Evening Complex of the
circadian clock. The circadian clock in this mutant becomes arrhythmic in constant-light conditions (Nozue et al.,
2007; Hicks et al., 1996; Covington et al., 2001), allowing the effects of the clock to be tested. The Ws-2 wildtype control behaved similarly to the Col-0 and exhibited periodic growth rhythms, with 10.7-fold higher growth
rates at the peak compared to the trough (Figure 2C). There was no periodic variation in growth rates in the elf3
mutant after transfer to continuous light (Figure 2D), consistent with its impaired clock function (Nozue et al.,
2007).
Creep measurements were conducted at pH5, where Col-0 exhibited large changes in creep rate in
continuous light. Ws-2 also showed a significant increase in creep rate between subjective dawn and subjective
dusk (P=0.015, Figure 3C). elf3 exhibited very high creep rates compared to the Ws-2 (Figure 3C); however,
there was no significant difference in creep rates between subjective dawn and dusk in elf3. Thus, a functional
circadian clock is required to modulate cell wall extensibility during periodic growth fluctuations and controls
plant expansion growth by modulating the activity of cell wall-modifying proteins active at pH5.
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Discussion
The ability to switch cell wall synthesis on and off should be of vital importance to plant cells given that
cell wall synthesis is the largest metabolic flux in growing cells, while resource supplies to support it can be scarce
and intermittent. Reports using isotope labelling techniques show that the incorporation of glucose units into cell
wall polymers decreases during the night in wild-type Arabidopsis plants, and is abolished from about 4 h into
the night in the starchless pgm mutant, showing a close temporal relationship to the depletion of sucrose in the
mutant (Boex-Fontvielle et al., 2014; Ishihara et al., 2015). These labelling experiments are in close agreement
with our results on diurnal changes in cellulose synthesis estimated from CSC dynamics. Based on our data and
the report from Ishihara et al. (2015), it is tempting to speculate that the production and secretion of other cell
wall components, like pectin and hemicelluloses, may be regulated in a similar manner. This would perhaps not
be surprising as they are also abundant components of cell walls, their synthesis is a significant resourceconsuming flux in growing cells and their deposition depends on vesicle trafficking and exocytosis (McFarlane
et al., 2014; Kim and Brandizzi, 2016).
It has been shown previously that cellulose synthesis is controlled by light through the action of the
photoreceptor Phytochrome B (Bischoff et al., 2011), in contrast to our findings indicating that light does not play
a part in diurnal CesA dynamics. However, that study focuses on and highlights the distinction between etiolated
plants growing in complete and constant darkness and light-grown seedlings growing in a long photoperiod.
Furthermore, there were no differences in CesA behavior between light- and dark-grown seedlings in wild-type
plants, the differences only becoming apparent when a subunit of the CesA complex was mutated, resulting in
slower CesA speeds in the dark. Taken together, our findings and those of Bischoff et al. (2011) suggest that there
may be functional differences in the CesA populations between etiolated seedlings and those growing in the light.
However, once growing in the light, the CesA population exhibits diurnal dynamics that do not depend on light
directly, but respond to the metabolic effects of light on photosynthesis and carbon assimilation.
The lack of correlation between UDP-glucose levels and apparent cellulose synthesis rates is intriguing.
However, it has been proposed, although never proven, that UDP-glucose may undergo metabolic channeling to
the active site of CesA by sucrose synthase (SuSy) associated with the CesA rosette as sucrose is broken down
(Haigler et al., 2001). Therefore, the relevant UDP-glucose concentration, i.e., that at the active site, may not be
apparent in UDPG measurements conducted on whole plant extracts, as was done here. Another possible
explanation is that a metabolic signalling process, rather than substrate levels directly, is responsible for the
observed CesA dynamics. While the identity of the metabolic sensor or signalling pathway responsible for the
observed patterns of CesA trafficking is not known at this point, several candidate sugar signalling pathways have
been reported, including sucrose/trehalose-6-phosphate signalling, and several kinases such as Target of
Rapamycin (TOR), Sucrose-NonFermenting (SNF1) and hexokinases (Moore et al., 2003; Cookson et al., 2005;
Ruan et al., 2010; Dobrenel et al., 2011; Eveland and Jackson, 2012; Yadav et al., 2014). It is not known whether
7
any of these pathways are involved in the regulation of vesicle trafficking; however, they are good candidates to
explore. Mechanisms linking cellular metabolic status to the regulation of vesicular secretion through Golgilocalised lipid-binding SEC14-domain proteins have been proposed in animal cells (Bankaitis et al., 2010).
Arabidopsis contains a family of these proteins and they are known to regulate vesicle trafficking (Huang et. al.,
2016). Investigation of the involvement of these pathways in the regulation of CesA trafficking and vesicle
trafficking in general may be a promising lead for future research.
Superimposed on the accumulation of cell wall material is cell expansion, resulting in increases in cell
and plant size. Regulation of expansion growth rate is known to be one of the chief outputs of the plant circadian
clock (Nozue et al., 2007; Dowson-Day and Millar, 1999). This regulation appears to work through the effect of
the clock on the expression of growth-promoting PHYTOCHROME INTERACTING FACTOR (PIF)
transcription factors, which further modulate auxin signalling (Nozue et al., 2011; Hornitschek et al., 2012; Seaton
et al., 2015). However, the final step in this regulatory cascade is not clear. The exclusive effect of the clock on
the activity of cell wall loosening proteins active at pH5 and high loads strongly suggests that expansins are the
ultimate targets in the regulation of expansion growth by the clock (Cosgrove, 2016b). This is also in line with
published observations that several expansin genes have rhythmic, circadian expression patterns (Harmer et al.,
2000; Bläsing et al., 2005). In addition, some of these have been implicated as targets of PIF transcription factors
(Leivar et al., 2009; Hornitschek et al., 2012), as well as being directly targeted by the clock transcription factor
CIRCADIAN CLOCK ASSOCIATED1 (Nagel et al., 2015). Interestingly, we observed that changes in cell wall
extensibility are also partly mediated by proteins active at pH6 and under low loads in short photoperiods (Figure
3A). Their activity is not controlled by the clock (Figure 3B), but may be dependent on light signalling and/or
metabolic inputs. Xyloglucan endotransglucosylase/hydrolases (XTHs) may be responsible for this activity
because of their characteristic pH-optima (Maris et al., 2011) and their ability to induce wall extension under low
loads (Miedes et al., 2013).
The EN vs. ED differences in cell wall extensibility were abolished by heat inactivation (Fig. 3A, B),
showing that they resulted from changes in the activity of cell wall loosening proteins active at pH 5 and pH 6
that are present in the wall. However, the diurnal regulation of cell expansion and the contribution of these proteins
to diurnal growth rhythms (Fig. 2) could depend not only on their relative abundance or activity, but also on
possible variations in turgor and pH. High turgor stimulates expansion by exerting a greater force on the wall,
while low pH has been shown to stimulate expansion by increasing the activity of expansins (Cosgrove 2000).
Higher turgor will favor the contribution of expansins, consistent with their more pronounced effects on creep
rate under higher loads (Miedes et al. 2013; Suslov et al., 2015). Diurnal and circadian variations in turgor have
been demonstrated in specialized cells such as stomata and pulvini (Hennesey & Field 1992). Although it has
never been shown to occur in Arabidopsis hypocotyls, diurnal changes in turgor there could reflect rhythmic
diurnal changes in the accumulation/mobilization of osmotically-active assimilates (Haydon et al., 2011) and
8
variation in plant water status brought on by stomatal opening and closing, known to affect turgor (Ache et. al.,
2010; Bramley et. al., 2015).
Changes in cell wall acidification may also play a role in the regulation of cell expansion. Cell wall
acidification mediated by plasma membrane H+-ATPases (Falhof et al., 2016) stimulates expansins due to their
low-pH optima (Cosgrove, 2016b) and is also the driving force for solute uptake to generate cell turgor. H+ATPases are major consumers of ATP, and their activity may be increased by light, which elevates the ATP level
(Liang et al., 2016). On the other hand, light strongly up- or downregulates H+-ATPase activity via
phosphorylation (Kinoshita and Shimazaki, 1999; Hohm et al., 2014). To our knowledge, no diurnal or circadian
rhythms in cell wall pH have ever been demonstrated. However, there is clear evidence for circadian rhythms in
auxin content in Arabidopsis shoots (Jouve et al., 1999). Auxin is known to promote cell wall acidification by
activating H+-ATPases (Spartz et al., 2014) and it is therefore likely that cell wall pH variation also contributes
to diurnal growth rhythms. These data show that the contribution of proteins active at pH 5 and pH 6 could vary
depending on the phase of the diurnal growth cycle and external conditions. The role of expansins may be
prevalent in conditions optimal for growth, i.e., in highly turgid cells having favorable energetic status. By
contrast, the contribution of proteins active at pH 6 may increase in more stressful conditions.
While we use the Arabidopsis hypocotyl as a simple model of one-directional cellular and organ growth,
it is possible that this system may be unique in certain aspects and may not reflect the regulation of growth in
other organs. For example, an investigation of rhythmic patterns of leaf growth and leaf movements in
Arabidopsis uncovered that light was an essential factor promoting leaf expansion during the day (Dornbusch et
al., 2014); however, light suppresses hypocotyl extension (Nozue et. al., 2007). Hypocotyls and leaves have
somewhat opposite growth responses, in that darkness, low light or short photoperiods stimulate hypocotyl
extension while inhibiting the growth and development of leaves in what is known as the etiolation response
(Fankhauser & Chory 1997). This serves to direct the plant’s resources to rapid expansion growth to “grow out”
of zones of darkness or shade, e.g., when seeds germinate underground. When light is sensed by photoreceptors,
hypocotyl growth is rapidly suppressed and leaf growth is stimulated to produce photosynthetic leaf area and
begin to assimilate carbon. Thus the regulation of leaf expansion growth is rather different to that of the hypocotyl.
On the other hand, biosynthesis is inevitably under metabolic constraints in any cell or organ and must be
regulated in accordance with the supply of available resources (Smith & Stitt. 2007). This regulation may differ
slightly in green organs that fix their own carbon (e.g., leaves) compared to non-green carbon-importing cells in
hypocotyls and roots; however, the constraints remain the same and unregulated biosynthesis in times of scarce
supply can exhaust the supply of carbon and energy and result in death. Therefore, unlike organ-specific
requirements for expansion growth in response to environmental cues, it is unlikely that hypocotyls are unique
with respect to the regulation of cellular biosynthesis.
This work demonstrates that cellulose biosynthesis and cell expansion are regulated by different
mechanisms, with the ensuing result that they can be uncoupled for short time periods. Of course, there cannot be
9
complete uncoupling between the two processes, as wall expansion cannot proceed limitlessly without the
synthesis and addition of new wall material to maintain the integrity of the cell wall. While we show that plant
carbon status defines the amount of cellulose the cell produces, Stewart et al., 2011 showed that sucrose also
affects hypocotyl expansion growth in Arabidopsis. In that study, growth became very slow, while still
maintaining diurnal rhythms, in short days in the absence of external carbon similar to our conditions, while the
addition of sucrose strongly stimulated expansion growth. This suggests that on longer timescales, the rate of cell
expansion adjusts to the amount of cell wall material that is able to be produced. This is reminiscent of the cell
wall integrity sensing mechanisms that have been shown to inhibit expansion growth in situations where cellulose
synthesis is inhibited using genetic or chemical means and under conditions of stress, known to involve several
receptor-like-kinases (RLKs) and wall-associated-kinases (WAKs) (Hematy et al., 2007; Wolf et al., 2012). These
kinases are worthy of investigation as the possible missing piece of the puzzle in the regulation of growth in
response to resource supply.
In summary, we show that cell wall synthesis and cell expansion can be uncoupled in Arabidopsis
hypocotyls on short timescales. We conclude that this uncoupling is based on different regulatory mechanisms of
cell wall biosynthesis and cell expansion. The biosynthesis of cellulose, and possibly other cell wall polymers, is
regulated through a relatively simple one-way control mechanism involving metabolic signals associated with the
carbon status of the plant. It is independent of the circadian clock or light signaling. On the other hand, cell
expansion is controlled by the plant circadian clock mediating protein-dependent changes in cell wall
extensibility. At the moment, these observations are only valid for the hypocotyl, which is a highly specialized
organ, and for a rather extreme photoperiod. Therefore, a much closer coupling between wall synthesis and cell
expansion may exist in other cell types or other conditions. Our finding that the two basic processes underlying
plant growth are regulated by different mechanisms enables their independent manipulation. This could open up
avenues to modulate plant growth and biomass production.
Materials and Methods
Plant material
YFP-CesA6 expressed under the native CesA6 promoter in the prc1-1 background was provided by D.
Ehrhardt (Paredez et al., 2006). elf3-4 mutant in Ws-2 background was provided by Prof. Andrew Millar (Hicks
et al., 1996).
Plant growth
Seeds were sterilised with 70% ethanol with a drop of Triton X detergent for 30 min and washed four
times with sterile water. Seeds were spread on MS agar plates with 0% or 2% sucrose and stratified at 4°C for 3
days. Plates were transferred to controlled environment chambers (Percival Scientific) running on a 4 h light/20
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h dark 20°C/18°C photoperiod and an irradiance of 100µmol m-2 s-1 provided by fluorescent lamps during the
day, and grown vertically. Confocal imaging was done on the 7th day after stratification. For continuous light
treatments plants were grown in 4 h photoperiods for 4 days and transferred to continuous low light at 20°C for
2 days. Lamps were darkened with plastic shades during continuous light to give an irradiance of 12 µE m-2 s-1.
For creep measurements, plants were harvested and frozen in liquid nitrogen on the 7th day in diurnal cycles and
on the second day of continuous light.
Microscopy
For live cell imaging, seedlings expressing YFP:CesA6 were mounted on glass slides under an agarose
gel pad and imaged on a Nikon Ti-E inverted confocal microscope with a CSU-X1 Yokogawa spinning disc head,
using a CFI APO TIRF 3100 N.A. 1.49 oil immersion objective, an evolve charge-coupled device camera
(Photometrics Technology), and a 1.2x lens between the spinning disc and camera. YFP was excited at 491 nm
using a multichannel dichroic and an ET525/50M band pass emission filter (Chroma Technology). Images were
acquired using Metamorph online premier, version 7.5. Exposure times were 400 ms.
Growth imaging
To image plant growth in real time, a modified Nikon DSLR camera was used, where the infra-red cutoff filter in front of the CCD sensor was removed. A long-pass near-infra-red filter was fitted to the objective so
that only infra-red radiation entered the camera, allowing the plants to be imaged day and night. Plants were
continuously illuminated from behind with infra-red light at 900 nm shown not to stimulate plant photoreceptor
systems. Images were taken every 5 minutes.
Image analysis
Confocal space-time stacks of fluorescently-labelled CesA were analysed using custom-written scripts
running in Fiji and R. For CesA particle density measurements, one biological replicate was a confocal time-lapse
movie of 30-frames and lasting 5 minutes, with each movie being collected on a different plant. Each movie was
analysed using our particle tracking algorithm, detecting tens to hundreds of CesA particles in each movie. Particle
density was calculated by dividing the number of detected particles by the area imaged in the movie in µm2.
Multiple movies were collected for each time point or treatment and the means and s.e.m. are reported in the
figures.
CesA Particle Tracking
The first 20 frames of each stack were used for analysis. The positions of CesA particles in each frame
were detected by applying a Laplacian filter with a smoothing kernel width of 1.5px. The particles were then
detected by finding local maxima with a noise tolerance of 800.
11
Tracks were detected in R. The detected particles in all frames were encoded as three-dimensional x-y-t points.
Particles in the top frame (first time-point) of each movie were used as starting points. Proceeding iteratively, unit
vectors from each starting point to all other points in all frames were computed. Vectors with a t-direction
component of at least 0.9 were then retained. This removes the majority of vectors which point to CesA particles
in other parts of the image (and thus probably not the same track).
The remaining points are then expected to contain points corresponding to the track of interest (the same CesA
particle moving over time) and a number of erroneous points corresponding to other CesAs moving into the search
region, or noise points. Due to the very regular movement of plasma membrane CesA particles (constant speed,
constant direction), the points in a track should have a very consistent direction through the x-y-t space, looking
from the starting point, while the erroneous points would deviate from this direction as outliers. A median
direction vector was calculated from the retained points using the L1 multivariate median (Vardi and Zhang,
2000), a measure with low sensitivity to outliers. All the points were then compared to the median direction using
cosine similarity, a measure that expresses the cosine of the angle between two direction vectors. Points with a
cosine similarity of at least 0.8 to the median direction were retained. The L1 median was then recalculated on
the remaining points and points with cosine similarity of at least 0.95 were retained. This way, only points with
highly similar directions are selected. These points are then treated as a track. Tracks with fewer than five points
were discarded.
Each track was then centred by subtracting its mean and a line of best fit in the x-y plane was fitted by extracting
the first principal component, a line which minimises the perpendicular distances between all points and itself.
The percentage of explained variance (similar to r2 in least squares regression) was used as a measure of linearity
for the track and tracks with linearities below 90% were discarded.
The tracks that satisfy the above criteria were then treated as a single CesA particle moving over time. For these
tracks, each point was projected onto the line of best fit to infer the “true” position of the particle, treating
deviations from the line as error. The position of the particle along the line was then regressed against time and
the slope taken as the speed of movement along the track. The number of tracks was divided by the area of the
region of interest to obtain particle density (CesA/area).
It must be noted that at a given time a proportion of SmaCCs move in a similar manner and with similar
speeds to plasma membrane CesA complexes and cannot be distinguished from them with certainty in all
instances. Thus speed and density measurements are only accurate in situations where few SmaCCs are present
and the vast majority of all particles are plasma membrane CesA complexes, or vice versa. In situations where
12
SmaCCs are in high abundance, plasma membrane CesA numbers are over-estimated, while speed distributions
are biased towards the movement properties of SmaCCs.
Analysis of hypocotyl length was done in ImageJ with subsequent data analysis in R. The length of the
plant was determined in each frame. The rate of extension growth was obtained by fitting cubic splines to the
growth curves and evaluating their first derivative at each time point.
Biomechanics measurements
Cell wall extensibility was estimated by the creep method in which frozen/thawed hypocotyls were
extended under a constant load. On the 7th day after germination, each Arabidopsis seedling was transferred to a
2 ml Eppendorf test tube, which was closed and plunged into liquid nitrogen. The frozen seedlings were stored at
–20°C and extended in creep tests within 2 weeks after freezing. Freeze/thawing disrupts the cells while leaving
the cell wall intact, making the measurements independent of turgor variations, while the constant load resembles
the action of turgor on the cell wall and allows the force applied to the wall to be controlled. Cell wall heatinactivation was performed at 90°C for 3 min as described in Suslov et al. (2015). The optimization of the heatinactivation procedure is detailed in Supplemental Figure 2. For each cell wall creep (time-dependent
deformation) measurement, one thawed Arabidopsis seedling was secured in a custom-built constant-load
extensiometer (Suslov and Verbelen, 2006) such that a 2 mm long hypocotyl segment located 1.5 mm below the
cotyledons was placed between the clamps of the set-up. This segment was first preincubated in a buffer (20 mM
MES-KOH, pH 6.0 or 20 mM Na-acetate, pH 5.0) in the relaxed state for 2 min, after which a constant load (1200,
1600 or 2000 mg) was applied, and cell wall extension was recorded for 15 min in the same buffer as during the
preincubation. Conducting the measurements in buffered conditions eliminates the role of wall pH variations that
may occur in vivo. The pH of buffers used corresponds to typical values in the plant apoplastic space (Felle,
2001). The early (0–5min after loading) cell wall deformation is known to contain large viscoelastic components
having no direct relation to cell wall extensibility (Suslov and Verbelen, 2006). This deformation was taken into
account to calculate the length of extending cell wall samples at 5 min after loading (Miedes et al., 2013). Then
the average creep rate was calculated during the interval 5 to 15 min after loading using the formula (ln L15min ln L5min)/T ´ 100%, where L15min and L5min indicate the length of an extending hypocotyl segment at 15 min and 5
min after loading, respectively, and T is the time during which the average creep rate is calculated (10 min)
(Miedes et al., 2013).
Metabolite analysis
Plants were grown as described above and harvested at relevant time points by quenching in liquid
nitrogen. Approximately 200 seedlings were pooled per biological replicate sample. Material was homogenised
while frozen in a ball mill (Retsch) and aliquoted into Eppendorf tubes. Samples were extracted in a 3:7
13
chloroform/methanol solution. Uridine diphosphate glucose (UDP-glucose) was measured using reverse-phase
LC-MS/MS as in Arrivault et al. (2009).
Statistical analysis
All statistical analyses was performed in R. Time-courses of particle densities were tested for a significant
effect of time using one-way parametric analysis of variance (ANOVA), testing the null hypothesis that particle
densities do not vary over time. Time courses of CesA speed were tested for a significant effect of time using the
non-parametric Kruskal-Wallis rank test as the CesA speed distributions were not normally distributed, testing
the null hypothesis that CesA speed does not vary over time. Pairwise comparisons of particle densities and creep
rates were conducted using Student’s t-test. Null hypotheses were rejected below the 0.05 P value threshold.
Accession Numbers
CesA6: At5g64740
ELF3: At2g25930
Supplemental Data
Supplemental Figure 1: Inhibition of photosynthesis by DCMU.
Supplemental Figure 2: Heat-inactivation of cell wall proteins under different conditions.
Supplemental Figure 3. Cellulose synthase localisation follows a diurnal pattern in living Arabidopsis cells
growing in very short days without exogenous sucrose.
Supplemental Figure 4. Cellulose synthase localisation to the plasma membrane in the light requires active
photosynthesis or an alternative carbon source.
Supplemental Figure 5. A favourable metabolic status is necessary and sufficient to promote localization of
cellulose synthase to the plasma membrane.
Supplemental Figure 6. Arabidopsis seedlings synthesise cellulose constantly under a favorable metabolic
status.
Supplemental Figure 7. Speed of moving particles in seedlings grown on medium without exogenous sucrose
and imaged throughout a diurnal cycle or after a 4-h extension of the night
Supplemental Movie 1: Cellulose synthase localization follows a diurnal pattern in living Arabidopsis cells
growing in very short days without exogenous sucrose.
Supplemental Movie 2: Cellulose synthase localization to the plasma membrane in the light requires active
photosynthesis or an alternative carbon source.
Supplemental Movie 3: A favorable metabolic status is necessary and sufficient to promote localization of
cellulose synthase to the plasma membrane.
Supplemental Movie 4: Arabidopsis seedlings synthesize cellulose constantly under a favorable metabolic status.
14
Acknowledgments
AI was funded by the Max-Planck Gesellschaft and University of Melbourne. AF was funded by the Max
Planck Gesellschaft. SP was funded by the Max-Planck Gesellschaft and by a R@MAP professorship at the
University of Melbourne, and acknowledges funding from an ARC Discovery grant (DP150103495), an ARC
Future fellowship grant (FT160100218) and a Hermon-Slade grant (Persson HSF 15/4). DS acknowledges Saint
Petersburg State University for research grants 1.38.233.2014 and 1.42.1025.2016, the Research park of Saint
Petersburg State University Center for Molecular and Cell Technologies, and RFBR for the grant No 15-0404075, and KV acknowledges the University of Antwerp and the National Research Foundation (FWOVlaanderen) for grants G.0.602.11.N.10, 1.5.091.11.N.00 and G039815N. We acknowledge Ross Dennis for
help with imaging.
Author contributions.
AI, DS, AF conceived the project and designed experiments. AI, DS, AF, FA, MF, US, FK conducted experiments
and interpreted results. AI, DS, MF analyzed data. MS, FK, KV, SP provided discussions on experiment design
and data interpretation.
Competing interests.
The authors declare no competing financial interests.
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Figure 1. Plasma membrane localization of cellulose synthase is driven by metabolic
signaling. (A) Density of moving particles of fluorescently labelled YFP-CesA6 in, or closeto, the plasma membrane in A) plants grown in 4 h day/20 h night diurnal cycles on medium
containing 0% sucrose and imaged throughout the day and night (black squares) or after a 4-h
extension of the night (open square) on the 7th day after germination using spinning disc
confocal microscopy (ANOVA Time effect P<0.001, Supplemental Table 1). (B) Particle
densities at the end of the night (ZT=0, EN) and end of the day (ZT=4, ED) in plants treated
with 1µM DCMU for 20 h in the presence (+Suc) or absence (-Suc) of sucrose. (C) particle
densities at the end of the night (EN, ZT=0) and after a 4-h extension of the night (ExN, ZT=28)
while being incubated in MS medium containing 2% sucrose or 2% mannitol. (D) Particle
densities in seedlings grown on medium containing 2% sucrose and imaged throughout a
diurnal cycle (black squares) or after a 4-h extension of the night (open square) (ANOVA Time
effect P=0.07, Supplemental Table 2). (E) Speed of moving particles in seedlings grown on
medium containing 2% sucrose and imaged throughout a diurnal cycle or after a 4-h extension
of the night (ExN), blue symbols represent all individual detected CesA tracks (Kruskal-Wallis
time effect P=0.11, Supplemental Table 3). (F) UDP-glucose (UDPG) content of seedlings
grown in 4 h day/20 h night diurnal cycles on sterile medium containing 0% sucrose (black
squares) or 2% sucrose (open squares) and sampled at different times during a diurnal cycle or
after a 4-h extension of the night (dark bar)(Supplemental Table 4). N=2–11 biological
replicates per timepoint, and each replicate is a time-lapse recording from a different seedling,
containing tens to hundreds of moving particles (A,B,C,D,E) and four biological replicates
containing 50–100 seedlings (F). Bars are standard error of the mean (A,B,C,D,F). Time is
indicated as Zeitgeber time, in hours after the dark-light transition. Light periods are indicated
with yellow bands. Night periods are indicated with grey bands. Extended night periods are
indicated with dark grey bands. Letters indicate significant differences at α=0.05 (Student’s ttest).
Figure 2. Rates of hypocotyl extension in Arabidopsis plants growing vertically on sterile
media. (A) Col-0 wild type growing in 4 h day/20 h night diurnal cycles. (B) Col-0 wild type
grown in 4 h day/20 h night diurnal cycles for 4 days and transferred to continuous light
conditions at ZT0. (C) Ws-2 wild type grown in 4 h day/20 h night diurnal cycles for 4 days
and transferred to continuous light conditions at ZT0. (D) elf3 mutant in the Ws-2 background
grown in 4 h day/20h night diurnal cycles for 4 days and transferred to continuous light
conditions at ZT0. Images were taken every 5 minutes. Time is indicated as Zeitgeber time, in
hours after the dark-light transition. Light periods are indicated with yellow bands. Light
periods in continuous light (B,C,D) indicate subjective day periods. N=5–8 plants. Dotted lines
are standard error.
Figure 3. Cell wall creep rates exhibit diurnal patterns and are regulated by the circadian
clock. Creep rates measured in vitro under controlled pH conditions at different loads in (A)
Col-0 wild type plants growing in 4 h day/20 h night diurnal cycles and harvested at the end of
the night (EN) and end of the day (ED) 7 days after germination, (B) Col-0 wild-type plants
grown in 4 h day/20 h night diurnal cycles for 4 days, transferred to continuous light and
harvested on the second day after transfer at the end of the subjective night (EN) and the end
of the subjective day (ED), and (C) Ws-2 wild-type and elf3 mutant plants grown in 4 h day/20
h night diurnal cycles for 4 days, transferred to continuous light and harvested on the second
day after transfer at the end of the subjective night (EN) and the end of the subjective day. Ws2 and elf3 were only measured at pH5 and under a 1600 mg load. In each creep test, a 2 mm
long hypocotyl segment of one thawed Arabidopsis seedling was extended. N=6–10. Bars are
s.e.m. Letters (bold for pH6 and italicized for pH5) indicate significant differences (Student’s
t-test, P < 0.05) between EN and ED under the same load.
Cellulose synthesis and cell expansion are regulated by different mechanisms in growing
Arabidopsis hypocotyls
Alexander A. Ivakov, Anna Flis, Federico Apelt, Maximilian Fuenfgeld, Ulrike Scherer, Mark Stitt,
Friedrich Kragler, Kris Vissenberg, Staffan Persson and Dmitry Suslov
Plant Cell; originally published online May 26, 2017;
DOI 10.1105/tpc.16.00782
This information is current as of July 31, 2017
Supplemental Data
/content/suppl/2017/05/26/tpc.16.00782.DC1.html
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