Download Pausing of Golgi Bodies on Microtubules Regulates

The Plant Cell, Vol. 21: 1141–1154, April 2009, www.plantcell.org ã 2009 American Society of Plant Biologists
Pausing of Golgi Bodies on Microtubules Regulates Secretion
of Cellulose Synthase Complexes in Arabidopsis
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Elizabeth Faris Crowell,a Volker Bischoff,a Thierry Desprez,a Aurélia Rolland,a York-Dieter Stierhof,b Karin
Schumacher,c Martine Gonneau,a Herman Höfte,a and Samantha Vernhettesa,1
a Laboratoire
de Biologie Cellulaire, Institut National de la Recherche Agronomique, 78026 Versailles cedex, France
for Plant Molecular Biology, Microscopy, University of Tübingen, 72074 Tübingen, Germany
c University of Heidelberg, Heidelberg Institute for Plant Sciences, Department VI Developmental Biology, 69120 Heidelberg,
Germany
b Center
Plant growth and organ formation depend on the oriented deposition of load-bearing cellulose microfibrils in the cell wall.
Cellulose is synthesized by plasma membrane–bound complexes containing cellulose synthase proteins (CESAs). Here, we
establish a role for the cytoskeleton in intracellular trafficking of cellulose synthase complexes (CSCs) through the in vivo
study of the green fluorescent protein (GFP)-CESA3 fusion protein in Arabidopsis thaliana hypocotyls. GFP-CESA3 localizes
to the plasma membrane, Golgi apparatus, a compartment identified by the VHA-a1 marker, and, surprisingly, a novel
microtubule-associated cellulose synthase compartment (MASC) whose formation and movement depend on the dynamic
cortical microtubule array. Osmotic stress or treatment with the cellulose synthesis inhibitor CGA 325’615 induces
internalization of CSCs in MASCs, mimicking the intracellular distribution of CSCs in nongrowing cells. Our results indicate
that cellulose synthesis is coordinated with growth status and regulated in part through CSC internalization. We find that
CSC insertion in the plasma membrane is regulated by pauses of the Golgi apparatus along cortical microtubules. Our data
support a model in which cortical microtubules not only guide the trajectories of CSCs in the plasma membrane, but also
regulate the insertion and internalization of CSCs, thus allowing dynamic remodeling of CSC secretion during cell expansion
and differentiation.
INTRODUCTION
Anisotropic cell expansion drives plant growth and organ formation and depends primarily on the properties of the cell wall
(Baskin, 2005). The load-bearing component of the plant cell wall
consists of cellulose microfibrils, which play a key role in
generating differential resistance to turgor pressure to allow
anisotropic expansion (Green and Selker, 1991). Shortly after the
discovery that cellulose microfibrils are deposited in precisely
oriented layers during development (Probine and Preston, 1961,
1962), it was proposed that long cytoplasmic elements sensitive
to colchicine could direct the orientation of cellulose microfibrils
in the wall (Green, 1962). The discovery of cortical microtubules
soon afterward, and the observation of their parallel alignment
with cellulose microfibrils, gave rise to the microtubule-microfibril
alignment hypothesis (Ledbetter and Porter, 1963). Subsequently, numerous studies reported correlations in the orientation of cortical microtubules and cellulose microfibrils (reviewed
in Baskin, 2001). However, the alignment hypothesis became
increasingly controversial as evidence for absence of correlation
1 Address
correspondence to [email protected].
The author responsible for the 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: Herman Höfte
([email protected]).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.108.065334
was found in certain cell types (Emons, 1982; Himmelspach et al.,
2003; Sugimoto et al., 2003).
In vascular plants, cellulose is synthesized by a plasma membrane–localized cellulose synthase complex (CSC), recognizable
as a hexameric rosette structure with a diameter of 25 to 30 nm
(Mueller and Brown, 1980) and a cytoplasmic component ;45 to
50 nm in diameter (Bowling and Brown, 2008). The only known
component of the CSC is the cellulose synthase catalytic subunit
(CESA) (Kimura et al., 1999). Identification of the genes encoding
CESAs in higher plants (reviewed in Somerville, 2006) and
advances in live-cell imaging techniques permitted visualization
of fluorescently tagged CSCs and microtubules simultaneously
in living cells and provided powerful evidence that cortical
microtubules define the trajectories of CSCs (Paredez et al.,
2006). Experimental evidence presented by Paredez et al. lends
strong support to the microtubule-microfibril alignment hypothesis. Nevertheless, the mechanism by which microtubules guide
CSC trajectories remains unknown and represents an important
focus of current research.
The importance of the actin cytoskeleton in cell expansion
in non-tip-growing cells is only beginning to be revealed (Smith
and Oppenheimer, 2005). Wightman and Turner (2008) recently
showed that actin participates in secondary cell wall synthesis by
delivering organelles containing fluorescently labeled CESA to
bands of secondary cell wall thickenings in developing xylem
cells. Thus, the role of actin in cellulose synthesis seems to be
related to the intracellular trafficking of cellulose synthases, but
this needs to be further confirmed.
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While it is generally accepted that the microtubule cytoskeleton influences patterns of deposition of cellulose, the actin and/
or microtubule cytoskeleton may also influence the rate of cell
wall synthesis by regulating the delivery of cell wall–synthesizing
enzymes to the plasma membrane. The regulation of cellulose
synthesis activity through cytoskeleton-dependent intracellular
trafficking has remained largely unexplored. Here, we studied the
intracellular trafficking of CESA3, an essential enzyme for primary cell wall cellulose synthesis. We demonstrate that Golgi
bodies carrying CSCs are distributed throughout the cell via the
acto-myosin system and that Golgi bodies pause on cortical
microtubules for the targeted delivery of CSCs to the plasma
membrane. Under conditions of growth cessation or treatment
with a cellulose synthesis inhibitor, CSCs are internalized into
compartments that show movement dependent on cortical
microtubule array dynamics. Together, our data establish novel
roles for the actin and microtubule cytoskeleton in regulating
cellulose synthesis in the primary cell wall.
RESULTS
CSC Density and Trajectories Are
Developmentally Regulated
To examine regulation of the localization of CESA proteins, we
used stably transformed Arabidopsis thaliana lines expressing
green fluorescent protein (GFP)-tagged CESA3 fusion protein
under the control of the CESA3 endogenous promoter. This
construct has been shown to complement the cesa3 mutant
phenotype (Desprez et al., 2007). We studied GFP-CESA3 on the
outer face of epidermal cells in 3-d-old etiolated hypocotyls, in
which epidermal cells show an acropetal developmental gradient
(Gendreau et al., 1997). Cells near the apical hook elongate
slowly and deposit a thick, external cell wall, cells in the middle of
the hypocotyl elongate rapidly, and cells at the base are fully
elongated (Refregier et al., 2004). As described previously,
discrete GFP-CESA3 particles were found aligned at the cell
surface (Figure 1A; Desprez et al., 2007). These particles exhibited steady, bidirectional movement in the focal plane of the
plasma membrane (Figures 1B and 1C; see Supplemental Movie
1 online). Immunogold labeling of cryosections confirmed the
plasma membrane localization of the GFP-CESA3 surface particles (Figure 1D), leading us to conclude that these particles
represent active CSCs in the plasma membrane. CSC density
was highest in cells in the apical hook and decreased toward the
base of the hypocotyl (Figure 2). The orientation of CSC trajectories and cortical microtubules visualized using a GFP fusion to
Arabidopsis a-tubulin6 (GFP-TUA6) varied from longitudinal to
transverse in the apical hook and hypocotyl top (Figures 2A and
2B), becoming strictly longitudinal at the base of the hypocotyl
(Figures 2C and 2D). Notably, the densities of cortical microtubules and CSCs were also correlated (Figure 2).
GFP-CESA3 Accumulates in Golgi Bodies and in the
VHA-a1 Compartment
GFP-CESA3 labels ring-shaped structures with bright, peripheral
punctae, previously shown to be Golgi bodies (Figure 1H;
Paredez et al., 2006). Golgi bodies circulate through the cytoplasm via the actomyosin system (Sparkes et al., 2008) and are
characterized by rapid, nonlinear movement (Figure 1I; see
Supplemental Movie 1 online). Electron microscopy (EM) images
of immunogold-labeled cryosections of hypocotyl epidermal
cells confirmed that GFP-CESA3 was present at the periphery
of the Golgi stack, consistent with their ring shape in confocal
images (Figures 1P and 1Q). In addition to Golgi bodies, we
identified a second, relatively rare population (<1 observed per
cell per 10 min imaging period) of smaller, homogeneously
labeled particles with the same rapid, nonlinear movement as
Golgi bodies (Figures 1J and 1K). These particles were invariably
labeled with the early endosome/trans-Golgi network (TGN)
marker VHA-a1-mRFP (Dettmer et al., 2006) in transgenic lines
expressing both markers (41 time series analyzed; Figures 1N
and 1O). VHA-a1 compartments rapidly associated and dissociated with Golgi bodies (Figures 1L to 1O; see Supplemental
Movie 2 online). Most often a single VHA-a1 compartment was
found centered beneath a Golgi stack (Figure 1L). The occasional
presence of GFP-CESA3 in the TGN was further confirmed in EM
micrographs (Figures 1Q and 1R). Thus, we will refer to these
particles as VHA-a1/GFP-CESA3–containing compartments.
Regulated Distribution of GFP-CESA3 in a Unique
Population of Intracellular Particles
We identified a third population of small, bright particles just
below the plasma membrane, which were distinguished from
CSCs by their fluctuating velocities and intracellular localization
(Figures 1E to 1G; see Supplemental Figures 2A to 2F online).
These particles were likewise distinguished from Golgi bodies
and VHA-a1/GFP-CESA3–containing compartments by their
smaller size and linear trajectories (Figures 1E to 1G). In basal
hypocotyl cells with low CSC density, GFP-CESA3 signal was
preferentially found in these bright intracellular compartments
(Figure 2D). Interestingly, treatment with 500 mM mannitol to
induce osmotic stress or treatment with 50 mM cycloheximide
both caused a rapid decrease in CSC density and a simultaneous
increase in the density of the same bright compartments (Figures
3A and 3B). To determine the response kinetics, we examined
the redistribution of GFP-CESA3 under mannitol treatment in the
root tip, a widely used, well-characterized system (Robinson
et al., 2008) where the absence of a thick cuticle allows rapid
penetration of treatment solutions. In the root tip, CSC redistribution was complete within 6 min (five independent experiments), suggesting that these bright intracellular compartments
result from internalization of CSCs as opposed to a block in
secretion (Figures 3C and 3D).
CGA 325’615 (CGA), a potent inhibitor of cellulose synthesis in
cotton fibers (Peng et al., 2001), causes reduced crystalline
cellulose content and isotropic cell expansion in Arabidopsis (see
Supplemental Figure 3 online). CGA treatment did not affect
microtubules labeled with either GFP-TUA6 or a GFP fusion to
the Arabidopsis microtubule end binding protein1A (EB1-GFP) or
actin filaments marked by an N-terminal GFP fusion to the
second actin binding domain of Arabidopsis fimbrin 1 (GFPFABD2) or alter the morphology of the Golgi apparatus or
organelles observed by EM (see Supplemental Figure 4 online).
Cellulose Synthase Trafficking
1143
Figure 1. Four Distinct Localizations of GFP-CESA3 in Epidermal Cells of Etiolated Arabidopsis Hypocotyls.
(A) to (D) Particles corresponding to CSCs in the plasma membrane.
(A) Maximum projected z-stack of first time point in a series showing CSC arranged in linear tracks in the plasma membrane focal plane.
(B) Average projection of the same time series showing organized CSC trajectories.
(C) Kymograph along the trajectory indicated by a line in (B). Time is represented along the vertical axis, and a 5-min scale bar is shown at left (see
Supplemental Figure 1 online for information on interpreting kymographs). CSC movement is steady and bidirectional (see Supplemental Movie
1 online).
(D) Ultrathin cryosection of epidermal cells in the upper hypocotyl of seedlings expressing GFP-CESA3, labeled with anti-GFP primary antibody and
a Nanogold-coupled secondary antibody. Control labeling of wild-type seedlings was negative. Discrete CSCs are labeled in the plasma membrane.
Bars = 500 nm.
(E) to (G) Small, bright, intracellular particles with fluctuating velocity (referred to as MASCs).
(E) Maximum projected z-stack of first time point in a series acquired in a cell near the apical hook; a small, bright, intracellular particle is indicated by an
arrowhead.
(F) Average projection of the time series.
(G) Kymograph along the trajectory indicated in (E). The movement of the small, bright particle is along a linear track, but different from that of the
surrounding CSCs. Initial unsteady translocation is followed by acceleration in the opposite direction.
(H) and (I) GFP-CESA3–labeled Golgi bodies.
(H) GFP-CESA3 is found exclusively at the periphery of Golgi bodies and is characterized by bright punctae (deconvolved image).
(I) Trajectories of four Golgi bodies indicated by colored lines overlaid on an image from the time series. Golgi body movement is nonlinear and
interrupted by pauses.
(J), (K), (N), and (O) VHA-a1/GFP-CESA3–containing compartments.
(J) Image from a time series showing a GFP-CESA3–labeled compartment (arrowhead) just below the plasma membrane.
(K) The white line illustrates the trajectory of the compartment marked in (I). The movement pattern is similar to that of Golgi. The particle associates with
a Golgi body before disappearing into a lower focal plane.
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Time-lapse imaging of GFP-CESA3 revealed that CGA treatment
resulted in clearance of CSC from the plasma membrane,
concomitant with an increase in the density of small, bright
intracellular compartments (Figure 3E) identical in appearance,
behavior, and velocity to those observed in untreated basal
hypocotyl cells and those induced by osmotic stress or cycloheximide treatment (cf. Figures 2D, 3A, 3B, and 3E; see Supplemental Figure 2G online). Mock treatment with an identical
dilution of DMSO did not observably alter GFP-CESA3 localization (see Supplemental Figure 5A online).
Characterization of CGA-Induced Compartments
CGA-induced compartments were observed to move with fluctuating velocities along linear tracks (Figure 3F; see Supplemental Movie 3 online). These linear tracks could be visualized in
average projections of time series, and their position was observed to change over time (Figure 3G). In image acquisitions of
10 to 15 min, individual compartments could be found that were
stationary, migrating steadily at 10 to 3000 nm/min, or moving as
fast as 8900 nm/min (Figure 3N; see Supplemental Movie 3
online). The linear trajectories of CGA-induced compartments
contrast with the movement of intracellular compartments
known to be regulated by the actomyosin system, such as Golgi
bodies. To test if an intact actin cytoskeleton is required for the
movement of CGA-induced compartments, we treated GFPCESA3–expressing seedlings first with CGA and then with the
actin-disrupting agent Cytochalasin D (CytD). CytD causes fragmentation of actin microfilaments and inhibits their dynamics
(see Supplemental Figures 5D and 5E online; Foissner and
Wasteneys, 2007). Treatment with CytD resulted in aggregation
of GFP-CESA3–labeled Golgi bodies within 25 min, but no
difference was found in the density or velocity distribution of
CGA-induced compartments compared with CGA treatment
alone (Figures 3H, 3I, and 3N).
To further study the nature of CGA-induced compartment
movement, seedlings were pretreated with the microtubulestabilizing drug taxol and then treated with taxol + CGA. The
resulting CGA-induced compartments showed a density similar
to that in CGA treatment alone (Figure 3J); however, the addition
of taxol greatly reduced the motility of the compartments (Figures
3K and 3N). Out of 989 individual particles measured, only 11
were found with velocities equal to or greater than the mean
velocity of 543 nm/min measured in CGA treatment (Figure 3N).
As a third test, seedlings were pretreated with 7 mM oryzalin,
which results in depolymerization of most cortical microtubules
within 4 h (see Supplemental Figures 5B and 5C online). The
subsequent treatment with oryzalin + CGA resulted in reduced
CSC density; however, we never observed characteristic CGAinduced compartments with normal density near the cell surface
(41 cells from two independent treatments; Figure 3L). In one cell,
we found a small group of aligned particles resembling CGAinduced compartments, but these particles were stationary
(Figure 3M), suggesting they were associated with fragments
of stable, resistant microtubules.
Based on these results, we conclude that an intact, dynamic
cortical microtubule array is essential for the movement of CGAinduced compartments, and the actin cytoskeleton does not
appear to have a role in their movement. Henceforth, we will refer
to CGA-induced compartments as microtubule-associated cellulose synthase compartments (MASCs). To verify that MASCs
observed in untreated cells were also associated with cortical
microtubules, we transiently transformed cotyledons of the GFPCESA3–expressing line with the microtubule binding domain
(MBD) of the microtubule-associated protein 4 fused to monomeric red fluorescent protein (mRFP). Although plants stably
overexpressing GFP-MBD and cultured on kanamycin were
observed to show growth defects in one study (Granger and
Cyr, 2001), MBD remains a widely used marker for microtubules
in plants and yields signals highly similar to that obtained with
immunolabeling of tubulin (Sainsbury et al., 2008). In the absence
of any drug treatments, we observed MASCs migrating along
cortical microtubules labeled by mRFP-MBD (Figure 3O; see
Supplemental Movie 4 online). Intriguingly, MASC particles
moving along linear tracks merge with one another, progressively
increasing in intensity and cotranslocating until a split occurs
(Figure 3P; see Supplemental Movie 3 online). This merging
behavior is reminiscent of that observed for the DAM1 ring
protein, known to move on depolymerizing microtubule ends
(Westermann et al., 2006). Indeed, MASC velocities (0 to 8.9 mm/
min) are within the range of velocities measured for depolymerization at leading or lagging microtubule ends (0 to 30 mm/min)
and polymerization at leading ends (0 to 15 mm/min) (Shaw et al.,
2003).
The Golgi Apparatus Pauses on Cortical Microtubules, and
Golgi Bodies and the VHA-a1 Compartment Physically
Interact with MASCs
High-frame-rate high-resolution imaging of GFP-CESA3 in combination with mRFP-MBD or VHA-a1-mRFP markers yielded
several salient observations. GFP-CESA3–labeled Golgi bodies
Figure 1. (continued).
(L) to (O) Confocal images from plants coexpressing GFP-CESA3 (green) and VHA-a1-mRFP (red). Bar = 5 mm.
(L) The VHA-a1 compartment was often found centered on the Golgi stack, without any detectable colocalization of VHA-a1-mRFP and GFP-CESA3.
(M) and (N) VHA-a1 compartments partially dissociated from Golgi show variable levels of colocalization with GFP-CESA3 (see Supplemental Movie 2
online).
(O) Two VHA-a1/GFP-CESA3–containing compartments physically separated from Golgi and labeled by GFP-CESA3. PM, plasma membrane; CP,
cytoplasm.
(P) to (R) Ultrathin cryosections of epidermal cells in the upper hypocotyl of seedlings expressing GFP-CESA3, labeled with anti-GFP primary antibody
and a Nanogold-coupled secondary antibody. Control labeling of wild-type seedlings was negative. G, Golgi stack; T, TGN. Bar = 500 nm.
(P) and (Q) Gold marker can be detected at the rims of many Golgi stacks and, less often, also surrounding the TGN ([Q] and [R]).
Bars = 10 mm except where noted.
Cellulose Synthase Trafficking
1145
were observed to pause at discrete sites on microtubules (Figures 4A to 4C; see Supplemental Movie 4 online). In one representative time series, 63 Golgi bodies were tracked and pause
events were quantified for each individual Golgi body. A pause
event was defined as a period of immobility lasting at least 8 s.
One or more pause events were observed for 48 of the 63 Golgi
bodies (76%), and in each instance, the pause event occurred on
a microtubule. The remaining 24% of the Golgi bodies moved
rapidly in and out of frame without undergoing a pause event. We
observed multiple Golgi bodies successively pausing at the
same positions (45 independent time series analyzed; presented
in the following section). Detailed analysis in untreated cells in the
hypocotyl revealed that Golgi pauses occurred in proximity to
MASCs (Figures 4D and 4E) and could last as long as 73 s (n = 34,
mean = 22 s). VHA-a1 compartments were also observed to
pause at MASCs, even in the absence of an associated Golgi
body (seven events from five independent experiments; Figure
4D). Visible connections between MASCs and Golgi bodies or
VHA-a1 compartments were prominent in CGA-, mannitol-, and
cycloheximide-treated cells (see Supplemental Movies 3 and 5
online; Figure 4E).
CSC Delivery to the Plasma Membrane Depends on
Interactions between the Golgi Apparatus and
the Cytoskeleton
The high density of CSCs in cells near the top of the hypocotyl
has thus far prevented visualization of individual CSC particle
insertions (DeBolt et al., 2007b). We circumvented this problem by focusing our attention on cells closer to the hypocotyl
base, where CSC density is lower (Figure 2C). Through the
rigorous analysis of time series acquired in these untreated
cells, we were able to identify seven CSC insertion events. Each
insertion event was analyzed both manually and through
kymographs to verify the absence of the CSC in the initial
frames, to monitor the movement of all particles in proximity to
the identified insertion site, and to measure the velocity of the
newly inserted CSC. The newly inserted CSCs all began translocating with a velocity approximating 270 nm/min, as expected
Figure 2. CSC Density and Trajectory Change with the Developmental
Gradient along the Length of Etiolated Hypocotyls.
(A) to (D) In the left-most and central column are maximum projected
z-stacks and average projections of time series, respectively, of CSCs
visualized in the outer face of epidermal cells in GFP-CESA3–expressing
plants. The right-most column shows maximum projected z-stacks of
GFP-TUA6–labeled microtubules in the same cell types.
(A) Mixed CSC trajectories and microtubule orientations in slowly
expanding cells of the apical hook. The high density of CSCs in these
cells correlates with the high rate of wall synthesis.
(B) A wide range of orientations from transverse to longitudinal is found in
rapidly expanding cells ;2 mm below the apical hook.
(C) Approximately 6 mm below the apical hook, in cells beginning to
cease elongation, CSC density is reduced and trajectories are consistently longitudinal.
(D) CSCs are nearly absent from cells 10 mm below the hook, and
cortical microtubules are equally sparse. Small, bright, intracellular
compartments, referred to as MASCs (arrowheads), are abundant in
these basal hypocotyl cells. Bar = 10 mm.
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Figure 3. Small, Bright, Intracellular Compartments Represent a MASC Whose Movement Is Dependent on Cortical Microtubule Dynamics.
(A) and (B) Maximum projected z-stacks showing GFP-CESA3 intracellular distribution in Arabidopsis epidermal hypocotyl cells. The ring-shaped
labeling of Golgi bodies is present in all treatments. Treatments with 50 mM cycloheximide (CHX) for 4 h (A) or 500 mM mannitol for 1.5 h (B) result in
disappearance of CSCs (cf. to Figure 1A) and accumulation of MASCs.
(C) and (D) Maximum projected z-stacks showing GFP-CESA3 intracellular distribution in the root tip upon treatment with 500 mM mannitol.
(C) After 2 min of treatment, CSCs are still visible in linear tracks in the plasma membrane, as they are in control treatments.
(D) After 6 min of treatment, CSCs have been internalized in MASCs. In five independent experiments, MASCs formed after ;5 to 6 min.
(E), (H), (J), and (L) Maximum projected z-stacks showing the GFP-CESA3 intracellular distribution in Arabidopsis epidermal hypocotyl cells. Labeling of
Golgi bodies is found in all treatments. Representative images are shown from treatments with 5 nM CGA for 5 h (E), 5 nM CGA followed by 5 nM CGA +
50 mM CytD (H), 4 mM taxol followed by 5 nM CGA + 4 mM taxol (J), and 7 mM oryzalin followed by 5 nM CGA + 7 mM oryzalin (L). In (E), (H), and (J),
MASCs formed at high densities near the cell surface, as they did in the presence of 5 nM CGA treatment alone, for an equal time duration. In (L), MASCs
did not form normally and are largely absent.
(F) Representative kymograph of a time series for 5 nM CGA treatment (corresponding to the region indicated by a line in [E]). Time is represented along
the vertical axis, and a 10-min scale bar is shown at left (see Supplemental Figure 1 online for information on interpreting kymographs). With 5 nM CGA
treatment alone, MASCs are highly motile and have inconstant velocities (cf. to Figure 1F; see Supplemental Movie 3 online).
(G) MASCs move along linear tracks that are remodeled over time. From left to right: the first time point in a series, average projection of the first 5 min of
the series, average projection of the last 6 min of the series, overlay of the two average projections. The red lines and arrowheads indicate new
trajectories that were not present at the 5 min time point.
(I), (K), and (M) Representative kymographs of time series for treatments with 5 nM CGA followed by 5 nM CGA + 50 mM CytD (I), 4 mM taxol followed by
Cellulose Synthase Trafficking
for GFP-CESA3–labeled CSCs (Desprez et al., 2007). In each
insertion event, appearance of new CSC particles was coincident with the pause of a Golgi body just beneath the plasma
membrane (Figures 5A to 5C). In two of these events, this pause
of the Golgi body resulted in the insertion of multiple particles in
a row (Figure 5B).
To further study the role of the Golgi apparatus and VHA-a1/
GFP-CESA3–containing compartments in CSC trafficking, we
inhibited the movement of these two compartments using
actin-depolymerizing agents. Incubation with CytD or Latrunculin B caused aggregation of Golgi bodies and VHA-a1/
GFP-CESA3–containing compartments and led to localized
reductions in CSC density (Figure 5D). Regions of the plasma
membrane containing high CSC density were found exclusively
above Golgi aggregates (seven independent experiments; Figures 5D to 5F).
We further explored the insertion of new CSC particles using
fluorescence recovery after photobleaching (FRAP) experiments
in untreated seedlings. We found that CSCs are not inserted
randomly, but in linear tracks resembling those present before
bleaching (Figure 5G). In these experiments, we could also
observe multiple CSCs inserted one after another in rows, and
these insertions were coincident with the passage of Golgi
bodies (13 experiments analyzed). Based on our observations
of the frequent pauses of Golgi bodies on microtubules (Figures
4A to 4C; see Supplemental Movie 4 online), and the insertion of
CSCs in rows along linear tracks, we hypothesized that interactions between Golgi bodies and microtubules were directing the
plasma membrane delivery of CSCs.
To test if cortical microtubules define the insertion sites of
CSCs, we examined the effects of treatment with oryzalin or taxol
on the organization of CSCs in the plasma membrane. Complete
removal of microtubules by treatment with 20 mM oryzalin
resulted in a uniform distribution of CSCs that lacked row
organization compared with controls (five independent experiments; Figure 5I). By contrast, microtubule stabilization through
taxol treatment resulted in an enhanced alignment of CSCs (five
independent experiments; Figure 5J). In light of our results, we
propose that disorganization of CSC insertion in the presence of
microtubule-disrupting drugs is partly due to deregulation of
interactions between the Golgi/VHA-a1/GFP-CESA3–containing
compartments and cortical microtubules.
1147
DISCUSSION
Synthesis of the plant cell wall requires secretion of cell wall
components and CSCs to specific locations at the cell periphery
by motile organelles. In this study, we establish a role for the actin
and cortical microtubule cytoskeletons and the Golgi apparatus
in the delivery of CSCs to the plasma membrane and shed light
on the mechanism of endocytosis of CSCs. A schematic model
of the intracellular trafficking of GFP-CESA3 is presented in
Figure 6.
Four Distinct Localizations of GFP-CESA3
It had been previously established using confocal microscopy
that GFP-CESA3 and GFP-CESA6 (another related primary cell
wall CSC isoform) are present as discrete particles at the cell
surface and in punctae at the periphery of Golgi stacks (Paredez
et al., 2006; Desprez et al., 2007). In this study, the plasma
membrane localization of GFP-CESA3 surface particles was
confirmed by EM (Figure 1D), lending further support to the idea
that these motile, discrete particles represent CSCs actively
synthesizing cellulose. We also confirmed the localization of
GFP-CESA3 at the periphery of medial- and trans-Golgi cisternae by EM (Figures 1P and 1Q). Early reports show that the CSC
is fully assembled upon arrival in the Golgi apparatus (Haigler and
Brown, 1986; Rudolph, 1987). The bright punctae observed at
the periphery of Golgi stacks may therefore correspond to
assembled CSCs, as represented in Figure 6.
In our experiments, we were able to identify two previously
unknown localizations for the CESA protein. We found GFPCESA3 in a subset of VHA-a1 compartments (Figures 1J to 1O)
and a unique compartment we refer to as a MASC (Figures 1E to
1G and 3; see Supplemental Figure 2 online). In our analyses,
pausing of the Golgi apparatus was always observed prior to
insertion of a new CSC in the plasma membrane, whereas VHAa1/GFP-CESA3–containing compartments were not detected in
proximity to these insertion sites (Figures 5A to 5C). Although we
cannot exclude that CSCs pass through VHA-a1/GFP-CESA3–
containing compartments on a time scale too rapid to resolve in
our experiments, it is more likely that GFP-CESA3 passes
through the VHA-a1 compartment only on the endocytic route,
as shown for PIN2 (Robert et al., 2008). Since VHA-a1 labels the
Figure 3. (continued).
5 nM CGA + 4 mM taxol (K), or 7 mM oryzalin followed by 5 nM CGA + 7 mM oryzalin (M). Time is represented along the vertical axis, and a 10 min scale
bar is shown at left (see Supplemental Figure 1 online for information on interpreting kymographs). The kymograph in (M) represents the region indicated
by a line in (L).
(N) Frequency histogram comparing MASC velocities calculated from kymographs in replicates of the experiments described in (C), (H), and (I) (CGA
alone, n = 278; CGA + taxol, n = 989; CGA + CytD, n = 342). The points indicate the percentage of particles with velocities in the range indicated on the x
axis. Lines have been added to facilitate visualization of each population distribution.
(O) Cotyledon cells of untreated GFP-CESA3–expressing plant transiently transformed with mRFP-MBD (red). The maximum projected z-stack (left
panel) shows a GFP-CESA3-labeled MASC particle (green) marked by an arrowhead. Right panel, the average projection of the time series showing
translocation of the MASC along the mRFP-MBD–labeled microtubule (see Supplemental Movie 4 online).
(P) Montage of a deconvolved time series illustrating representative merging behavior of four aligned MASCs in cells treated with 50 mM cycloheximide
for 4 h. At 8.8 s, the fourth particle (from the top) merges with the third, and both continue cotranslocating. A second encounter at 19.8 s results in a rapid
split of the particle. The resulting three particles then remain relatively stable in their positions.
Bars = 10 mm.
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The Plant Cell
Figure 4. Golgi Bodies Pause along Microtubules.
Golgi bodies and the VHA-a1 compartment exhibit physical interactions with MASCs. Bars = 10 mm.
(A) to (C) Deconvolved images of a cotyledon cell from an untreated GFP-CESA3–expressing plant transiently transformed with mRFP-MBD (red).
(A) CSCs are observed along the cortical microtubules. The arrowhead shows a future pause site of the Golgi body marked by an arrow.
(B) Average projection of the time series. The yellow color indicates colocalization between motile CSCs and microtubules.
(C) A blue line indicating the trajectory of the Golgi body is overlaid on the average projection. The Golgi body follows this microtubule and pauses
directly on it. See Supplemental Movie 4 online for a second example.
(D) Deconvolved time series from untreated plants coexpressing GFP-CESA3 (green) and RFP-VHA-a1 (red). An associated Golgi/VHA-a1
compartment (arrow) moves rapidly into view and pauses at an immobile MASC (arrowhead) for ;20 s. At the 52 s time point, the Golgi dissociates,
but the VHA-a1 compartment remains colocalized with the MASC. On the right-most image, the blue line indicates the trajectory of the VHA-a1
compartment.
(E) Connections are visible between Golgi and MASCs during interactions. Representative deconvolved time series (same series as in Figure 3L) from a
50-mM cycloheximide-treated seedling showing a GFP-CESA3–labeled Golgi body (blue arrow) changing trajectory and pausing at a MASC (red
arrowhead).
TGN (Dettmer et al., 2006), these combined data question the
role of the TGN in secretion in plants.
Cellulose Synthesis Is Regulated by Rapid Internalization
into MASCs
Under conditions of osmotic stress, protein synthesis inhibition,
or cellulose synthesis inhibition, we find GFP-CESA3 present in a
novel compartment we refer to as a MASC (Figures 3A, 3B, and
3E). Lower densities of CSCs in fully elongated basal hypocotyl
cells correlated with a higher abundance of MASCs, consistent
with a reduced rate of primary cell wall cellulose synthesis in
these mature cells (Figure 2D). We conclude that MASCs result
from internalization of CSCs, rather than a block in secretion,
based on the following three observations: formation of MASCs
correlates with disappearance of CSCs at the plasma membrane
(Figure 2D), redistribution of CSCs into MASCs occurs within only
6 min in mannitol-treated roots (Figures 3C and 3D), and MASCs
still form despite prolonged inhibition of protein synthesis using
the drug cycloheximide (Figure 3A). Clathrin-mediated endocytosis has recently been shown to be a predominant endocytic
pathway in plants (Dhonukshe et al., 2007). Given that the 45- to
50-nm diameter cytoplasmic domain of the CSC (Bowling and
Brown, 2008) would be expected to interfere with the formation
of the clathrin coat, it is likely that a different mechanism operates
in the endocytosis of CSCs (Figure 6). Indeed, the reduced
formation of MASCs in the presence of oryzalin suggests that
microtubules may have an active role in the internalization of
CSCs under CGA treatment. Oryzalin also prevented the redistribution of YFP-CESA6 into similar small, bright compartments
Cellulose Synthase Trafficking
1149
in seedlings treated with Morlin, an inhibitor of CESA and
microtubule dynamics (DeBolt et al., 2007a). Together, these
results show that cellulose synthesis is regulated in part through
the internalization of CSCs in intracellular compartments and that
internalization of CSCs into MASCs is environmentally regulated.
The higher abundance of MASCs in fully elongated basal hypocotyl cells compared with growing apical hypocotyl cells suggests that internalization in MASCs is also developmentally
programmed.
MASC Movement Is Dependent on an Intact, Dynamic
Cortical Microtubule Array
The unusual movement pattern of MASCs (see Supplemental
Movies 3 and 5 online) has not been reported for any other plant
Figure 5. CSC Delivery to the Plasma Membrane Depends on Interactions between the Golgi Apparatus and the Cytoskeleton.
(A) to (C) Images from times series showing three examples of de novo
CSC insertion in the plasma membrane focal plane by GFP-CESA3–
labeled Golgi bodies in untreated seedlings. Kymographs of the time
series are at the far right, with time represented along the vertical axis as
indicated by the scale bars (see Supplemental Figure 1 online for
information on interpreting kymographs).
(A) and (B) In the left-most image, no particle can be detected at or near
the future position of insertion (corresponding to black region above the
trace in the kymograph). The central image shows a Golgi body pausing
beneath the plasma membrane, which can also be detected on the
kymograph at right (white smear marked by arrow). In (A), a single CSC is
inserted and begins moving nearly immediately, as evidenced by the
angled white trace on the kymograph. In (B), three new CSC are inserted
in a row and begin moving steadily (only one is kymographed).
(C) Image 1 shows a first Golgi body pausing at a specific site marked by
an arrowhead. Image 2 shows this pause does not result in an insertion
event. In image 3, a new Golgi body moves into frame and pauses at the
same site (arrowhead), resulting in insertion of a CSC, marked by the
arrowhead in image 4. The white smears on the kymograph (marked by a
gray arrow and black arrow) illustrate the pauses of successive Golgi and
the newly inserted CSC that begins moving with a steady velocity.
(D) to (F) GFP-CESA3 intracellular distribution in epidermal hypocotyl
cells treated with 1 mM Latrunculin B.
(D) Maximum projected z-stack in the plasma membrane focal plane
shows localized reductions in CSC density.
(E) Average projection of the time series illustrates that CSC are migrating normally along linear trajectories.
(F) Maximum projected z-stack ;600 nm below the plasma membrane
showing Golgi body aggregates (arrows). Positions of Golgi body aggregates correspond to the areas of highest CSC density in (A).
(G) FRAP in untreated epidermal hypocotyl cells. The prebleach image is
an average of 10 frames showing CSCs arranged in obliquely oriented
rows. The central image was acquired ;500 ms after bleaching the
indicated area. In the average projection of the postbleach frames, newly
inserted CSCs are arranged in oblique rows and migrate along linear
tracks similar to those present in the prebleach image.
(H) to (J) Images of a single time point showing the organization of GFPCESA3–labeled CSCs in epidermal hypocotyl cells.
(H) Untreated seedling showing CSCs arranged in rows.
(I) After an 8-h treatment with 20 mM oryzalin, row organization is lost and
CSCs are scattered uniformly throughout the plasma membrane.
(J) A 7-h treatment with 4 mM taxol increases the level of CSC row
organization.
Bars = 10 mm.
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The Plant Cell
Figure 6. Schematic Illustration of CESA Trafficking.
Objects were drawn to scale based on published data and EM micrographs obtained in this study. CSCs may be secreted directly to the plasma
membrane from the Golgi apparatus or may transit rapidly through the TGN. CSC insertion is accompanied by interactions between the Golgi apparatus
and cortical microtubules. We define MASCs as a microtubule-associated compartment containing the CSC, which may encompass both short-lived
secretory vesicles and more stable endocytic vesicles. Although we were not able to directly observe CSC secretory vesicles, we hypothesize that they
may associate with microtubules if the CSC itself is competent for interactions with the microtubule array. It is possible that CSC secretory vesicles do
not associate with microtubules or that secretion occurs in the absence of a vesicular intermediate, though we find these hypotheses less likely. Once
fused with the plasma membrane, CSCs are propelled by the polymerization of glucan chains into cellulose microfibrils, and their trajectories may be
guided by CSC-cortical microtubule interactions of a transient nature. Internalization of CSCs is promoted under certain conditions (e.g., osmotic
stress, cell growth cessation, and cellulose synthesis inhibitor treatment). Consideration of the size of the CSC rosette (;25-nm diameter; Mueller and
Brown, 1980) and the CSC cytoplasmic domain (45 to 50-nm diameter; Bowling and Brown, 2008) with respect to the size of clathrin-coated vesicles
(CCV) (30-nm diameter excluding coat; Dhonukshe et al., 2007) suggests that endocytosis of CSC does not occur via conventional clathrin-coated
vesicle–mediated endocytosis. When internalized, CSCs show surprisingly stable associations with microtubules, which may be due to an interaction
between a microtubule-associated protein and the CSC itself or a coat protein specifically recruited to MASC vesicles. Together, our results support a
model in which interactions with cortical microtubules have a role both in plasma membrane delivery and internalization of CSCs. These interactions
provide a regulatory mechanism for the dynamic remodeling of CSC secretion during cell expansion and differentiation.
intracellular compartments or markers thus far, to our knowledge. The merging behavior of MASCs and their movement
along linear tracks clearly distinguish them from the actomyosinbased movement of Golgi bodies (Sparkes et al., 2008) and the
nonlinear movement of the VHA-a1 compartment (cf. Supplemental Movies 2 and 3 online). MASC movement is not significantly changed by the fragmentation of actin or the inhibition of
cytoplasmic streaming (Figure 3I). Both the velocity distribution
and the qualitative characteristics of MASC movement remain
the same in controls and CytD-treated cells (Figure 3N). By
contrast, the depolymerization of the cortical microtubule array
or the inhibition of array dynamics by treatment with taxol has a
striking effect on the formation and velocity of MASCs (Figures 3J
to 3N). The fact that stabilization of microtubule array dynamics
also immobilizes MASCs indicates that their movement may not
be kinesin driven, but rather depends on intrinsic properties of
the microtubule array dynamics itself.
Our findings lend support to the hypothesis that CSCs transiently interact with cortical microtubules, rather than merely
passively diffusing between them. For the microtubule-dependent
Cellulose Synthase Trafficking
movement of MASCs to be possible, either an interaction exists
between a microtubule-associated protein and a coat protein
specifically recruited to MASC vesicles or an interaction exists
between a microtubule-associated protein and a component of
the CSC itself. It will be interesting to determine if microtubuleassociated proteins can be copurified with CSCs under CGA
treatment. The discovery of proteins able to mediate interactions
between CSC and microtubules will be essential to reveal
the mechanism underlying microtubule-microfibril alignment in
growing cells.
Trafficking of CSCs Is Dependent on Actin and
Microtubule Cytoskeletons
Our combined results strongly support a role for the microtubule
cytoskeleton in targeting CSC secretion. CSC density and cortical microtubule density are correlated during cellular differentiation in the hypocotyl (Figure 2), suggesting that cortical
microtubules may be involved in regulating CSC delivery to the
plasma membrane. We demonstrate that new CSC insertions are
preceded by pauses of the Golgi apparatus at specific sites
beneath the plasma membrane (Figures 5A to 5C). The so-called
stop-and-go behavior of Golgi bodies was proposed to be due to
pauses at ER export sites (Nebenführ et al., 1999); however,
recent evidence suggests that ER export sites and Golgi stacks
may form mobile secretory units (daSilva et al., 2004). Our data
reveal that Golgi bodies pause at cortical microtubules, and
these pauses often result in insertion of CSCs at the plasma
membrane (Figures 5A to 5C).
Evidence of Golgi bodies pausing at cortical microtubules has
also been observed in cells undergoing high rates of secondary
cell wall synthesis. In developing xylem vessels, secondary cell
wall synthesis is nonuniform and characterized by cell wall
thickenings in a banded pattern. Within these bands, CSCs
are more abundant in the plasma membrane (Herth, 1985),
and microtubules are found in organized bundles (Hepler and
Newcomb, 1964). It was recently shown that organelles labeled
by YFP-CESA7, of which a subset colocalizes with the Golgi
marker mannosidase I, pause at bands marked by microtubules
(Wightman and Turner, 2008). These organelles appear to pause
less frequently following depolymerization of the microtubule
bands by oryzalin, although the statistical significance of this
observation can be questioned. Thus, there is limited evidence
that delivery of secondary cell wall CSCs to the plasma membrane in developing xylem vessels may also be related to the
pausing of Golgi bodies on microtubules.
In plants, Golgi bodies, mitochondria, and peroxisomes are
three organelles known to move along actin filaments via myosins (Sparkes et al., 2008). Although the actomyosin system may
drive the movement of these organelles, there is increasing
evidence showing their proper function also depends on interactions with cortical microtubules. The plant peroxisomal multifunctional protein localizes to peroxisomes and decorates the
cortical microtubule array, thus mediating interactions between
peroxisomes and cortical microtubules (Chuong et al., 2005).
The Arabidopsis internal motor kinesin, Kinesin-13A, and the
homologous Gossypium hirsutum Kinesin-13A have been shown
to localize to Golgi bodies (Lu et al., 2005). Arabidopsis kinesin-
1151
13a mutants show aggregation of Golgi bodies in trichomes and
have defects in trichome morphogenesis, suggesting a role for
Kinesin-13A in regulating secretion of cell wall components by
the Golgi apparatus (Lu et al., 2005). Similarly, mutants in the
kinesin FRAGILE FIBER1 show alterations in cellulose deposition
and reduced fiber strength (Zhong et al., 2002). Finally, cortical
microtubules were found in close association with electrondense secretory vesicles targeted to pectic mucilage pockets in
Arabidopsis seed coat cells (McFarlane et al., 2008). Thus, the
actin filament and microtubule components of the cytoskeleton
are highly interdependent, and each plays an important role in
secretion and organelle movement.
Our data support a model in which the actomyosin system
serves to distribute Golgi bodies throughout the cell, and sites of
secretion are defined through interactions with the microtubule
cytoskeleton. This finding explains the diffuse, randomized appearance of CSCs at the plasma membrane following oryzalin
treatment and is also supported by observations of enhanced CSC
alignment under taxol treatment. In conclusion, we propose that in
growing cells, cortical microtubules not only direct the trajectories
of CSCs in the plasma membrane, but also regulate the delivery of
CSCs to the plasma membrane and possibly their internalization.
This allows the dynamic remodeling of CSC secretion patterns
during the expansion and differentiation of plant cells.
METHODS
Plant Material and in Vitro Growth Conditions
Seeds of Arabidopsis thaliana ecotype Columbia (Col) were provided by
K. Feldman (University of Arizona, Tucson, AZ), and GFP-CESA3 and
VHA-a1-mRFP lines were described previously (Dettmer et al., 2006;
Desprez et al., 2007). GFP-TUA6 (Ueda et al., 1999) and EB1-GFP (Chan
et al., 2003) lines were a generous gift of J. Chan (John Innes Centre,
Norwich, UK). GFP-FABD2 was a gift of M. Pastuglia and has been
described previously (Voigt et al., 2005). For imaging, seedlings were
cultured in chambers as described (Chan et al., 2007). For hypocotyl
measurements, seedlings were grown at 208C in Petri dishes on Estelle
and Somerville medium (Estelle and Somerville, 1987) without sucrose.
Seeds were cold-treated at 48C for 48 h, exposed to fluorescent white
light (150 mmol m22 s21, True Light; Philips) for 6 h, and then wrapped in
aluminum foil. The age of the seedlings was defined with respect to the
end of the cold treatment.
Hypocotyl Length Measurements
Etiolated seedlings were fixed in 0.2% paraformaldehyde. The lengths of
at least 50 hypocotyls per treatment were measured using Optimas 5.2
software as described (Refregier et al., 2004).
Generation of the GFP-CESA3/RFP-VHA-a1 Transgenic Line
Pollen from GFP-CESA3–expressing plants in je5cesa3 background
(Desprez et al., 2007) was used to fertilize plants expressing VHA-a1mRFP (Dettmer et al., 2006). F1 seeds were collected and amplified. F2
seedlings were screened by PCR for the presence of both markers
and the cesa3 mutation. F3 selected seedlings were used for imaging.
Drug Treatments
For hypocotyl measurements, Col-0 seeds were germinated on media
containing CGA 325’615 (CGA) in the range of 0 (mock treatment DMSO
1152
The Plant Cell
dilution) to 5 nM and measured after 4 d. For Fourier transform infrared
microspectroscopy, Col-0 seeds were germinated on media plates
containing drugs at the indicated concentrations, and mutant seeds
were germinated on normal media. Measurements were taken after 4 d.
For live-cell imaging, treatment and mock treatment solutions were
injected into chambers using a fine-needle syringe under green light.
Seedlings expressing markers to verify drug treatment efficacy were
cultured in the same chambers as GFP-CESA3 seedlings. Single treatments were performed with 5 nM CGA for 5 to 6 h, 1 mM Latrunculin B for 6
h, 20 mM oryzalin for 8 h, or 500 mM mannitol for 3 h. Treatments combining
two different inhibitors were performed as follows: 7 mM oryzalin for 7 h,
then 5 nM CGA + 7 mM oryzalin for 5 h before analysis; 4 mM taxol for 8 h,
then 5 nM CGA + 4 mM taxol for 6 h; and 5 nM CGA for 5.5 h followed by 5
nM CGA + 50 mM CytD for 30 min. All chemicals (Sigma-Aldrich) were
dissolved in DMSO stock solutions and kept at 2208C for no longer than 6
months. Stocks were diluted at least 1000-fold in buffered water before
use. CGA 325’615 was kindly provided by Bayer CropScience.
For the experiment represented in Supplemental Figures 2A to 2F
online, 3-d-old light-grown seedlings were first incubated in a solution of
dilute DMSO (mock treatment) or 5 nM CGA for 10 to 15 min and then
mounted in a solution of 50 mM FM4-64 and imaged within 5 min.
Transient Expression in Arabidopsis Seedlings
The mRFP-MBD vector was constructed by M. Pastuglia (Institut JeanPierre Bourgin-Institut National de la Recherche Agronomique). The MBD
of microtubule-associated protein 4 was amplified from the GFP-MBD
construct (Marc et al., 1998) and cloned into Gateway vector pDONR207
(Invitrogen). To subclone the mRFP-MBD fusion downstream of the 35S
promoter, pDONR207:MBD was used in an LR reaction with destination
vector pH7WGR2 (Karimi et al., 2002). mRFP-MBD and GFP-CESA3
expression vectors were electroporated in Agrobacterium tumefaciens.
For transient expression studies, A. tumefaciens cultures were cultured
overnight at 288C and diluted to OD600 = 1 in 5% sucrose with 200 mM
acetosyringone. Infiltration of 4-d-old Arabidopsis seedlings was performed as described (Marion et al., 2008).
Spinning Disk Microscopy and Image Analysis
During all experiments, cell viability was verified by monitoring cytoplasmic streaming. Hypocotyls of 3-d-old etiolated seedlings or root tips of
3-d-old light-grown seedlings were analyzed on an Axiovert 200M
microscope (Zeiss) equipped with a Yokogawa CSU22 spinning disk,
Zeiss 100/1.4 numerical aperture oil objective, and Andor EMCCD iXon
DU 895 camera (Plateforme d’Imagerie Dynamique, Institut Pasteur,
Paris, France). A 488- or 560-nm diode-pumped solid-state laser was
used for excitation, and emission was collected using band-pass 488/25
and 568/25 filters (Semrock) for GFP and mRFP, respectively. Projections
presented in figures were acquired with a minimum of three z-slices using
a 300-nm step interval. For analysis of particle interactions, time series
were acquired at a frame rate of 2 to 4 frames/second.
FRAP experiments were performed using an inverted microscope
(Nikon TE2000) equipped with a cooled interline CCD detector (Roper
Coolsnap HQ2) and a piezoelectric stage driver (LVDT; Physik Instrument)
and operated with MetaMorph 7.1.6 software (Molecular Devices). Series
were acquired at 2 frames/second with a FRAP duration of 20 ms and an
xy resolution of 130 nm/pixel (2 3 2 binning).
When required, fluorescent images were deconvolved using a classic
maximum likelihood estimation deconvolution algorithm (Huygens Professional v. 3.1; Scientific Volume Imaging). The integrated morphometry
analysis function in Metamorph Offline (V. 7.0; Molecular Devices) was
used to detect and measure the orientations of microtubules. Particle
velocities were calculated from kymographs created in Image J (W.
Rasband, National Institutes of Health). Kymograph montages were
generated with the help of an Image J plug-in developed by F. Cordelières
(Plateforme d’Imagerie Cellulaire et Tissulaire , Unité Mixte de Recherche
146, Institut Curie). Image size and contrast were adjusted when needed
using Adobe Photoshop CS v. 8.0.1 (Adobe Systems).
Confocal Laser Scanning Microscopy
The images presented in Supplemental Figures 2A to 2F online were
obtained using a spectral Leica SP2 AOBS confocal microscope (Leica
Microsystems). GFP-CESA3 and the sterol dye FM4-64 (Molecular
Probes) were excited at 488 nm, and emission was collected at 492 to
551 nm and 645 to 833 nm, respectively. A 363 water immersion
objective (numerical aperture 1.2) and a zoom factor of eight were used
to collect z-series in a 512-scan format.
Fourier Transform Infrared Microspectroscopy
Four-day-old seedlings were crushed between two barium fluoride
windows and rinsed abundantly with distilled water for 2 min, before
drying at 378C for 20 min. For each mutant, 20 spectra were collected
from hypocotyls from four independent cultures (five seedlings from each
culture) as described (Mouille et al., 2003). The collected spectra were
baselined and normalized as described, and statistical analysis was
performed by a Student’s t test (Robin et al., 2003).
Crystalline Cellulose Measurements
Wild-type Col-0 plants were grown in darkness on plates containing 0.625
nM CGA or a DMSO mock treatment solution. Four-day-old seedlings
were harvested and stored in 70% ethanol at 48C. Crude cell wall extracts
were prepared as described (Reiter et al., 1993) with minor changes. Seed
coats were removed and plant material was incubated twice during 1 h at
708C in 70% ethanol. Pellets were washed in methanol for 5 min and
acetone for 2 min and were vacuum-dried. The dry cell wall pellets were
weighed in 2-mL reaction tubes, suspended in 3-mL Updegraff reagent
(acetic acid:nitric acid:water; 8:1:2; v/v), and further incubated in a boiling
water bath for 30 min. Crystalline cellulose was determined as described
(Scott and Melvin, 1953; Updegraff, 1969). Absorbance was determined
at 630 nm (Labsystems iEMS Reader MF). Measurements were performed in duplicate using independent seed batches.
Transmission Electron Microscopy
Degassed seedlings were high-pressure frozen, freeze-substituted, and
rehydrated (Ripper et al., 2008) before samples were infiltrated with the
cryoprotectant polyvinylpyrrolidone and sucrose, mounted on a specimen holder, and frozen in liquid nitrogen for ultrathin cryosectioning as
described before (Reichardt et al., 2007). Ultrathin thawed cryosections
were treated with 0.5% BSA, 1% milk powder in PBS (20 min) before
labeling with rabbit anti-GFP antibodies (Torrey Pines Biolabs; 1:500)
and silver-enhanced Nanogold coupled to goat anti-rabbit IgG (1:60)
(HQSilver, 8.5 min; Nanoprobes) for 60 min each.
Accession Numbers
Sequence data from this article can be found in the Arabidopsis Genome
Initiative or GenBank/EMBL databases under the following accession
numbers: CESA3, At5g05170; VHA-a1, At2g28520; EB1a, At3g47690;
TUA6, At4g14960; MBD from MAP4, M72414; and FABD2 from At FIM1,
At4g26700.
Supplemental Data
The following materials are available in the online version of this article.
Cellulose Synthase Trafficking
Supplemental Figure 1. Image Analysis Methods.
Supplemental Figure 2. Localization of GFP-CESA3 following CGA
Treatment in FM4-64–Labeled Cells.
Supplemental Figure 3. CGA Causes Specific Reductions in Crystalline Cellulose and Inhibits Cellular Elongation.
Supplemental Figure 4. CGA Does Not Have an Observable Effect
on the Cytoskeleton or Golgi.
Supplemental Figure 5. Effects of Drug Treatments Used in This
Study.
Supplemental Movie 1. CSCs Move Bidirectionally with Constant
Velocities.
Supplemental Movie 2. GFP-CESA3–Labeled Golgi Bodies and the
VHA-a1 Compartment Rapidly Associate and Dissociate.
Supplemental Movie 3. CGA-Induced MASCs Have a Wide Range of
Velocities and Merge with One Another along Linear Tracks.
Supplemental Movie 4. GFP-CESA3–Labeled Golgi Bodies Pause on
Microtubules.
Supplemental Movie 5. Physical Interactions Occur between
Mannitol-Induced MASCs and GFP-CESA3–Labeled Golgi Bodies.
ACKNOWLEDGMENTS
We thank Fabrice Cordelières (Plateforme d’Imagerie Cellulaire et
Tissulaire, Unité Mixte de Recherche 146, Institut Curie, Gif-sur-Yvette,
France) for development of plug-ins to facilitate image analysis; Vincent
Fraisier (Institut Curie, PICT-IBiSA Imaging Facility, Paris, France) for
assistance with FRAP experiments; Nicolas Chenouard (Unité Analyse
d’Images Quantitative, Institut Pasteur, Paris, France), Anne Danckaert,
and Christophe Machu for helpful advice and image analysis tools
(Plateforme d’Imagerie Dynamique, Institut Pasteur, Paris, France);
Olivier Grandjean (Plateforme de Cytologie et d’Imagerie Végétale,
Institut Jean-Pierre Bourgin, Institut National de la Recherche Agronomique) for technical assistance with the Leica SP2; and Jordi Chan
(John Innes Centre, Norwich, UK) and Martine Pastuglia (Institut JeanPierre Bourgin, Institut National de la Recherche Agronomique) for
contributing fluorescently tagged cytoskeleton markers. E.F.C. was
supported by the National Agency for Research Project ‘‘IMACEL’’
ANR-06-BLAN-0262. V.B. was supported by the European Union
Framework Program 6 (FP6) “CASPIC” NEST-CT-2004-028974 and
“Wallnet” RTN 512265. Part of this work was supported by FP6 program
037704 “AGRON-OMICS.” The Plateforme de Cytologie et d’Imagerie
Végétale is supported in part by the Région Ile-de-France.
Received December 19, 2008; revised March 10, 2009; accepted March
19, 2009; published April 17, 2009.
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Arabidopsis
Elizabeth Faris Crowell, Volker Bischoff, Thierry Desprez, Aurélia Rolland, York-Dieter Stierhof, Karin
Schumacher, Martine Gonneau, Herman Höfte and Samantha Vernhettes
Plant Cell 2009;21;1141-1154; originally published online April 17, 2009;
DOI 10.1105/tpc.108.065334
This information is current as of June 16, 2017
Supplemental Data
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References
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