Download Microtubules Regulate Dynamic Organization of Vacuoles in

Microtubules Regulate Dynamic Organization of Vacuoles in
Physcomitrella patens
Yoshihisa Oda1,8, Aiko Hirata1, Toshio Sano1,2, Tomomichi Fujita3, Yuji Hiwatashi4,7, Yoshikatsu Sato5,
Akeo Kadota6, Mitsuyasu Hasebe4,5,7 and Seiichiro Hasezawa1,2,*
1Department
Eukaryotic cells have developed several essential
membrane components. In flowering plants, appropriate
structures and distributions of the major membrane
components are predominantly regulated by actin
microfilaments. In this study, we have focused on the
regulatory mechanism of vacuolar structures in the moss,
Physcomitrella patens. The high ability of P. patens to
undergo homologous recombination enabled us stably to
express green fluorescent protein (GFP) or red fluorescent
protein (RFP) fusion proteins, and the simple body
structure of P. patens enabled us to perform detailed
visualization of the intracellular vacuolar and cytoskeletal
structures. Three-dimensional analysis and high-speed
time-lapse observations revealed surprisingly complex
structures and dynamics of the vacuole, with inner sheets
and tubular protrusions, and frequent rearrangements by
separation and fusion of the membranes. Depolymerization
of microtubules dramatically affected these structures
and movements. Dual observation of microtubules and
vacuolar membranes revealed that microtubules induced
tubular protrusions and cytoplasmic strands of the
vacuoles, indicative of interactions between microtubules
and vacuolar membranes. These results demonstrate
a novel function of microtubules in maintaining the
Regular Paper
of Integrated Biosciences, Graduate School of Frontier Sciences, The University of Tokyo, Kashiwanoha 5-1-5, Kashiwa,
Chiba, 277-8562 Japan
2Institute for Bioinformatics Research and Development (BIRD), Japan Science and Technology Agency (JST), Chiyoda-ku, Tokyo,
102-8666 Japan
3Department of Biological Sciences, Faculty of Science, Hokkaido University, Kita-ku Kita-10-jo-Nishi-8, Sapporo, 060-0810, Hokkaido,
Japan
4Division of Evolutionary Biology, National Institute for Basic Biology, Okazaki, 444-8585 Japan
5ERATO, JST, Okazaki, 444-8585 Japan
6Department of Biological Sciences, Graduate School of Science and Engineering, Tokyo Metropolitan University, Minami-Ohsawa 1-1,
Hachioji, Tokyo, 192-0397 Japan
7Department of Basic Biology, School of Life Science, The Graduate University for Advanced Studies, Okazaki, 444-8585 Japan
distribution of the vacuole and suggest a functional
divergence of cytoskeletal functions in land plant
evolution.
Keywords: Actin microfilament • Cytoskeleton • Microtubule
• Physcomitrella patens • Vacuolar membrane • Vacuole.
Abbreviations: BCECF-AM, 2,7-bis-(2-carboxyethyl)-5(6)carboxyfluoresceine-acetoxymethylester; CLSM, confocal
laser scanning microscopy; DMSO, dimethylsulfoxide; GFP,
green fluorescent protein; ER, endoplasmic reticulum; FM464, N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)
phenyl)hexatrienyl pyridinium dibromide; mRFP, monomeric
red fluorescent protein.
Introduction
Eukaryotic cells have developed several essential membrane
components, such as the Golgi body, endoplasmic reticulum
(ER), lysosome and vacuole. Their integrated functions,
through membrane traffic, are indispensable for cellular
activity, growth and proliferation. Each membrane component is regulated by a cytoskeleton that maintains its appropriate structure, function, distribution and inheritance in a
cell. In animal cells, the structure and distribution of both
8Present address: Department of Biological Science, Graduate School of Sciences, The University of Tokyo, Hongo 7-3-1, Bunkyo-ku, Tokyo,
113-0033 Japan.
*Corresponding author: E-mail, [email protected]. Fax, +81-4-7136-3706
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031, available online at www.pcp.oxfordjournals.org
© The Author 2009. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
855
Y. Oda et al.
the ER (Du et al. 2004, Borgese et al. 2006, Vedrenne and
Hauri 2006) and the Golgi apparatus (Allen et al. 2002, Egea
et al. 2006) are regulated mainly by microtubules. In contrast, flowering plant cells predominantly utilize actin microfilaments for the spatial regulation of their major membrane
components, including the ER (Sheahan et al. 2004, Runions
et al. 2005), Golgi (Boevink et al. 1998, Nebenführ et al. 1999)
and vacuoles (Higaki et la. 2006).
Vacuoles perform essential roles in metabolism, homeostasis and storage in plant cells. In particular, their large volumes
and ability to regulate osmotic pressures are indispensable
for plant cell morphogenesis. Recently, the visualization of
vacuoles in higher plant cells by use of fluorescent dyes or
green fluorescent protein (GFP) fusion proteins together
with reconstruction of their three-dimensional structures
has revealed the complexity of these structures (Yoneda et al.
2007). For instance, spherical membranes and sheet-like
membranes have been reported within the vacuolar lumen
of Arabidopsis epidermal cells (Saito et al. 2002), and dynamic
changes of vacuolar structures have been observed during
cell cycle progression in tobacco BY-2 cells (Kutsuna and
Hasezawa 2002, Kutsuna et al. 2003). Several reports have
suggested that these vacuolar structures co-localize with
actin microfilaments, rather than microtubules, and that
their maintenance and distribution are actin dependent
(Ovecka et al. 2005, Higaki et al. 2006).
To investigate vacuolar morphology and its regulatory
mechanisms, we have, in this study, established a new visualization system using the moss, Physcomitrella patens. This
tiny plant has a very simple body with filamentous tissue,
protonemata and rhizoids, composed of linearly linked single
cells, which can be observed directly by microscopy. Furthermore, since these tissues contain an apical meristematic cell
that repeatedly divides and produces stably matured cells,
we have been able to monitor the sequential events of cell
division, differentiation and elongation. In addition, we have
taken advantage of the feasible gene targeting technique
(Kammerer and Cove 1996, Schaefer and Zrÿd 1997, Schaefer
2001) to insert marker genes into the genome for visualization of intracellular structures. Finally, by using this moss,
one of the model land plants most distantly related to flowering plants, we can study the divergence and evolution of
endomembrane regulatory mechanisms.
By exploiting these advantages of P. patens, we have
succeeded in visualizing both the vacuolar membrane and
the cytoskeleton, and monitoring their interactive behaviors.
The vacuolar structures were found to be highly complex
and dynamic. There were tubular protrusions around the
inner surface of the cells and the elongating cell tips. Unexpectedly, we found that microtubules, rather than actin
microfilaments, were needed to maintain these vacuolar
structures, and that the microtubules and vacuolar membranes were co-localized. Furthermore, we observed that the
856
elongating microtubule could tug the vacuolar membranes,
and that the interaction between microtubules and vacuolar
membranes could regulate vacuolar distribution. Thus, this
study demonstrates a novel function for microtubules in
land plants.
Results
Visualization of vacuolar membranes in living
chloronema cells
Although several fluorescent dyes are available for observing
vacuoles, they are generally unsuitable for detailed analysis
of vacuolar structures because of fluorescence fading upon
multi-image acquisition. We thus employed a fusion
construct of GFP and an Arabidopsis tonoplast t-SNARE,
encoded by AtVAM3/SYP22, that is an excellent tonoplast
marker and has been used successfully to visualize vacuolar
membranes in flowering plant cells (Uemura et al. 2002).
Physcomitrella patens transformants with constitutive
expression of GFP at an appropriate level for observation
were selected (Fig. 1A, B). The morphology and growth of
the transformants were not distinguishable from those of
the wild type (see Supplementary Fig. S1).
To examine the localization of the GFP–AtVAM3 proteins, chloronema cells, the primary cell type in protonema
filaments, were stained with N-(3-triethylammoniumpropyl)4-(6-(4-(diethylamino)phenyl)hexatrienyl pyridinium dibromide (FM4-64). Although this dye is known as a useful
endosome marker, FM4-64 is first incorporated into the
plasma membrane and is then transferred to the vacuolar
membrane via endosomes (Bolte et al. 2004). Soon after
chloronema cells were stained with FM4-64, the dye became
localized to the plasma membrane and endosomes. Although
the GFP signals also closely localized to the plasma membrane, they did not co-localize with the FM4-64 signals
at the plasma membrane (Fig. 1C). At 3 h after staining, the
FM4-64 dye moved to the vacuolar membrane and became
localized to the endosomes and also the vacuolar membrane
where it almost overlapped with the GFP signals (Fig. 1D).
We thus concluded that the GFP–AtVAM3 fusion proteins
localized only to the vacuolar membrane, and that these
transgenic moss plants were suitable for investigations into
vacuolar structures.
Three-dimensional structures of vacuoles
in chloronema cells
We next examined the vacuolar structures of chloronema
apical cells with tip growth (Menand et al. 2007) and cell
division to produce a new apical cell and a subapical cell. In
order to investigate the three-dimensional (3-D) structure
of the vacuole, we took Z-series images by confocal laser
scanning microscopy (CLSM) and established the maximum
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
(Fig. 2I–M). The sheet-like membranes in the inner side of
the vacuole and the fine tubular structures on the outer side
of the vacuole were rarely observed (Fig. 2N, O). Instead,
numerous indentations of the vacuole made by the chloroplasts were apparent (Fig. 2K, I). When a subapical cell began
to grow by branching, complex vacuoles with tubules also
began to develop (Supplementary Fig. S2). Thus, these
complex vacuolar structures appear to be a feature of cells
with apical growth.
B
Fluorescent Bright field
A
D
C
3h
Dynamics of vacuolar structures in the
chloronema cell cortex
Merged
FM4-64
GFP
0h
Fig. 1 Visualization of vacuolar membranes by stable expression of
GFP–AtVAM3 fusion protein. (A and B) Bright field and fluorescent
images of a chloronema apical cell (A) and a non-apical cell (B) of a
transgenic line constitutively expressing GFP–AtVAM3 fusion proteins.
Green and red signals represent GFP and autofluorescence of chloroplasts, respectively. (C) Confocal laser scanning microscopy (CLSM) of
a chloronema cell. The plasma membrane is labeled with FM4-64. The
GFP signals do not co-localize with FM4-64 signals.
(D) The vacuolar membrane is labeled with FM4-64. GFP signals
almost overlap with FM4-64 signals except for in the endosomes.
Bars = 20 µm.
intensity projections (Fig. 2A, B). Interestingly, we observed
an inner vacuolar membrane, which penetrated the cellapex side of the large vacuole (Fig. 2A, arrow), and also fine
structures on the outer side of the large vacuole (Fig. 2B,
arrowhead). Cross-sections of these structures indicated
that these were sheet-like (Fig. 2C) and tubular (Fig. 2D)
components of the vacuoles, respectively. A sequential view
of the Z-series images showed the tubular structure to
be derived from a part of the large vacuole (Fig. 2E). To
understand better the overall structure of the vacuole, we
performed surface modeling of the vacuolar membrane
(Fig. 2F–H). The tubular vacuole (Fig. 2F–H, arrowheads)
extended from a region of the central vacuole to the cytoplasm in between chloroplasts. The sheet-like membrane,
which denotes a cytoplasmic strand, developed from a site
near the nucleus and spread towards the apex (Fig. 2H, red
broken line and red arrow).
The vacuolar structures of a subapical cell, which shows
limited growth and no further division except for side
branching, were examined by the same process. In contrast
to apical cells, the subapical cell had a much simpler vacuole
To obtain information about how such tubular vacuolar
membranes developed in the chloronema cell cortex, timesequential observations of the cortex of chloronema cells
were performed (Fig. 3). The vacuolar membrane underwent dynamic and repeated wave movements, with the
tubular vacuoles sometimes splitting and fusing with other
parts of the vacuolar membrane (Fig. 3A, B; see Supplementary Video 1). Although we cannot exclude the possibility
that these phenomena were the result of the vacuole being
out of focus, we consider that they were in fact the result of
vacuolar membrane rearrangements, since in many cases
the chloroplasts placed spatial limits on the vacuolar movements. These observations revealed the amazing flexibility of
the vacuolar structures despite the vacuolar membrane
encompassing a single large vacuole.
To examine the relationship between vacuolar membrane
structures and chloroplasts, we induced chloroplast movement by a microscopic light system (Sato et al. 2001). When
chloroplasts were dispersed around the cell by blue light
irradiation, no tubular structures were observed (Fig. 3C,
0–36 min). However, once the chloroplasts gathered at the
center of the cell following weak blue light irradiation, many
tubular vacuoles were induced between the chloroplasts
(Fig. 3C, 48–60 min, see Supplementary Video 2). During
chloroplast movement, tubular vacuoles also developed on
the chloroplasts (Fig. 3D). These results suggest that the
vacuolar membrane would rarely interfere with organelle
movement due to its flexibility, despite the vacuole filling a
large part of the cell.
Effect of cytoskeletal inhibitors on vacuolar
structures in chloronema cells
To examine the relationship between these observed vacuolar membrane structures and the cytoskeleton, the strong
actin depolymerization agent, bistheonellide A (Saito et al.
1998), was applied to the chloronema cells. Bistheonellide
A had little effect on vacuolar morphology (Fig. 4A, B) while
it almost completely depolymerized actin microfilaments
(Supplementary Fig. S4). In contrast, application of the
microtubule depolymerization herbicide, oryzalin, clearly
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
857
N
O
Number of tubules
2.0
1.5
1.0
0.5
0
Apical Subapical
Number of inner sheets
Y. Oda et al.
1.5
1.0
0.5
0
Apical Subapical
Fig. 2 Vacuolar 3-D structures of a chloronema apical cell and a chloronema subapical cell. (A and I) Mid-planes of apical (A) and subapical (I)
cells observed by CLSM. The arrow in (A) shows a cytoplasmic strand running from the nucleus towards the cell tip. (B and J) Maximum intensity
projections of the vacuolar membrane and chloroplasts. The arrowhead in (B) shows a tubular vacuolar structure at the cell cortex. (C) A crosssectional image along the line (p–q) in (B). The arrow shows the sheet-like structure of the vacuolar membrane. (D) A cross-sectional image along
the line (r–s) in (B). The arrowhead shows the outer fine tubule of the vacuole. (E) Z-series images of the blue box in (B). The numbers represent the
depth of the section. The arrowheads show the tubular vacuolar structure. (F–H) Computationally reconstructed 3-D structure of the vacuole and
chloroplasts. (F and G) Top view of the vacuole (green) and chloroplasts (red). Only the vacuole is shown in (G). (H) The back view of the vacuole.
The inner surface of the vacuolar membrane is shown in gray. The arrowheads show the protruding tubular vacuolar membrane. The arrow in (H)
shows the cytoplasmic strand shown in (A). The red dashed line indicates the sheet-like structure of the vacuolar membrane extending from the
nucleus to the cell tip. (K–M) The 3-D structures of the chloronema subapical cell. Compared with the apical cell, the vacuole has a simple structure
without inner sheets and outer tubules. (N) Number of vacuolar tubules in apical and subapical cells. Data represent the mean ± SE (n = 20, P <0.01,
t-test). (O) Number of inner sheets of the vacuolar membrane. Data represent the mean ± SE (n = 20, P <0.01, t-test). Bars = 20 µm.
858
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
A
C
B
D
Fig. 3 Dynamics of tubular structures of vacuoles in the cortex of chloronema cells. (A) Time-sequential observations of the tubular structure of
a vacuole (green signals) between chloroplasts (red signals). The vacuolar tube divides (arrows) and then fuses with a tubule that has extended
from another region of the large vacuole (arrowheads). Bar = 5 µm. (B) A fine tube of the vacuole (arrowhead) projects from the vacuole and fuses
with another region of the vacuolar membrane. Bar = 5 µm. (C) Observation of vacuolar dynamics during chloroplast movement. Arrows show
some of the chloroplasts observed in a chloronema cell. During irradiation of the center of the cell with blue light (100 Wm–2), the chloroplasts
move to both edges of the cell (12–36 min). Note that the vacuolar structure is smooth at the center of the cell. Subsequent irradiation of the
center of the cell with a weak blue light (1 Wm–2, 48–60 min) causes movement of the chloroplasts toward the center of the cell (54–60 min). The
arrowheads show the tubular vacuolar structures that appear between the densely gathered chloroplasts. Bar = 20 µm. (D) During movement of
a gourd-shaped chloroplast (arrows), a tubular vacuolar structure appears along the groove of the chloroplast (arrowheads). Bar = 4 µm.
affected the vacuolar structures (Fig. 4C). To investigate the
effects of microtubule depolymerization on these vacuolar
structures more precisely, we performed time-sequential
observations of oryzalin-treated cells (Fig. 4D). The tubular
vacuole around the cell cortex somehow expanded and
became spherical although the continuity of the membrane
was retained (Fig. 4E). At the cell tip, the straight development of the vacuolar membrane became disorganized and
there was irregular accumulation of cytoplasm in this region
(Fig. 4F). However, the possibility still remained that
repositioning of numerous chloroplasts caused this vacuolar
deformation. To observe chloroplast-independent effects,
cells were treated with ampicillin for 2 d, which inhibited
chloroplast division and resulted in chloronema cells that
had only a few large chloroplasts (Kasten and Resk, 1997).
In these cells, disorganization of the vacuolar structures at
the cell tip was clearly observed after addition of oryzalin
(Fig. 4G, H), and small vacuolar fragments remained at the
cell tip (Fig. 4F, H, arrowheads). Since removal of the inhibitor allowed repair of the vacuolar structures, these effects
were not the result of irreversible cell damage but rather the
result of microtubule depolymerization.
To confirm the specific effect of microtubule depolymerization, we treated chloronema cells with oryzalin, bistheonellide A and latrunculin B, and observed the vacuolar structures
in the cell tip. Oryzalin treatment induced spherically
deformed vacuoles and cytoplasmic accumulation, whereas
actin depolymerization agents did not affect vacuolar structures or the distribution of the cytoplasm (Fig. 4I, J).
Vacuolar dynamics and effects of a microtubule
inhibitor in rhizoid cells
The moss possesses another filamentous cell type, called the
rhizoid, which grows from a stem of a gametophore, a leafy
shoot. To examine the vacuolar structures and their dependency on the cytoskeletons, we performed time-sequential
observations of the vacuolar membrane in a rhizoid cell.
Here, the vacuole showed highly dynamic and rapid movements (Fig. 5A), and several vacuolar protrusions towards
the cell apex repeatedly contracted and extended. At times
the vacuolar membrane appeared to reach the plasma membrane (Fig. 5A, arrowheads; see Supplementary Video 3),
indicative of possible interactions between the vacuolar
membrane and plasma membrane. Inhibition of microtubule depolymerization by oryzalin caused the dramatic
disappearance of these structures and stopped these
movements (Fig. 5B; see Supplementary Video 4). Thus the
vacuolar structures and their dynamics are also dependent
on microtubules. To analyze the vacuolar dynamics precisely,
the distance between the vacuolar tip, which is indicated
by arrowheads in Fig. 5A, and the rhizoid tip was plotted
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
859
Y. Oda et al.
B
C
E
D
F
G
J
I
Control
Oryzalin
BA
Lat B
Cytoplasmic accumulation (µm)
A
H
2.5
2.0
1.5
1.0
0.5
0
Control Oryzalin
BA
Lat B
Fig. 4 Effects of cytoskeletal inhibitors on chloronema apical cells. (A–C) Chloronema cells treated with DMSO (A), bistheonellide A (B) or
oryzalin (C) for 2 h. Aberrant vacuolar membranes (arrows) were induced only in the oryzalin-treated cells (C). (D–F) Chloronema cells transiently treated with oryzalin. The vacuole shows tubular structures at the cell cortex [yellow box, magnified in (E), arrowheads] and strands [blue
box, magnified in (F), which is merged with the bright field image]. At 1 h after oryzalin application (10 µM), the tubular structures expanded
(E, 60 min arrowheads) and the strands disappeared. Note the accumulation of cytoplasm at the cell tip (60 min, arrows). After removal of oryzalin, both the tubular and strand structures reappeared (150 min). (G and H) Effect of oryzalin (10 µM) on an ampicillin-treated chloronema cell.
(H) Magnified image of the magenta box in (G). The image was merged with the bright field image. Vacuolar strands disappeared and cytoplasm
accumulated abnormally (arrows). The arrowhead shows a small vacuole remaining at the cell tip. After removal of oryzalin, the strand structures
recovered (240 min). (I) Vacuolar structures of the chloronema apex after treatment with oryzalin, bistheonellide A (BA) and latrunculin B
(Lat B). Note that the vacuolar structure is deformed and cytoplasmic accumulation appeared in the cell treated with oryzalin. The yellow dotted
line indicates the positions of cell tips, and the double-headed arrow indicates the depth of cytoplasmic accumulation. (J) Depth of cytoplasmic
accumulations after inhibitor treatment. Values are the mean ± SE (n = 20). Oryzalin treatment induced cytoplasmic accumulation (P <0.01,
t-test). Bars = 20 µm.
(Fig. 5C). During the plotting period (100 s), the cellular tip
scarcely grew and stayed in the same position. The vacuolar
tip exhibited oscillatory motion, while such a dynamic
motion of the vacuole was not observed in oryzalin-treated
cell (Fig. 5C). This strongly suggests microtubule-dependent
vacuolar dynamics. The periods in which the vacuolar tip
contacted the plasma membrane were overlaid with
magenta color. To analyze the vacuole movement further,
the rate of vacuolar movement was summarized in Table 1.
We also observed microtubule dynamics using a transgenic
line with GFP–tubulin (GTU193, Hiwatashi et al. 2008) to
compare the vacuolar movement and microtubule dynamics. Interestingly, the contraction rate just after contact with
860
the plasma membrane (mean ± SD = 699 ± 271 nm s–1) was
more rapid than normal contraction (261 ± 165 nm s–1) and
microtubule shrinkage (324 ± 123 nm s–1). These data suggest interactions between the vacuolar membrane and
plasma membrane.
Localization and dynamics of microtubules and their
relationship with the vacuole
The above pharmacological experiments suggested that the
microtubule was a major cytoskeleton regulating vacuolar
structures and dynamics in protonema and rhizoid cells, and
we furthermore attempted to visualize microtubules and the
vacuolar membrane simultaneously. In the transformants
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
A
C
Distance of vacuole from
rhizoid tip (µm)
10
8
6
4
2
0
0
D
1.0
Rate of vacuole movement
(µm/sec)
B
Oryzalin
Control
20
40
60
Time (sec)
80
100
0.5
0
0.5
1.0
2.0
Fig. 5 Time-sequential observations of vacuoles in rhizoid cell tips. (A) A highly dynamic vacuolar structure observed in the tip of a rhizoid apical
cell. The tubular vacuolar structure repeatedly extends towards the cell apex and appears to touch the plasma membrane (arrowheads). See
Supplementary Video 3 for a high-speed video. (B) Effect of oryzalin on vacuolar dynamics. Time-sequential observations of a rhizoid cell after
application of oryzalin (10 µM, 0 min). The tubular protrusion of the vacuole (arrows) retreats from the cell tip (5 and 7 min) and disappears
(10 min). Vacuolar dynamics are also lost (see Supplementary Video 4). Bars = 5 µm. (C) Time course of the distance between the vacuole and
rhizoid tip in control and oryzalin-treated cells. Magenta color indicates the contact between the vacuolar tip and plasma membrane. (D) The
rate of movements of the vacuolar tip of the control cell shown in C. Asterisks indicate shrinking of the vacuolar tip just after the contact. Data
shown in C and D are individual traces for representative cells.
Table 1 Rate of vacuole movement and microtubule dynamics
Event
Rate (nm s–1)
No. of observations
Vacuole extension
266 ± 164
164 events
Vacuole contraction
311 ± 228
156 events
Vacuole contraction after
PM contact
699 ± 271
18 events
Vacuole contraction in
cytoplasm
261 ± 165
138 events
Microtubule growth rate
86 ± 16
8 microtubules
Microtubule shrinkage rate
324 ± 123
12 microtubules
Values are means ± SD. Vacuolar events are collected from three different rhizoid
cells including the cell shown in Fig. 5. Note that the rate of vacuole contraction
after PM (plasma membrane) contact is higher than the rate of contraction in
cytoplasm (P <0.01, t-test). Microtubule dynamics were observed in cortex of
three different chloronema cells.
with both GFP–AtVAM3 and mRFP–PpTUA1 genes, fibrous
structures of the microtubules were found to lie along the
periphery of the plasma membrane (Fig. 6A). The tubular
vacuolar structures were closely associated with some of
these microtubules (Fig. 6A). Elongating microtubules occasionally collided with the tubular vacuole, and caused movement and stretching of the vacuole (Fig. 6B; see also
Supplementary Video 5). We next examined the motions
of the vacuolar membrane and microtubules at the tip of a
chloronema apical cell. Here, we observed several longitudinally arranged microtubules, some of which were associated
with vacuolar strands and small protrusions of the vacuole
towards the cell apex (Fig. 7A). Furthermore, by time-sequence observations, we found that the elongating microtubule penetrated the vacuole and created the cytoplasmic
strand (Fig. 7B, arrowheads), and that the vacuolar membrane protruded towards the cell apex as if it was being
tugged by the elongating microtubule (Fig. 7B, arrows; see
Supplementary Video 6). Fig. 7C shows the intensity of
monomeric red fluorescent protein (mRFP)–tubulin and
GFP–AtVAM3 along the yellow broken lines in Fig. 7B.
Co-localization of cytoplasmic strands and microtubules
(asterisks), and the concomitant appearance of vacuolar
membranes and microtubules (arrows), could be confirmed.
Microtubules that penetrate the vacuole and associate with
protruding vacuolar membrane were also observed in a
GTU193 transgenic line (Fig. 7D, E). These observations suggest that the vacuolar structures, and their distribution in
the cell apex, are accompanied by elongating microtubules
that interact with and drag the vacuolar membrane. Furthermore, to confirm the interaction between the vacuolar
membrane and microtubules, we fixed chloronema cells by
high-pressure freezing and performed transmission electron
microscopy. Fig. 8 shows the ultrastructures around
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
861
Y. Oda et al.
A
mRFP-tubulin
GFP-AtVAM3
Merged
B
Fig. 6 Dual observations of vacuolar membranes and microtubules in the chloronema cell cortex. (A) Several microtubules (mRFP–tubulin,
arrowheads) and tubular vacuolar structures (GFP–AtVAM3, arrowheads) are observed at the chloronema cell cortex. Note that the tubular
vacuoles are co-aligned with microtubules (merged, arrowheads). The arrowheads in each panel show the same locations. (B) A time-sequential
observation of tubular vacuolar structures and microtubules. The arrows show the putative plus end of a microtubule. As the microtubule elongates, the tubular vacuolar structure is stretched as it follows the end of the microtubule. Bars = 5 µm.
D
Merged
GFP-tubulin
FM4-64
Merged
40 sec
GFP-AtVAM3
mRFP-tubulin
* *
80 sec
**
FM4-64
E
C
GFP-tubulin
GFP-AtVAM3
Intensity (a. u.) Intensity (a. u.)
B
mRFP-tubulin
Merged
A
Position
Fig. 7 Dual observations of vacuolar membranes and microtubules at the mid-planes of chloronema apexes. (A) Several longitudinal microtubules are observed (mRFP–tubulin). The vacuole shows strand structures (GFP–AtVAM3, arrowheads) directed towards the cell apex and small
projections (GFP–AtVAM3, arrows). Note that the vacuolar strands (merged, arrowheads) and the projection from the vacuolar membranes
(merged, arrows) are associated with microtubules. (B) Time-sequential observations of a chloronema apex. Elongation of a microtubule towards
the apex (mRFP–tubulin, arrowheads) and concomitant vacuolar strand movement (GFP–AtVAM3, and merged, arrowheads). Asterisks indicate
the cytoplasmic strands which appeared. Arrows show a projection of the vacuolar membrane (GFP–AtVAM3). Note that the vacuolar projection follows the elongating microtubules (mRFP–tubulin, arrows). (C) Intensities (a.u. = arbitrary units) along the yellow lines in B (40 and 80 s).
The green line indicates GFP signals and the red line indicates mRFP signals. Asterisks indicate cytoplasmic strands. Arrows indicate mRFP–tubulin signals accompanying the vacuolar membrane. (D) Dual observations of microtubules and the vacuolar membrane by GFP–tubulin and
FM4-64 staining. Microtubules are developed through the transvacuolar strand (arrowheads) and co-localized with the vacuolar membrane
at the tip of the large vacuole (arrows). (E) Time-lapse observation of microtubules (GFP–tubulin) and the vacuolar membrane labeled with
FM4-64. Protrusion of the vacuole towards the cell tip is maintained with sequential microtubule association (arrowheads). Bars = 5 µm.
862
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
Fig. 8 Ultrastructures of the vacuolar membrane and microtubules.
(A) Chloronema cells of the wild-type strain were cryofixed and
observed under a transmission electron microscope. A part of the
large vacuole and cytoplasm are shown. (B and C) The blue frame (B)
and yellow frame (C) of A are magnified. The microtubules and vacuolar membranes are localized close together. Green arrows indicate
vacuolar membranes. Magenta arrowheads indicate microtubules.
Bars = 100 nm.
the vacuolar membrane in the chloronema cell cortex
(Fig. 8A–C). Microtubules were observed in the region very
close to the vacuole. These observations suggested the
possibility of interaction between microtubules and vacuolar membrane and supported the microtubule-dependent
regulatory system of vacuolar organization in P. patens.
Discussion
Visualization and analysis of vacuolar structures
and dynamics in Physcomitrella patens
There has been significant recent progress in the visualization and structural analysis of every organelle in living higher
plant cells. Such progress has been due, in large part, to the
ease of Agrobacterium-mediated transformation of Arabidopsis and other familiar plants or their cultured cells with
marker genes. Numerous studies have reported on the visualization of microtubules, important for primary (Paradez et al.
2006) and secondary (Oda et al. 2005, Oda and Hasezawa
2006) cell wall synthesis and cell cycle progression (Hasezawa
et al. 1999, Kumagai et al. 2001), as well as on the visualization
of the vacuolar membrane (Kutsuna and Hasezawa 2002,
Saito et al. 2002, Uemura et al. 2002, Kutsuna et al. 2003,
Reisen et al. 2005). Our current study, however, precisely
analyzes and demonstrates the close relationship between
the vacuole and microtubules.
In this study, we succeeded in establishing several transformants expressing GFP–AtVAM3 and mRFP–tubulin. As
the growth of these transformants was not distinguishable
from that of the wild type, the expression of these marker
genes probably had little or no detrimental effects on the
cells (Supplementary Fig. S1A, B). In addition, the cell phenotypes of each transgenic line were indistinguishable from
those of the wild-type cells (Supplementary Fig. S1C), and
the characteristic vacuolar structures observed in these
transformants were also recognized in wild-type cells stained
with FM4-64 or BCECF-AM (Fig. 7D, E; Supplementary Fig. 3).
We therefore conclude that the observed structures are
indeed authentic.
An interesting feature of the vacuole found in this study,
by precise observations of sequential optical sections and
examination of the 3-D structures, was a tubular structure of
the vacuole at the chloronema cell cortex (Fig. 2). These
vacuolar tubules actively underwent wave motions and rearrangements by separation and fusion with other vacuolar
regions between chloroplasts (Fig. 3A, B). Such dynamics
and the flexibility of the vacuolar membrane would facilitate
the smooth motility of organelles in the cell, perhaps through
cooperation between chloroplasts and the vacuolar membrane via the cytoskeleton. In fact, the tubular structures of
the vacuole were frequently observed when chloroplast
movement was induced (Fig. 3C, D).
We also found that rhizoid cells of P. patens have a
dynamic vacuole with fine tubular structures that repeatedly extend towards the cell apex (Fig. 5A). Dynamic tubular
vacuoles have also been observed in root hair cells and pollen
tubules of higher plants (Hepler et al. 2001, Hicks et al. 2004,
Ovecka et al. 2004), suggesting that these vacuolar structures may be a feature of rapidly tip-growing cells. Interestingly, our high-speed time-sequential observations revealed
that the vacuole extended to, and even touched, the plasma
membrane of the rhizoid cell apex (Fig. 5A; Supplementary
Video 4), suggesting some interaction between the plasma
membrane and vacuole. The rapid motions of the vacuolar
tip observed just after this touching might be caused by tension generated by a direct or indirect physical interaction
between the vacuolar membrane and plasma membrane.
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
863
Y. Oda et al.
In chloronema cells, the vacuole developed along the apical
region and also created small protrusions that made contact
with the plasma membrane (Fig. 7B). The remaining small
vacuoles at the cell tip, when vacuolar deformation was
induced by a microtubule inhibitor, also support this possibility (Fig. 4E, G). Such interactions may facilitate membrane
traffic between vacuoles and the plasma membranes at the
dense endosomal region, and/or play a role in anchoring the
vacuole for inheritance by the next apical daughter cell.
Regulatory mechanisms of vacuolar structures by
microtubules
In this study, we have demonstrated the characteristic structures and dynamics of vacuoles in P. patens. Depolymerization of microtubules dramatically affected these structures
as well as the dynamics of the vacuoles (Figs. 4, 5), indicating
that these features are dependent on microtubules. To clarify these observations, we performed dual observations of
the vacuolar membrane and microtubules by expressing
mRFP–tubulin in transformants in which the vacuolar
membrane had been visualized, or by labeling vacuolar membranes with FM4-64 in the transformant strain that expresses
GFP–tubulin. We were therefore able to demonstrate that
the tubular vacuoles around the cortex, the protruding vacuoles at the cell tip and the invaginations of cytoplasmic strands
were closely associated with microtubules (Figs. 6A, 7A).
Transmission electron microscopy confirmed their close
association (Fig. 8). Although we could not observe ultrastructures of the apical region due to technical difficulties,
such associations might mediate vacuolar motion along
microtubules at the cell apex. These observations strongly
suggest that the vacuolar structures are supported by
microtubules.
These results were rather unexpected, since several
reports previously suggested that vacuolar structures and
movements in higher plant cells are dependent on actin
microfilaments. When actin microfilaments were destroyed,
the vacuoles deformed, fragmented and lost their dynamics
in tobacco BY-2 cells and root hair cells (Kutsuna et al. 2003,
Ovecka et al. 2005). Furthermore, actin microfilaments were
found to be closely localized with the vacuolar membrane,
and an actomyosin-dependent regulation of vacuolar
dynamics was suggested in tobacco BY-2 cells (Higaki et al.
2006). In this context, the regulation of vacuolar structures
and dynamics in P. patens is more similar to that of fungal
hyphal tips, that have a motile tubular vacuolar system regulated by microtubules (Ashford 1998), than to that of flowering plants. This difference between the moss and flowering
plants could not be explained by growth mode such as tip
growth and diffuse growth, because the vacuolar structures
and dynamics are also dependent on actin microfilaments in
root hairs, which exhibit tip growth (Ovecka et al. 2005).
Lovy-Wheeler et al. (2007) reported that movements of
864
organelles including the vacuole also depended on actin
microfilaments in lily pollen tubes. These suggest the possibility of a divergence in the regulatory system of vacuolar
structures by cytoskeletons during land plant evolution.
Further investigations over a wide range of species and cell
types will be needed to confirm this possibility.
A question that remains is how the vacuolar structures are
regulated by microtubules in P. patens. Our time-sequential
observations suggested that the vacuolar membrane
protruded as if being tugged by elongating microtubules
(Figs. 6B, 7B). These observations indicate that the vacuolar
membrane interacts tightly with the polymerizing microtubule. In animal cells, ER tubules move towards the microtubule plus or minus ends through the activities of kinesins
and dyneins. Concomitant movement of the ER tubule with
elongating microtubule plus ends also suggested that polymerization of microtubules functions as a force for ER tubule
dynamics (Terasaki and Reese 1994, Feiguain et al. 1994, Steffen
et al. 1997, Waterman-Storer and Salmon, 1998). Similar
mechanisms using microtubule motors may regulate vacuolar
structures and dynamics in P. patens.
Our analysis of the vacuolar dynamics at the rhizoid
cell tip revealed rapid extension (mean ± SD = 266 ± 164
nm s–1) and contraction (261 ± 165 nm s–1 in cytoplasm,
699 ± 271 nm s–1 after contact with the plasma membrane).
They are more rapid than the previously reported microtubule growth rate (60–80 nm s–1) and shrinkage rate
(80–300 nm s–1) in higher plant cells (Chan et al. 2003,
Dhonukshe and Gadella 2003, Shaw et al. 2003, Vos et al.
2004). We also found that the microtubule growth rate
(86 ± 16 nm s–1) and shrinkage rate (335 ± 123 nm s–) in chloronema cells were both slower than vacuolar dynamics in
the rhizoid cell (Table 1). Thus tugging by microtubules is
not a sufficient explanation for the vacuolar dynamics. Perhaps the vacuole could be rapidly extended along pre-existing microtubules as if it was being zippered by some
microtubule-associated linkers or as if being slid by motor
proteins. Rapid shrinkage could be caused by accumulated
tension of the vacuolar membrane or by the sudden disappearance of microtubules after collision with the plasma
membrane, which might induce depolymerization of microtubules. In addition to microtubule dynamics, the balance
between binding to microtubules and vacuolar tension
might be involved in maintaining the vacuolar dynamics.
Actin microfilaments are unlikely to be involved in this
event, because disruption of actin microfilaments by inhibitors did not cause vacuolar retraction in protonema cells
(Fig. 4) and the actin-dependent organelle movement is
even faster (>3 µm s–1 at maximum, Nebenführ et al. 1999)
than the vacuolar movement we observed here.
The significance of the regulation of vacuolar distribution
and dynamics by microtubules is still unclear. Although
microtubules are known to function in the maintenance of
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
polarity and normal growth in moss protonema cells
(Doonan et al. 1988), the precise mechanisms mediating
such regulation are still unknown. Doonan et al (1988)
showed that long-term depolymerization of microtubules
induced swelling of protonema tips and suggested that
microtubules are essential to impose tubular shape and
directionality upon expansion. In this study, we demonstrated that microtubule depolymerization induces vacuolar
deformation, and, simultaneously, induced irregular accumulation of cytoplasm at the cell tip (Fig. 4). Such mislocalization of the vacuole and cytoplasm might prevent
homeostasis and appropriate osmotic pressure from being
maintained at the cell tip. Proper cytoplasmic distribution
might be necessary to maintain pointed growth of protonema cells. The novel microtubule-dependent regulation
of vacuolar structures and dynamics, which we have demonstrated here, could therefore be one aspect of normal tip
growth regulation by microtubules.
contains an E7113 promoter (Mitsuhara et al. 1996, kindly
provided by Dr. Y. Ohashi, NIAS) for constitutive expression,
and non-coding P. patens genomic DNA fragments BS213
(Schaefer and Zrÿd 1997; kindly provided by Dr. D. G. Schaefer,
INRA Versailles and Dr. J.-P. Zrÿd, Lausanne University) for
targeting.
To observe vacuolar membranes in caulonema and
rhizoid cells, in which the rice actin promoter sometimes
weakly induced GFP–AtVAM3, the GFP–AtVAM3 fusion
construct was PCR-amplified with appropriate primers
(5′-ctcgagatggtgagcaagggcgag-3′ and 5′-gggccctcaagctgcgagt
actat-3′) and inserted into the pCMAK1 vector.
Staining of vacuolar membranes with FM4-64 dye
To label vacuolar membranes, protonema cells were treated
with 36 µM FM4-64 (Invitrogen) for 30 min. Subsequently,
the cells were washed three times with fresh liquid medium
and incubated for 3 h with gentle agitation.
Microscopy
Materials and Methods
Plant culture
Physcomitrella patens Gransden2004 was cultured on BCDAT
agar plates (Nishiyama et al. 2000) at 23°C under a 16 h
light/8 h dark photoperiod (50 µmol photons m–2 s–1). To
observe chloronema cells, a colony was homogenized with a
Polytron (Kinematica, Littau, Switzerland), suspended in
40 ml of BCDAT liquid medium containing 0.1% glucose, and
agitated on a rotary shaker at 140 r.p.m. under the same lighting conditions as the agar culture. Every week, this culture
was homogenized and diluted 8-fold in 40 ml of fresh liquid
medium. Chloronema cells grew well in 3–6 d, and were used
for cell staining, observations and inhibitor treatments.
Establishment of stable transformants expressing
GFP–AtVAM3 and mRFP–tubulin
To visualize vacuolar membranes in chloronema cells, a GFP
(S65T)–AtVAM3 fragment (kindly supplied by Dr. M. H. Sato,
Kyoto Prefectural University) was PCR-amplified with appropriate primers (5′-ggcgcgccatggtgagcaagggcgag-3′ and 5′gggccctcaagctgcgagtactat-3′) and placed under the control
of the rice actin promoter (McElroy et al. 1990) in vector
pTFH15.3 (http://moss.nibb.ac.jp/). This vector contains
Pphb7 genomic fragments (Sakakibara et al. 2003), and thus
allows the GFP–AtVAM3 fragment to be inserted into the
Pphb7 locus. Polyethylene glycol (PEG)-mediated transformation was performed as described by Nishiyama et al.
(2000), and transformants were selected on BCDAT medium
supplemented with 20 mg l–1 G418 (Invitrogen Corporation,
Carlsbad, CA, USA). For dual observations of the vacuolar
membrane and microtubules, the fusion gene of mRFP
(Campbell et al. 2002) and PpTUA1, a P. patens α-tubulin, in
vector pCMAK1 was used (Hiwatashi et al. 2008). This vector
Aliquots of the protonema cell culture were transferred
onto 35 mm Petri dishes with a 27 mm coverslip window at
the bottom (Matsunami Glass Industries, Osaka, Japan).
For simultaneous observations of GFP, FM4-64 and chloroplasts, the dish was set on an inverted microscope (IX, Olympus, Tokyo, Japan) equipped with a confocal scanning unit
GB200 (Olympus) and a PlanApo × 60 oil immersion lens
(Olympus). GFP, FM4-64 and chlorophyll were excited by a
488 nm argon laser. When FM4-64 signals were detected, a
band-pass absorption filter was used to eliminate autofluorescence of chloroplasts. The maximum intensity projection
of the scanned Z-series image stacks was created with ImageJ
software (http://rsb.info.nih.gov/ij/). For surface modeling,
the vacuolar membranes and the border of chloroplasts on
each confocal image were manually tracked on Photoshop
(Adobe Systems, San Jose, CA, USA) and reconstructed on
Amira 2.3 (Indeed-Visual Concepts GmbH, Berlin, Germany).
To construct the 3-D structure of the vacuole, the tracked
border of the vacuole was recognized with the Label/
Voxel tool on Amira. The 3-D models were constructed from
the border information with default settings. Finally, the 3-D
models were simplified by reducing the number vertices.
For dual observations of GFP and mRFP, the dish was set
on an inverted microscope (IX, Olympus) equipped with a
cooled CCD camera head system (Cool-SNAP HQ, PhotoMetrics, Huntington Beach, Canada). When mRFP signals
were detected, a 604–644 nm band-pass filter (Semrock,
Rochester, NY, USA) was used to eliminate autofluorescence
of chloroplasts. Acquired images were sharpened in Image J
with the Fourier transform band-pass filter to remove high
and low spatial frequency signals.
To observe GFP–tubulin and FM4-64 simultaneously,
GFP and FM4-64 were excited by a 488 nm argon laser, and
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
865
Y. Oda et al.
detected through a confocal unit (CSU10, Yokogawa Electric
Corporation, Tokyo, Japan) with a 524–546 nm band-pass
filter (Semrock) for GFP, and a 575–625 nm band-pass filter
(Olympus) for FM4-64
Inhibitor treatments
For actin depolymerization, protonema cells were treated
with 1 µM bistheonellide A (Wako Pure Chemical Industries,
Osaka, Japan), and cells were observed after a 2 h incubation.
For microtubule depolymerization, cells were treated with
10 µM oryzalin (Wako), and observed after a 2 h culture
period. As control, a 1 ml suspension culture was incubated
with 0.1% (v/v) dimethylsulfoxide (DMSO). After inhibitor
addition, the protonema cells were agitated on a rotary
shaker at 140 r.p.m. under light. For transient and real-time
inhibition of microtubules, cells were immobilized by the
agar–gelatin method (Wada et al. 1987). Cells were transferred onto 35 mm Petri dishes with a 27 mm coverslip
window at the bottom (Matsunami Glass Industries). BCDAT
medium containing 0.8% (v/w) agar and 0.05% (v/w) gelatin
(Wako) was warmed to 45°C and a thin medium layer, created in a stainless wire ring, was immediately dropped onto
the cells. After 3 min, liquid BCDAT medium was transferred
into the dish and pre-incubated for >1 h. The cells could be
observed directly, and inhibitors were added after the beginning of observations. For removal of inhibitors, the medium
was carefully exchanged more than four times with fresh
liquid medium.
High-pressure freezing and transmission electron
microscopy
Protonema cells were frozen in a high-pressure freezing
machine (HPM010; Bal-Tec, Balzers, Liechtenstein) that had
been cooled with liquid nitrogen (–196°C). The samples were
transferred to anhydrous acetone containing 2% OsO4 at
–80°C in a Leica EM AFS automatic freeze-substitution apparatus (Leica Mikrosysteme GmbH, Wien, Austria), held at
–80°C for 78 h, warmed gradually to 0°C over 11.4 h, held at
0°C for 1.5 h, warmed gradually to 23°C over 3.9 h, and at 23°C
for 2 h. Then, the samples were washed three times with dry
acetone at room temperature and infiltrated with increasing
concentrations of Spurr's resin (Spurr 1969) in dry acetone,
and finally with 100% Spurr's resin. Ultrathin sections (0.06–
0.07 µm) were cut with a diamond knife on an ultramicrotome (Leica Ultracut UCT, Leica Mikrosystems) and mounted
on formvar-coated copper grids. The sections were stained
with 3% uranyl acetate for 2 h at room temperature, and with
lead citrate for 10 min at room temperature, and then examined using an electron microscope (H-7600; Hitachi High
Technologies, Tokyo, Japan) at 100 kV.
Supplementary data
Supplementary data are available at PCP online.
866
Funding
The Japan Society for the Promotion of Science Scientific
(Research Grant No. 02718 to Y.O.); the Japanese Ministry
of Education, Science, Culture, Sports and Technology
Grant-in-Aid for Scientific Research on Priority Areas
(Nos. 19039008 and 20061008 to S.H.); the Institute for
Bioinformatics Research and Development (BIRD) from the
Japan Science and Technology Agency (to S.H.)
Acknowledgments
We are grateful to Dr. Yuko Ohashi of NIAS for providing the
E7113 promoter sequence, Dr. Jean-Pierre Zrÿd of Lausanne
University and Dr. Didier Schaefer of INRA Versailles for providing the BS213 site sequence, Dr. Masa Hiko Sato of Kyoto
Prefectural University for the GFP–AtVAM3 fusion gene fragment, Dr. Natsumaro Kutsuna of The University of Tokyo for
technical advice on 3-D reconstruction, and Hiroko Yamashita
of Tokyo Metropolitan University for technical assistance.
References
Allen, V.J., Thompson, H.M. and McNiven, M.A. (2002) Motoring
around the Golgi. Nat. Cell Biol. 4: 226–242.
Ashford, A.E. (1998) Dynamic pleiomorphic vacuole systems: are
they endosomes and transport compartments in fungal hyphae.
Adv. Bot. Res. 28: 199–159.
Boevink, P., Oparka, K., Santa Cruz, S., Martin, B., Betteridge, A. and
Hawes, C. (1998) Stacks on tracks: the plant Golgi apparatus traffics
on an actin/ER network. Plant J. 15: 441–447.
Bolte, S., Talbot, C., Boutte, Y., Catrice, C., Read, N.D. and SatiatJeunemaitre, B. (2004) FM-dyes as experimental probes for dissecting
vesicle trafficking in living plant cells. J. Microsc. 214: 159–173.
Borgese, N., Francolini, M. and Snap, E.L. (2006) Endoplasmic reticulum
architecture: structures in flux. Curr. Opin. Cell Biol. 18: 358–364.
Campbell, R.E., Tour, O., Palmer, A.E., Steinbach, P.A., Baird, G.S.,
Zacharias, D.A., et al. (2002) A monomeric red fluorescent protein.
Proc. Natl Acad. Sci. USA 99: 7877–7882.
Chan, J., Calder, G.M., Doonan, J.H. and Lloyd, C.W. (2003) EB1 reveals
mobile microtubule nucleation sites in Arabidopsis. Nat. Cell Biol. 5:
967–971.
Dhonukshe, P. and Gadella, T.W., Jr., (2003) Alteration of microtubule
dynamic instability during preprophase band formation revealed by
yellow fluorescent protein–CLIP170 microtubule plus-end labeling.
Plant Cell 15: 597–611.
Doonan, J.H., Cove, D.J. and Lloyd, C.W. (1988) Microtubules and
microfilaments in tip growth: evidence that microtubules impose
polarity on protonemal growth in Physcomitrella patens. J. Cell Sci.
89: 533–540.
Du, Y., Ferro-Novick, S. and Novick, P. (2004) Dynamics and inheritance
of the endoplasmic reticulum. J. Cell Sci. 117: 2871–2878.
Egea, G., Lazaro-Dieguez, F. and Vilella, M. (2006) Actin dynamics at the
Golgi complex in mammalian cells. Curr. Opin. Cell Biol. 18: 168–178.
Feiguin, F., Ferreira, A., Kosik, K.S. and Caceres, A. (1994) Kinesinmediated organelle translocation revealed by specific cellular
manipulations. J. Cell Biol. 127: 1021–1039.
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
Microtubules regulate organization of vacuoles
Hasezawa, S., Ueda, K. and Kumagai, F. (2000) Time-sequence
observations of microtubule dynamics throughout mitosis in living
cell suspensions of stable transgenic Arabidopsis--direct evidence
for the origin of cortical microtubules at M/G1 interface. Plant Cell
Physiol. 41: 244–250.
Hepler, P.K., Vidali, L. and Cheung, A.Y. (2001) Polarized cell growth in
higher plants. Annu. Rev. Cell Dev. Biol. 17: 159–187.
Hicks, G.R., Rojo, E., Hong, S., Carter, D.G. and Raikhel, N.V. (2004)
Germinating pollen has tubular vacuoles, displays highly dynamic
vacuole biogenesis, and requires VACUOLESS1 for proper function.
Plant Physiol. 134: 1227–1239.
Higaki, T., Kutsuna, N., Okubo, E., Sano, T. and Hasezawa, S. (2006)
Actin microfilaments regulate vacuolar structures and dynamics:
dual observation of actin microfilaments and vacuolar membrane in
living tobacco BY-2 cells. Plant Cell Physiol. 47: 839–852.
Hiwatashi, Y., Obara, M., Sato, Y., Fujita, T., Murata, T. and Hasebe, M.
(2008) Kinesins are indispensable for interdigitation of phragmoplast
microtubules in the moss Physcomitrella patens. Plant Cell 20:
3094–3106.
Kammerer, W. and Cove, D.J. (1996) Genetic analysis of the effects of
re-transformation of transgenic lines of the moss Physcomitrella
patens. Mol. Gen. Genet. 250: 380–382.
Kasten, B. and Reski, R. (1997) β-Lactam antibiotics inhibit chloroplast
division in a moss (Physcomitrella patens) but not in tomato
(Lycopersicon esculentum). J. Plant Physiol. 150: 137–140.
Kumagai, F., Yoneda, A., Tomida, T., Sano, T., Nagata, T. and
Hasezawa, S. (2001) Fate of nascent microtubules organized at the
M/G1 interface, as visualized by synchronized tobacco BY-2 cells
stably expressing GFP-tubulin: time-sequence observations of the
reorganization of cortical microtubules in living plant cells. Plant
Cell Physiol. 42: 723-732.
Kutsuna, N. and Hasezawa, S. (2002) Dynamic organization of vacuolar
and microtubule structures during cell cycle progression in
synchronized tobacco BY-2 cells. Plant Cell Physiol. 43: 965–973.
Kutsuna, N., Kumagai, F., Sato, M.H. and Hasezawa, S. (2003) Threedimensional reconstruction of tubular structure of vacuolar
membrane throughout mitosis in living tobacco cells. Plant Cell
Physiol. 44: 1045–1054.
Lovy-Wheeler, A., Cárdenas, L., Kunkel, J.G. and Hepler, P.K. (2007)
Differential organelle movement on the actin cytoskeleton in lily
pollen tubes. Cell Motil. Cytoskel. 64: 217–232.
McElroy, D., Zhang, W., Cao, J. and Wu, R. (1990) Isolation of an
efficient actin promoter for use in rice transformation. Plant Cell 2:
163–171.
Menand, B., Calder, G. and Dolan, L. (2007) Both chloronemal and
caulonemal cells expand by tip growth in the moss Physcomitrella
patens. J. Exp. Bot. 58: 1843–1849.
Mitsuhara, I., Ugaki, M., Hirochika, H., Ohshima, M., Murakami, T.,
Gotoh, Y., et al. (1996) Efficient promoter cassettes for enhanced expression of foreign genes in dicotyledonous and monocotyledonous
plants. Plant Cell Physiol. 37: 49–59.
Nebenführ, A., Gallagher, L.A., Dunahay, T.G., Frohlick, J.A.,
Mazurkiewicz, A.M., Meehl, J.B., et al. (1999) Stop-and-go movements
of plant Golgi stacks are mediated by the acto-myosin system. Plant
Physiol. 121: 1127–1142.
Nishiyama, T., Hiwatashi, Y., Sakakibara, I., Kato, M. and Hasebe, M.
(2000) Tagged mutagenesis and gene-trap in the moss, Physcomitrella patens by shuttle mutagenesis. DNA Res. 7: 9–17.
Oda, Y. and Hasezawa, S. (2006) Cytoskeletal organization during
xylem cell differentiation. J. Plant Res. 119: 167–177.
Oda, Y., Mimura, T. and Hasezawa, S. (2005) Regulation of secondary
cell wall development by microtubules during tracheary element
differen-tiation in Arabidopsis suspensions. Plant Physiol. 137:
1027–1036.
Ovecka, M., Lang, I., Baluska, F., Ismail, A., Illes, P. and Lichtscheidl, I.K.
(2005) Endocytosis and vesicle trafficking during tip growth of root
hairs. Protoplasma 226: 39–54.
Paradez, A., Wright, A. and Ehrhardt, D.W. (2006) Microtubule cortical
array organization and plant cell morphogenesis. Curr. Opin. Plant.
Biol. 9: 571–578.
Reisen, D., Marty, F. and Leborgne-Castel, N. (2005) New insights into
the tonoplast architecture of plant vacuoles and vacuolar dynamics
during osmotic stress. BMC Plant Biol. 5: 13.
Runions, J., Brach, T., Kuhner, S. and Hawes, C. (2005) Photoactivation
of GFP reveals protein dynamics within the endoplasmic reticulum
membrane. J. Exp. Bot. 57: 43–50.
Saito, C., Ueda, T., Abe, H., Wada, Y., Kuroiwa, T., Hisada, A., et al. (2002)
Complex and mobile structure forms a distinct subregion within
the continuous vacuolar membrane in young cotyledons of
Arabidopsis. Plant J. 29: 245–255.
Saito, S., Watabe, S., Ozaki, H., Kobayashi, M., Suzuki, T., Kobayashi, H.,
et al. (1998) Actin-depolymerizing effect of dimeric macrolides,
bistheonellide A and swinholide A. J. Biochem. 123: 571–578.
Sakakibara, K., Nishiyama, T., Sumikawa, N., Kofuji, R., Murata, T. and
Hasebe, M. (2003) Involvement of auxin and a homeodomainleucine zipper I gene in rhizoid development of the moss
Physcomitrella patens. Development 130: 4835–4846.
Sato, Y., Wada, M. and Kadota, A. (2001) Choice of tracks, microtubules
and/or actin filaments for chloroplast photo-movement is
differentially controlled by phytochrome and a blue light receptor.
J. Cell Sci. 114: 269-279.
Schaefer, D.G. (2001) Gene targeting in Physcomitrella patens. Curr.
Opin. Plant Biol. 4: 143–150.
Schaefer, D.G. and Zrÿd, J.P. (1997) Efficient gene targeting in the moss
Physcomitrella patens. Plant J. 11: 1196–1206.
Shaw, S.L., Kamyar, R. and Ehrhardt, D.W. (2003) Sustained microtubule
tread-milling in Arabidopsis cortical arrays. Science 300: 1715–1718.
Sheahan, M.B., Rose, R.J. and McCurdy, D.W. (2004) Organelle
inheritance in plant cell division: the actin cytoskeleton is required
for unbiased inheritance of chloroplasts, mitochondria and
endoplasmic reticulum in dividing protoplasts. Plant J. 37: 379–390.
Spurr, A.R. (1969) A low-viscosity epoxy resin embedding medium for
electron microscopy. J. Ultrastruct. Res. 26: 31–43.
Steffen, W., Karki, S., Vaughan, K.T., Vallee, R.B., Holzbaur, E.L., Weiss, D.G.,
et al. (1997) The involvement of the intermediate chain of
cytoplasmic dynein in binding the motor complex to membranous
organelles of Xenopus oocytes. Mol. Biol. Cell 8: 2077–2088.
Terasaki, M. and Reese, T.S. (1994) Interactions among endoplasmic
reticulum, microtubules, and retrograde movements of the cell
surface. Cell Motil. Cytoskel. 29: 291–300.
Uemura, T., Yoshimura, S.H., Takeyasu, K. and Sato, M.H. (2002)
Vacuolar membrane dynamics revealed by GFP–AtVam3 fusion
protein. Genes Cells 7: 743–753.
Vedrenne, C. and Hauri, H.P. (2006) Morphogenesis of the endoplasmic
reticulum: beyond active membrane expansion. Traffic 7:
639–646.
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.
867
Y. Oda et al.
Vos, J.W., Dogterom, M. and Emons, A.M. (2004) Microtubules become
more dynamic but not shorter during preprophase band formation:
a possible ‘search-and-capture’ mechanism for microtubule
translocation. Cell Motil. Cytoskel. 57: 246–258.
Wada, M., Shimizu, H. and Kondo, N. (1987) A model system to study
the effect of SO2 on plant cells II. Effect of sulfite on fern spore
germination and rhizoid development. Bot. Mag. Tokyo 100: 51-62.
Waterman-Storer, C.M. and Salmon, E.D. (1998) Endoplasmic reticulum
membrane tubules are distributed by microtubules in living cells
using three distinct mechanisms. Curr. Biol. 8: 798–806.
Yoneda, A., Kutsuna, N., Higaki, T., Oda, Y., Sano, T. and Hasezawa, S.
(2007) Recent progress in living cell imaging of plant cytoskeleton
and vacuole using fluorescent-protein transgenic lines and threedimensional imaging. Protoplasma 230: 129–139.
(Received October 30, 2008; Accepted February 19, 2009)
868
Plant Cell Physiol. 50(4): 855–868 (2009) doi:10.1093/pcp/pcp031 © The Author 2009.