Microtubules Regulate Dynamic Organization of Vacuoles in Physcomitrella patens

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1 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, * 1 Department of Integrated Biosciences, Graduate School of Frontier Sciences, The University of Tokyo, Kashiwanoha 5-1-5, Kashiwa, Chiba, Japan 2 Institute for Bioinformatics Research and Development (BIRD), Japan Science and Technology Agency (JST), Chiyoda-ku, Tokyo, Japan 3 Department of Biological Sciences, Faculty of Science, Hokkaido University, Kita-ku Kita-10-jo-Nishi-8, Sapporo, , Hokkaido, Japan 4 Division of Evolutionary Biology, National Institute for Basic Biology, Okazaki, Japan 5 ERATO, JST, Okazaki, Japan 6 Department of Biological Sciences, Graduate School of Science and Engineering, Tokyo Metropolitan University, Minami-Ohsawa 1-1, Hachioji, Tokyo, Japan 7 Department of Basic Biology, School of Life Science, The Graduate University for Advanced Studies, Okazaki, Japan 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 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 ; FM4-64, 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 Regular Paper 8 Present address: Department of Biological Science, Graduate School of Sciences, The University of Tokyo, Hongo 7-3-1, Bunkyo-ku, Tokyo, Japan. *Corresponding author: , hasezawa@k.u-tokyo.ac.jp. Fax, Plant Cell Physiol. 50(4): (2009) doi: /pcp/pcp031, available online at The Author Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please journals.permissions@oxfordjournals.org 855

2 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 ) 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 ), Golgi ( Boevink et al. 1998, Nebenführ et al ) 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 ). For instance, spherical membranes and sheet-like membranes have been reported within the vacuolar lumen of Arabidopsis epidermal cells ( Saito et al ), 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 ). 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 ). 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 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 ). 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 ). 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 ) 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 856

3 Microtubules regulate organization of vacuoles Fluorescent Bright field Merged FM4-64 GFP A C 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 B D 0 h 3 h ( 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. Dynamics of vacuolar structures in the chloronema cell cortex 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, 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 ), 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 857

4 Y. Oda et al. N 2.0 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. Number of tubules Apical Subapical O Number of inner sheets Apical Subapical 858

5 Microtubules regulate organization of vacuoles A C B 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 µ m. (B) A fine tube of the vacuole (arrowhead) projects from the vacuole and fuses with another region of the vacuolar membrane. Bar = 5 µ m. (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, 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 µ m. (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 D 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 859

6 Y. Oda et al. A B C D E F G H I 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 µ M), 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 µ M) 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 ) to compare the vacuolar movement and microtubule dynamics. Interestingly, the contraction rate just after contact with J Cytoplasmic accumulation (µm) Control Oryzalin BA Lat B 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 860

7 Microtubules regulate organization of vacuoles A C 10 Distance of vacuole from rhizoid tip (µm) Oryzalin Control B 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 µ M, 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 µ m. (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 ± events Vacuole contraction 311 ± events Vacuole contraction after 699 ± events PM contact Vacuole contraction in 261 ± events cytoplasm Microtubule growth rate 86 ± 16 8 microtubules Microtubule shrinkage rate 324 ± 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 D Rate of vacuole movement (µm/sec) Time (sec) 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 861

8 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. A mrfp-tubulin GFP-AtVAM3 Merged D FM4-64 GFP-tubulin Merged B C Intensity (a. u.) Intensity (a. u.) GFP-AtVAM3 mrfp-tubulin * * * * Position 40 sec 80 sec E Merged GFP-tubulin FM4-64 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

9 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 ) 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 ). 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. 863

10 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 ). 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 ). 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 ). Lovy-Wheeler et al. (2007) reported that movements of 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 ( nm s 1 ) in higher plant cells ( Chan et al. 2003, Dhonukshe and Gadella 2003, Shaw et al. 2003, Vos et al ). 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 ) 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 864

11 Microtubules regulate organization of vacuoles polarity and normal growth in moss protonema cells ( Doonan et al ), 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. Materials and Methods Plant culture Physcomitrella patens Gransden2004 was cultured on BCDAT agar plates ( Nishiyama et al ) at 23 C under a 16 h light/8 h dark photoperiod (50 µ mol 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 ) in vector ptfh15.3 ( ). This vector contains Pphb7 genomic fragments ( Sakakibara et al ), 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 ) and PpTUA1, a P. patens α-tubulin, in vector pcmak1 was used ( Hiwatashi et al ). This vector 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 µ M 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 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 ( ). 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, Photo- Metrics, Huntington Beach, Canada). When mrfp signals were detected, a 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 865

12 Y. Oda et al. detected through a confocal unit (CSU10, Yokogawa Electric Corporation, Tokyo, Japan) with a nm band-pass filter (Semrock) for GFP, and a nm band-pass filter (Olympus) for FM4-64 Inhibitor treatments For actin depolymerization, protonema cells were treated with 1 µ M 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 µ M 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% OsO 4 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 ( µ m) 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. Funding The Japan Society for the Promotion of Science Scientific (Research Grant No to Y.O.); the Japanese Ministry of Education, Science, Culture, Sports and Technology Grant-in-Aid for Scientific Research on Priority Areas (Nos and 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 : Ashford, A.E. 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