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First published online June 26, 2008; 10.1104/pp.108.119842 Plant Physiology 147:1699-1709 (2008) © 2008 American Society of Plant Biologists OPEN ACCESS ARTICLE
Synthetic Lipid (DOPG) Vesicles Accumulate in the Cell Plate Region But Do Not Fuse1,[W],[OA]Laboratory of Plant Cell Biology, Department of Plant Sciences, Wageningen University, 6703–BD Wageningen, The Netherlands (A.E.-O., J.W.V., A.A.M.v.L., A.M.C.E.); and FOM Institute for Atomic and Molecular Physics, 1009–DB Amsterdam, The Netherlands (A.M.C.E.)
The cell plate is the new cell wall, with bordering plasma membrane, that is formed between two daughter cells in plants, and it is formed by fusion of vesicles (approximately 60 nm). To start to determine physical properties of cell plate forming vesicles for their transport through the phragmoplast, and fusion with each other, we microinjected fluorescent synthetic lipid vesicles that were made of 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (DOPG) into Tradescantia virginiana stamen hair cells. During interphase, the 60-nm wide DOPG vesicles moved inside the cytoplasm comparably to organelles. During cytokinesis, they were transported through the phragmoplast and accumulated in the cell plate region together with the endogenous vesicles, even inside the central cell plate region. Because at this stage microtubules are virtually absent from that region, while actin filaments are present, actin filaments may have a role in the transport of vesicles toward the cell plate. Unlike the endogenous vesicles, the synthetic DOPG vesicles did not fuse with the developing cell plate. Instead, they redistributed into the cytoplasm of the daughter cells upon completion of cytokinesis. Because the redistribution of the vesicles occurs when actin filaments disappear from the phragmoplast, actin filaments may be involved in keeping the vesicles inside the developing cell plate region.
In plant cells, the cell plate constitutes the new cell wall with plasma membranes that separates the cytoplasm of the two daughter cells during cytokinesis. It is formed by the fusion of membrane vesicles of approximately 60 nm in diameter that contain a variety of hemicelluloses and pectins, and have callose and cellulose synthesizing enzyme complexes in their membrane (Zuo et al., 2000  ; Verma, 2001; Yokoyama and Nishitani, 2001).
The cell plate is built up in the middle of a structure called the phragmoplast, often at an equatorial plane in the cell. The phragmoplast is a cytoplasmic dense area containing microtubules, actin filaments, endoplasmic reticulum (ER), and cell plate forming vesicles (Schopfer and Hepler, 1991
The process of cell plate formation with its intermediate stages is well studied (Seguí-Simarro et al., 2004
With the help of dynamin-like molecules, the fusion of vesicles leads to the formation of membrane fusion tubes with a diameter of 20 nm (Samuels et al., 1995
For years it was believed that cell plate forming vesicles are only Golgi derived (Whaley and Mollenhauer, 1963 Here, we describe results obtained with the injection of synthetic phospholipid vesicles into plant cells to start to determine possible necessary and sufficient physical properties of cell plate forming vesicles for their transport through the phragmoplast and for their fusion with each other and with the existing cell plate. We used phospholipid vesicles made of 1,2-dioleoyl-sn-glycero-3-[phospho-rac-(1-glycerol)] (DOPG) that partly mimic the behavior of endogenous vesicles during cell plate formation. We show that upon injection, DOPG vesicles are transported to and through the phragmoplast and accumulate in the region of the existing growing cell plate. However, unlike endogenous vesicles, synthetic vesicles redistribute into the cytoplasm of the daughter cells upon completion of cytokinesis. This suggests that the phragmoplast is not selective for lipid membrane composition or for integral membrane proteins. Interestingly, the synthetic vesicles are kept in the cell plate region during its formation. In this process the CPAM could be involved. Because vesicles keep moving to the center of the cell plate during late telophase when the microtubules have left that area but actin filaments are still present, we envision a role for the actin cytoskeleton in this process.
Synthetic Lipid Vesicles (DOPG) Distribute throughout the Cytoplasm of Interphase Tradescantia virginiana Stamen Hair Cells We made synthetic vesicles from negatively charged (anionic) DOPG, labeled with 2% fluorescent phosphocholine Bodipy FC12-HPC, with a size of 60 nm (±7 nm; Fig. 1 ) to study physical properties of the vesicles that are required for transport through the phragmoplast and the formation of the cell plate. The vesicles did not fuse or aggregate with each other, even after 7 d in the buffer used for microinjection (data not shown). Synthetic lipid (DOPG) vesicles injected into young elongating interphase stamen hair cells of T. virginiana distributed evenly in the cytoplasm within 5 to 10 min after microinjection and were visible as individual fluorescent speckles (Fig. 2, A and B ). The injection of these vesicles was not lethal to the cells, nor did it affect cytoplasmic streaming or the cytoarchitecture (Fig. 2A; differential interference contrast [DIC] images). Inside the cells, the vesicles also seemed to be stable because their fluorescence was not incorporated into the plasma membrane, tonoplast, ER, or other organelles (mitochondria and plastids) that can be seen with DIC microscopy. In addition, the vesicles did not fuse together in the cytoplasm, as no obvious conglomerates were formed and the number of speckles did not markedly drop over time. Microinjected fluorescent vesicles moved through the cytoplasm for at least 1.5 h after injection.
The average velocity of vesicles in the cytoplasm was 0.74 ± 0.04 (SE) µm/s, which is comparable to, but consistently slower than, that of visible organelles with a size of approximately 1 to 2 µm in these cells (1.12 ± 0.06 [SE] µm/s) and in cells that were not injected (1.24 ± 0.12 [SE] µm/s; Fig. 2C). The movement of DOPG vesicles and organelles in injected cells was similar (Supplemental Fig. S1). This suggests that the injected vesicles may be coated with motor proteins and move on cytoskeletal tracks like organelles do, or else, move by hydrodynamic flow produced by the moving organelles.
Upon microinjection into cells in early-to-late anaphase, synthetic lipid vesicles moved from the microinjection site and distributed evenly in the cytoplasm. Within 5 to 10 min after injection into anaphase cells, synthetic lipid vesicles spread throughout the cytoplasm and between the chromosomes in the spindle (Fig. 3 ). When injected cells entered telophase, the vesicles accumulated at the spindle midzone and were seen as a broad band of fluorescence. In DIC microscopy, a cell plate could not yet be observed at this stage (Fig. 3A; 22 min). This band of accumulated synthetic vesicles subsequently narrowed at the same time when the young cell plate became visible with DIC microscopy. This was seen in all injected cells (n = 24 cells). Occasionally, an additional accumulation of lipid vesicles was seen around the edges of the growing cell plate (data not shown).
The accumulation of injected synthetic lipid vesicles in the cell plate region was specific; vesicles did not just fill the accessible volume like coinjected fluorescent dextran did (Fig. 3C).
We observed a uniform distribution of fluorescence throughout the cell plate region, i.e. there was no black line in the middle, which suggests that injected vesicles accumulated not only on the surface of the developing cell plate, but also within the fenestrated cell plate. We compared the width of the vesicle accumulation zone with that of the cell plate itself, as measured with FM4-64 labeling. FM4-64 is a fluorescent lipophilic styryl dye that labels the developing cell plate when applied from the outside of the cell (Bolte et al., 2004
The injection of synthetic lipid vesicles did not hinder or delay cell plate formation. In all vesicle injected cells, cell plate formation took on average 27 min ± 3 s, which did not differ from the experimental controls (Fig. 5 ). Furthermore, the initial appearance, the growth, and the attachment of the cell plate to the parental cell wall were all normal. Interestingly, the moment of injection did not have any influence on the accumulation of vesicles in the cell plate region. Injected vesicles accumulated in the whole cell plate region when injected at different stages of cell division (Table I ), even in cells in which a cell plate was already visible at the time of injection (Fig. 6A ). Further, synthetic vesicles were not redistributed back into the cell as long as the phragmoplast existed. Because microtubules are mostly present at the periphery of the torus-shaped phragmoplast, they are not likely to be instrumental in bringing the vesicles to the center or keeping them there at this later stage of cell plate formation (Fig. 6B).
To exclude that fluorescently labeled Bodipy FC12-HPC phosphocholine lipid or free Bodipy dye could label the cell plate without being part of the synthetic vesicles, we injected the vesicles into the interphase cell, neighboring a cell in anaphase (Fig. 7 ). Upon injection, some fluorescence was transferred from the injected cell into the neighboring cells. This could be free Bodipy FC12-HPC, which has a molecular mass of 0.895 kD and is close to the exclusion limit (0.8 kD) of plasmodesmata (Pheasant and Hepler, 1987 ). In the dividing cell, no labeling of the cell plate occurred, which further underpins that the observed fluorescence in the cell plate region comes from the accumulation of injected fluorescent vesicles, and not from free dye.
Synthetic Lipid Vesicles Redistribute inside the Daughter Cells after Cell Plate Attachment At the end of cytokinesis, when the cell plate attached to the parental plasma membrane and cell wall, the lipid vesicles redistributed completely into the cytoplasm, while the FM4-64 labeling remained in the plasma membranes lining the young cell wall (Fig. 8 ). The fluorescent band of lipid vesicles first broadened (Fig. 8A; 2 min), then two clouds of fluorescence were formed one on each side of the new cell wall (Fig. 8A; 4–8 min), and finally the fluorescence completely redistributed in the cytoplasm of the cells. This shows that although synthetic lipid vesicles accumulate in the fenestrated cell plate, they do not fuse together with endogenous vesicles to form the cell plate.
We analyzed the fluorescence intensities of a region of 1.5 x 5 µm of the phragmoplast over time, including the cell plate, in cells injected with fluorescent synthetic vesicles and labeled with FM4-64 (Fig. 9 ). During the initial phase of cell plate growth and expansion, injected DOPG vesicles and FM4-64 labeled membranes showed the same distribution (Fig. 4B). The thickness of the cell plate of synthetic vesicles and FM4-64 labeling, which was measured as the width at half-height of a Gauss curve fitted to the fluorescence intensity profile (see also Fig. 4C), stabilized after 10 to 15 min at 0.5 to 0.7 µm (Fig. 9A). About 30 to 35 min after initiation, the cell plate attached to the parental plasma membrane. At this moment the DOPG vesicles lost their confinement to the cell plate region and concomitantly the width of the fluorescence peak broadened. On the other hand, the thickness of cell plate as measured with FM4-64 did not broaden and stayed approximately 0.5 µm. The disappearance of the synthetic vesicles from the new cell wall between the two daughter cells is also evident from the fluorescence intensity measurements over time (Fig. 9B). Starting at 25 to 30 min, the maximum fluorescence intensity at the cell plate decreased steadily to the initial levels, whereas the FM4-64 labeling remained constant during the whole period. The decrease in fluorescence of the synthetic vesicles in the cell plate region was not caused by the bleaching of Bodipy; the fluorescence intensity of synthetic vesicles in the cytoplasm of the same cells did not change over time. This shows that injected vesicles only accumulate in the cell plate region, but do not fuse with each other, with the endogenous cell plate forming vesicles, or with the developing cell plate. They are again redistributed throughout the cytoplasm when the cell plate attaches to the parental plasma membrane and cell wall, and the maturation of the young cell wall begins, at the moment that the typical phragmoplast actin cytoskeleton also disappears.
Injected Synthetic Vesicles Move through the Cytoplasm with Velocities Comparable But Slightly Less Than Organelles
We used synthetic DOPG vesicles to study basic physical properties that vesicles require to enter, move through, and accumulate in the cell plate region. We prepared 60-nm DOPG vesicles with the same size as endogenous Golgi vesicles that form the cell plate (Reichardt et al., 2007 The injected lipid vesicles did not fuse together or with endogenous membranes, but moved through the cytoplasm of interphase T. virginiana stamen hair cells at slightly lower velocities than large endogenous organelles. The presence of the synthetic DOPG membrane did not affect the movement of these endogenous organelles in the cytoplasm (Figs. 2 and 3).
Although we injected uncoated lipid vesicles, these vesicles may be coated in the cells with cytoplasmic proteins. The fast distribution of the vesicles and their movement inside the cytoplasm could be caused by motor protein binding and subsequent transport activity. It is known that myosin-I prefers to bind anionic lipids (Adams and Pollard, 1989
Although their membrane composition and content are different from endogenous vesicles, microinjected synthetic DOPG vesicles passed through the phragmoplast structure of microtubules, actin filaments, and ER membranes, and accumulated at the developing cell plate. They did not disturb the transport of endogenous vesicles toward the cell plate because the initiation and the subsequent development of the cell plate was not disturbed in any of the injected cells (as visualized with DIC and FM4-64 and with regard to time; Figs. 4–6
The specific accumulation of synthetic vesicles in the cell plate region of the phragmoplast, instead of filling the accessible volume (Supplemental Fig. S3), points to two processes, namely, the directional transport of these vesicles inside the phragmoplast toward the cell plate and the inhibition of transport away from the cell plate. Because the vesicles were transported through the cytoplasm after they were injected, it could well be that these vesicles were, as we discussed above, coated with cytoplasmic motor proteins that caused their movement along the phragmoplast microtubule and/or actin cytoskeleton. However, the mode of transport of endogenous vesicles through the phragmoplast is still unclear. With electron microscopy (EM), it has been observed that some vesicles are associated with phragmoplast microtubules, suggesting the microtubules as the cytoskeletal element along which the vesicles are transported, but some other vesicles apparently were not associated with microtubules (Otegui et al., 2001
Although all our data suggest a role for the actin cytoskeleton in keeping vesicles in the cell plate region, we cannot exclude the possibility that the negative charge of DOPG vesicles contribute to its accumulation in the cell plate region.
Although the injected synthetic vesicles were transported through the phragmoplast and accumulate in the cell plate region, they did not fuse with the developing cell plate, but redistributed again throughout the cytoplasm upon attachment of the cell plate to the parental cell wall (Fig. 8). This redistribution strongly suggests that these vesicles cannot fuse with endogenous vesicles/cell plate or with each other. The latter was already indicated by the fact that the vesicles did not fuse in vitro, even after 7 d. In injected cells, the DOPG vesicles probably reside in the tubulo-vesicular network, the tubular network, and finally in the fenestrae (Samuels et al., 1995
Our observation that injected vesicles cannot fuse with endogenous vesicles in the developing cell plate was to be expected. Firstly, the injected vesicles lack trans-membrane SNARE proteins and are probably not able to acquire them after injection. SNAREs are necessary for the fusion of cell plate forming vesicles because the Arabidopsis SNARE mutants knolle and keule have unfused cytokinetic vesicles in the cell plate region (Assaad et al., 1996
During the centrifugal growth of the cell plate, the centrally located microtubules depolymerize and reassemble at the periphery of the phragmoplast, which then becomes torus shaped (second phase; Valster and Hepler, 1997
The breakdown of actin filaments in the phragmoplast occurs uniformly along the whole width at the moment when the cell wall is completed, rather than in a centrifugal pattern (Cleary et al., 1992 We have shown that DOPG vesicles are a useful tool to study the processes involved in cell plate formation. Using these synthetic vesicles, we uncovered what to our knowledge is a possible new role for the phragmoplast actin cytoskeleton in cell plate formation. The next challenge is to elucidate the exact origin(s) and the nature of the endogenous vesicles that form the cell plate. As one of the direct experimental approaches, we propose to inject vesicles with experimentally defined surfaces (e.g. specific phospho- and/or glycolipid compositions and embedded and/or [covalently] attached proteins) to analyze the vesicle transport mechanism and the requirements for vesicles to fuse and form the cell plate.
Plant Material
Tradescantia virginiana plants were grown in a growth chamber with a 16-h photoperiod at 25°C and 8-h dark period at 18°C and 75% to 80% relative humidity. Stamen hairs with dividing cells in the apical region were dissected from immature flower buds with a length of approximately 5 mm. For microinjection experiments, we immobilized these stamen hairs in a thin layer of 1% low-temperature gelling agarose (BDH Laboratory Supplies) in culture medium (5 mM HEPES, 1 mM MgCl2, and 0.1 mM CaCl2, pH 7.0) and 0.025% Triton X-100 (BDH Laboratory Supplies), following the procedure described by Vos et al. (1999)
Synthetic vesicles consisted for 98% of the anionic nonfluorescent phospholipid DOPG (Avanti Polar Lipids) and for 2% of the fluorescent phosphocholine Bodipy FC12-HPC (excitation maximum at 503 nm, emission maximum at 512 nm; Molecular Probes). Bodipy FC12-HPC was especially chosen because it labels the acyl chain rather than the hydrophilic head of the phospholipid. It therefore prevents the movement of the fluorescent probe between lipid layers (jumping). DOPG was purchased as a chloroform solution of a sodium salt. Bodipy FC12-HPC was purchased as a powder and was than dissolved in ethanol. Phospholipids were mixed together and dried onto a glass surface under a stream of nitrogen, followed by at least 2 h under vacuum to remove the last traces of solvent. The dried lipid mixture was hydrated with injection buffer (5 mM HEPES, 0.1 mM KCl, pH 7.0) to a concentration of 0.5 mg/mL; this was an optimal concentration to observe vesicles in the microscope and to measure them with dynamic light scattering. The lipids were freeze-thawed with liquid nitrogen for five cycles to disperse them and pushed through an extruder with a polycarbonate filter with a 50-nm pore size to yield vesicles with a diameter of approximately 60 nm. The vesicle diameter was determined using dynamic light scattering and cryo-transmission electron microscopy (cryo-TEM). For dynamic light scattering, the instrumental setup consisted of an ALV-5000 correlator and a scattering device with an ALV-125 goniometer and a multiline Lexel AR-laser source. Data for monodispersity were collected at scattering angles of 60°, 90°, and 120° at a wavelength of 513 nm. After preparation, vesicle fluorescence was checked under a confocal laser scanning microscope. Vesicles were stored at 4°C and used for microinjection within 4 d, although they were stable in microinjection buffer for more than 7 d as determined with dynamic light scattering. For coinjection experiments, vesicles were mixed with fluorescent dextran (Alexa-568 dextran, 10 kD; Invitrogen). Dextran was dissolved in microinjection buffer as a 5 mg/mL stock and diluted to 0.5 mg/mL before the microinjection experiment in the microinjection buffer containing the DOPG vesicles. For labeling of the cell plate with FM4-64 (Invitrogen), the dye was dissolved in DMSO as a stock of 200 µM and further diluted in the preparation medium or water and used at a final concentration of 2 µM.
For observation of synthetic DOPG vesicles in cryo-TEM, 200-mesh copper grids covered with holey carbon film were submerged in the vesicle solution in microinjection buffer. The excess fluid from the film was removed with filter paper. The grids were plunge-frozen in liquid propane and after short storage in liquid nitrogen placed in a cryo-holder (Gatan 626). The vesicles were observed with a transmission electron microscope JEOL 1200 EX II. Images were made with a KeenView digital camera (SIS).
The microinjection experiments were conducted according to Vos et al. (1999)
Microinjections were performed on inverted microscopes. Images were collected with a Cell Map IC (Bio-Rad) confocal laser-scanning microscope, coupled to an Eclipse TE2000-S (Nikon) or with an LSM 5 Pascal confocal laser-scanning microscope coupled to an Axiovert 200 microscope (Zeiss). For Bodipy/Alexa-568 dextran or FM4-64 dual scanning we used the excitation/emission combination of 488/520 to 540 nm BP and 532/560 nm LP (Cell Map IC) or the combination of 488/BP 505 to 550 nm BP and 543/560 nm LP (HFT 488/NFT 545/HFT 543; LSM 5 Pascal). Images were obtained with a 1.4 NA 60x or 1.4 NA 63x oil immersion objective, collected by Kalman averaging of two to three full scans (Cell Map IC) or with scan speed 7 (LSM 5 Pascal). Images were taken at 2- or 3-min intervals, which allowed observation of developing cell plates for long periods of time without disturbing the cell plate formation process. Images were processed and analyzed with the software programs Confocal Assistant 4.02 (written by Todd Clark Brelje), Adobe Photoshop 5.0 and 8.0 (Adobe Systems), and Image J (version 1.32j; National Institutes of Health). For fluorescence intensity plots, rectangles (10 x 50 pixels) of images at different time points were cut out and reduced to 1 x 50 pixels to average the fluorescence. The resulting pixel intensities were saved as text files with Image J, and plotted and curve-fitted in Origin (version 7.5 SR5; OriginLab) using standard Gauss equations. Images of actin filaments in phragmoplast of BY-2 cells transformed with GFP::FABD were made with a spinning disc confocal microscope (PerkinElmer) coupled to an Eclipse TE2000-S (Nikon).
The following materials are available in the online version of this article.
We gratefully thank Richard Kik, Mieke Kleijn, and Frans Leermakers of the Laboratory of Physical Chemistry and Colloid Science, Wageningen University, for help with designing the vesicles and for useful discussions. We thank Magdalena Szechy  ska-Hebda for initial help with the vesicle microinjections. We also thank Adriaan van Aelst for help with cryo-TEM and John Esseling for critical reading of the manuscript. Received March 26, 2008; accepted June 19, 2008; published June 26, 2008.
1 This work was supported by the FOM Institute for Atomic and Molecular Physics, Amsterdam (to A.M.C.E.). 
2 Present address: Department of Tumor Immunology, Nijmegen Centre for Molecular Life Sciences, Geert Grooteplein zuid 28, 6500–HB Nijmegen, The Netherlands. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Anne Mie C. Emons (annemie.emons{at}wur.nl).
[W] The online version of this article contains Web-only data.
[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.108.119842 * Corresponding author; e-mail annemie.emons{at}wur.nl.
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ka F, Schlicht M, Hlavacka A, 
amaj J, Friml J, Gadella WJ Jr (2006) Endocytosis of cell surface material mediates cell plate formation during plant cytokinesis. Dev Cell 10: 137–150
