TY - JOUR
AU - Emons, Anne Mie C.
AB - Abstract In plant cells, Golgi vesicles are transported to the division plane to fuse with each other, forming the cell plate, the initial membrane-bordered cell wall separating daughter cells. Vesicles, but not organelles, move through the phragmoplast, which consists of two opposing cylinders of microtubules and actin filaments, interlaced with endoplasmic reticulum membrane. To study physical aspects of this transport/inhibition process, we microinjected fluorescent synthetic 1,2-dioleoyl-sn-glycero-3-phospho-rac-1-glycerol (DOPG) vesicles and polystyrene beads into Tradescantia virginiana stamen hair cells. The phragmoplast was nonselective for DOPG vesicles of a size up to 150 nm in diameter but was a physical barrier for polystyrene beads having a diameter of 20 and 40 nm and also when beads were coated with the same DOPG membrane. We conclude that stiffness is a parameter for vesicle transit through the phragmoplast and discuss that cytoskeleton configurations can physically block such transit. Cells and their constituents are physical entities, and next to chemical interactions, cell structures are determinants of cell behavior. Therefore, apart from techniques to image living cells at the subcellular level, experiments are needed that probe physical parameters important in cell function in vivo. We took the plant phragmoplast structure to answer the question whether the physical aspect “stiffness” is a factor in the inhibition of transport through this structure by microinjecting synthetic vesicles and polystyrene beads in Tradescantia virginiana stamen hair cells during cytokinesis, when the phragmoplast is essential for partitioning the cytoplasm between two daughter cells. Plant cells partition by producing a cell plate made of fused 60- to 80-nm-diameter vesicles (Staehelin and Hepler, 1996; Juݶrgens, 2005) proven to be Golgi vesicles (Reichardt et al., 2007). Their content becomes the new cell wall and their membranes become the daughter cell plasma membranes. The phragmoplast consists of two opposing cylinders of microtubules and actin filaments, interlaced with similarly aligned endoplasmic reticulum (ER) membranes. This phragmoplast cytoskeleton is the transport vehicle for Golgi vesicles to the plane where the cell plate is being formed (Staehelin and Hepler, 1996; Valster et al., 1997), keeps them in this plane (Esseling-Ozdoba et al., 2008b), where they fuse with each other (Samuels et al., 1995; Otegui et al., 2001; Seguiݩ-Simarro et al., 2004), and assists in the proper attachment of the cell plate to the parental cell wall (Valster et al., 1997; Molchan et al., 2002). Transit of organelles, including Golgi bodies, is inhibited (Staehelin and Hepler, 1996; Nebenfuݶhr et al., 2000; Seguiݩ-Simarro et al., 2004). Most of these data are known from static electron microscopy images. Electron microscopy after high-pressure freezing and freeze substitution (Thijsen et al., 1998) and electron tomography studies (Otegui et al., 2001; Seguiݩ-Simarro et al., 2004; Austin et al., 2005) show that, in the early stage of cell plate formation in the center and later at the phragmoplast border, microtubules are aligned parallel to each other at distances of 20 to 100 nm. Keeping in mind that also actin filaments and ER membranes, aligned in the same orientation, are present between the microtubules, this leaves little room for the cell plate-forming vesicles during their transport through this phragmoplast. Clearly, during the past decade, significant progress has been made in the elucidation of the structural organization of cell plate-forming phragmoplasts, which has set the stage for studies to elucidate physical properties of phragmoplasts. The experimental approach we use is injecting particulate and vesicular fluorescent probes into living and dividing cells and observing the extent to which such probes can enter the phragmoplast and can be transported to the cell plate region. We have shown before that synthetic lipid 1,2-dioleoyl-sn-glycero-3-phospho-rac-1-glycerol (DOPG) vesicles of 60 nm diameter are transported through the phragmoplast, accumulate, and are kept in the cell plate region but do not fuse (Esseling-Ozdoba et al., 2008b). Now, we asked whether similar, flexible, synthetic lipid (DOPG) vesicles of various sizes, smaller and larger than endogenous vesicles, as well as stiff polystyrene beads, and such beads coated with the DOPG membrane, are transported through the phragmoplast and enter the plane where the cell plate is being formed, a question pertaining to a physical property of the phragmoplast. Our principal finding is that injected synthetic vesicles up to 150 nm diameter can enter and be transported to the cell plate region, where they accumulate but do not become incorporated into the cell plate. In contrast, polystyrene beads, the noncoated ones and those coated with the same lipid as the vesicles with diameters of 20 and 40 nm, can enter phragmoplasts but cannot be transported to the cell plate region, and the 40-nm beads slow cell plate formation, possibly by interfering with the delivery of normal, cell plate-forming vesicles to the cell plate. RESULTS Injected Synthetic Lipid Vesicles and Polystyrene Beads Move in Interphase Cells of T. virginiana Stamen Hairs Fluorescently labeled synthetic vesicles made of DOPG and fluorescent polystyrene beads injected into interphase cells of T. virginiana stamen hairs were visible as individual fluorescent speckles and distributed evenly in the complete cytoplasm within 5 to 10 min (Fig. 1, A and B Figure 1. Open in new tabDownload slide A and B, Fluorescently labeled synthetic lipid vesicles of 60 nm (A) and polystyrene beads of 40 nm (B) distribute in the cytoplasm of interphase stamen hair cells of T. virginiana upon injection. Cells in A and B are 10 min after microinjection. Bars = 10 μm (A) and 5 μm (B). C and D, Time series of moving vesicles (C; moving vesicle indicated by white arrowheads or asterisks) and polystyrene beads (D; moving bead indicated by white arrowheads or asterisks) in the cytoplasm. Time intervals are 1.9 s. Bars = 5 μm. E, Average velocity of organelles, injected vesicles, and polystyrene beads in interphase cells of T. virginiana. The average velocity was calculated from the displacement of vesicles or organelles over the time that they were followed. Data shown with sd. Figure 1. Open in new tabDownload slide A and B, Fluorescently labeled synthetic lipid vesicles of 60 nm (A) and polystyrene beads of 40 nm (B) distribute in the cytoplasm of interphase stamen hair cells of T. virginiana upon injection. Cells in A and B are 10 min after microinjection. Bars = 10 μm (A) and 5 μm (B). C and D, Time series of moving vesicles (C; moving vesicle indicated by white arrowheads or asterisks) and polystyrene beads (D; moving bead indicated by white arrowheads or asterisks) in the cytoplasm. Time intervals are 1.9 s. Bars = 5 μm. E, Average velocity of organelles, injected vesicles, and polystyrene beads in interphase cells of T. virginiana. The average velocity was calculated from the displacement of vesicles or organelles over the time that they were followed. Data shown with sd. ). Their injection did not affect cytoplasmic streaming or cytoarchitecture. They moved through the cytoplasm for at least 1.5 h after injection with average velocities similar to each other but slightly slower than those of visible organelles with a size of approximately 1 to 2 μm (Fig. 1, C–E). Polystyrene beads moved with velocities of 0.64 μm s™1 (sd = 0.34) and vesicles with 0.75 μm s™1 (sd = 0.28), whereas the velocity of the organelles mentioned is 0.95 μm s™1 (sd = 0.20). In Mitotic Cells, Synthetic Vesicles of Various Sizes Are Transported through the Phragmoplast, Delivered to the Cell Plate Region, and Kept in This Region, But They Redistribute into the Cytoplasm after Cell Plate Completion Vesicles ranging from 45 nm, which are 0.75 times the diameter of endogenous vesicles, up to 150 nm, a diameter of 2.5 times that of endogenous vesicles, all accumulated in the plane of the cell plate (Fig. 2A Figure 2. Open in new tabDownload slide Synthetic vesicles accumulate in the cell plate region but polystyrene beads of 40 nm do not. Time course of T. virginiana stamen hair cells at anaphase with injected fluorescently labeled synthetic (DOPG) lipid vesicles of 150 nm (A) and polystyrene beads of 40 nm (B). Synthetic lipid vesicles of 45, 60, 75, and 150 nm (shown here) accumulate in the cell plate region (white arrows). Polystyrene beads do not accumulate in the cell plate region (white arrowheads). Numbers indicate time in minutes after injection. Bars = 10 μm. Figure 2. Open in new tabDownload slide Synthetic vesicles accumulate in the cell plate region but polystyrene beads of 40 nm do not. Time course of T. virginiana stamen hair cells at anaphase with injected fluorescently labeled synthetic (DOPG) lipid vesicles of 150 nm (A) and polystyrene beads of 40 nm (B). Synthetic lipid vesicles of 45, 60, 75, and 150 nm (shown here) accumulate in the cell plate region (white arrows). Polystyrene beads do not accumulate in the cell plate region (white arrowheads). Numbers indicate time in minutes after injection. Bars = 10 μm. ; Table I Table I. Synthetic vesicles of various sizes accumulate in the cell plate region Size of Injected Vesicles . Percentage of Cells with Vesicle Accumulation . 45 nm 100 (n = 5) 60 nm 100 (n = 29) 75 nm 100 (n = 8) 150 nm 100 (n = 7) Size of Injected Vesicles . Percentage of Cells with Vesicle Accumulation . 45 nm 100 (n = 5) 60 nm 100 (n = 29) 75 nm 100 (n = 8) 150 nm 100 (n = 7) Open in new tab Table I. Synthetic vesicles of various sizes accumulate in the cell plate region Size of Injected Vesicles . Percentage of Cells with Vesicle Accumulation . 45 nm 100 (n = 5) 60 nm 100 (n = 29) 75 nm 100 (n = 8) 150 nm 100 (n = 7) Size of Injected Vesicles . Percentage of Cells with Vesicle Accumulation . 45 nm 100 (n = 5) 60 nm 100 (n = 29) 75 nm 100 (n = 8) 150 nm 100 (n = 7) Open in new tab ) without any delay or disruption of cell plate formation (Fig. 3 Figure 3. Open in new tabDownload slide A, Line profiles through the phragmoplast of cells, injected either with vesicles (150 nm) or polystyrene beads (40 nm). The average fluorescence intensity of each horizontal row of pixels was plotted versus its vertical position. The fluorescence of vesicles is highest in the cell plate region (1.5–2.0 μm), while fluorescence of beads is lowest in the cell plate region. Polystyrene beads do not accumulate in the cell plate region, while vesicles accumulate there specifically. The same was observed for any size of vesicles or beads. B, Duration of cell plate formation in control noninjected cells and cells injected with vesicles of 45 or 150 nm and 20- or 40-nm polystyrene beads. Polystyrene beads of 40 nm delay cell plate formation, while synthetic vesicles and beads with diameter of 20 nm do not. Data shown with sd. Figure 3. Open in new tabDownload slide A, Line profiles through the phragmoplast of cells, injected either with vesicles (150 nm) or polystyrene beads (40 nm). The average fluorescence intensity of each horizontal row of pixels was plotted versus its vertical position. The fluorescence of vesicles is highest in the cell plate region (1.5–2.0 μm), while fluorescence of beads is lowest in the cell plate region. Polystyrene beads do not accumulate in the cell plate region, while vesicles accumulate there specifically. The same was observed for any size of vesicles or beads. B, Duration of cell plate formation in control noninjected cells and cells injected with vesicles of 45 or 150 nm and 20- or 40-nm polystyrene beads. Polystyrene beads of 40 nm delay cell plate formation, while synthetic vesicles and beads with diameter of 20 nm do not. Data shown with sd. ). Similar to the vesicles of 60 nm that we injected before (Esseling-Ozdoba et al., 2008b), the vesicles of different sizes also only accumulated and did not fuse with the cell plate. This was deduced from the observations that the endogenous FM4-64-stained cell plate is thinner than the synthetic beads-labeled plate and that at the time point at which the cell plate membrane fuses with the parental plasma membrane when two separate cells have been formed, the injected vesicles redistributed away from the cell plate into the two daughter cells, while the FM4-64-stained endogenous cell plate membrane stayed intact (Esseling-Ozdoba et al., 2008b). Polystyrene Beads of 20 and 40 nm Diameter Enter the Phragmoplast But Are Not Released from it into the Cell Plate Region, and Beads of 40 nm Diameter Block the Transport of Endogenous Vesicles Fluorescent polystyrene beads with sizes of 20 and 40 nm diameter, which are much smaller than endogenous vesicles and even smaller than the smallest injected DOPG vesicles, entered the phragmoplast but did not traverse it. Their accumulation in the plane of the developing cell plate was never observed (Figs. 2B and 3A; Table II Table II. Polystyrene beads (20 and 40 nm) not coated and coated with DOPG do not accumulate in the cell plate region Injected Polystyrene Beads . Percentage of Cells without Beads Accumulation . 20 nm 100 (n = 6) 40 nm 100 (n = 8) Beads (40 nm) coated with DOPG 100 (n = 5) Injected Polystyrene Beads . Percentage of Cells without Beads Accumulation . 20 nm 100 (n = 6) 40 nm 100 (n = 8) Beads (40 nm) coated with DOPG 100 (n = 5) Open in new tab Table II. Polystyrene beads (20 and 40 nm) not coated and coated with DOPG do not accumulate in the cell plate region Injected Polystyrene Beads . Percentage of Cells without Beads Accumulation . 20 nm 100 (n = 6) 40 nm 100 (n = 8) Beads (40 nm) coated with DOPG 100 (n = 5) Injected Polystyrene Beads . Percentage of Cells without Beads Accumulation . 20 nm 100 (n = 6) 40 nm 100 (n = 8) Beads (40 nm) coated with DOPG 100 (n = 5) Open in new tab ). They distributed within the phragmoplast (Figs. 2B and 3A). Thus, they did not exit the phragmoplast toward the cell plate. In addition, the polystyrene beads of 40 nm delayed cell plate formation, while the smaller polystyrene beads of 20 nm did not (Fig. 3B), even when the total number of 20-nm beads injected into a cell was 20 times higher than that of the 40-nm beads. Although we have not shown this in a direct way, this suggests that polystyrene beads of 40 nm block the transport of endogenous vesicles to the cell plate. Polystyrene Beads Coated with a DOPG Membrane Do Not Accumulate in the Cell Plate Region To test if the presence of a lipid surface causes DOPG vesicles to enter the plane of the cell plate, while beads are excluded, we coated 40-nm polystyrene beads with DOPG in such a way that their outside surface was the same as that of the synthetic vesicles. The polystyrene beads that we used for microinjection have negatively charged carboxyl groups on their surface. To coat such beads with DOPG, which is also negatively charged, beads first were coated with the positively charged surfactant cetyl trimethyl ammonium bromide (CTAB) and then with DOPG (Fig. 4 Figure 4. Open in new tabDownload slide Characteristics of bead coating. A, The mean radius r of particles and/or aggregates determined by cumulant fitting of dynamic light-scattering results as a function of the CTAB concentration in μ m (white circles) and of the CTAB + DOPG concentration in μ m (black circles), where C CTAB = 100 μ m and C DOPG = C ™ C CTAB. Adding more CTAB improves the colloidal stability caused by the adsorption of additional surfactant or DOPG molecules to the CTAB-coated polystyrene beads, resulting in the formation of a densely packed monolayer, where the hydrophilic groups are directed to the aqueous solution. B, The size distribution of the beads (continuous line), the beads in a 100 μ m CTAB solution (long dashed line), and the beads in a solution of 100 μ m CTAB and 400 μ m DOPG (short dashed line). The size of beads did not change significantly upon coating with DOPG. These results confirm the formation of a densely packed lipid/surfactant monolayer on the outside of the CTAB-coated polystyrene beads. Figure 4. Open in new tabDownload slide Characteristics of bead coating. A, The mean radius r of particles and/or aggregates determined by cumulant fitting of dynamic light-scattering results as a function of the CTAB concentration in μ m (white circles) and of the CTAB + DOPG concentration in μ m (black circles), where C CTAB = 100 μ m and C DOPG = C ™ C CTAB. Adding more CTAB improves the colloidal stability caused by the adsorption of additional surfactant or DOPG molecules to the CTAB-coated polystyrene beads, resulting in the formation of a densely packed monolayer, where the hydrophilic groups are directed to the aqueous solution. B, The size distribution of the beads (continuous line), the beads in a 100 μ m CTAB solution (long dashed line), and the beads in a solution of 100 μ m CTAB and 400 μ m DOPG (short dashed line). The size of beads did not change significantly upon coating with DOPG. These results confirm the formation of a densely packed lipid/surfactant monolayer on the outside of the CTAB-coated polystyrene beads. ). The adsorption of DOPG on CTAB-coated beads is entirely driven by hydrophobic interaction, resulting in the formation of a surfactant/lipid monolayer on the outside of the beads. In this monolayer, the tails are oriented toward the bead and the hydrophilic heads are directed outward, such that the outer monolayer resembles the outside of a normal DOPG layer. The coating with DOPG did not significantly change the diameter of the beads (Fig. 5A Figure 5. Open in new tabDownload slide Characteristics of the lipid DOPG layer around the beads. A, The volume fraction (φ) profile of a tensionless bilayer consisting of DOPG lipids. The z axis is given in lattice layers. The ionic strength is shown by φ z = 1.0 × 10™4 (corresponding to approximately 5 mm). B, The volume fraction profile of DOPG and CTAB in the presence of a polystyrene surface. The polystyrene surface is located at z = 0. The outer part of an adsorbed CTAB/DOPG layer on the negatively charged polystyrene beads corresponds to the outside of a DOPG vesicle. DOPC, 1,2-Dioleoyl-sn-glycero-3-phosphocholine. Figure 5. Open in new tabDownload slide Characteristics of the lipid DOPG layer around the beads. A, The volume fraction (φ) profile of a tensionless bilayer consisting of DOPG lipids. The z axis is given in lattice layers. The ionic strength is shown by φ z = 1.0 × 10™4 (corresponding to approximately 5 mm). B, The volume fraction profile of DOPG and CTAB in the presence of a polystyrene surface. The polystyrene surface is located at z = 0. The outer part of an adsorbed CTAB/DOPG layer on the negatively charged polystyrene beads corresponds to the outside of a DOPG vesicle. DOPC, 1,2-Dioleoyl-sn-glycero-3-phosphocholine. ), and the layer of DOPG on the surface of the beads had the same biophysical properties as the bilayer of DOPG vesicles (Fig. 5B). Injected DOPG-coated polystyrene beads into dividing cells at anaphase behaved like 40-nm uncoated beads: they entered the phragmoplast but did not accumulate in the plane of the cell plate (Fig. 6 Figure 6. Open in new tabDownload slide Polystyrene beads coated with DOPG do not accumulate in the cell plate region of T. virginiana stamen hair cell injected at anaphase with polystyrene beads of 40 nm coated with DOPG. Polystyrene beads coated with DOPG were observed in the cytoplasm and the phragmoplast but did not accumulate in the cell plate region (white arrowheads) at 30 min after injection. Bar = 10 μm. Figure 6. Open in new tabDownload slide Polystyrene beads coated with DOPG do not accumulate in the cell plate region of T. virginiana stamen hair cell injected at anaphase with polystyrene beads of 40 nm coated with DOPG. Polystyrene beads coated with DOPG were observed in the cytoplasm and the phragmoplast but did not accumulate in the cell plate region (white arrowheads) at 30 min after injection. Bar = 10 μm. ; Table II). These results show that it is their stiffness that inhibits transit through/release from the phragmoplast and entry into the plane of the cell plate, and not their surface properties. DISCUSSION During plant cytokinesis, the phragmoplast inhibits the transport of micrometer-sized organelles (e.g. mitochondria, Golgi bodies, and plastids) toward the cell plate region but allows the transport of cell plate-forming vesicles (approximately 60 nm in diameter; Seguiݩ-Simarro et al., 2004). Using microinjection of fluorescent synthetic (DOPG) lipid vesicles, fluorescent polystyrene beads, and the same beads coated with DOPG, we show that the phragmoplast is nonselective for synthetic lipid vesicles, tested up to 150 nm diameter, but is a barrier for stiff polystyrene beads of 20 and 40 nm diameter to exit from the phragmoplast or enter into the cell plate region. What is particularly interesting is the finding that the 40-nm beads slow cell plate formation. We conclude that flexibility/stiffness is an important parameter for vesicle exit from the phragmoplast or entry into the cell plate region and that occluding the phragmoplast with stiff structures inhibits its function. What we show is that physical aspects determine which structures can pass through a cytoskeleton region, indicating that a mutation that would make the content of the Golgi vesicles passing the phragmoplast, or their membrane, more stiff would cause a cytokinesis problem. Are Vesicles Squeezed through the Phragmoplast, and Where Is the Bead Barrier? Microtubules are thought to be the transport vehicles for motor molecule-coated endogenous vesicles during cell plate formation (Otegui et al., 2001; Smith, 2002). The largest injected vesicles (150 nm in diameter) were transported through the phragmoplast similarly to the smallest vesicles (45 nm). This is an interesting observation, taking into account the dense arrays of microtubules, actin filaments, and ER membranes in the phragmoplast, in which the measured distance between the phragmoplast microtubules (20–100 nm; Thijsen et al., 1998; Seguiݩ-Simarro et al., 2004; Austin et al., 2005) is smaller than the diameter of these vesicles. Are the vesicles squeezed through the phragmoplast? The force for pushing/pulling these vesicles through the phragmoplast is possibly created by motor proteins that are attached to the surface of injected vesicles. The motor proteins could pull the vesicular membrane along the phragmoplast cytoskeleton, deforming the vesicle to go through the phragmoplast. This would be comparable to the results from in vitro experiments shown by Koster et al. (2003), where kinesin motor proteins could pull thin tubes from phospholipid (1,2-dioleoyl-sn-glycero-3-phosphocholine) vesicles along microtubules. An alternative for squeezing the vesicles is buckling of microtubules and actin filaments, thus making way for bigger vesicles. Individual microtubules (Dogterom and Yurke, 1997; Janson and Dogterom, 2004) and actin filaments (Kovar and Pollard, 2004) can buckle in vitro. We do not know whether the injected vesicles and beads are coated with motor proteins and actively moved or whether they are dragged by hydrodynamic flow created by motor-driven transport of endogenous vesicles (Houtman et al., 2007; Esseling-Ozdoba et al., 2008a). Where exactly is the physical barrier excluding the beads from the plane of the cell plate? Is it a phragmoplast transit barrier or a phragmoplast exit barrier? It is not a phragmoplast entry barrier. If the actin/microtubule cytoskeleton becomes more and more finely structured toward the cell plate, the phragmoplast cytoskeleton could be a physical transit barrier, acting at the side of the growing cell plate. A splaying out into finer and finer actin filament bundles is seen in the subapex of growing root hairs (Miller et al., 1999, Ketelaar et al., 2003) and is induced in legume root hairs by bacterial signal molecules that promote cell elongation by first promoting fine filamentous actin (de Ruijter et al., 1999). This could be the case for the phragmoplast cytoskeleton as well and needs higher resolution studies. However, the distribution of the beads, filling the whole accessible volume of the phragmoplast, without a visible accumulation gradient, does not favor this possibility. The results, therefore, suggest that there is a phragmoplast exit barrier. The so-called cell plate assembly matrix of unidentified molecular makeup (Seguiݩ-Simarro et al., 2004) could be (part of) this selective barrier. Synthetic Lipid (DOPG) Vesicles and Polystyrene Beads: Tools to Study Transport through and Properties of Cell Areas with Different Cytoskeletal Configurations Several types of in vivo cell biological questions become amenable to experimentation with our approach. If motor molecules indeed attach to the vesicles and beads (Romagnoli et al., 2003), bead injection studies in higher plant cells can reveal which motor proteins are involved in organelle movement through cytoplasmic strands and vesicular movement through the phragmoplast. A second type of question that could be solved using experiments with injected beads and synthetic vesicles of various sizes pertains to the physical properties of cell areas with specific cytoskeleton configurations in plant as well as animal cells. Injection of beads and synthetic vesicles in growing root hairs with their dense subapical fine F-actin in pollen tubes of lily (Lilium longiflorum; Geitmann et al., 2000) and root hairs of Vicia sativa (Miller et al., 1999) and Arabidopsis (Arabidopsis thaliana; Ketelaar et al., 2003), and a similar configuration with in addition a dense network of endoplasmic microtubules in growing Medicago truncatula root hairs (Sieberer et al., 2002), will elucidate physical properties of these cytoskeleton configurations. In addition, specific coating of vesicles with a protein prior to injection, or insertion of specific proteins into the vesicle membrane, could help to reveal the function of these proteins in various cell processes, notably cell plate formation. In interphase cells of T. virginiana stamen hair cells, microinjected synthetic lipid vesicles of 60 nm (as shown before; Esseling-Ozdoba et al., 2008b) and polystyrene beads of 40 nm move comparably to organelles (Fig. 1), although a bit slower, which makes them excellent tools to study transport processes in these cells. The movement of synthetic lipid vesicles and polystyrene beads might be an active process caused by cytoplasmic motor proteins that possibly attach to their surfaces after injection into the cells. This is in line with results obtained with uncoated polystyrene beads introduced into internodal cells of the green algae Chara (Chaen et al., 1995) and BSC-1 cells, a continuous renal epithelial cell line derived from the African green monkey (Beckerle, 1984). In these studies, it has been shown that polystyrene beads move in the cytoplasm along the cytoskeleton as a result of the attachment of cytoplasmic motor proteins to the bead surface. If motor molecules indeed attach to the beads and vesicles, similar studies in higher plant cells can reveal which motor proteins are involved in organelle movement through cytoplasmic strands and vesicular movement through the phragmoplast. It cannot be excluded at the moment, however, that the injected beads and synthetic vesicles are moved by hydrodynamic flow caused by the active transport of endogenous organelles, which we have shown to be expected theoretically (Houtman et al., 2007) and experimentally to be the case for free GFP in the cytoplasm (Esseling-Ozdoba et al., 2008a). Using experiments with injected beads and synthetic vesicles of various sizes into living cells, for instance with specific coating of vesicles with a protein prior to injection or insertion of specific proteins into the vesicle membrane, would combine the two types of approaches and help to reveal the functions of these proteins in various cell processes, notably cell plate formation. MATERIALS AND METHODS Plant Material Tradescantia virginiana plants were grown in a growth chamber with a 16-h photoperiod at 25°C and an 8-h dark period at 18°C and 75% to 80% relative humidity. Stamen hair cells with dividing and elongating cells in the apical region were collected from immature, unopened flower buds with a length of approximately 5 mm. For microinjection experiments, we dissected and immobilized 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) and Esseling-Ozdoba et al. (2008a, 2008b). Preparation of Synthetic Vesicles, Polystyrene Beads, and Dextran Synthetic vesicles were made from 98% anionic nonfluorescent phospholipid (DOPG; Avanti Polar Lipids) and 2% fluorescent phosphocholine BodipyFC12-HPC (excitation maximum at 503 nm, emission maximum at 512 nm; Molecular Probes) as described by Esseling-Ozdoba et al. (2008b). Vesicles of various sizes were made with a miniextruder using a polycarbonate membrane of different pore sizes: 30 nm (45-nm vesicles), 50 nm (60-nm vesicles), 80 nm (75-nm vesicles), and 400 nm (150-nm vesicles). The size of the vesicles after preparation was measured with dynamic light scattering. Polystyrene beads of 40 nm, carboxylated-modified FluoroSpheres (Fluorescent Microspheres), were purchased from Molecular Probes (F10720). We used yellow-green beads with an excitation maximum at 505 nm and an emission maximum at 515 nm and red beads with an excitation maximum at 580 nm and an emission maximum at 605 nm. The solution contained 5% (v/v) beads (1.4 × 1015 beads mL™1) without preservatives. For microinjection, polystyrene beads were diluted 1:500 in microinjection buffer (5 mm HEPES, 0.1 mm KCl, pH 7.0). With a scanning electron microscope and dynamic light scattering, we observed that the beads did not cluster in microinjection buffer. Because the beads solution contained small numbers of larger beads (95 nm, 2%–5%), the solution was sonicated (2 × 30 min) and centrifuged at 25,000g for another 30 min before microinjections to remove the large beads that could block the needle during microinjection. After centrifugation, the supernatant was sonicated for 30 min in order to have beads well dispersed in the solution. Besides this, we added 1% bovine serum albumin (albumin fraction V; Merck) to the bead solution to prevent clustering of the beads and possible needle blockage during microinjection. Bovine serum albumin in this concentration did not disturb cell plate formation. Small polystyrene beads were purchased from Micromod Partikeltechnologie (Micromer, GreenF). These beads were also carboxylate modified. The solution contained 5.5 × 1015 beads mL™1 without any preservatives. Although they were described by the manufacturer as beads of 15 nm, most of the beads (95%; scanning electron microscopy data) in the solution appeared to have a size of 20 nm. For this reason, we call these beads 20-nm polystyrene beads. For microinjection, polystyrene beads were diluted 1:100 in microinjection buffer. The solution used for microinjection contained 5.5 × 1013 beads mL™1. Before the beads were injected into the cells, they were sonicated (2 × 30 min) and 1% bovine serum albumin was added to prevent clustering of the beads during microinjection. For coinjection experiments, vesicles or polystyrene beads were mixed with fluorescent dextran (0.5 mg mL™1; Alexa 568-dextran, 10 kD; Molecular Probes, Invitrogen). For coinjection experiments with dextran, yellow-green beads were used. Until microinjection experiments, vesicles and polystyrene beads were stored on ice. Coating of Polystyrene Beads with DOPG Membrane Negatively charged (carboxylated) polystyrene beads were first coated with the positively charged surfactant CTAB, in concentration of 100 μ m, and then with the negatively charged DOPG, in concentration of 400 μ m. CTAB was dissolved in microinjection buffer as 1 mm stock. DOPG vesicles of 200 nm were prepared (by sonication) in microinjection buffer, with DOPG concentration of 4 mm. The coating of polystyrene beads was done by mixing the beads with CTAB and DOPG and sonicating for 10 min with a bath sonicator (Laboratory Supplies). The quality of the coating was checked with dynamic light scattering. To gain more insight into the molecular organization of the adsorbed layer, we performed a number of calculations using a molecularly realistic self-consistent field model. We will explain briefly how the model works; interested readers can refer to the literature for full details (Evers et al., 1990; Meijer et al., 1999; Kik et al., 2005). The self-consistent field method is based on the reduction of the many-molecule problem (i.e. each molecule in a system has interactions with all other molecules) to the problem of one molecule in the field of mean force of all the others. In general, this field depends on the distributions and conformations of all molecules and their interactions. In the model used here, the molecules are divided into segments (“united atoms,” for example, a CH2 or CH3 group forms a segment C), and these segments are placed on a lattice (one segment per lattice site). The segment-type-dependent potential fields depend on the way in which the segments are distributed over the lattice. These potentials have contributions due to the short-range interactions between the segments, the electrostatic interactions, and a contribution linked to packing constraints. If a surface is present in the system, the segment-specific interactions with this surface are also included in the potential fields. The equilibrium state of a system is obtained by determining the distribution corresponding to the minimal mean field free energy in an iterative way. We routinely find this distribution with a very high accuracy by a numerical algorithm. In the calculations discussed below, we consider the structure of a free lipid bilayer and the adsorbed CTAB/DOPG layer on the polystyrene beads in one direction only, namely the direction perpendicular to the layers. This enables us to reduce the problem to a one-dimensional one (using a one-dimensional lattice), which saves a lot of calculation time. It implies that lateral structural fluctuations in the layers are averaged out. The polystyrene surface is taken to be hydrophobic, and 10% of this surface is covered with carboxyl groups, which are partly uncharged and partly negatively charged, depending on the pH of the solution. In the calculations, the pKa of the COOH groups was taken to be 4.5 and a pH value of 7 was used. There are three molecular species in the system that consist of just one segment, namely water (W), of which a small part occurs as a positive species (H3O+) or a negative one (hydroxyl) depending on the pH of the solution and the local electrostatic potential. In addition, there are two small salt ions, a positively charged ion (K) and a negatively charged ion (S). The surfactant CTAB is modeled as a molecule consisting of 19 segments, X3C16. The hydrophilic head group segments X each carry a charge of 1/3, and the tail is composed of 16 hydrophobic segments (united atoms, C). The negatively charged lipid N3C3P3(C18)2, which represents DOPG, consists of two hydrophobic tails, each having 18 C segments connected through a side chain to a head group that consists of nine united atoms: three negatively charged hydrophilic segments (P), each having a partial charge of ™1/3, three hydrophobic segments (C), and three uncharged hydrophilic segments (N). The short-range interactions between the segments are accounted for by so-called Flory-Huggins nearest neighbor interaction parameters. These parameters have been chosen in such a way that the critical micellization concentration values of CTAB and DOPG in the model closely agree with experimental data. The electrostatic potential depends on the local charge density and follows from the solution of the Poisson equation. Microinjection The microinjection experiments were conducted according to the detailed description of the process and equipment published by Vos et al. (1999) and for vesicles described by Esseling-Ozdoba et al. (2008b). Microscopy, Imaging, and Data Analysis Microinjections were performed on inverted microscopes. Images were collected with a MRC600 confocal laser-scanning unit (Bio-Rad) coupled to a Nikon Labophot microscope, with a Cell Map IC confocal laser-scanning unit (Bio-Rad) coupled to a Nikon Eclipse TE 2000-S microscope, or with a LSM 5 Pascal confocal laser-scanning unit coupled to a Zeiss Axiovert 200 microscope. For imaging vesicles or beads coinjected with dextran (Bodipy/yellow-green beads/Alexa-568 dextran dual scanning), we used excitation/emission combinations of 488 nm/520 to 540 (Cell Map IC) or 488 nm/BP 505 to 550 with the HFT 488 primary and NFT 545 secondary dichroic mirrors (LSM 5 Pascal) for Bodipy and 532 nm/560 LP (Cell Map IC) or 543 nm/LP 560 with the HFT 543 primary dichroic mirror (LSM 5 Pascal) for Alexa-568 dextran. For imaging of yellow-green or red beads with the MRC600, neutral density filters were set to obtain 1% transmission intensity from the laser beam, using the 488-nm wavelength for yellow-green beads with DM 488 and BA 522 and the 568-nm wavelength for red fluorescence of beads with DM 560 long-pass BA 585. Images were obtained with 1.4 numerical aperture 60× and 1.4 numerical aperture 63× oil-immersion objectives collected by Kalman averaging of two to three full scans (MRC600, Cell Map IC) or with scan speed 7 (LSM 5 Pascal). Images were taken in 2- or 3-min intervals. This choice of intervals allowed observation of the developing cell plate for a long period of time without disturbing the cell plate formation process. Images were acquired and processed with software programs including Comos (MRC600), LaserSharp 2000 (Cell Map IC), LSM 5 Pascal (version 3.5 SP1.1; Carl Zeiss), Confocal Assistant 4.02 (Todd Clark Brelje), and Adobe Photoshop 5.0 and 8.0 (Adobe Systems). 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[OA] Open Access articles can be viewed online without a subscription. www.plantphysiol.org/cgi/doi/10.1104/pp.109.150417 © 2010 American Society of Plant Biologists © The Author(s) 2010. Published by Oxford University Press on behalf of American Society of Plant Biologists. This is an Open Access article distributed under the terms of the Creative Commons Attribution License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited.
TI - Flexibility contra Stiffness: The Phragmoplast as a Physical Barrier for Beads But Not for Vesicles
JF - Plant Physiology
DO - 10.1104/pp.109.150417
DA - 2010-02-03
UR - https://www.deepdyve.com/lp/oxford-university-press/flexibility-contra-stiffness-the-phragmoplast-as-a-physical-barrier-Ir8iOA8Isq
SP - 1065
EP - 1072
VL - 152
IS - 2
DP - DeepDyve
ER -