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Plant Cell Physiol. 49(10): 1508–1521 (2008) doi:10.1093/pcp/pcn122, available online at www.pcp.oxfordjournals.org ß The Author 2008. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists. All rights reserved. For permissions, please email: [email protected] FM Dyes Label Sterol-Rich Plasma Membrane Domains and are Internalized Independently of the Cytoskeleton in Characean Internodal Cells Andreas Klima and Ilse Foissner * Department of Cell Biology, Division of Plant Physiology, University of Salzburg, Hellbrunnerstrasse 34, A-5020 Salzburg, Austria We applied the endocytic markers FM1-43, FM4-64 and filipin to internodal cells of the green alga Chara corallina. Both FM dyes stained stable, long-living plasma membrane patches with a diameter of up to 1 mm. After 5 min, FM dyes labeled cortical, trembling structures up to 500 nm in size. After 15 min, FM dyes localized to endoplasmic organelles up to 1 mm in diameter, which migrated actively along actin bundles or participated in cytoplasmic mass streaming. After 30–60 min, FM fluorescence appeared in the membrane of small, endoplasmic vacuoles but not in that of the central vacuole. Some of the FM-labeled organelles were also stained by neutral red and lysotracker yellow, indicative of acidic compartments. Filipin, a sterol-specific marker, likewise labeled plasma membrane domains which co-localized with the FM patches. However, internalization of filipin could not be observed. KCN, cytochalasin D, latrunculin B and oryzalin had no effect on size, shape and distribution of FM- and filipin-labeled plasma membrane domains. Internalization of FM dyes was inhibited by KCN but not by drugs which interfere with the actin or microtubule cytoskeleton. Our data indicate that the plasma membrane of characean internodal cells contains discrete domains which are enriched in sterols and probably correspond to clusters of lipid rafts. The inhibitor experiments suggest that FM uptake is active but independent of actin filaments, actin polymerization and microtubules. The possible function of the sterol-rich, FM labeled plasma membrane areas and the significance of actin-independent FM internalization (via endocytosis or energy-dependent flippases) are discussed. Keywords: Chara — Cytoskeleton — Endocytosis — Lipid raft — Plasma membrane domain — Sterol. Abbreviations: AFW, artificial fresh water; BDM, 2,3butanedione monoxime; CD, cytochalasin D; CLSM, confocal laser scanning microscope/microscopy; DIC, differential interference contrast; DiOC6(3), 3,30 , dihexyloxacarbocyanine iodide; DMSO, dimethylsulfoxide; FRAP, fluorescence recovery after photobleaching; LatB, latrunculin B; PBS, phosphate-buffered saline; PIPES, piperazineN,N0 -bis (2-ethanesulfonic acid). Introduction Endocytosis is the process by which cells internalize plasma membrane and extracellular material. It is important for the recycling of plasma membrane components, for nutrient uptake and for signaling (for recent reviews, see Geldner 2004, Murphy et al. 2005, Samaj et al. 2005, Polo and Di Fiore 2006, Samaj et al. 2006a, Samaj et al. 2006b). A variety of endocytic pathways have been described in animal and fungal cells which differ mainly in the participation of proteins and in the size of endocytic vesicles (Conner and Schmid 2003, Soldati and Schliwa 2006). Until a few years ago, little was known about endocytosis in plant cells. Although earlier work proved uptake of electron-opaque markers from the external medium (Hawes et al. 1991), the existence of endocytosis in plant cells was even questioned because of the high turgor pressure (references in Marcote et al. 2000, Aniento and Robinson 2005). This changed rapidly with the introduction of FM dyes and filipin as endocytic tracers and with the development of new molecular biological techniques (Betz et al. 1996, Grebe et al. 2003, Meckel et al. 2004). Today, the existence of endocytotic processes in plant cells is beyond debate, but our understanding of plasma membrane internalization and uptake of extracellular material is still very limited. Currently, research focuses on clathrin-dependent endocytic pathways and clathrinindependent mechanisms via lipid rafts (Grebe et al. 2003, Geldner 2004, Aniento and Robinson 2005, Murphy et al. 2005, Samaj et al. 2005, Ovecka and Lichtscheidl 2006, Verma et al. 2006). In this study, we wanted to investigate constitutive endocytosis in mature, non-elongating internodal cells of characean algae which are considered to be close relatives of higher plants (Graham and Kaneko 1991, Tanabe et al. 2005). We were especially interested in the role of the actin cytoskeleton which was reported to be involved in plasma membrane internalization in various types of cells including those of higher plants (Engqvist-Goldstein and Drubin 2003, Baluska et al. 2004, Ayscough 2005, Ovecka et al. 2005, Voigt et al. 2005, Samaj et al. 2006b, Sun et al. 2006). The characean internodes are well known for their rapid *Corresponding author: E-mail, [email protected]; Fax, þ43-662-8044-619. 1508 FM internalization in characean algae cytoplasmic streaming which depends on interaction of myosin-coated organelles with parallel, subcortical actin bundles (Grolig and Pierson 2000, Shimmen and Yokota 2004) and are a valuable experimental tool for investigating the plant cytoskeleton (Foissner and Wasteneys 2000). In our study, we applied FM1-43 and FM4-64 which are widely used as fluorescent markers of endocytosis in animal, fungal and plant cells (Betz et al. 1996, FischerParton et al. 2000, Bolte et al. 2004, Aniento and Robinson 2005, Samaj 2006). It has also been suggested that FM internalization is due to the activity of energy-dependent flippases (Fischer-Parton et al. 2000) or mechanosensitive cation channels (Nishikawa and Sasaki 1996), and that the differential distribution of FM dyes to organelle membranes is due to a lipid sorting mechanism. An argument against endocytosis-independent uptake is, however, that FM dyes injected into growing pollen tubes and other cells became homogeneously distributed and did not localize to specific organelles (Parton et al. 2003, Van Gisbergen et al. 2007). This observation and recent data obtained by fluorescence resonance energy transfer (FRET) analysis (Johnson and Griffing 2007) also point to FM uptake via endocytic vesicles. In plant cells, the FM dyes insert into the plasma membrane and gradually appear in compartments where they partially co-localize with endosomal (Ueda et al. 2004), pre-vacuolar (Tse et al. 2004), trans-Golgi (Dettmer et al. 2006) and vacuolar marker proteins (Kutsuna and Hasezawa 2002). Therefore, irrespective of the uptake mechanism, the FM dyes are at least suited for monitoring the distribution and dynamics of endocytic organelles. We also applied filipin which was used to follow steroldependent endocytosis in higher plant cells (Grebe et al. 2003, Ovecka and Lichtscheidl 2006). We reported previously at a meeting that FM dyes label distinct plasma membrane areas in characean internodal cells and that FM internalization cannot be inhibited by cytochalasin D (CD; Foissner and Klima 2008). Here we prove that the FM-stained plasma membrane areas are enriched in sterols and that their distribution is independent of the cytoskeleton. Observations in cytoplasmic droplets, additional inhibitor experiments and quantification of FM internalization in inhibitor-treated and control cells provide further evidence that constitutive endocytosis, or at least FM internalization and transport of FM fluorescent organelles in characean internodes, is not only independent of an intact actin (and microtubule) cytoskeleton but also independent of actin polymerization. Results The staining results for FM1-43 and FM4-64 were similar, and co-labeling with both dyes resulted in identical images (Fig. 1A–C, see below). However, FM1-43 was 1509 preferred because of its brighter fluorescence and its green emission which is easier to discriminate from the red autofluorescence of chloroplasts. Furthermore, FM1-43 could be applied for several days without any obvious harmful effect, whereas cells treated with the same concentration of FM4-64 died after 1 d. Heterogeneous labeling of the plasma membrane by FM dyes and filipin The plasma membrane of various cell types have been reported to be homogeneously labeled by FM dyes (Samaj et al. 2006a) and filipin (Grebe et al. 2003, Ovecka and Lichtscheidl 2006). In internodal cells of Chara corallina, the periphery of the cytoplasm was heterogeneously stained. Bright patches became visible within 5 min of treatment with FM dyes and alternated with weakly fluorescent regions (Fig. 1A–D). Most of these patches were disc-like (Fig. 1A–C), but elongate (Fig. 1D) or bowl-shaped structures (not shown) were also observed. Optical crosssections through Z-series revealed that some of these patches protruded slightly into the cell interior (Fig. 1E). The FM-stained plasma membrane domains were present along the whole cytoplasmic surface except for the cross-walls between adjacent cells (Fig. 1F, G). They were also largely absent from the neutral line which separates regions of oppositely streaming endoplasm (Fig. 1H) and from those regions where the chloroplast surface abutted the plasma membrane (asterisk in Fig. 1A). The number and (numerical) density of the FM-stained patches varied greatly between cells. The highest densities were found in internodes of the branchlets, with up to 200 disc-shaped patches per 100 mm2 (Fig. 1A–C). There was also considerable variability in density of FM-stained patches along the surface of individual internodes (not shown). In contrast to these spatial variations, both the size and distribution of the FM-stained patches were rather stable. Although we investigated several dozens of cells over time periods up to 3 h, detachment, migration or gross shape changes of patches were never observed. The distribution of the FM patches was similar in elongating (not shown) and in mature internodal cells and therefore independent of the orientation of cortical actin filaments or microtubules, which are transverse in elongating and random in mature cells (Wasteneys and Williamson 1987). There was also no correlation between the distribution of FM patches and that of the subcortical actin bundles. For instance, FM patches were absent from the neutral line which lacks subcortical bundles, but they were also absent from the cross-walls where the subcortical actin bundles are in close contact with the plasma membrane (see above). Neither CD nor oryzalin affected the pattern of FM fluorescence at the cell surface (Fig. 1I–K), suggesting that 1510 FM internalization in characean algae Fig. 1 Heterogeneous labeling of the plasma membrane by FM dyes and filipin, and comparison with the distribution of cortical organelles in internodes of Chara corallina. (A–C) Double staining of the plasma membrane with FM4-64 (A), FM1-43 (B) and merged image (C). The asterisk in A indicates the position of an underlying chloroplast. (D) Spot-like and elongate, branched structures after staining with FM1-43. (E) Optical cross-section through the plasma membrane (stack of four single sections). The FM-stained patches alternate with weakly fluorescent areas and occasionally project into the cytoplasm (arrow). The plasma membrane appears up to 1 mm thick due to an optical slice thickness of 550 nm (pinhole size 1.4 Airy units) and imperfect optical cross-sectioning. (F, G) Oblique view on the cross-wall between a nodal (N) and internodal cell (I). Note the absence of plasma membrane patches (arrows in the fluorescence image F and the corresponding bright field image G). The asterisk in G indicates a chloroplast. (H) Plasma membrane patches stained by FM1-43 are less abundant at the neutral line (NL). (I–K) FM1-43-stained patches are stable and are not affected by inhibitors of the cytoskeleton. Plasma membrane before addition of inhibitors (I), after 30 min in 5 mM cytochalasin C (CD; J) and after another 30 min in 5 mM CD and 10 mM oryzalin (K). (L) FM1-43-stained plasma membrane patches (green) are clearly different from cortical mitochondria visualized by mitotracker orange (red). (M, N) Secretory vesicles that migrated towards the plasma membrane after local UV irradiation are seen between and at the surface of cortical chloroplasts at the left side of the DIC image (M, arrow) and are not labeled in the corresponding FM1-43 image (N, arrow indicates the same area as in M). (O, P) FM patches co-localize with sterol-rich plasma membrane areas. The internodal cell was simultaneously labeled with FM1-43 (O) and filipin (P). All images were captured within 30 min after pulse labeling. Bar in E is 4 mm (E), 8 mm (F, G, I–K, M, N), 10 mm (A–D, L, O, P) and 15 mm (H). the morphology and distribution of the FM patches are independent of the cytoskeleton. The plasma membrane remained fluorescent for several hours even when dyes were washed out after 10 min incubation. The staining patterns of pulse-labeled cells were therefore similar to those of cells permanently immersed in dye-containing artificial fresh water (AFW), and the patches occasionally remained visible until the day after pulse labeling. FRAP (fluorescence recovery after photobleaching) experiments should clarify whether the FM internalization in characean algae permanent labeling of patches was due to consistent recycling of labeled plasma membrane, but laser intensities sufficient for bleaching induced a wound response, i.e. the rapid deposition of cell wall material and the deposition of a new, evenly stained plasma membrane (to be described elsewhere). Therefore, we can neither prove nor exclude this possibility. The persistent staining of the plasma membrane could also be due to the storage of FM dye within the cell wall, as suggested for stomatal guard cells (Meckel et al. 2004). In order to prove that the peripheral FM patches were indeed localized to the plasma membrane and were not due to staining of cortical organelles or heterogeneous labeling of the cell wall, we performed a number of experiments. The cortical cytoplasm of characean internodal cells is dominated by files of stationary chloroplasts but also harbors other organelles, such as cortical endoplasmic reticulum, mitochondria, secretory vesicles and peroxisomes. The reticulate, 3,30 -dihexyloxacarbocyanine iodide [DiOC6(3)]stained cortical endoplasmic reticulum was clearly distinct from the mostly punctate FM pattern (Supplementary Fig. S1A). Co-labeling of cells with FM1-43 and mitotracker orange revealed considerable differences between the distribution of the FM patches and that of mitochondria as long as the observation time did not exceed 2 h (Fig. 1L). In pollen tubes, FM dyes are rapidly transferred to secretory vesicles which accumulate at the growing tip (see, for example, Hörmanseder et al. 2005). In Chara, the accumulation of Golgi-derived secretory vesicles can be induced by mechanical damage or local irradiation with UV. These vesicles are large enough to be visualized in the light microscope, have characteristic refractile properties and dynamic behavior, and their involvement in wound wall deposition has been documented by electron microscopy and video-enhanced contrast microscopy (Foissner 1988, Foissner et al. 1996). The comparison of the differential interference contrast (DIC) with the FM fluorescence image showed that these organelles were not stained and were mostly much smaller than the FM patches (Fig. 1M, N). Immunofluorescence of peroxisomes with an antibody against catalase revealed roundish organelles with a diameter of up to 2 mm. Their location and distribution were clearly different from that of the FM patches (not shown). Finally, no FM fluorescence was detected in cell walls isolated from FM-stained cells (not shown). Obviously, FM dyes accumulated (or fluoresced stronger) in stable plasma membrane domains, which differed from their surroundings by an altered composition of structural components. Additional support for this view came from the staining pattern obtained with filipin, a dye supposed to be specific for cholesterol in animal cells and sterol-like substances in plant cells (Grebe et al. 2003). Filipin labeled punctate or elongate structures in the 1511 peripheral cytoplasm of live (Fig. 1P) and fixed cells (Supplementary Fig. S1C) which had a similar size and distribution to those of the FM patches. Double staining with FM1-43 and filipin revealed that both dyes labeled the same areas (Fig. 1O, P). Taken together, these data strongly suggest that the FM patches correspond to sterol-rich plasma membrane domains. Time course of FM internalization In contrast to the rather stationary FM patches described above, we sometimes observed considerable dynamics in their periphery (3D environment) and between them (Fig. 2A–C). Tiny structures appeared to detach from or to fuse with the immobile FM patches, while others appeared or disappeared from the focal plane, suggesting movement from or into deeper regions of the cytoplasm. These dynamic FM-stained structures were of diffractionlimited size and only occasionally seen because their visualization requires optimal optical conditions, i.e. superior staining and a thin cell wall free of epiphytes. They may represent the first detectable stages of plasma membrane internalization in characean cells (compare Meckel et al. 2004). Large organelles which formed at and pinched off the plasma membrane as described in root cells or in protoplasts and guard cells after hyperosmotic treatment (Baluska et al. 2002, Meckel et al. 2005) were not observed. A few minutes after dye addition, FM-fluorescent spotlike structures of diffraction-limited size and larger, vesicular organelles with a diameter of up to 500 nm were also detected beneath the plasma membrane (Fig. 2D). The FM-stained organelles were clearly different from cortical mitochondria and performed oscillating, probably Brownian motion in the clefts between the chloroplast files (Fig. 2D–F). Some of them trembled back towards the plasma membrane, moved along straight or curved tracks between the stationary FM patches and then returned back into deeper regions of the cortical cytoplasm (not shown). In the endoplasm, FM-labeled organelles became visible about 15 min after dye addition (Fig. 2G). Most of them participated in cytoplasmic mass streaming which reached up to 60 mm s–1 in our studies (Fig. 2H, right trajectory). Those which were close to the subcortical actin bundles travelled with variable and considerably slower velocity (Fig. 2H, left trajectory; mean ¼ 11. 2 m 4.9 mm s–1; n ¼ 10). Double labeling with mitotracker orange excluded the possibility that the larger FM-stained organelles represent mitochondria which have a similar dynamic behavior (Fig. 2G; Kachar 1985, Foissner 2004). The FM-stained endoplasmic organelles were also different from the tubular inner endoplasmic reticulum (Supplementary Fig. S1B), from peroxisomes and secretory vesicles (see above and section on cytoplasmic droplets below). 1512 FM internalization in characean algae Fig. 2 FM-stained organelles in internodal cells of Chara corallina. Time course of FM internalization (A–L) and co-localization with acidotropic dyes (M–W). (A–C) Dynamics of FM1-43-labeled structures at the plasma membrane. Note dynamic extensions (thick arrows) of the stable FM patches. Weakly fluorescent, minute structures (thin arrows) appear and disappear between the patches. The time series was taken 5 min after pulse labeling; the time interval between images is 4 s. (D) At 10 min after pulse labeling, FM1-43-stained organelles (green) are seen in the cortical cytoplasm underneath the plasma membrane, and are clearly different from mitochondria identified by mitotracker orange (red). (E, F) FM1-43-stained organelles (arrows in E) between cortical chloroplasts perform trembling motions typical for Brownian movement. The time interval between positions of organelles indicated by dots in F is 500 ms. (G) At 20 min after pulse labeling, FM1-43-stained organelles (green) are seen in the endoplasm. Red-labeled organelles are mitochondria. (H) The FM-stained organelles move slowly and with variable velocity along the subcortical actin bundles (left trajectory in H with 500 ms time interval). In deeper regions of the endoplasm, FM-stained organelles participate in continuous mass streaming with a considerably faster velocity of about 60 mm s–1 (right trajectory in H with 500 ms time interval). (I–L) Optical sections through internodal cells about 30 min after pulse labeling with FM1-43. (I, J) The membrane of a small vacuole, which is located at the periphery of the cytoplasm at the chloroplast-free neutral line and protrudes into the large central vacuole (V), is stained with FM1-43 (white arrows in the fluorescence image I and in the DIC image J; the cytoplasm is overexposed). The black arrow indicates the cell wall. (K, L) The tonoplast of the large central vacuole (V; arrow in the DIC image L) is not labeled. The small arrow in the fluorescence image K points to an FM-stained organelle. A file of small chloroplasts typical of elongating cells is visible in L; the black arrow indicates the cell wall. (M–S) Dual labeling with FM1-43 and neutral red. (M O) Cortical organelles are stained by FM1-43 (M) or by neutral red (N). Two of them co-localize (arrows and merged image O). (P–S) The membrane of small vacuoles (asterisks) is stained by FM1-43 (P) and neutral red (Q). Note also organelles at the surface of the vacuoles which are stained by both dyes. R is the merged image, S is the DIC image. (T–U) Dual labeling with FM4-64 (T) and lysotracker yellow (U). Lysotracker yellow accumulates in vacuoles (asterisks) the periphery of which is stained with FM or covered by FM-stained organelles. V is the merged image; W is the DIC image; C ¼ chloroplast. Bar in F is 3 mm (A–C, I–W) and 5 mm (D–H). Characean internodal cells contain a huge central vacuole and numerous smaller vacuoles located at the periphery of the streaming endoplasm (Fig 2I–L). Occasionally, these vacuoles are also found in the stagnant cytoplasm between the chloroplast files where the imaging conditions are better than in deeper regions (Fig. 2P–W). At 30–60 min after dye addition, the membrane of some of these vacuoles became labeled by FM internalization in characean algae 1513 Fig. 3 Cytoplasmic droplets prepared from FM-labeled and unstained internodal cells of Chara corallina. (A and B) Spot-like, fluorescent organelles are present in the cytoplasm isolated from a cell which was stained with FM4-64 for 30 min (A, DIC image; B, fluorescence). Some of them (arrows) associate with an actin bundle, which is coated with numerous, unlabeled secretory vesicles. (C, D) The higher magnification shows that fluorescent organelles have a spherical or disk-like shape (left arrow) or an irregular outline (right arrows). The membrane of the nucleus (N) is not stained (C, DIC image; D, FM4-64 image). (E, F) After 1 d staining, FM dyes also localize to secretory vesicles (arrows) and other organelles. The membrane of the cytoplasmic droplet is of tonoplast origin and does not fluoresce (arrowheads in the DIC image E and the FM1-43 image F). (G, H) A cytoplasmic droplet of an unstained cell squeezed into perfusion solution containing FM4-64 does not internalize the dye (G, DIC; H. fluorescence image taken 15 min after preparation). The asterisk indicates an autofluorescent chloroplast outside the droplet. Bar in H is 2.5 mm (C, D) and 5 mm (all other images). FM dyes (Fig. 2I, J). The fluorescence was probably delivered by punctate FM-labeled organelles which attached to the surface (membrane) of these vacuoles and were often unevenly distributed or appeared clumped (compare Fig. 2P and T). Fusion processes, however, could not be documented, suggesting that these are longlasting processes. The tonoplast of the central vacuole never received FM label (Fig. 2K, L) even when cells were incubated in the dye solution for 24 h (compare Fig. 3E and F). After 2–3 h, FM dyes gradually appeared in all cortical and endoplasmic mitochondria, in secretory vesicles and in various other organelles, except the endoplasmic reticulum and the tonoplast (see below). These results suggest that FM dyes can be used for monitoring endocytosis in Chara internodes as long as the observation time does not exceed 2 h. In order to prove the endosomal character of FM-fluorescent organelles, we applied the acidotropic dyes neutral red and lysotracker yellow. Neutral red stains the membrane of endosomal compartments and accumulates in lytic vacuoles (Beilby and Shepherd 1989, Guttenberger et al. 2000, Dubrovsky et al 2006). In Chara internodal cells, neutral red labeled small organelles in the cortex and in the endoplasm and some of them co-localized with FM1-43-stained particles (Fig. 2M–O). It also stained the membrane of the FMlabeled vacuoles (Fig. 2P–S). A faint pink color indicated that neutral red was trapped inside these vacuoles, but the concentration was not sufficient for documentation. We also applied lysotracker yellow which is another suitable probe for lytic compartments (Guttenberger 2000). It accumulated in vacuoles with FM-stained membranes or in vacuoles which were surrounded by FM-labeled organelles (Fig. 2T–W). Both neutral red and lysotracker yellow became trapped in the large central vacuole although the tonoplast was not labeled by FM (Fig. 2K) or by neutral red (not shown). In order to improve the visualization of endoplasmic membranes and organelles, we used cytoplasmic droplets squeezed out from internodal cells (Kamiya and Kuroda 1957, Jarosch 1961). They contain cortical and endoplasmic organelles as well as dynamic actin filaments which associate into bundles and rotating rings and which have a strong affinity for secretory vesicles (Fig. 3A–H). Observations on cytoplasmic droplets from FM-stained cells confirmed the staining patterns described above. In droplets prepared from internodes 30 min after pulse labeling, some of the FM-stained organelles associated with actin bundles but were clearly different from the FM internalization in characean algae majority of secretory vesicles visualized by DIC (Fig. 3A, B) and from mitochondria identified with mitotracker orange (not shown). The FM fluorescent structures had either a punctate, roughly spherical or disc-like appearance, or an irregular shape. They corresponded to roundish or lobed organelles which were hardly recognizable with DIC microscopy (Fig. 3C, D). No fluorescence was detected in the membrane of the nucleus or that of the endoplasmic reticulum. If cells were squeezed out 1 d after dye addition, FM fluorescence also localized to secretory vesicles (Fig. 3E, F), to mitochondria and to the nuclear membrane (not shown). The membrane of the cytoplasmic droplets is of tonoplast origin (Sakano and Tazawa 1986) and did not fluoresce (Fig. 3E, F), consistent with the observations in intact cells. Neither the tonoplast-derived membrane nor the content of droplets became labeled when the cytoplasm of unstained cells was squeezed into perfusion solution containing FM dyes (Fig. 3G, H). These observations indicate that FM1-43 and FM4-64 are internalized via the plasma membrane and localize to organelles putatively involved in endocytosis as long as the observation time does not exceed 2 h. In contrast, filipinlabeled organelles were not observed in intact cells or in cytoplasmic droplets prepared from filipin-stained internodes, but this is probably due to difficult imaging conditions (Grebe et al. 2003). Constitutive FM internalization is not affected by inhibitors of the cytoskeleton In most fungal, animal and plant cells investigated so far endocytosis has been reported to depend on actin– myosin interaction and/or actin polymerization (see Introduction). In this study, we applied CD, latrunculin B (LatB) and 2,3-butanedione monoxime (BDM) in order to investigate the role of the actin cytoskeleton in FM internalization in characean algae. We also used oryzalin to evaluate a possible contribution of microtubules. CD reversibly reorganizes the cortical actin cytoskeleton of characean internodes into actin rods and rapidly inhibits cytoplasmic streaming without disassembling the subcortical actin bundles (Williamson 1978, Collings et al. 1995; Supplementary Fig. S1D, E). We first treated internodal cells with 5 mM CD for 30 min and subsequently with a solution containing CD and FM1-43. After another 30 min treatment, cells were washed in AFW, fixed, mounted and investigated in the confocal laser scanning microsope (CLSM). FM patches at the plasma membrane were well preserved, and FM-stained organelles were present in the cortex and in the endoplasm. The proportion of fluorescent organelles in the endoplasm of CD-treated internodes was slightly lower than in control cells, but means were not significantly different (Fig. 4). In order to exclude the possibility that fixation caused redistribution of Fluorescence in endoplasm (vol%) 1514 15 10 5 0 + CD − CD Fixed cells − CD + CD Live cells Fig. 4 Effect of cytochalasin D (CD) on FM internalization in internodal cells of Chara corallina. Cells were treated with 5 mM CD for 30 min before addition of FM1-43. After another 30 min, CD-treated and control cells were fixed, mounted and studied in the CLSM or investigated live. Volume of FM-fluorescent organelles is given as a percentage of total volume of endoplasm. Means (SD) are based on Z-series of 16 fixed and four live cells, respectively, and are not significantly different. For further details, see text. plasma membrane-bound FM to cytoplasmic organelles, we also investigated internodes grown under low light conditions. These cells had thin and weakly autofluorescent chloroplasts, a prerequisite for live imaging of FM-labeled organelles in the endoplasm. Internodes treated with 5 mM CD for 30 min and subsequently with CD and FM1-43 contained abundant fluorescent organelles after 30 min. In the fast streaming endoplasm of untreated internodes, individual organelles could not be resolved with conventional CLSM. Therefore, cytoplasmic streaming in control cells was also arrested with CD but only immediately prior to observation. Again, we found no significant effect of the inhibitor on FM internalization in internodal cells (Fig. 4). Latrunculins bind actin monomers and inhibit their polymerization into filaments (Spector et al. 1999). At a concentration of 50 mM, LatB disassembles cortical actin filaments of Chara internodal cells although subcortical bundles remain intact and support cytoplasmic streaming (Supplementary Fig. S1F; compare Foissner and Wasteneys 2007). In combination with CD, latrunculins are most effective in arresting cytoplasmic streaming and depolymerizing cortical actin in characean internodes (Foissner and Wasteneys 2007). However, neither LatB alone (Fig. 5A, B) FM internalization in characean algae 1515 Fig. 5 Inhibitor effects on FM internalization in internodal cells of Chara corallina. (A, B) Cytoplasmic droplets prepared from internodal cells which were treated with 50 nM latrunculin B (LatB) for 30 min and stained with LatB/FM1-43 for 20 min contain numerous FMfluorescent organelles. The asterisk indicates an autofluorescent chloroplast. (C, D) Effect of combined treatment with CD and LatB. The density of fluorescent organelles in the endoplasm of internodes which were treated with 5 mM CD and 50 mM Lat A before and during staining with FM1-43 (30 min each, C) is similar to that observed in control cells (D). (E) FM1-43-fluorescent organelles are present in internodal cells treated with 100 mM 2,3-butanedione monoxime (BDM) for 30 min before and during dye addition. (F–H) FM1-43 internalization is completely inhibited by treatment with 1 mM KCN. FM-stained plasma membrane patches are seen at the left and right side of the image but the endoplasm in the middle of the image is devoid of fluorescent organelles (F). Cytoplasmic droplets from cells treated with KCN 30 min before and 30 min after dye addition show no FM1-43 fluorescence (G, DIC; H, fluorescence image). The asterisk indicates an autofluorescent chloroplast outside a droplet. Bar in H is 5 m (A–D, G, H) and 20 m (E, F). nor the combined treatment with LatB and CD (Fig. 5C, D) prevented the appearance of FM-stained organelles in intact internodes and in cytoplasmic droplets prepared from these cells. BDM is used as a general myosin inhibitor in spite of many contradictory data (e.g. McCurdy 1999). In Chara, concentrations of about 100 mM are required for inhibition of cytoplasmic streaming and organelle movement, and this effect is not reversible (McCurdy 1999, Funaki et al. 2004). Even this concentration did not prevent FM internalization in internodal cells; it caused, however, clumping of FMfluorescent organelles (Fig. 5E) Oryzalin at a concentration of 10 mM disassembles microtubules of characean internodal cells within 30 min (Supplementary Fig. S1G, H; compare Wasteneys and Williamson 1987). Experiments with oryzalin-treated cells ( CD) showed that FM internalization was also independent of microtubules (Supplementary Fig. S1I). FM dyes are assumed to enter the cytoplasm via energy-dependent vesicle-mediated endocytosis (Betz et al. 1996). The unexpected insensitivity of FM internalization in characean cells against cytoskeletal inhibitors could be due to passive uptake (diffusion) independent of cellular metabolism. We therefore applied KCN at a concentration sufficient to arrest ATP-dependent cytoplasmic streaming and found that this treatment completely inhibited the appearance of FM fluorescent organelles in the cytoplasm (Fig. 5F) and in droplets obtained from such cells (Fig. 5G, H). Staining of plasma membrane patches was not affected (Fig. 5F). The same results were obtained when KCN solutions were supplemented by dimethylsulfoxide (DMSO) up to a concentration of 1%, i.e. the plasma membrane became stained but internalization of FM dyes was not observed. These data suggest that FM internalization in unwounded cells of characean algae is active but independent of the cytoskeleton and not mediated by up to 1% DMSO which was used as solvent for dyes and inhibitors in this study. They do, however, not prove that internalization occurs via vesicular transport. Discussion Sterol-rich plasma membrane domains in Chara—clusters of lipid rafts? To our knowledge, characean internodes are the only plant cells in which the plasma membrane is heterogeneously labeled by FM1-43 and FM4-64. Both dyes accumulated or fluoresced more brightly in stable, mostly circular patches with a diameter of up to 1 mm. In this study, we prove that these plasma membrane patches are also labeled by filipin, which is specific for cholesterol and 1516 FM internalization in characean algae sterol-like substances in plants (Robinson and Karnovsky 1980, Grebe et al. 2003), suggesting that the FM-labeled membrane domains are enriched in sterols. In yeast, filipin used at high concentrations was shown to cause clustering of sterols into plasma membrane domains which also colocalized with FM4-64 for unknown reasons (ValdezTaubas and Pelham 2003; see also Robinson and Karnovsky 1980, Miller 1984 for filipin-induced artifacts). In Chara, FM dyes labeled plasma membrane patches in the absence of filipin and, on the other hand, plasma membrane patches were stained by filipin after fixation of cells, which excludes the possibility of a staining artifact. Sterol-like substances that could be stained by filipin (Grebe et al. 2003) have been isolated from various characean species (Patterson et al. 1991). Such lipids are enriched in detergent-resistant plasma membrane microdomains, called lipid rafts (Simons and Van Meer 1988). Typical lipid rafts are small (in the nm range), quite unstable (Pralle et al. 2000, Pike 2006), and can therefore not be resolved in a laser scanning microscope (Grebe et al. 2003). In the Characeae, lipid rafts appear to be clustered into larger, stable aggregates as reported from Saccharomyces and other fungi (Malinska et al. 2004, Alvarez et al. 2007) and from plant cells during pathogen attack (Bhat and Panstruga 2005). Our observations thus prove that sterol-rich domains are indeed present in intact plant cells, consistent with biochemical data (Mongrand et al. 2004, Shahollari et al. 2004, Borner et al. 2005, see also Kirkham and Parton 2005, Martin et al. 2005). Possible function of the plasma membrane domains The fact that the plasma membrane domains of characean internodal cells are labeled by endocytic markers as well as reports about the involvement of lipid rafts in sterol-dependent endocytosis suggests a participation in plasma membrane recycling (Nichols and LippincottSchwartz 2001, Cheng et al. 2006). Indeed, minute FMstained particles have been observed at the periphery of the FM patches, suggesting detachment or fusion of tiny organelles, but these observations may simply reflect the overlap of signals from closely associated, separate organelles or structures. Filipin can be used to follow constitutive recycling of sterols in plant cells (Grebe et al. 2003, Geldner 2004, Murphy et al. 2005, Ovecka and Lichtscheidl 2006). In our study, filipin labeled the plasma membrane domains, but uptake of filipin into the cytoplasm was never observed. Whether this is due to insufficient imaging conditions, to the presence of quenching substances in the cytoplasm or to sterol recycling which excludes the fluorescent marker remains to be investigated. Lipid rafts and sterol-rich plasma membrane domains have also been implicated in actin cytoskeleton organization (Bhat and Panstruga 2005, Alvarez et al. 2007). The comparison of the actin cytoarchitecture with the distribution of the FM patches as well as the inhibitor experiments clearly argue against such a possibility in characean internodal cells. It is possible that the FM- and filipin-stained patches in Chara internodal cells correspond to complex plasma membrane invaginations, known as charasomes. Charasomes also show a non-uniform distribution, are absent from cross-walls and the neutral line and may protrude deeply into the cytoplasm (Barton 1965, Crawley 1965, Franceschi and Lucas 1980, Lucas and Franceschi 1981). They are probably involved in enhanced transport of ions or inorganic carbon (Franceschi and Lucas 1982, Price and Whitecross 1983, Price et al. 1985, Bisson et al. 1991, Chau et al. 1994). Charasomes develop from coated pits, and coated vesicles appear to pinch off their surface (Lucas and Franceschi 1981), which corroborates our findings about a possible detachment of FM-stained organelles from these plasma membrane domains. However, FM patches, although at lower densities, are also present in internodes of the genus Nitella, which lack charasomes (own unpublished observations). Therefore, further research is needed to clarify the identity of FM-labeled, sterol-enriched plasma membrane domains in characean algae. Whatever their function is, the possibility of visualizing these domains in the living state by the rather non-toxic FM dyes offers new perspectives for in vivo observation of plasma membrane organization and metabolism. Actin-independent FM internalization and transport of putative endosomes There is a large body of evidence from yeast and mammalian cells implicating the actin cytoskeleton in several endocytic pathways (see reviews by EngqvistGoldstein and Drubin 2003, Ayscough 2005). In plant cells as well, both actin nucleation and myosin motor activity along actin filaments have been described to be required for plasma membrane internalization (for references, see Baluska et al. 2002, Grebe et al. 2003, Meckel et al. 2004, Ovecka et al. 2005, Voigt et al. 2005, Ovecka and Lichtscheidl 2006, Samaj et al. 2006b). In Chara internodal cells, neither CD, BDM nor LatB arrested uptake of FM dyes and transport of FM-stained organelles, suggesting that both processes are largely independent of an intact actin cytoskeleton, independent of acto-myosin interaction and independent of actin polymerization (actin comets). Experiments with oryzalin excluded a possible participation of microtubules. We cannot fully exclude the possibility that the insensitivity of FM uptake against cytoskeleton inhibitors is due to non-vesicular internalization of the dyes. Our KCN experiments prove that FM1-43 and FM4-64 were actively internalized and did not enter the cytoplasm FM internalization in characean algae by diffusion. However, FM uptake in characean internodes could be mediated by the activity of energy-dependent flippases (Fischer-Parton et al. 2000; see also Introduction). Irrespective of the mechanism for FM internalization, the inhibitor experiments definitely show that transport of FMstained organelles, putative endosomes, from the cortex to the endoplasm requires neither intact actin filaments nor actin polymerization (actin comets). These findings are consistent with the special cytoarchitecture of the nonelongating, mature internodes where cortical and subcortical actin cytoskeleton are widely separated from each other and rarely linked by actin filaments which traverse the chloroplast layer (Foissner and Wasteneys 2000). Consistent with actin-independent motility, FM-stained organelles performed trembling movements within the clefts between the chloroplast files even in non-treated cells, suggesting that Brownian movement is sufficient to account for the transport of putative endosomes from the cortex towards the endoplasm. In the subcortex, however, fluorescent organelles migrated slowly and with variable velocity along tracks that corresponded to the orientation of subcortical actin bundles. These observations indicate that at least some of the FM-stained organelles carry myosin at their surface and are able to move actively along actin filaments if possible and required (compare Kachar 1985, Foissner et al. 1996 for a similar dynamic behavior of other organelles). Actin-independent plasma membrane internalization and transport of FM-stained organelles may be regarded as an adaptation to the special cytoarchitecture of characean internodes. However, there exist several reports about actin-independent endocytosis and transport of endosomes in yeast and mammalian cells (Fujimoto et al. 2000, Gachet and Hyams 2005, Boucrot et al. 2006). The current evidence in support of a role for the actin cytoskeleton in plant endocytosis is circumstantial and is mainly based on drug studies (Samaj et al. 2006b). Endocytosis is usually coupled to exocytosis in order to maintain plasma membrane tension and the surface area of the cell (Murphy et al. 2005, Samaj et al. 2005). In plant cells, transport of exocytotic vesicles towards the plasma membrane is usually actin based. The frequently observed arrest of endocytosis by inhibitors of the actin cytoskeleton may therefore be a secondary effect of disturbed exocytosis. The results of our study and the discovery that in maize root cells PIN proteins may be retrieved from the plasma membrane via a non-actin dependent process (Boutte et al. 2006) suggest that actin-independent plasma membrane internalization is more widespread than thought. However, we cannot exclude the possibility that FM internalization in Chara (and other cells) is not due to endocytosis but to the activity of flippases and ion channels (see Introduction). 1517 FM dyes as endocytic markers in characean algae In characean algae, FM dyes stained the plasma membrane, cytoplasmic organelles and vacuoles with kinetics similar to those reported from various plant and fungal cells (Samaj et al. 2006a). Double staining with other markers and the comparison with electron microscopic images makes it likely that the FM-labeled organelles correspond to various types of endosomes (see below). Preliminary data indicate further that FM-stained organelles can be labeled with antibodies against various endocytic markers (own unpublished data). We found that FM1-43 and FM4-64 are both suited for studying endocytosis, or at least the distribution of endocytic organelles in characean algae (compare Ovecka et al. 2005). After pulse labeling, FM dyes gradually disappear from the plasma membrane of most plant cells and are transported to the tonoplast, where the majority of the dye molecules remain (Parton et al. 2001, Kutsuna and Hasezawa 2002, Ovecka et al. 2005). In the characean algae, the plasma membrane remained fluorescent up to 24 h after pulse labeling just as in stomatal guard cells (Meckel et al. 2004). So far it is not clear whether this is due to rapid recycling of labeled plasma membrane or to dye storage in the cell wall. FM dyes are widely accepted as markers for non-sterol, clathrin-mediated endocytosis, although FM internalization via coated vesicles has not yet been demonstrated due to their small size (Holstein 2002, Bolte et al. 2004, Meckel et al. 2004). On electron micrographs of characean algae, coated pits and coated vesicles are distributed over the plasma membrane of unwounded cells (Pickett-Heaps 1967, Lucas and Franceschi 1981, Pesacreta and Lucas 1984, McLean and Juniper 1986, own unpublished results) and probably contribute to constitutive endocytosis. The identity of the cortical FM-stained organelles, which are obviously larger than coated vesicles, remains to be established. Both clathrin- and lipid raft-mediated pathways in plants are reported to meet at the partially coated reticulum, the equivalent of early endosomes in other cell types (Geldner 2004, Murphy et al. 2005, Samaj et al. 2005). In mature, unwounded characean internodal cells, the partially coated reticulum and organelles involved in late stages of endocytosis, i.e. multivesicular bodies and Golgi bodies, are located within the streaming endoplasm, up to several micrometers away from the plasma membrane where internalization occurs (Pickett-Heaps 1967, Pesacreta and Lucas 1984, Foissner 1988). They may correspond to the lobed and roundish FM-fluorescent organelles described from cytoplasmic droplets. Characean internodal cells contain various vacuoles which differ in size, shape and content (compare Neuhaus and Paris 2006 for vacuoles in higher plants). This diversity 1518 FM internalization in characean algae was also reflected by the staining results. The membrane of small vacuoles located at the inner periphery of the endoplasm and occasionally between the chloroplast files became labeled with FM dyes 30–60 min after dye addition. These vacuoles sequestered the acidotropic dyes neutral red and lysotracker yellow, and probably have lytic functions. Our staining results indicate that the big central vacuole is likewise an acidic compartment, but its membrane, the tonoplast, was never labeled by FM dyes, suggesting that this huge compartment has mainly storage function. Consistent with this observation, Schulte et al. (1994) found that the central vacuole of characean internodes accumulates huge amounts of sugar. Materials and Methods Plant material and culture conditions Shoots of C. corallina Klein ex Willd., em. R.D.W. were grown in a substrate of soil, peat and sand in 10–50 liter aquaria filled with distilled water. The temperature was about 208C, and fluorescent lamps provided a 16/8 h light/dark cycle. Internodal cells of the main axis or the branchlets were harvested 1 d prior to experiments, trimmed of neighboring internodal cells and left overnight in AFW (10–3 M NaCl, 10–4 M KCl, 10–4 M CaCl2). In this study we used mainly mature, non-elongating internodes. Younger, elongating cells with small and weakly fluorescent chloroplasts were preferred for the study of endoplasmic organelles and vacuolar membranes. In order to improve the visualization of FM-stained membranes and organelles, we also prepared cytoplasmic droplets as described in Kamiya and Kuroda (1957) and Jarosch (1961). Long internodal cells were blotted dry, and cut open with fine scissors after loss of turgor. The cell sap and the cytoplasm were extruded with forceps, collected on a slide and covered by a coverslip. For some experiments, the cytoplasm was squeezed into isotonic perfusion solution containing 200 mM sucrose, 70 mM KCl, 4.5 mM MgCl2, 5 mM EGTA, 1.48 mM CaCl2, 10 mM piperazine-N,N0 -bis(2-ethanesulfonic acid) (PIPES; pH 7) and 10 mM FM1-43 or FM4-64 (see below). In vivo staining and inhibitor treatments FM4-64 [N-(3-triethylammoniumpropyl)-4-(6-(4-(diethylamino)phenyl)hexatrienyl)-pyridinium dibromide] and the fixable analog of FM1-43 [N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide], which retains its fluorescence after fixation with glutaraldehyde, were purchased from Invitrogen (Carlsbad, CA, USA) and used at 10 mM diluted from 10 mM stock solutions in DMSO or from 500 mM stock solutions in distilled water. The stock solution of filipin III (Sigma, St Louis, MO, USA; 25 mg ml–1) was prepared in DMSO. Aliquots were frozen at 708C, thawed immediately before use and diluted 1: 4,000 in AFW for labeling live cells or in phosphate-buffered saline (PBS: 140 mM NaCl, 3 mM KCl, 10 mM phosphate buffer, pH 7.4, 0.12% NaN3) for labeling cells fixed in 1% (v/v) glutaraldehyde/ PBS. Lysotracker yellow HCK-123 (Invitrogen) was used at 10 mM diluted from a 1 mM stock solution in DMSO, and neutral red (Invitrogen) was used at 10–4% diluted from a 10–1% stock solution in distilled water. Cells were loaded with lysotracker yellow and neutral red for at least 2 h before addition of FM dyes. Mitochondria were stained with 1 mM mitotracker orange (Invitrogen; 1 mM stock solution in DMSO) and the endoplasmic reticulum was labeled with 1 mM DiOC6(3) (Invitrogen, 1 mM stock solution in DMSO). Cells were exposed to 5 mM CD (Sigma; stock solution 10 mM in DMSO), 50 mM LatB (Calbiochem, San Diego, CA, USA; stock solution 10 mM in DMSO), 50 mM BDM (Sigma), 10 mM oryzalin (Riedel-de Haen, Seelze, Germany; stock solution 10 mM in acetone) and 1 mM KCN (Sigma). All dyes and inhibitors were diluted with AFW which has a pH of about 6. The pH of the AFW used for dissolving neutral red and lysotracker yellow was adjusted to pH 8 with sodium bicarbonate in order to achieve loading into acidic compartments (Dubrovsky et al. 2006). We used internodal cells of the same culture and of similar age as controls and treated them with the corresponding amounts of solvent, which never exceeded 1% DMSO or 0.1% acetone, respectively. These concentrations had no visible effect on cytoplasmic streaming or internalization of FM dyes. Actin filaments and microtubules were visualized by perfusion of cells with fluorescent phalloidin (Alexa 488 phalloidin; Invitrogen) and fluorescent taxol (Flutax-2; Calbiochem) as described in Foissner (2004). Confocal scanning microscopy The CLSMs used in this study were a Zeiss LSM 510 and a Zeiss LSM Meta coupled to Zeiss Axiovert inverted microscopes (Jena, Germany). Images produced by the software were further processed with Adobe Photoshop. Organelle dynamics were studied by analyzing time series taken at minimum laser intensity and pixel time in order to avoid photobleaching and stress response. For co-localization of FM dyes, FM1-43 was excited with the 488 nm argon laser line at an intensity insufficient to excite FM464, and the resulting emission wavelengths were collected between 505 and 530 nm. FM4-64 was excited with the 543 nm helium–neon laser line, and emission wavelengths were recorded between 560 and 615 nm. The same settings were used for the dual labeling with FM1-43 and mitotracker orange and for the dual labeling with FM1-43 and neutral red. Lysotracker yellow and FM4-64 were excited with 488 and 543 nm, respectively, and their emission wavelengths were collected between 505 and 550 nm and between 560 and 615 nm. Filipin was excited with 351 nm, and the emission was collected between 470 and 490 nm. All images presented in this study are single sections. Images of cells are positioned with vertical sides parallel to the long axes of the internodes. Quantification of FM internalization The effect of CD on FM internalization was quantified in living and in fixed cells. Internodes were treated with 5 mM CD 30 min before and 30 min after addition of FM1-43, whereas control cells were incubated in FM1-43 alone for 30 min. In order to obtain comparable images of the stagnant endoplasm, cytoplasmic streaming in control cells was also arrested with 5 mM CD but only immediately prior to observation in the CLSM. The fluorescence of endoplasmic organelles in intact cells is attenuated by the cortical chloroplasts. Therefore, we also quantified FM internalization in fixed cells. Internodes were treated with 5 mM CD 30 min before and 30 min after addition of FM1-43 as described above. CD-treated cells were fixed for 30 min FM internalization in characean algae in a solution containing 1% glutaraldehyde and 5 mM CD dissolved in PBS. Control cells were fixed in the absence of CD. Fixed cells were washed in PBS and cut into cylinders which were split longitudinally. Cell fragments were mounted in PBS/glycerol (1:1) with the cytoplasmic side facing the coverslip so that the endoplasm could be studied without the disturbing chloroplast autofluorescence. Endoplasmic volume fractions occupied by FM1-43-fluorescent organelles (volume of fluorescent organelles as a percentage of total cytoplasmic volume) were calculated from Z-series of at least four cells using the Zeiss 3D-software. Differences between means were analysed by t-test and considered to be significant if P 0.01 (Sachs 1984). Supplementary material Supplementary material are available at PCP Online. Acknowledgments We are grateful to Richard Trelease (Arizona State University), to Ted Farmer (University of Lausanne) and to Takashi Ueda (University of Tokyo) for antibodies. We thank Beatrice Satiat-Jeunemaitre (CNRS, Gif-sur-Yvette) and Markus Grebe (Umea Plant Science Centre) for stimulating discussions. References Alvarez, F.J., Douglas, L.M. and Konopka, J.B. (2007) Sterol-rich plasma membrane domains in fungi. Eukaryot. Cell 6: 755–763. Aniento, F. and Robinson, D.G. (2005) Testing for endocytosis in plants. Protoplasma 226: 3–11. Ayscough, K.R. (2005) Coupling actin dynamics to the endocytic process in Saccharomyces cerevisiae. Protoplasma 226: 81–88. Baluska, F., Hlavacka, A., Samaj, J., Palme, K., Robinson, D.G., Matoh, T., McCurdy, D.W., Menzel, D. and Volkmann, D. (2002) F-actin-dependent endocytosis of cell wall pectins in meristematic root cells. Insights from brefeldin A-induced compartments. Plant Physiol. 130: 422–431. Baluska, F., Samaj, J., Hlavacka, A., Kendrick-Jones, J. and Volkmann, D. (2004) Actin-dependent fluid-phase endocytosis in inner cortex cells of maize root apices. J. Exp. Bot. 55: 463–473. Barton, R. (1965) An unusual organelle in the peripheral cytoplasm of Chara cells. Nature 205: 201. Beilby, M.J. and Shepherd, V.A. (1989) Cytoplasm-enriched fragments of Chara: structure and electrophysiology. Protoplasma 148: 150–163. Betz, W.J., Mao, F. and Smith, C.B. (1996) Imaging exocytosis and endocytosis. Curr. Opin. Neurobiol. 6: 365–371. Bhat, R. and Panstruga, R. (2005) Lipid rafts in plants. Planta 223: 5–19. Bisson, M.A., Siegel, A., Chau, R., Gelsomino, S.A. and Herdic, S.L. (1991) Distribution of charasomes in Chara: banding pattern and effect of photosynthetic inhibitors. Aust. J. Plant Physiol. 18: 81–93. Bolte, S., Talbot, C., Boutte, Y., Catrice, O., Read, N.D. and SatiatJeunemaitre, A. (2004) FM-dyes as experimental probes for dissecting vesicle trafficking in living plant cells. J. Microsc. 214: 159–173. Borner, G.H.H., Sherrier, D.J., Weimar, T., Michaelson, L.V., Hawkins, N.D., MacAskill, A., Napier, J.A., Beale, M.H., Lilley, K.S. and Dupree, P. (2005) Analysis of detergent-resistant membranes in Arabidopsis. Evidence for plasma membrane lipid rafts. Plant Physiol. 137: 104–116. Boucrot, E., Saffarian, S., Massol, R., Kirchhausen, T. and Ehrlich, M. (2006) Role of lipids and actin in the formation of clathrin-coated pits. Exp. Cell Res. 312: 4036–4048. Boutte, Y., Crosnier, M.T., Carraro, N., Traas, J. and SatiatJeunemaitre, B. (2006) The plasma membrane recycling pathway and 1519 cell polarity in plants: studies on PIN proteins. J. Cell Sci. 119: 1255–1265. Chau, R., Bisson, M.A., Siegel, A., Elkin, G., Klim, P. and Straubinger, R.M. (1994) Distribution of charasomes in Chara: re-establishment and loss in darkness and correlation with banding and inorganic carbon uptake. Funct. Plant Biol. 21: 113–123. Cheng, Z.J., Singh, R.D., Sharma, D.K., Holicky, E.L., Hanada, K., Marks, D.L. and Pagano, R.E. (2006) Distinct mechanisms of clathrinindependent endocytosis have unique sphingolipid requirements. Mol. Biol. Cell 17: 3197–3210. Collings, D.A., Wasteneys, G.O. and Williamson, R.E. (1995) Cytochalasin rearranges cortical actin of the alga Nitella into short, stable rods. Plant Cell Physiol. 36: 765–772. Conner, S.D. and Schmid, S.L. (2003) Regulated portals of entry into the cell. Nature 422: 37–44. Crawley, J.C.W. (1965) An unusual organelle in the peripheral cytoplasm of Chara cells. Nature 205: 200–201. Dettmer, J., Hong-Hermesdorf, A., Stierhof, Y.D. and Schumacher, K. (2006) Vacuolar Hþ-ATPase activity is required for endocytic and secretory trafficking in Arabidopsis. Plant Cell 18: 715–730. Dubrovsky, J.G., Guttenberger, M., Saralegui, A., Napsucialy-Mendivil, S., Voigt, B., Baluska, F. and Menzel, D. (2006) Neutral red as a probe for confocal laser scanning microscopy studies of plant roots. Ann. Bot. 97: 1127–1138. Engqvist-Goldstein, A.E.Y. and Drubin, D.G. (2003) Actin assembly and endocytosis: from yeast to mammals. Annu. Rev. Cell Dev. Biol. 19: 287–332. Fischer-Parton, S., Parton, R.M., Hickey, P.C., Dijksterhuis, J., Atkinson, H.A. and Read, N.D. (2000) Confocal microscopy of FM464 as a tool for analysing endocytosis and vesicle trafficking in living fungal hyphae. J. Microsc. 198: 246–259. Foissner, I. (1988) The relationship of echinate inclusions and coated vesicles on wound healing in Nitella flexilis (Characeae). Protoplasma 142: 164–175. Foissner, I. (2004) Microfilaments and microtubules control the shape, motility, and subcellular distribution of cortical mitochondria in characean internodal cells. Protoplasma 224: 145–157. Foissner, I. and Klima, A. (2008) Constitutive endocytosis in characean internodal cells is independent of an intact actin cytoskeleton. Cell Biol. Int. 32: 579–589. Foissner, I., Lichtscheidl, I.K. and Wasteneys, G.O. (1996) Actin-based vesicle dynamics and exocytosis during wound wall formation in characean internodal cells. Cell Motil. Cytoskel. 35: 35–48. Foissner, I. and Wasteneys, G.O. (2000) Actin in characean internodal cells. In Actin: A Dynamic Framework for Multiple Plant Cell Functions. Edited by Staiger, C., Baluska, D.F., Volkmann, D. and Barlow, P. pp. 259–274. Kluwer Academic Publisher, Dordrecht, The Netherlands. Foissner, I. and Wasteneys, G. (2007) Wide-ranging effects of eight cytochalasins and latrunculin A and B on intracellular motility and actin filament reorganization in characean internodal cells. Plant Cell Physiol. 48: 585–597. Franceschi, V.R. and Lucas, W.J. (1980) Structure and possible function(s) of charasomes; complex plasmalemma–cell wall elaborations present in some characean species. Protoplasma 104: 253–271. Franceschi, V.R. and Lucas, W.J. (1982) The relationship of the charasome to chloride uptake in Chara corallina: physiological and histochemical investigations. Planta 154: 525–537. Fujimoto, L.M., Roth, R., Heuser, J.E. and Schmid, S.L. (2000) Actin assembly plays a variable, but not obligatory role in receptor-mediated endocytosis in mammalian cells. Traffic 1: 161–171. Funaki, K., Nagata, A., Akimoto, Y., Shimada, K., Ito, K. and Yamamoto, K. (2004) The motility of Chara corallina myosin was inhibited reversibly by 2,3-butanedione monoxime (BDM). Plant Cell Physiol. 45: 1342–1345. Gachet, Y. and Hyams, J.S. (2005) Endocytosis in fission yeast is spatially associated with the actin cytoskeleton during polarised cell growth and cytokinesis. J. Cell Sci. 118: 4231–4242. Geldner, N. (2004) The plant endosomal system—its structure and role in signal transduction and plant development. Planta 219: 547–560. 1520 FM internalization in characean algae Graham, L. and Kaneko, Y. (1991) Subcellular structures of relevance to the origin of land plants (embryophytes) from green algae. CRC Crit. Rev. Plant Sci. 10: 323–342. Grebe, M., Xu, J., Mobius, W., Ueda, T., Nakano, A., Geuze, H.J., Rook, M.B. and Scheres, B. (2003) Arabidopsis sterol endocytosis involves actin-mediated trafficking via ARA6-positive early endosomes. Curr. Biol. 13: 1378–1387. Grolig, F. and Pierson, E.S. (2000) Cytoplasmic streaming: From flow to track. In Actin: A Dynamic Framework for Multiple Plant Cell Functions. Edited by Staiger, C., Baluska, D.F., Volkmann, D. and Barlow, P. pp. 165–181. Kluwer Academic Publisher, Dordrecht, The Netherlands. Guttenberger, M. (2000) Arbuscules of vesicular-arbuscular mycorrhizal fungi inhabit an acidic compartment within plant roots. Planta 211: 299–304. Hawes, C.R., Coleman, J.O.D. and Evans, D.E. (1991) Endocytosis, Exocytosis and Vesicle Traffic in Plants. Cambridge: Cambridge University Press. Holstein, S.E.H. (2002) Clathrin and plant endocytosis. Traffic 3: 614–620. Hörmanseder, K., Obermeyer, G. and Foissner, I. (2005) Disturbance of endomembrane trafficking by brefeldin A and calyculin A reorganizes the actin cytoskeleton of Lilium longiflorum pollen tubes. Protoplasma 227: 25–36. Jarosch, R. (1961) Das Characeen-Protoplasma und seine Inhaltskörper. Protoplasma 53: 34–56. Johnson, C. and Griffing, L.R. (2007) FRET analysis of trans-membrane flipping of FM4-64 in living root hairs: is FM4-64 a robust marker for endocytosis? 8th International Botanical Microscopy Meeting, Salzburg. Kachar, B. (1985) Direct visualization of organelle movement along actin filaments dissociated from characean algae. Science 227: 1355–1357. Kamiya, N. and Kuroda, K. (1957) Cell operation in Nitella. II. Behaviour of isolated endoplasm. Proc. Jpn. Acad. 33: 201–215. Kirkham, M. and Parton, R.G. (2005) Clathrin-independent endocytosis: new insights into caveolae and non-caveolar lipid raft carriers. Biochim. Biophys. Acta 1745: 273–286. Kutsuna, N. and Hasezawa, S. (2002) Dynamic organization of vacuolar and microtubule structures during cell cycle progression in synchronized tobacco BY-2 cells. Plant Cell Physiol. 43: 965–973. Lucas, W.J. and Franceschi, V.R. (1981) Characean charasome-complex and plasmalemma vesicle development. Protoplasma 107: 255–267. Malinska, K., Malinsky, J., Opekarova, M. and Tanner, W. (2004) Distribution of Can1p into stable domains reflects lateral protein segregation within the plasma membrane of living S. cerevisiae cells. J. Cell Sci. 117: 6031–6041. Marcote, M.J., Gu, F., Gruenberg, J. and Aniento, F. (2000) Membrane transport in the endocytic pathway: animal versus plant cells. Protoplasma 210: 123–132. Martin, S.W., Glover, B.J. and Davies, J.M. (2005) Lipid microdomains— plant membranes get organized. Trends Plant Sci. 10: 263–265. McCurdy, D.W. (1999) Is 2,3-butanedione monoxime an effective inhibitor of myosin-based activities in plant cells? Protoplasma 209: 120–125. McLean, B. and Juniper, B.E. (1986) The plasma membrane of young Chara internodal cells revealed by rapid freezing. Planta 169: 153–161. Meckel, T., Hurst, A.C., Thiel, G. and Homann, U. (2004) Endocytosis against high turgor: intact guard cells of Vicia faba constitutively endocytose fluorescently labelled plasma membrane and GFP-tagged Kþ-channel KAT1. Plant J. 39: 182–193. Meckel, T., Hurst, A.C., Thiel, G. and Homann, U. (2005) Guard cells undergo constitutive and pressure-driven membrane turnover. Protoplasma 226: 23–29. Miller, R.G. (1984) The use and abuse of filipin to localize cholesterol in membranes. Cell Biol. Int. Rep. 8: 519–535. Mongrand, S., Morel, J., Laroche, J., Claverol, S., Carde, J.P., Hartmann, M.A., Bonneu, M., Simon-Plas, F., Lessire, R. and Bessoule, J.J. (2004) Lipid rafts in higher plant cells—purification and characterization of Triton X-100-insoluble microdomains from tobacco plasma membrane. J. Biol. Chem. 279: 36277–36286. Murphy, A.S., Bandyopadhyay, A., Holstein, S.E. and Peer, W.A. (2005) Endocytotic cycling of plasma membrane proteins. Annu. Rev. Plant Biol. 56: 221–251. Neuhaus, J.M. and Paris, S. (2006) Plant vacuoles: from biogenesis to function. In Plant Endocytosis. Edited by Samaj, J., Baluska, D.F. and Menzel, D. pp. 63–82. Springer, Berlin. Nichols, B.J. and Lippincott-Schwartz, J. (2001) Endocytosis without clathrin coats. Trends Cell Biol. 11: 406–412. Nishikawa, S. and Sasaki, F. (1996) Internalization of styryl dye FM1-43 in the hair cells of lateral line organs in Xenopus larvae. J. Histochem. Cytochem. 44: 733–741. Ovecka, M., Lang, I., Baluska, F., Ismail, A., Illes, P. and Lichtscheidl, I.K. (2005) Endocytosis and vesicle trafficking during tip growth of root hairs. Protoplasma 226: 39–54. Ovecka, M. and Lichtscheidl, I. (2006) Sterol endocytosis and trafficking in plant cells. In Plant Endocytosis. Edited by Samaj, J., Baluska, D.F. and Menzel, D. pp. 117–138. Springer, Berlin. Parton, R.M., Fischer-Parton, S., Trewavas, A.J. and Watahiki, M.K. (2003) Pollen tubes exhibit regular periodic membrane trafficking events in the absence of apical extension. J. Cell Sci. 116: 2707–2719. Parton, R.M., Fischer-Parton, S., Watahiki, M.K. and Trewavas, A.J. (2001) Dynamics of the apical vesicle accumulation and the rate of growth are related in individual pollen tubes. J. Cell Sci. 114: 2685–2695. Patterson, G.W., McKenna, K.C., Lusby, W.R. and Bisson, M.A. (1991) Sterols of the charophyceae. J. Nat. Prod. 54: 1141–1142. Pesacreta, T.C. and Lucas, W.J. (1984) Plasma membrane coat and a coated vesicle-associated reticulum of membranes: their structure and possible interrelationship in Chara corallina. J. Cell Biol. 98: 1537–1545. Pickett-Heaps, J.D. (1967) Ultrastructure and differentiation in Chara sp. I. Vegetative cells. Austr. J. Biol. Sci. 20: 539–551. Pike, L.J. (2006) Rafts defined: a report on the Keystone symposium on lipid rafts and cell function. J. Lipid Res. 47: 1597–1598. Polo, S. and Di Fiore, P.P. (2006) Endocytosis conducts the cell signaling orchestra. Cell 124: 897–900. Pralle, A., Keller, P., Florin, E.L., Simons, K. and Horber, J.K.H. (2000) Sphingolipid–cholesterol rafts diffuse as small entities in the plasma membrane of mammalian cells. J. Cell Biol. 148: 997–1007. Price, G.D., Badger, M.R., Bassett, M.E. and Whitecross, M.I. (1985) Involvement of plasmalemmasomes and carbonic anhydrase in photosynthetic utilization of bicarbonate in Chara corallina. Aust. J. Plant Physiol. 12: 241–256. Price, G.D. and Whitecross, M.I. (1983) Cytochemical localisation of ATPase activity on the plasmalemma of Chara corallina. Protoplasma 116: 65–74. Robinson, J.M. and Karnovsky, M.J. (1980) Evaluation of the polyene antibiotic filipin as a cytochemical probe for membrane cholesterol. J. Histochem. Cytochem. 28: 161–168. Sachs, L. (1984) Angewandte Statistik. Springer, Berlin. Sakano, K. and Tazawa, M. (1986) Tonoplast origin of the envelope membrane of cytoplasmic droplets prepared from Chara internodal cells. Protoplasma 131: 247–249. Samaj, J. (2006) Methods and molecular tools for studying endoytosis in plants—an overview. In Plant Endocytosis. Edited by Samaj, J., Baluska, D.F. and Menzel, D. pp. 1–18. Springer, Berlin. Samaj, J., Baluska, D.F. and Menzel, D., eds. (2006a) Plant Endocytosis. Springer, Berlin. Samaj, J., Baluska, D.F., Voigt, B., Volkmann, D. and Menzel, D. (2006b) Endocytosis and actomyosin cytoskeleton. In Plant Endocytosis. Edited by Samaj, J., Baluska, D.F. and Menzel, D. pp. 233–244. Springer, Berlin. Samaj, J., Read, N.D., Volkmann, D., Menzel, D. and Baluska, F. (2005) The endocytic network in plants. Trends Cell Biol. 15: 425–433. Schulte, C., Kirst, G.O. and Winter, U. (1994) Source–sink characteristic of photoassimilate transport in fertile and sterile plants of Chara vulgaris L. Bot. Acta 107: 362–368. Shahollari, B., Peskan-Berghofer, T. and Oelmuller, R. (2004) Receptor kinases with leucine-rich repeats are enriched in Triton X-100 insoluble plasma membrane microdomains from plants. Physiol. Plant. 122: 397–403. FM internalization in characean algae Shimmen, T. and Yokota, E. (2004) Cytoplasmic streaming in plants. Curr. Opin. Cell Biol. 16: 68–72. Simons, K. and Van Meer, G. (1988) Lipid sorting in epithelial cells. Biochemistry 27: 6197–6202. Soldati, T. and Schliwa, M. (2006) Powering membrane traffic in endocytosis and recycling. Nat. Rev. Mol. Cell Biol. 7: 897–908. Spector, I., Braet, F., Shochet, N.R. and Bubb, M.R. (1999) New anti-actin drugs in the study of the organization and function of the actin cytoskeleton. Microsc. Res. Tech. 47: 18–37. Sun, Y.D., Martin, A.C. and Drubin, D.G. (2006) Endocytic internalization in budding yeast requires coordinated actin nucleation and myosin motor activity. Dev. Cell 11: 33–46. Tanabe, Y., Hasebe, M., Sekimoto, H., Nishiyama, T., Kitani, M., Henschel, K., Munster, T., Theissen, G., Nozaki, H. and Ito, M. (2005) Characterization of MADS-box genes in charophycean green algae and its implication for the evolution of MADS-box genes. Proc. Natl Acad. Sci. USA 102: 2436–2441. Tse, Y.C., Mo, B.X., Hillmer, S., Zhao, M., Lo, S.W., Robinson, D.G. and Jiang, L.W. (2004) Identification of multivesicular bodies as prevacuolar compartments in Nicotiana tabacum BY-2 cells. Plant Cell 16: 672–693. 1521 Ueda, T., Uemura, T., Sato, M.H. and Nakano, A. (2004) Functional differentiation of endosomes in Arabidopsis cells. Plant J. 40: 783–789. Valdez-Taubas, J. and Pelham, H.R.B. (2003) Slow diffusion of proteins in the yeast plasma membrane allows polarity to be maintained by endocytic cycling. Curr. Biol. 13: 1636–1640. Van Gisbergen, P., Esseling-Ozdoba, A. and Vos, J.W. (2007) Microinjecting FM4-64 supports its specifity for the endocytic pathway. 8th International Botanical Microscopy Meeting, Salzburg Verma, D.P.S., Hong, Z. and Menzel, D. (2006) Dynamin-related proteins in plant endocytosis. In Plant Endocytosis. Edited by Samaj, J., Baluska, D.F. and Menzel, D. pp. 217–232. Springer, Berlin. Voigt, B., Timmers, A.C.J., Samaj, J., Hlavacka, A., Ueda, T. et al. (2005) Actin-based motility of endosomes is linked to the polar tip growth of root hairs. Eur. J. Cell Biol. 84: 609–621. Wasteneys, G.O. and Williamson, R.E. (1987) Microtubule orientation in developing internodal cells of Nitella: a quantitative analysis. Eur. J. Cell Biol. 43: 14–22. Williamson, R.E. (1978) Cytochalasin B stabilises the sub-cortical actin bundles of Chara against a solution of low ionic strength. Cytobiologie 18: 107–113. (Received July 21, 2008; Accepted August 14, 2008)