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Hyperosmotic Stress Induces Formation of Tubulin Macrotubules in Root-Tip Cells of Triticum turgidum: Their Probable Involvement in Protoplast Volume Control

Hyperosmotic Stress Induces Formation of Tubulin Macrotubules in Root-Tip Cells of Triticum... Abstract Treatment of root-tip cells of Triticum turgidum with 1 M mannitol solution for 30 min induces microtubule (Mt) disintegration in the plasmolyzed protoplasts. Interphase plasmolyzed cells possess many cortical, perinuclear and endoplasmic macrotubules, 35 nm in mean diameter, forming prominent arrays. In dividing cells macrotubules assemble into aberrant mitotic and cytokinetic apparatuses resulting in the disturbance of cell division. Putative tubulin paracrystals were occasionally observed in plasmolyzed cells. The quantity of polymeric tubulin in plasmolyzed cells exceeds that in control cells. Root-tip cells exposed for 2–8 h to plasmolyticum recover partially, although the volume of the plasmolyzed protoplast does not change detectably. Among other events, the macrotubules are replaced by Mts, chromatin assumes its typical appearance and the cells undergo typical cell divisions. Additionally, polysaccharidic material is found in the periplasmic space. Oryzalin and colchicine treatment induced macrotubule disintegration and a significant reduction of protoplast volume in every plasmolyzed cell type examined, whereas cytochalasin B had only minor effects restricted to differentiated cells. These results suggest that Mt destruction by hyperosmotic stress, and their replacement by tubulin macrotubules and putative tubulin paracrystals is a common feature among angiosperms and that macrotubules are involved in the mechanism of protoplast volume regulation. (Received February 15, 2002; Accepted May 30, 2002) Introduction Evidence accumulated so far shows that in various organisms, the cytoskeleton is sensitive to osmotic stress. Osmotic stress induces microtubule (Mt) disintegration and/or disorganization in the following higher plant cell types: (a) epidermal cells of Pisum sativum (Roberts et al. 1985), Allium cepa (Lang-Pauluzzi and Gunning 2000) and Tradescantia virginiana (Cleary 2001), (b) leaf cells of Spinacea oleracea (Bartolo and Carter 1991), (c) differentiating root cells of Zea mays (Blancaflor and Hasenstein 1995). Besides, hyperosmotic treatment affects Mts in the yeast Saccharomyces cerevisiae (Slaninova et al. 2000) and in different animal systems, as in withdrawing reticulopodia of the protozoan Allogromia (Welnhofer and Travis 1996) and dividing PtK-1 cells (Mullins and Snyder 1989). Recent work has revealed that non-ionic hyperosmotic stress dramatically affects the organization of the Mt cytoskeleton in the leaf cells of Chlorophyton comosum (Komis et al. 2001). In mitotic and cytokinetic cells Mts are disorganized and tubulin is polymerized into macrotubules, 32 nm in mean diameter, and into paracrystals. The former assemble atypical spindles and phragmoplasts accompanied by disturbed cell division. Malformed spindles and phragmoplasts have also been observed in plasmolyzed leaf epidermal cells of T. virginiana (Cleary 2001). In interphase plasmolyzed cells of C. comosum the cortical and endoplasmic Mts are also disintegrated and numerous tubulin paracrystals are assembled. Macrotubules were absent from those cells (Komis et al. 2001). From the above it is evident that although the effects of hyperosmotic stress on the organization of the Mt cytoskeleton are well documented, it was found to induce the formation of macrotubules and putative tubulin paracrystals only in leaf cells of C. comosum. In order to test whether the formation of atypical tubulin polymers is a general response of higher plant cells in hyperosmotic stress, we studied the organization of tubulin cytoskeleton in plasmolyzed root-tip cells of Triticum turgidum. Since there are some reports implicating Mts in certain aspects of animal cell volume regulation (Moustakas et al. 1998), we also studied the possible involvement of tubulin polymers in the regulation of the plasmolyzed protoplast volume through the use of anti-Mt drugs. Results Control root-tip cells Mt organization in meristematic root-tip cells of T. turgidum has been previously described (Zachariadis et al. 2000, Frantzios et al. 2000, Frantzios et al. 2001) and is similar to that described in root-tip cells of other angiosperms. Interphase cortical Mt systems, preprophase Mt bands (PPBs), mitotic spindles, interzonal Mt systems, as well as phragmoplasts at different stages of development were observed in the meristematic cells examined (Fig. 1A–E). Differentiating root-tip cells exhibit cortical Mts and an endoplasmic Mt array, which in some cells radiates from the perinuclear area towards the cell cortex. Plasmolyzed root-tips Pattern of plasmolytic cycle The 1 M mannitol solution induced extensive changes in the organization of most root-tip cells. Water loss results in a reduction of the protoplast volume and the detachment of the plasmalemma from the cell wall (plasmolysis). The extent of plasmolysis varied among different cell types as well as among cells from different root regions. Study of serial semithin sections of root-tips subjected to plasmolysis for 30 min revealed that every cell was plasmolyzed. However, in meristematic cells, the detachment of the protoplast from the cell wall was minimal and restricted to cell corners. Convex plasmolysis was the dominant plasmolysis pattern observed (Fig. 2A, B). Examination of living root-tips with DIC optics revealed that, at least in rhizodermal and root-cap cells, plasmolysis commences immediately after immersion of the root-tips in mannitol solution and is completed within 5 min. Afterwards, the protoplast volume does not change significantly, even in root-tips that remained in the plasmolytic solution for 2–8 h (Fig. 2A, B). The same plasmolysis pattern was induced by 0.5 and 0.7 M mannitol solutions. However, the plasmolyzed cells in these solutions were fewer, while the course of plasmolysis was completed in longer times compared with those induced by 1 M mannitol. In root-tips that had undergone plasmolysis, all the rhizodermal cells deplasmolyzed within 2–3 min when they were placed in distilled water. Root-tip cells following a short mannitol treatment The changes observed in root-tip cells were identical among all the mannitol solutions used (0.5, 0.7 and 1 M). To avoid duplication of the results the organization of cells plasmolyzed with 1 M mannitol will be described only. Transmission electron microscope (TEM) examination of root-tip cells following a short (up to 30 min) mannitol treatment revealed that the plasmolyzed protoplasts were characterized by: (a) an increase in the electron density of the cytoplasm (Fig. 3A), (b) an intense dictyosome activity and the stacking of the endoplasmic reticulum membranes, (c) the ‘coalescence’ of chromatin as well as of chromosomes into a condensed mass, which in interphase cells is largely separated from the nuclear envelope (Fig. 3A). The chromatin changes were also conspicuous at the light microscope level after toluidine blue staining as well as in fluorescence microscopy after staining with Hoechst 33258 (Fig. 4E inset, 6E). In the periplasmic space, i.e. in the space between the protoplast and the cell wall, Hechtian strands and a loosely arranged granular material were found (Fig. 3B). Observation of tubulin immunolabeled cells that were plasmolyzed for 30 min, revealed that they displayed many thick and intensely fluorescing tubulin strands, the organization of which varied according to the cell type and the stage of the cell cycle (Fig. 4A–F). Notably, the quantity of polymeric tubulin per cell in plasmolyzed cells was 30–35% higher than in control cells as assessed by digital image analysis of the specimens (Fig. 5). In interphase meristematic cells many tubulin strands traversed the cortical cytoplasm transversely to the long cell axis or in different directions (Fig. 4A, B) as well as the subcortical cytoplasm and the endoplasm (Fig. 4 C). The subcortical tubulin strands were usually arranged parallel to the long cell axis, while the endoplasmic ones were mainly localized in the perinuclear cytoplasm (Fig. 4C). The preprophase-prophase cells exhibited aberrant PPBs, tubulin strands running through the cortical cytoplasm outside the PPB cortical zone and perinuclear tubulin strands (Fig. 6A, B). In mitotic cells the chromosomes appeared coalesced into one or two chromosome masses (Fig. 6E, F inset). These cells assembled highly elongated and sharply focused atypical spindles (Fig. 6C, D) similar to those of the plasmolyzed leaf cells of C. comosum (Komis et al. 2001). Frequently, tubulin immunoreactivity was localized into chromatin masses (Fig. 6F). The cytokinetic cells formed atypical phragmoplasts made of prominent tubulin polymer bundles (Fig. 6G–I), which accompanied atypical cell plates (Fig. 6J, K). Often the latter appeared to comprise ‘vacuole-like’ dilations (Fig. 6K, 8A), like those previously described in plasmolyzed epidermal cells of Tradescantia (Cleary 2001). Some plasmolyzed cytokinetic root-tip cells of T. turgidum did not form cell plates. Differentiating plasmolyzed root cells possessed numerous endoplasmic tubulin strands, which in some cases were arranged in a radial perinuclear system reaching up the cell cortex (Fig. 4D, E). In addition, many thick cortical tubulin strands were frequently focussed on cortical sites (Fig. 4F). TEM examination revealed that all cells that had been plasmolyzed for 30 min, including the rhizodermal ones, lacked Mts, but possessed macrotubules of diameter ranging between 26 nm and 41 nm. Their mean diameter was 35 nm, while the mean value of Mt diameter in control cells was 22 nm. These values were derived from measurement of 450 macrotubules and 200 Mts. The macrotubule and Mt diameter was measured as described in Komis et al. (2001). In interphase cells single or bundles of macrotubules traversed the cortical cytoplasm and the endoplasm (Fig. 7A, B). Macrotubules traversed some of the Hechtian strands along the whole of their length (Fig. 3B). In dividing cells aberrant mitotic and cytokinetic apparatuses consist of macrotubules identical to those found in interphase cells (Fig. 8A, B). Therefore, the tubulin strands observed in the immunofluorescent specimens consist of macrotubules. The plasmolyzed cells also displayed some elongated electron dense paracrystals, made of fine sheets or filaments (Fig. 7C). The structural similarity of these paracrystals to one category of those found in the plasmolyzed leaf cells of C. comosum (Komis et al. 2001), favors the hypothesis that they consist of tubulin. Root-tip cells following a prolonged mannitol treatment Protoplasts of root-tip cells, which remained in the hypertonic solution for 8 h, showed some structural recovery, although their volume did not increase detectably. The interphase and mitotic cells displayed typical nuclei and chromosomes respectively (Fig. 9A, 10B inset), low electron density of the cytoplasm and rarely endoplasmic reticulum stacks. Moreover, the periplasmic space contained material (Fig. 9B) positive for PAS, calcofluor and toluidine blue staining (Fig. 9C). These tests demonstrate that this material, which is secreted by the protoplast, is of polysaccharidic nature. The majority of the interphase cells that had undergone 2–8 h treatment with mannitol solution were devoid of the thick fluorescent tubulin strands formed during the first 30 min of treatment and possessed fluorescent structures resembling the appearance of Mts of control cells after tubulin immunolabelling (Fig. 10A; cf. Fig 1A). TEM examination confirmed the absence of macrotubules and the appearance of cortical and endoplasmic Mt arrays in those cells. The dividing cells found in the root tips that had undergone long hyperosmotic treatment (up to 8 h) were classified into two groups. The first group includes cells that entered cell division during plasmolysis. The appearance of the chromosomes and the organization of the mitotic spindle and the phragmoplast were similar to those of control cells (Fig. 10B–D; cf. Fig. 1C–E). The second group includes atypical mitotic cells, in which chromosomes appeared normal, but were either dispersed throughout the cytoplasm or were arranged in rosette conformations. These cells exhibited atypical mono-, bi- or multipolar and probably non-functional spindles. They give rise to polyploid or multinucleate cells. TEM examination showed that mitotic and cytokinetic apparatuses of cells plasmolyzed for 8 h consisted of Mts. Anti-Mt drug and cytochalasin B-treated plasmolyzed cells Following tubulin immunofluorescence of oryzalin- and colchicine-treated plasmolyzed cells it was found that thick fluorescing strands were disintegrated. These cells showed fluorescing fragments, which probably represent remnants of the macrotubule strands (Fig. 10E). It is important to note that macrotubule disorganization following the anti-Mt-drug treatment affected plasmolysis. In non-treated rhizodermal cells, plasmolysis was completed in 5 min after the immersion of the root-tips in the mannitol solution, while in the oryzalin treated ones, plasmolysis continued for at least 1 h (Fig. 11A, B; cf. Fig. 2A, B). The protoplast volume after 30 min of plasmolysis was measured in oryzalin-treated and non-treated rhizodermal cells. It was found that in the oryzalin-treated cells, the protoplast volume was 40–45% less than that in the non-treated plasmolyzed cells (Fig. 12). Additionally, in many oryzalin-treated cells the protoplast frequently exhibited concave patterns of plasmolysis (Fig. 11D), while in others the protoplast was divided into subprotoplasts (Fig. 11E), phenomena not observed in the untreated plasmolyzed cells. All oryzalin-treated cells showed more Hechtian strands than the untreated cells (Fig. 11C). Cells pretreated with oryzalin for 24 h displayed a greater protoplast volume reduction than those pretreated for 1 h (Fig. 11D, E; cf. Fig. 11A–C). The response of the plasmolyzed cells to colchicine was similar to that of cells treated with oryzalin. In contrast, cytochalasin B (CB) treatment did not have any appreciable effect on the protoplast shape and volume of meristematic and differentiating plasmolyzed rhizodermal cells (Fig. 12). However, in fully differentiated plasmolyzed rhizodermal cells CB brought about an approximately 20% reduction in protoplast volume (Fig. 12). Discussion General remarks This work shows that root-tip cells of T. turgidum exposed to non-ionic hyperosmotic stress, undergo extensive structural changes and that the affected cells have the ability to overcome certain of the stress effects in the presence of the plasmolytic solution. The main observations of this study are given below. (a) Hyperosmotic stress induced, among other effects, Mt disintegration, formation of tubulin macrotubules and putative tubulin paracrystals and highly condensed chromatin. (b) In interphase cells the macrotubules are numerous and form well-organized cortical and endoplasmic systems, while in the mitotic and cytokinetic cells they assemble aberrant mitotic and cytokinetic apparatuses, disturbing chromosome separation and cell division. (c) During prolonged treatments (2–8 h) of the root-tips with the plasmolytic solution the plasmolyzed protoplasts recover partially, restoring Mt arrays in place of the macrotubules. (d) Macrotubule disassembly after anti-Mt drug treatment extends the duration of plasmolysis, induces a significant decrease of the protoplast volume compared with the untreated plasmolyzed cells and affects the shape of the plasmolyzed protoplasts. Some of these phenomena will be discussed below. Mt-disassembly The present work on T. turgidum, as well as previous results on the plasmolyzed cells of C. comosum (Komis et al. 2001), support the hypothesis that the replacement of Mts by aberrant tubulin polymers is an immediate response of angiosperm cells to hyperosmotic stress. Changes in Mt dynamics and/or organization induced by osmotic stress have also been observed: (a) in higher plant cells (Bartolo and Carter 1991, Blancaflor and Hasenstein 1995, Lang-Pauluzzi and Gunning 2000, Cleary 2001), (b) in fungal cells (Slaninova et al. 2000) and (c) in animal cells (Welnhofer and Travis 1996, Mullins and Snyder 1989). The rapid response of the Mt cytoskeleton to hyperosmotic stress is probably due to rapid and profound changes in cell metabolism. In plant cells the osmotic stress induces: (a) changes in the cytosolic Ca2+ concentration (Bush 1995, Knight et al. 1997, Knight et al. 1998, Cessna et al. 1998, Brownlee et al. 1999, Knight 2000), (b) activation of a mitogen-activated protein kinase cascade (Popping et al. 1996, Mizoguchi et al. 1997, Hirt 2000), (c) triggering of the inositol-signaling pathway (Munnik et al. 2000, Meijer et al. 2001). All these phenomena are directly related to mechanisms controlling Mt dynamics (Desai and Mitchison 1997, Quader 1998, Vaughn and Harper 1998, Bögre et al. 2000) and therefore they can potentially explain Mt disorganization in plasmolyzed root-tip cells of T. turgidum. Alternatively, the mechanical forces imposed upon the protoplast due to volume reduction upon plasmolysis may lead to Mt disruption. The sensitivity of the Mt cytoskeleton to mechanical stress is fairly well documented in plant (Wymer et al. 1996) and animal (Ingber 1997) systems. Moreover, in plants, stomatal pore closure, a process depending on reduction of the guard cell volume, is accompanied by disintegration of the cortical Mt arrays (Jiang et al. 1996, Huang and Wang 1997, Fukuda et al. 1998, Marcus et al. 2001, Yu et al. 2001). Plasmalemma ion channels, including the Ca2+ ones, can be activated by stretch (Cosgrove and Hedrich 1991, Ding and Pickard 1993, Janmey 1998). Their activation by hyperosmotic stress induces Ca2+ transportin the protoplast and triggers processes controlled by Ca2+ as a response to osmotic stress (Brownlee et al. 1999). These processes, among others, may induce Mt disintegration. Finally, Mt disassembly might also be attributed to disruption of the cell wall-to-plasmalemma continuum (Fowler and Quatrano 1997). Formation of abnormal tubulin polymers In the plasmolyzed root tip-cells of T. turgidum, similarly to leaf cells of C. comosum (Komis et al. 2001), Mt disorganization is accompanied by the formation of tubulin macrotubules and putative tubulin paracrystals. Their assembly is probably a general response of plant cells exposed to non-ionic hyperosmotic stress. In plants tubulin macrotubules, probably consisting of more than 13 protofilaments, have also been observed in oil-body cells of Marchantia paleacea (Galatis and Apostolakos 1976, Apostolakos and Galatis 1998) and in Al-treated root-tip cells of T. turgidum (Frantzios et al. 2000, Frantzios et al. 2001). Komis et al. (2001) suggested that in plasmolyzed leaf cells of C. comosum the cytosolic Ca2+ concentration that prevails at the beginning of plasmolysis leads to Mt disorganization, and that later this is changed, leading to tubulin macrotubule and paracrystal formation. The same mechanism may function in the plasmolyzed root-tip cells of T. turgidum. The changes of cytosolic Ca2+ concentration estimated in different plant systems which underwent hyperosmotic stress (Takahashi et al. 1997, Cessna et al. 1998) support the above hypothesis. Moreover, the formation of macrotubules and tubulin paracrystals in vitro is defined by interactions between tubulin protofilaments, regulated by Ca2+ concentration (Vater et al. 1997). In hyperosmotically treated root-tip cells the quantity of polymeric tubulin is 30–35% higher when compared with control cells, an observation also made in C. comosum (Komis et al. 2001). Therefore, the induction of excessive tubulin polymerization may be considered as a consistent response of the plant cells to hyperosmotic stress. In contrast to the plasmolyzed interphase leaf cells of C. comosusm, interphase T. turgidum root-tip cells form cortical macrotubules. In the interphase root-tip cells of the examined plant the macrotubules form a very prominent cortical cytoskeletal system. This difference may reflect response differences between leaf cells and root-tip cells to hyperosmotic treatment. The cytoskeletal response could be coupled to different signaling pathways between these systems. Dividing T. turgidum root-tip cells form atypical spindles and phragmoplasts, a phenomenon also observed in hyperosmotically treated leaf cells of C. comosum (Komis et al. 2001) and T. virginiana (Cleary 2001). Therefore, the organization of atypical mitotic and cytokinetic apparatuses is a general cell reaction to hyperosmotic stress (see also Mullins and Snyder 1989). Probable macrotubule function(s) Following destruction of macrotubules by treatment with anti-Mt drugs, the protoplast volume diminution lasted for 2–3 h. Further exposure of roots to plasmolytic solutions supplemented with oryzalin resulted in excessive necrosis. These observations indicate a direct involvement of macrotubules in the regulatory mechanism of plasmolyzed protoplast volume in the root-tip cells. This was somewhat unexpected, given that most animal cells under anisosmotic conditions retain their volume mainly by the activity of actin filaments (Papakonstanti et al. 2000, Szaszi et al. 2000). In plasmolyzed leaf cells of C. comosum the mechanism by which the plasmolyzed protoplasts control their volume is also based on actin filaments (Komis et al. 2002). The minimal contribution of actin filaments in this process in root-tip cells of T. turgidum was demonstrated by the treatment of the plasmolyzed cells with CB. This treatment does not affect the volume of the plasmolyzed protoplast, nor does it affect the pattern of plasmolysis in meristematic and differentiating cells. In fully differentiated cells, however, the role of actin filaments in protoplast volume regulation is somewhat increased. In those cells the application of CB affects the extent of protoplast volume reduction by 20%. Nevertheless, in this case macrotubules play a dominant role in volume regulation as well, since oryzalin exerts drastic effects in protoplast volume reduction. There are some reports implicating Mts in many aspects of animal cell volume regulation. Mt arrays have been shown to activate both ion channels, generally responsible for short-term volume regulatory events and osmolyte transporters, implicated in long-term volume regulation events (Moustakas et al. 1998). The role of Mts in aquaporin clustering is well documented in animal systems often exposed to anisosmotic conditions (Sabolic et al. 1995). In plants, Mts are required upstream to the ionic events (i.e. H+ efflux and K+ influx), leading to stomatal pore opening (Marcus et al. 2001, Yu et al. 2001). In other plant cell types Mts also appear to be involved in the control of the activity of the plasmalemma ion channels (Thion et al. 1996, Thion et al. 1998). Considering the above-mentioned reports it may be suggested that in the root-tip cells of T. turgidum the plasmolyzed protoplast principally controls its volume by the formation of macrotubules. They may regulate the plasmalemma and tonoplast ion channel activity under hyperosmotic conditions and/or the distribution of aquaporins, water transporters that might be responsible for the rehydration of root-tip cells during prolonged hypertonic conditions. The macrotubules in T. turgidum may offer mechanical support to the protoplast to resist forces exerted on it during plasmolysis, a hypothesis also made for tubulin paracrystals in plasmolyzed leaf cells of C.comosum (Komis et al. 2001). Animal cells under mechanical stress reinforce the cortical cytoplasm at the sites of extensive force application with additional cytoskeletal elements (Ingber 1997, Chicurel et al. 1998, Ko and McCulloch 2000). Considering that during plasmolysis stretching forces are generated, which might produce an injury event at the plasmalemma (Oparka 1994) and that the Hechtian strands are under high tensional forces (Buer et al. 2000), the cortical macrotubules as well as the endoplasmic ones may also function as a strong supportive system. Recovery of the plasmolyzed protoplasts Notably root-tip cells of T. turgidum are able to recover from the severe effects on their structure and function caused by high extracellular osmolarity (see also Blancaflor and Hasenstein 1995). Further more, this ability developed during the course of the plasmolytic treatment. Although this became mostly evident after 8 h of treatment, early signs of recovery were recognizable as early as 1.5–2 h after the exposure of roots to the plasmolytic medium. The ability of the plasmolyzed protoplast to counteract, at least partially, the hyperosmotic stress can safely be attributed to cortical macrotubule formation. Indeed, oryzalin-treated cells are necrotized after a few hours in the hypersmoticum. Among the prominent changes of the recovering protoplasts are the chromatin reorganization, the decrease in electron density of the cytoplasm, the disappearance of macrotubules and the reappearance of Mts, and the return of normal cell divisions. The data of this work show that the formation of abnormal tubulin polymers is a reversible phenomenon. The recovery of the plasmolyzed protoplasts can be explained assuming the activation of a mechanism regulating the protoplast volume and the cytoplasm osmolarity. Deposition of polysaccharidic material in the periplasmic space may prevent protoplast enlargement in the recovering cells during prolonged plasmolysis. Deposition of wall materials in the periplasmic space has also been described in other plasmolyzed cell types (Robinson and Cummins 1976, Schnepf et al. 1986). Material and Methods Plant material and treatments Caryopses of T. turgidum L. var. durum Raddi were imbibed with distilled water for 36–48 h in darkness, at 25°C. The seedlings were then treated with an aqueous 0.5 M or 0.7 M or 1 M mannitol solution on moistened cotton for 5 min, 30 min, 2, 4, 8 and 12 h. In addition, roots treated with 10 µM oryzalin for 1–24 h or 2 mM colchicine for 1–24 h or 100 µM CB for 1 h, were subjected to 1 M mannitol treatment further supplemented with anti-Mt drugs or CB. To minimize anoxic effects on the root segments immersed for a long time in the plasmolytic solution, the following precautions were taken. The examined root segments were excised from seedlings exposed to the plasmolyticum under aerated conditions, i.e. from plants that were kept in cotton saturated with the mannitol solution. Root segments from each treatment (control, plasmolysis, oryzalin and CB) were sampled at 30 min intervals and measurements never lasted longer than 5 min. Some samples from each treatment were also fixed in 3% v/v glutaraldehyde for 20 min before measurements. Protoplast volume measurements For protoplast volume measurements of non-treated, oryzalin-treated and CB-treated plasmolyzed cells, Hoffler’s numerical approach was followed (see Strugger 1949). According to this method the protoplast is considered as an almost cylindrical body with two hemispherical caps. This approach applies best to the convex plasmolysis pattern, which is exhibited by the vast majority of rhizodermal and root-cap cells used for measurements. Rhizodermal cells included in the measurements were classified according to the length of their longitudinal walls, i.e. parallel to the root axis. Mean protoplast volumes per experimental approach and per cell classes were plotted as a histogram. Measurements from different experimental procedures were compared using the Student’s t-test to identify statistically significant differences. Tubulin immunolabeling Control and mannitol-treated root tips were fixed in 8% w/v paraformaldehyde (PFA) in MSB (50 mM PIPES, 5 mM EGTA, 5 mM MgSO4) pH 6.8 made in 0.25 M glycerol for 1 h. Fixation was followed by washing in MSB. Afterwards, root-tip cell walls were digested in an enzyme cocktail containing 2% w/v cellulysin, 2% w/v driselase, 1% w/v cellulase, 1% w/v pectinase and 0.5% w/v macerozyme for 2 h in MSB made up in 0.25 M glycerol at pH 5.6. In some samples, glycerol was omitted from both the fixative and the enzyme solution. Following brief washing in MSB pH 6.8, root-tips were gently forced through a Pasteur pipette to separate the cells. The cell suspension was then filtered through 200 µm mesh to remove unmacerated tissue. Then it was mildly centrifuged and resuspended in glycerol-free MSB for three times to remove cell debris. Aliquots of the cell suspension were spread on acid-washed, poly-l-lysine-coated coverslips and were allowed to dry. Following that, coverslips were immersed into ice-cold (–20°C) anhydrous acetone supplemented with 5 mM EGTA for 1 h. Acetone treatment rendered the highly condensed cytoplasm much more permeable to the antibodies used for tubulin immunolocalization and labeling. The coverslips were then rapidly transferred to small beakers containing 5% v/v Triton X-100, 2% w/v bovine serum albumin (BSA) and 0.05% v/v Tween-20 in MSB for an additional extraction and blocking step for 1 h. After thorough washing in phosphate-buffered saline (PBS) pH 7.4, the specimens were incubated sequentially with a rat monoclonal anti-α-tubulin antibody (clone YOL 1/34; Harlan Seralab) overnight at 25°C followed by a FITC-conjugated anti-rat IgG (Sigma) at 37°C for 1 h. Both antibodies were used at a 1 : 40 dilution in PBS pH 7.4 supplemented with 1% w/v BSA. Chromatin was stained with 1 µg ml–1 Hoechst 33258 (Sigma) in PBS and finally, specimens were mounted in 0.1% w/v p-phenylenediamine in PBS made up in 90% v/v glycerol at pH 8. Specimens were observed in a Zeiss Axioplan microscope equipped with standard filters for immunofluorescence. Photomicrographs were captured on T-MAX 400 film pushed at 1600 ASA. Electron microscopy For fixation, root tips were immersed in 5% v/v glutaraldehyde, 1–2% w/v tannic acid, 0.2% v/v DMSO (Fassel et al. 1997), in the presence or absence of 4% w/v PFA in 100 mM Na-cacodylate buffer at pH 7.4 for 2–3 h at room temperature. Following extensive washing in Na-cacodylate buffer soluble polysaccharides were extracted in some samples by incubation of the root tips in either 2% w/v EDTA or 1% v/v DMSO in Na-cacodylate buffer. EDTA extraction lasted overnight whereas DMSO extraction was carried out for 6 h (Roland 1978). Afterwards, root tips were washed in Na-cacodylate buffer and postfixed in 1% w/v OsO4 for 5–12 h, at 4°C, in Na-cacodylate buffer. Root tips were subsequently washed in buffer and dehydrated in a graded series of acetone solutions followed by two changes in anhydrous propylene oxide and finally infiltrated in a graded series of Spurr’s resin in propylene oxide. Prior to resin polymerization, root tips were exhaustively incubated in complete resin mixture, at least for 6 d, and specimens were left at 60°C for at least 4 d for polymerization. Semithin and ultrathin sectioning was done using an LKB ultramicrotome. Semithin sections were stained with 1% w/v toluidine blue made in 1% w/v aqueous borax solution. Thin sections were stained with 4% w/v uranyl acetate in 70% v/v ethanol followed by Reynold’s lead citrate. The sections were examined with a Philips 300 TEM. Histochemical reactions For section histochemistry, polysaccharide extraction was minimized by modifying the TEM fixation protocol. In detail, glutaraldehyde fixation was carried out in ice, for less than 30 min. Wash out steps were also minimized, while DMSO and/or EDTA treatments were omitted. OsO4 post-fixation was reduced to 1 h on ice and root-tips were dehydrated in acetone series on ice. In some experiments, root-tips were fixed in OsO4 solely. Insoluble polysaccharides were localized in semithin sections by PAS staining according to Galatis et al. (1978) and in isolated fixed cells after staining with 0.001% w/v calcofluor white in PBS for 10 min. In the latter case the cells were separated using pectinase. Calcofluor staining was also applied in intact PFA-fixed root segments. Assessment of the polymerized tubulin content To compare the quantity of polymeric tubulin between control and plasmolyzed cells, a microscopy-based qualitative assay was used. For this purpose, photomicrographic negatives of anti-tubulin stained control and plasmolyzed cells, were scanned through an Agfa DuoScan scanner, and images were reversed and digitized using Agfa FotoLook software. For further analysis of digital images a suitable macro built on Image Pro-Plus (Media Cybernetics) was used. Through this macro, background fluorescence was subtracted and rectangular non-overlapping areas of a defined magnitude (which was kept the same for every cell examined) were drawn within the cells so that to include as much of the cell as possible. Following background subtraction, all the fluorescing structures were automatically tracked and summed per cell. Since background was selected from tubulin polymer free areas, the sum of all grey levels above background represents the quantity of polymeric tubulin per cell. The results were then plotted as the average quantity of tubulin in the polymeric form per cell versus cell categories (interphase, mitotic, cytokinetic). Similar digital image analysis approaches have been used by other authors to quantify either Mts (Schwarzerová et al. 2002), or actin filaments (Cramer 1999). Acknowledgments We thank Dr M. Issidorides for allowing access facilities to the digital image analysis of the Cozzica Foundation. Thanks are also extended to Mr S. Yietos for developing the suitable macro for automation of tubulin polymer tracking and fluorescence intensity measurements. G. Komis was awarded a scholarship by the State Scholarship Foundation. 1 Corresponding author: E-mail, bgalatis@biol.uoa.gr; Fax, +003-010-7274702. View largeDownload slide Fig. 1 Dividing control root-tip cells after tubulin immunolabeling. Bar, 10 µm for all figures. (A) Cortical Mt array in an interphase cell. (B) Preprophase cell. The arrow marks the PPB. (C, D) Metaphase (C) and anaphase (D) spindle. (E) Phragmoplast. View largeDownload slide Fig. 1 Dividing control root-tip cells after tubulin immunolabeling. Bar, 10 µm for all figures. (A) Cortical Mt array in an interphase cell. (B) Preprophase cell. The arrow marks the PPB. (C, D) Metaphase (C) and anaphase (D) spindle. (E) Phragmoplast. View largeDownload slide Fig. 2 Living rhizodermal cells following 1 min (A) and 120 min (B) plasmolysis with 1 M mannitol, as they appear with DIC optics. The asterisks mark the periplasmic space. Bar, 10 µm. View largeDownload slide Fig. 2 Living rhizodermal cells following 1 min (A) and 120 min (B) plasmolysis with 1 M mannitol, as they appear with DIC optics. The asterisks mark the periplasmic space. Bar, 10 µm. View largeDownload slide Fig. 3 Electron micrographs of cells following plasmolysis with 1 M mannitol for 30 min. (A) The nucleus of a plasmolyzed interphase cell. Note the condensed chromatin. The arrows indicate the nuclear envelope. Bar, 1 µm. (B) Transverse section of a Hechtian strand (large arrow). The small arrows show the macrotubules, while the arrowheads point to the electron dense material of the periplasmic space. Bar, 0.2 µm. View largeDownload slide Fig. 3 Electron micrographs of cells following plasmolysis with 1 M mannitol for 30 min. (A) The nucleus of a plasmolyzed interphase cell. Note the condensed chromatin. The arrows indicate the nuclear envelope. Bar, 1 µm. (B) Transverse section of a Hechtian strand (large arrow). The small arrows show the macrotubules, while the arrowheads point to the electron dense material of the periplasmic space. Bar, 0.2 µm. View largeDownload slide Fig. 4 Tubulin immunolabeled root-tip cells following plasmolysis with 1 M mannitol for 30 min. Bar, 10 µm for all figures. (A) Cortical tubulin strands in an interphase meristematic cell. (B, C) Interphase meristematic cell in a cortical (B) and a median (C) plane. Numerous tubulin strands traverse the cortical (B) and perinuclear (C) cytoplasm. (D) Differentiating root-tip cell displaying numerous endoplasmic tubulin strands. N, nucleus. (E) Differentiating root-tip cell, in which endoplasmic tubulin strands form an impressive radial perinuclear system. N, nucleus. Inset. The nucleus after Hoechst 33258 staining. (F) Optical section through the cortex of differentiating root-tip cell. Note the convergence of tubulin strands on certain sites. View largeDownload slide Fig. 4 Tubulin immunolabeled root-tip cells following plasmolysis with 1 M mannitol for 30 min. Bar, 10 µm for all figures. (A) Cortical tubulin strands in an interphase meristematic cell. (B, C) Interphase meristematic cell in a cortical (B) and a median (C) plane. Numerous tubulin strands traverse the cortical (B) and perinuclear (C) cytoplasm. (D) Differentiating root-tip cell displaying numerous endoplasmic tubulin strands. N, nucleus. (E) Differentiating root-tip cell, in which endoplasmic tubulin strands form an impressive radial perinuclear system. N, nucleus. Inset. The nucleus after Hoechst 33258 staining. (F) Optical section through the cortex of differentiating root-tip cell. Note the convergence of tubulin strands on certain sites. View largeDownload slide Fig. 5 Histograms showing the quantity of tubulin in the polymeric form in control and plasmolyzed root-tip cells, as assessed by digital image analysis. View largeDownload slide Fig. 5 Histograms showing the quantity of tubulin in the polymeric form in control and plasmolyzed root-tip cells, as assessed by digital image analysis. View largeDownload slide Fig. 6 Tubulin immunolabeling in dividing root-tip cells following 30 min plasmolysis with 1 M mannitol. Bar, 10 µm for all figures. (A) Preprophase cell encompassing an atypical PPB (arrow) and numerous cortical tubulin strands outside the PPB region. (B–D) Atypical prophase (B) prometaphase (C) and metaphase (D) spindle. (E) The metaphase chromosomes of the cell shown in D after Hoechst 33258 staining. (F) Atypical anaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (G, H) Early cytokinetic cells possessing an atypical phragmoplast in the interzonal region. (I) Late cytokinetic cell displaying an atypical phragmoplast at the end of the cell plate. (J) The cytokinetic cell shown in I, as it appears with DIC optics. The arrow points to the cell plate. (K) Cytokinetic cell showing an atypical cell plate (arrow) as it appears with DIC optics. View largeDownload slide Fig. 6 Tubulin immunolabeling in dividing root-tip cells following 30 min plasmolysis with 1 M mannitol. Bar, 10 µm for all figures. (A) Preprophase cell encompassing an atypical PPB (arrow) and numerous cortical tubulin strands outside the PPB region. (B–D) Atypical prophase (B) prometaphase (C) and metaphase (D) spindle. (E) The metaphase chromosomes of the cell shown in D after Hoechst 33258 staining. (F) Atypical anaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (G, H) Early cytokinetic cells possessing an atypical phragmoplast in the interzonal region. (I) Late cytokinetic cell displaying an atypical phragmoplast at the end of the cell plate. (J) The cytokinetic cell shown in I, as it appears with DIC optics. The arrow points to the cell plate. (K) Cytokinetic cell showing an atypical cell plate (arrow) as it appears with DIC optics. View largeDownload slide Fig. 7 (A, B) Electron micrographs of macrotubules (arrows) in longitudinal (A) and transverse (B) sections. The arrowheads in B indicate the plasmalemma. P, periplasmic space. Bars, 0.2 µm. (C) Electron micrograph of a putative tubulin paracrystal. Bar, 0.2 µm. View largeDownload slide Fig. 7 (A, B) Electron micrographs of macrotubules (arrows) in longitudinal (A) and transverse (B) sections. The arrowheads in B indicate the plasmalemma. P, periplasmic space. Bars, 0.2 µm. (C) Electron micrograph of a putative tubulin paracrystal. Bar, 0.2 µm. View largeDownload slide Fig. 8 (A) Electron micrograph of a cytokinetic root-tip cell following 30 min plasmolysis with 1 M mannitol. The large arrows show an aberrant cell plate, the small ones the endoplasmic reticulum membrane arrays, while the arrowheads show macrotubule bundles of the atypical phragmoplast. N, nucleus. Bar, 2 µm. (B) Higher magnification of a macrotubule bundle of the atypical phragmoplast shown in A. Bar, 0.2 µm. View largeDownload slide Fig. 8 (A) Electron micrograph of a cytokinetic root-tip cell following 30 min plasmolysis with 1 M mannitol. The large arrows show an aberrant cell plate, the small ones the endoplasmic reticulum membrane arrays, while the arrowheads show macrotubule bundles of the atypical phragmoplast. N, nucleus. Bar, 2 µm. (B) Higher magnification of a macrotubule bundle of the atypical phragmoplast shown in A. Bar, 0.2 µm. View largeDownload slide Fig. 9 Electron (A, B) and light (C) micrographs of root-tip cells following 8 h plasmolysis with 1 M mannitol. (A) Interphase nuclei. The chromatin has resumed the normal organization (cf. with Fig. 3A). Bar, 2 µm. (B) Periplasmic space region in a higher magnification. Electron dense material occupies the whole periplasmic space. CW, cell wall. Bar, 2 µm. (C) Differentiating root-tip cells after PAS staining. The periplasmic spaces (arrows) are filled with material positive to PAS. Bar, 10 µm. View largeDownload slide Fig. 9 Electron (A, B) and light (C) micrographs of root-tip cells following 8 h plasmolysis with 1 M mannitol. (A) Interphase nuclei. The chromatin has resumed the normal organization (cf. with Fig. 3A). Bar, 2 µm. (B) Periplasmic space region in a higher magnification. Electron dense material occupies the whole periplasmic space. CW, cell wall. Bar, 2 µm. (C) Differentiating root-tip cells after PAS staining. The periplasmic spaces (arrows) are filled with material positive to PAS. Bar, 10 µm. View largeDownload slide Fig. 10 (A–D) Tubulin immunolabeling in root-tip cells treated with 1 M mannitol for 8 h. Bar, 10 µm for all figures. (A) Interphase cell showing cortical Mts. (B) Typical metaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (C) Anaphase spindle. (D) Phragmoplast. (E) Oryzalin-treated plasmolyzed root-tip cell. The tubulin strands have been disorganized. View largeDownload slide Fig. 10 (A–D) Tubulin immunolabeling in root-tip cells treated with 1 M mannitol for 8 h. Bar, 10 µm for all figures. (A) Interphase cell showing cortical Mts. (B) Typical metaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (C) Anaphase spindle. (D) Phragmoplast. (E) Oryzalin-treated plasmolyzed root-tip cell. The tubulin strands have been disorganized. View largeDownload slide Fig. 11 Oryzalin-treated living plasmolyzed rhizodermal cells observed with DIC optics. Bar, 10 µm for all figures. (A, B) The same rhizodermal cell after different times of plasmolysis with 1 M mannitol. The number in each figure shows the time of plasmolysis in min. The asterisks indicate the periplasmic space. Oryzalin pre-treatment 1 h. (C) Oryzalin-affected cell displaying numerous Hechtian strands (arrows) in the periplasmic space. Oryzalin pre-treatment 1 h. (D, E) Oryzalin-treated plasmolyzed cells. The number in each figure indicates the time of plasmolysis in min. Note the intense reduction of the protoplast volume. The arrow in D points to a protoplast region, completely detached from the cell wall, while the arrows in E point to subprotoplasts. Oryzalin pre-treatment 24 h. View largeDownload slide Fig. 11 Oryzalin-treated living plasmolyzed rhizodermal cells observed with DIC optics. Bar, 10 µm for all figures. (A, B) The same rhizodermal cell after different times of plasmolysis with 1 M mannitol. The number in each figure shows the time of plasmolysis in min. The asterisks indicate the periplasmic space. Oryzalin pre-treatment 1 h. (C) Oryzalin-affected cell displaying numerous Hechtian strands (arrows) in the periplasmic space. Oryzalin pre-treatment 1 h. (D, E) Oryzalin-treated plasmolyzed cells. The number in each figure indicates the time of plasmolysis in min. Note the intense reduction of the protoplast volume. The arrow in D points to a protoplast region, completely detached from the cell wall, while the arrows in E point to subprotoplasts. Oryzalin pre-treatment 24 h. View largeDownload slide Fig. 12 Histograms showing the volume of the plasmolyzed protoplast, as it has been estimated with Hoeffler’s method in untreated, oryzalin-treated and CB-treated rhizodermal cells, following 30 min plasmolysis with 1 M mannitol. The first three groups of histograms correspond to meristematic and differentiating cells, while the fourth corresponds to mature cells. View largeDownload slide Fig. 12 Histograms showing the volume of the plasmolyzed protoplast, as it has been estimated with Hoeffler’s method in untreated, oryzalin-treated and CB-treated rhizodermal cells, following 30 min plasmolysis with 1 M mannitol. The first three groups of histograms correspond to meristematic and differentiating cells, while the fourth corresponds to mature cells. 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Hyperosmotic Stress Induces Formation of Tubulin Macrotubules in Root-Tip Cells of Triticum turgidum: Their Probable Involvement in Protoplast Volume Control

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Oxford University Press
ISSN
0032-0781
eISSN
1471-9053
DOI
10.1093/pcp/pcf114
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Abstract

Abstract Treatment of root-tip cells of Triticum turgidum with 1 M mannitol solution for 30 min induces microtubule (Mt) disintegration in the plasmolyzed protoplasts. Interphase plasmolyzed cells possess many cortical, perinuclear and endoplasmic macrotubules, 35 nm in mean diameter, forming prominent arrays. In dividing cells macrotubules assemble into aberrant mitotic and cytokinetic apparatuses resulting in the disturbance of cell division. Putative tubulin paracrystals were occasionally observed in plasmolyzed cells. The quantity of polymeric tubulin in plasmolyzed cells exceeds that in control cells. Root-tip cells exposed for 2–8 h to plasmolyticum recover partially, although the volume of the plasmolyzed protoplast does not change detectably. Among other events, the macrotubules are replaced by Mts, chromatin assumes its typical appearance and the cells undergo typical cell divisions. Additionally, polysaccharidic material is found in the periplasmic space. Oryzalin and colchicine treatment induced macrotubule disintegration and a significant reduction of protoplast volume in every plasmolyzed cell type examined, whereas cytochalasin B had only minor effects restricted to differentiated cells. These results suggest that Mt destruction by hyperosmotic stress, and their replacement by tubulin macrotubules and putative tubulin paracrystals is a common feature among angiosperms and that macrotubules are involved in the mechanism of protoplast volume regulation. (Received February 15, 2002; Accepted May 30, 2002) Introduction Evidence accumulated so far shows that in various organisms, the cytoskeleton is sensitive to osmotic stress. Osmotic stress induces microtubule (Mt) disintegration and/or disorganization in the following higher plant cell types: (a) epidermal cells of Pisum sativum (Roberts et al. 1985), Allium cepa (Lang-Pauluzzi and Gunning 2000) and Tradescantia virginiana (Cleary 2001), (b) leaf cells of Spinacea oleracea (Bartolo and Carter 1991), (c) differentiating root cells of Zea mays (Blancaflor and Hasenstein 1995). Besides, hyperosmotic treatment affects Mts in the yeast Saccharomyces cerevisiae (Slaninova et al. 2000) and in different animal systems, as in withdrawing reticulopodia of the protozoan Allogromia (Welnhofer and Travis 1996) and dividing PtK-1 cells (Mullins and Snyder 1989). Recent work has revealed that non-ionic hyperosmotic stress dramatically affects the organization of the Mt cytoskeleton in the leaf cells of Chlorophyton comosum (Komis et al. 2001). In mitotic and cytokinetic cells Mts are disorganized and tubulin is polymerized into macrotubules, 32 nm in mean diameter, and into paracrystals. The former assemble atypical spindles and phragmoplasts accompanied by disturbed cell division. Malformed spindles and phragmoplasts have also been observed in plasmolyzed leaf epidermal cells of T. virginiana (Cleary 2001). In interphase plasmolyzed cells of C. comosum the cortical and endoplasmic Mts are also disintegrated and numerous tubulin paracrystals are assembled. Macrotubules were absent from those cells (Komis et al. 2001). From the above it is evident that although the effects of hyperosmotic stress on the organization of the Mt cytoskeleton are well documented, it was found to induce the formation of macrotubules and putative tubulin paracrystals only in leaf cells of C. comosum. In order to test whether the formation of atypical tubulin polymers is a general response of higher plant cells in hyperosmotic stress, we studied the organization of tubulin cytoskeleton in plasmolyzed root-tip cells of Triticum turgidum. Since there are some reports implicating Mts in certain aspects of animal cell volume regulation (Moustakas et al. 1998), we also studied the possible involvement of tubulin polymers in the regulation of the plasmolyzed protoplast volume through the use of anti-Mt drugs. Results Control root-tip cells Mt organization in meristematic root-tip cells of T. turgidum has been previously described (Zachariadis et al. 2000, Frantzios et al. 2000, Frantzios et al. 2001) and is similar to that described in root-tip cells of other angiosperms. Interphase cortical Mt systems, preprophase Mt bands (PPBs), mitotic spindles, interzonal Mt systems, as well as phragmoplasts at different stages of development were observed in the meristematic cells examined (Fig. 1A–E). Differentiating root-tip cells exhibit cortical Mts and an endoplasmic Mt array, which in some cells radiates from the perinuclear area towards the cell cortex. Plasmolyzed root-tips Pattern of plasmolytic cycle The 1 M mannitol solution induced extensive changes in the organization of most root-tip cells. Water loss results in a reduction of the protoplast volume and the detachment of the plasmalemma from the cell wall (plasmolysis). The extent of plasmolysis varied among different cell types as well as among cells from different root regions. Study of serial semithin sections of root-tips subjected to plasmolysis for 30 min revealed that every cell was plasmolyzed. However, in meristematic cells, the detachment of the protoplast from the cell wall was minimal and restricted to cell corners. Convex plasmolysis was the dominant plasmolysis pattern observed (Fig. 2A, B). Examination of living root-tips with DIC optics revealed that, at least in rhizodermal and root-cap cells, plasmolysis commences immediately after immersion of the root-tips in mannitol solution and is completed within 5 min. Afterwards, the protoplast volume does not change significantly, even in root-tips that remained in the plasmolytic solution for 2–8 h (Fig. 2A, B). The same plasmolysis pattern was induced by 0.5 and 0.7 M mannitol solutions. However, the plasmolyzed cells in these solutions were fewer, while the course of plasmolysis was completed in longer times compared with those induced by 1 M mannitol. In root-tips that had undergone plasmolysis, all the rhizodermal cells deplasmolyzed within 2–3 min when they were placed in distilled water. Root-tip cells following a short mannitol treatment The changes observed in root-tip cells were identical among all the mannitol solutions used (0.5, 0.7 and 1 M). To avoid duplication of the results the organization of cells plasmolyzed with 1 M mannitol will be described only. Transmission electron microscope (TEM) examination of root-tip cells following a short (up to 30 min) mannitol treatment revealed that the plasmolyzed protoplasts were characterized by: (a) an increase in the electron density of the cytoplasm (Fig. 3A), (b) an intense dictyosome activity and the stacking of the endoplasmic reticulum membranes, (c) the ‘coalescence’ of chromatin as well as of chromosomes into a condensed mass, which in interphase cells is largely separated from the nuclear envelope (Fig. 3A). The chromatin changes were also conspicuous at the light microscope level after toluidine blue staining as well as in fluorescence microscopy after staining with Hoechst 33258 (Fig. 4E inset, 6E). In the periplasmic space, i.e. in the space between the protoplast and the cell wall, Hechtian strands and a loosely arranged granular material were found (Fig. 3B). Observation of tubulin immunolabeled cells that were plasmolyzed for 30 min, revealed that they displayed many thick and intensely fluorescing tubulin strands, the organization of which varied according to the cell type and the stage of the cell cycle (Fig. 4A–F). Notably, the quantity of polymeric tubulin per cell in plasmolyzed cells was 30–35% higher than in control cells as assessed by digital image analysis of the specimens (Fig. 5). In interphase meristematic cells many tubulin strands traversed the cortical cytoplasm transversely to the long cell axis or in different directions (Fig. 4A, B) as well as the subcortical cytoplasm and the endoplasm (Fig. 4 C). The subcortical tubulin strands were usually arranged parallel to the long cell axis, while the endoplasmic ones were mainly localized in the perinuclear cytoplasm (Fig. 4C). The preprophase-prophase cells exhibited aberrant PPBs, tubulin strands running through the cortical cytoplasm outside the PPB cortical zone and perinuclear tubulin strands (Fig. 6A, B). In mitotic cells the chromosomes appeared coalesced into one or two chromosome masses (Fig. 6E, F inset). These cells assembled highly elongated and sharply focused atypical spindles (Fig. 6C, D) similar to those of the plasmolyzed leaf cells of C. comosum (Komis et al. 2001). Frequently, tubulin immunoreactivity was localized into chromatin masses (Fig. 6F). The cytokinetic cells formed atypical phragmoplasts made of prominent tubulin polymer bundles (Fig. 6G–I), which accompanied atypical cell plates (Fig. 6J, K). Often the latter appeared to comprise ‘vacuole-like’ dilations (Fig. 6K, 8A), like those previously described in plasmolyzed epidermal cells of Tradescantia (Cleary 2001). Some plasmolyzed cytokinetic root-tip cells of T. turgidum did not form cell plates. Differentiating plasmolyzed root cells possessed numerous endoplasmic tubulin strands, which in some cases were arranged in a radial perinuclear system reaching up the cell cortex (Fig. 4D, E). In addition, many thick cortical tubulin strands were frequently focussed on cortical sites (Fig. 4F). TEM examination revealed that all cells that had been plasmolyzed for 30 min, including the rhizodermal ones, lacked Mts, but possessed macrotubules of diameter ranging between 26 nm and 41 nm. Their mean diameter was 35 nm, while the mean value of Mt diameter in control cells was 22 nm. These values were derived from measurement of 450 macrotubules and 200 Mts. The macrotubule and Mt diameter was measured as described in Komis et al. (2001). In interphase cells single or bundles of macrotubules traversed the cortical cytoplasm and the endoplasm (Fig. 7A, B). Macrotubules traversed some of the Hechtian strands along the whole of their length (Fig. 3B). In dividing cells aberrant mitotic and cytokinetic apparatuses consist of macrotubules identical to those found in interphase cells (Fig. 8A, B). Therefore, the tubulin strands observed in the immunofluorescent specimens consist of macrotubules. The plasmolyzed cells also displayed some elongated electron dense paracrystals, made of fine sheets or filaments (Fig. 7C). The structural similarity of these paracrystals to one category of those found in the plasmolyzed leaf cells of C. comosum (Komis et al. 2001), favors the hypothesis that they consist of tubulin. Root-tip cells following a prolonged mannitol treatment Protoplasts of root-tip cells, which remained in the hypertonic solution for 8 h, showed some structural recovery, although their volume did not increase detectably. The interphase and mitotic cells displayed typical nuclei and chromosomes respectively (Fig. 9A, 10B inset), low electron density of the cytoplasm and rarely endoplasmic reticulum stacks. Moreover, the periplasmic space contained material (Fig. 9B) positive for PAS, calcofluor and toluidine blue staining (Fig. 9C). These tests demonstrate that this material, which is secreted by the protoplast, is of polysaccharidic nature. The majority of the interphase cells that had undergone 2–8 h treatment with mannitol solution were devoid of the thick fluorescent tubulin strands formed during the first 30 min of treatment and possessed fluorescent structures resembling the appearance of Mts of control cells after tubulin immunolabelling (Fig. 10A; cf. Fig 1A). TEM examination confirmed the absence of macrotubules and the appearance of cortical and endoplasmic Mt arrays in those cells. The dividing cells found in the root tips that had undergone long hyperosmotic treatment (up to 8 h) were classified into two groups. The first group includes cells that entered cell division during plasmolysis. The appearance of the chromosomes and the organization of the mitotic spindle and the phragmoplast were similar to those of control cells (Fig. 10B–D; cf. Fig. 1C–E). The second group includes atypical mitotic cells, in which chromosomes appeared normal, but were either dispersed throughout the cytoplasm or were arranged in rosette conformations. These cells exhibited atypical mono-, bi- or multipolar and probably non-functional spindles. They give rise to polyploid or multinucleate cells. TEM examination showed that mitotic and cytokinetic apparatuses of cells plasmolyzed for 8 h consisted of Mts. Anti-Mt drug and cytochalasin B-treated plasmolyzed cells Following tubulin immunofluorescence of oryzalin- and colchicine-treated plasmolyzed cells it was found that thick fluorescing strands were disintegrated. These cells showed fluorescing fragments, which probably represent remnants of the macrotubule strands (Fig. 10E). It is important to note that macrotubule disorganization following the anti-Mt-drug treatment affected plasmolysis. In non-treated rhizodermal cells, plasmolysis was completed in 5 min after the immersion of the root-tips in the mannitol solution, while in the oryzalin treated ones, plasmolysis continued for at least 1 h (Fig. 11A, B; cf. Fig. 2A, B). The protoplast volume after 30 min of plasmolysis was measured in oryzalin-treated and non-treated rhizodermal cells. It was found that in the oryzalin-treated cells, the protoplast volume was 40–45% less than that in the non-treated plasmolyzed cells (Fig. 12). Additionally, in many oryzalin-treated cells the protoplast frequently exhibited concave patterns of plasmolysis (Fig. 11D), while in others the protoplast was divided into subprotoplasts (Fig. 11E), phenomena not observed in the untreated plasmolyzed cells. All oryzalin-treated cells showed more Hechtian strands than the untreated cells (Fig. 11C). Cells pretreated with oryzalin for 24 h displayed a greater protoplast volume reduction than those pretreated for 1 h (Fig. 11D, E; cf. Fig. 11A–C). The response of the plasmolyzed cells to colchicine was similar to that of cells treated with oryzalin. In contrast, cytochalasin B (CB) treatment did not have any appreciable effect on the protoplast shape and volume of meristematic and differentiating plasmolyzed rhizodermal cells (Fig. 12). However, in fully differentiated plasmolyzed rhizodermal cells CB brought about an approximately 20% reduction in protoplast volume (Fig. 12). Discussion General remarks This work shows that root-tip cells of T. turgidum exposed to non-ionic hyperosmotic stress, undergo extensive structural changes and that the affected cells have the ability to overcome certain of the stress effects in the presence of the plasmolytic solution. The main observations of this study are given below. (a) Hyperosmotic stress induced, among other effects, Mt disintegration, formation of tubulin macrotubules and putative tubulin paracrystals and highly condensed chromatin. (b) In interphase cells the macrotubules are numerous and form well-organized cortical and endoplasmic systems, while in the mitotic and cytokinetic cells they assemble aberrant mitotic and cytokinetic apparatuses, disturbing chromosome separation and cell division. (c) During prolonged treatments (2–8 h) of the root-tips with the plasmolytic solution the plasmolyzed protoplasts recover partially, restoring Mt arrays in place of the macrotubules. (d) Macrotubule disassembly after anti-Mt drug treatment extends the duration of plasmolysis, induces a significant decrease of the protoplast volume compared with the untreated plasmolyzed cells and affects the shape of the plasmolyzed protoplasts. Some of these phenomena will be discussed below. Mt-disassembly The present work on T. turgidum, as well as previous results on the plasmolyzed cells of C. comosum (Komis et al. 2001), support the hypothesis that the replacement of Mts by aberrant tubulin polymers is an immediate response of angiosperm cells to hyperosmotic stress. Changes in Mt dynamics and/or organization induced by osmotic stress have also been observed: (a) in higher plant cells (Bartolo and Carter 1991, Blancaflor and Hasenstein 1995, Lang-Pauluzzi and Gunning 2000, Cleary 2001), (b) in fungal cells (Slaninova et al. 2000) and (c) in animal cells (Welnhofer and Travis 1996, Mullins and Snyder 1989). The rapid response of the Mt cytoskeleton to hyperosmotic stress is probably due to rapid and profound changes in cell metabolism. In plant cells the osmotic stress induces: (a) changes in the cytosolic Ca2+ concentration (Bush 1995, Knight et al. 1997, Knight et al. 1998, Cessna et al. 1998, Brownlee et al. 1999, Knight 2000), (b) activation of a mitogen-activated protein kinase cascade (Popping et al. 1996, Mizoguchi et al. 1997, Hirt 2000), (c) triggering of the inositol-signaling pathway (Munnik et al. 2000, Meijer et al. 2001). All these phenomena are directly related to mechanisms controlling Mt dynamics (Desai and Mitchison 1997, Quader 1998, Vaughn and Harper 1998, Bögre et al. 2000) and therefore they can potentially explain Mt disorganization in plasmolyzed root-tip cells of T. turgidum. Alternatively, the mechanical forces imposed upon the protoplast due to volume reduction upon plasmolysis may lead to Mt disruption. The sensitivity of the Mt cytoskeleton to mechanical stress is fairly well documented in plant (Wymer et al. 1996) and animal (Ingber 1997) systems. Moreover, in plants, stomatal pore closure, a process depending on reduction of the guard cell volume, is accompanied by disintegration of the cortical Mt arrays (Jiang et al. 1996, Huang and Wang 1997, Fukuda et al. 1998, Marcus et al. 2001, Yu et al. 2001). Plasmalemma ion channels, including the Ca2+ ones, can be activated by stretch (Cosgrove and Hedrich 1991, Ding and Pickard 1993, Janmey 1998). Their activation by hyperosmotic stress induces Ca2+ transportin the protoplast and triggers processes controlled by Ca2+ as a response to osmotic stress (Brownlee et al. 1999). These processes, among others, may induce Mt disintegration. Finally, Mt disassembly might also be attributed to disruption of the cell wall-to-plasmalemma continuum (Fowler and Quatrano 1997). Formation of abnormal tubulin polymers In the plasmolyzed root tip-cells of T. turgidum, similarly to leaf cells of C. comosum (Komis et al. 2001), Mt disorganization is accompanied by the formation of tubulin macrotubules and putative tubulin paracrystals. Their assembly is probably a general response of plant cells exposed to non-ionic hyperosmotic stress. In plants tubulin macrotubules, probably consisting of more than 13 protofilaments, have also been observed in oil-body cells of Marchantia paleacea (Galatis and Apostolakos 1976, Apostolakos and Galatis 1998) and in Al-treated root-tip cells of T. turgidum (Frantzios et al. 2000, Frantzios et al. 2001). Komis et al. (2001) suggested that in plasmolyzed leaf cells of C. comosum the cytosolic Ca2+ concentration that prevails at the beginning of plasmolysis leads to Mt disorganization, and that later this is changed, leading to tubulin macrotubule and paracrystal formation. The same mechanism may function in the plasmolyzed root-tip cells of T. turgidum. The changes of cytosolic Ca2+ concentration estimated in different plant systems which underwent hyperosmotic stress (Takahashi et al. 1997, Cessna et al. 1998) support the above hypothesis. Moreover, the formation of macrotubules and tubulin paracrystals in vitro is defined by interactions between tubulin protofilaments, regulated by Ca2+ concentration (Vater et al. 1997). In hyperosmotically treated root-tip cells the quantity of polymeric tubulin is 30–35% higher when compared with control cells, an observation also made in C. comosum (Komis et al. 2001). Therefore, the induction of excessive tubulin polymerization may be considered as a consistent response of the plant cells to hyperosmotic stress. In contrast to the plasmolyzed interphase leaf cells of C. comosusm, interphase T. turgidum root-tip cells form cortical macrotubules. In the interphase root-tip cells of the examined plant the macrotubules form a very prominent cortical cytoskeletal system. This difference may reflect response differences between leaf cells and root-tip cells to hyperosmotic treatment. The cytoskeletal response could be coupled to different signaling pathways between these systems. Dividing T. turgidum root-tip cells form atypical spindles and phragmoplasts, a phenomenon also observed in hyperosmotically treated leaf cells of C. comosum (Komis et al. 2001) and T. virginiana (Cleary 2001). Therefore, the organization of atypical mitotic and cytokinetic apparatuses is a general cell reaction to hyperosmotic stress (see also Mullins and Snyder 1989). Probable macrotubule function(s) Following destruction of macrotubules by treatment with anti-Mt drugs, the protoplast volume diminution lasted for 2–3 h. Further exposure of roots to plasmolytic solutions supplemented with oryzalin resulted in excessive necrosis. These observations indicate a direct involvement of macrotubules in the regulatory mechanism of plasmolyzed protoplast volume in the root-tip cells. This was somewhat unexpected, given that most animal cells under anisosmotic conditions retain their volume mainly by the activity of actin filaments (Papakonstanti et al. 2000, Szaszi et al. 2000). In plasmolyzed leaf cells of C. comosum the mechanism by which the plasmolyzed protoplasts control their volume is also based on actin filaments (Komis et al. 2002). The minimal contribution of actin filaments in this process in root-tip cells of T. turgidum was demonstrated by the treatment of the plasmolyzed cells with CB. This treatment does not affect the volume of the plasmolyzed protoplast, nor does it affect the pattern of plasmolysis in meristematic and differentiating cells. In fully differentiated cells, however, the role of actin filaments in protoplast volume regulation is somewhat increased. In those cells the application of CB affects the extent of protoplast volume reduction by 20%. Nevertheless, in this case macrotubules play a dominant role in volume regulation as well, since oryzalin exerts drastic effects in protoplast volume reduction. There are some reports implicating Mts in many aspects of animal cell volume regulation. Mt arrays have been shown to activate both ion channels, generally responsible for short-term volume regulatory events and osmolyte transporters, implicated in long-term volume regulation events (Moustakas et al. 1998). The role of Mts in aquaporin clustering is well documented in animal systems often exposed to anisosmotic conditions (Sabolic et al. 1995). In plants, Mts are required upstream to the ionic events (i.e. H+ efflux and K+ influx), leading to stomatal pore opening (Marcus et al. 2001, Yu et al. 2001). In other plant cell types Mts also appear to be involved in the control of the activity of the plasmalemma ion channels (Thion et al. 1996, Thion et al. 1998). Considering the above-mentioned reports it may be suggested that in the root-tip cells of T. turgidum the plasmolyzed protoplast principally controls its volume by the formation of macrotubules. They may regulate the plasmalemma and tonoplast ion channel activity under hyperosmotic conditions and/or the distribution of aquaporins, water transporters that might be responsible for the rehydration of root-tip cells during prolonged hypertonic conditions. The macrotubules in T. turgidum may offer mechanical support to the protoplast to resist forces exerted on it during plasmolysis, a hypothesis also made for tubulin paracrystals in plasmolyzed leaf cells of C.comosum (Komis et al. 2001). Animal cells under mechanical stress reinforce the cortical cytoplasm at the sites of extensive force application with additional cytoskeletal elements (Ingber 1997, Chicurel et al. 1998, Ko and McCulloch 2000). Considering that during plasmolysis stretching forces are generated, which might produce an injury event at the plasmalemma (Oparka 1994) and that the Hechtian strands are under high tensional forces (Buer et al. 2000), the cortical macrotubules as well as the endoplasmic ones may also function as a strong supportive system. Recovery of the plasmolyzed protoplasts Notably root-tip cells of T. turgidum are able to recover from the severe effects on their structure and function caused by high extracellular osmolarity (see also Blancaflor and Hasenstein 1995). Further more, this ability developed during the course of the plasmolytic treatment. Although this became mostly evident after 8 h of treatment, early signs of recovery were recognizable as early as 1.5–2 h after the exposure of roots to the plasmolytic medium. The ability of the plasmolyzed protoplast to counteract, at least partially, the hyperosmotic stress can safely be attributed to cortical macrotubule formation. Indeed, oryzalin-treated cells are necrotized after a few hours in the hypersmoticum. Among the prominent changes of the recovering protoplasts are the chromatin reorganization, the decrease in electron density of the cytoplasm, the disappearance of macrotubules and the reappearance of Mts, and the return of normal cell divisions. The data of this work show that the formation of abnormal tubulin polymers is a reversible phenomenon. The recovery of the plasmolyzed protoplasts can be explained assuming the activation of a mechanism regulating the protoplast volume and the cytoplasm osmolarity. Deposition of polysaccharidic material in the periplasmic space may prevent protoplast enlargement in the recovering cells during prolonged plasmolysis. Deposition of wall materials in the periplasmic space has also been described in other plasmolyzed cell types (Robinson and Cummins 1976, Schnepf et al. 1986). Material and Methods Plant material and treatments Caryopses of T. turgidum L. var. durum Raddi were imbibed with distilled water for 36–48 h in darkness, at 25°C. The seedlings were then treated with an aqueous 0.5 M or 0.7 M or 1 M mannitol solution on moistened cotton for 5 min, 30 min, 2, 4, 8 and 12 h. In addition, roots treated with 10 µM oryzalin for 1–24 h or 2 mM colchicine for 1–24 h or 100 µM CB for 1 h, were subjected to 1 M mannitol treatment further supplemented with anti-Mt drugs or CB. To minimize anoxic effects on the root segments immersed for a long time in the plasmolytic solution, the following precautions were taken. The examined root segments were excised from seedlings exposed to the plasmolyticum under aerated conditions, i.e. from plants that were kept in cotton saturated with the mannitol solution. Root segments from each treatment (control, plasmolysis, oryzalin and CB) were sampled at 30 min intervals and measurements never lasted longer than 5 min. Some samples from each treatment were also fixed in 3% v/v glutaraldehyde for 20 min before measurements. Protoplast volume measurements For protoplast volume measurements of non-treated, oryzalin-treated and CB-treated plasmolyzed cells, Hoffler’s numerical approach was followed (see Strugger 1949). According to this method the protoplast is considered as an almost cylindrical body with two hemispherical caps. This approach applies best to the convex plasmolysis pattern, which is exhibited by the vast majority of rhizodermal and root-cap cells used for measurements. Rhizodermal cells included in the measurements were classified according to the length of their longitudinal walls, i.e. parallel to the root axis. Mean protoplast volumes per experimental approach and per cell classes were plotted as a histogram. Measurements from different experimental procedures were compared using the Student’s t-test to identify statistically significant differences. Tubulin immunolabeling Control and mannitol-treated root tips were fixed in 8% w/v paraformaldehyde (PFA) in MSB (50 mM PIPES, 5 mM EGTA, 5 mM MgSO4) pH 6.8 made in 0.25 M glycerol for 1 h. Fixation was followed by washing in MSB. Afterwards, root-tip cell walls were digested in an enzyme cocktail containing 2% w/v cellulysin, 2% w/v driselase, 1% w/v cellulase, 1% w/v pectinase and 0.5% w/v macerozyme for 2 h in MSB made up in 0.25 M glycerol at pH 5.6. In some samples, glycerol was omitted from both the fixative and the enzyme solution. Following brief washing in MSB pH 6.8, root-tips were gently forced through a Pasteur pipette to separate the cells. The cell suspension was then filtered through 200 µm mesh to remove unmacerated tissue. Then it was mildly centrifuged and resuspended in glycerol-free MSB for three times to remove cell debris. Aliquots of the cell suspension were spread on acid-washed, poly-l-lysine-coated coverslips and were allowed to dry. Following that, coverslips were immersed into ice-cold (–20°C) anhydrous acetone supplemented with 5 mM EGTA for 1 h. Acetone treatment rendered the highly condensed cytoplasm much more permeable to the antibodies used for tubulin immunolocalization and labeling. The coverslips were then rapidly transferred to small beakers containing 5% v/v Triton X-100, 2% w/v bovine serum albumin (BSA) and 0.05% v/v Tween-20 in MSB for an additional extraction and blocking step for 1 h. After thorough washing in phosphate-buffered saline (PBS) pH 7.4, the specimens were incubated sequentially with a rat monoclonal anti-α-tubulin antibody (clone YOL 1/34; Harlan Seralab) overnight at 25°C followed by a FITC-conjugated anti-rat IgG (Sigma) at 37°C for 1 h. Both antibodies were used at a 1 : 40 dilution in PBS pH 7.4 supplemented with 1% w/v BSA. Chromatin was stained with 1 µg ml–1 Hoechst 33258 (Sigma) in PBS and finally, specimens were mounted in 0.1% w/v p-phenylenediamine in PBS made up in 90% v/v glycerol at pH 8. Specimens were observed in a Zeiss Axioplan microscope equipped with standard filters for immunofluorescence. Photomicrographs were captured on T-MAX 400 film pushed at 1600 ASA. Electron microscopy For fixation, root tips were immersed in 5% v/v glutaraldehyde, 1–2% w/v tannic acid, 0.2% v/v DMSO (Fassel et al. 1997), in the presence or absence of 4% w/v PFA in 100 mM Na-cacodylate buffer at pH 7.4 for 2–3 h at room temperature. Following extensive washing in Na-cacodylate buffer soluble polysaccharides were extracted in some samples by incubation of the root tips in either 2% w/v EDTA or 1% v/v DMSO in Na-cacodylate buffer. EDTA extraction lasted overnight whereas DMSO extraction was carried out for 6 h (Roland 1978). Afterwards, root tips were washed in Na-cacodylate buffer and postfixed in 1% w/v OsO4 for 5–12 h, at 4°C, in Na-cacodylate buffer. Root tips were subsequently washed in buffer and dehydrated in a graded series of acetone solutions followed by two changes in anhydrous propylene oxide and finally infiltrated in a graded series of Spurr’s resin in propylene oxide. Prior to resin polymerization, root tips were exhaustively incubated in complete resin mixture, at least for 6 d, and specimens were left at 60°C for at least 4 d for polymerization. Semithin and ultrathin sectioning was done using an LKB ultramicrotome. Semithin sections were stained with 1% w/v toluidine blue made in 1% w/v aqueous borax solution. Thin sections were stained with 4% w/v uranyl acetate in 70% v/v ethanol followed by Reynold’s lead citrate. The sections were examined with a Philips 300 TEM. Histochemical reactions For section histochemistry, polysaccharide extraction was minimized by modifying the TEM fixation protocol. In detail, glutaraldehyde fixation was carried out in ice, for less than 30 min. Wash out steps were also minimized, while DMSO and/or EDTA treatments were omitted. OsO4 post-fixation was reduced to 1 h on ice and root-tips were dehydrated in acetone series on ice. In some experiments, root-tips were fixed in OsO4 solely. Insoluble polysaccharides were localized in semithin sections by PAS staining according to Galatis et al. (1978) and in isolated fixed cells after staining with 0.001% w/v calcofluor white in PBS for 10 min. In the latter case the cells were separated using pectinase. Calcofluor staining was also applied in intact PFA-fixed root segments. Assessment of the polymerized tubulin content To compare the quantity of polymeric tubulin between control and plasmolyzed cells, a microscopy-based qualitative assay was used. For this purpose, photomicrographic negatives of anti-tubulin stained control and plasmolyzed cells, were scanned through an Agfa DuoScan scanner, and images were reversed and digitized using Agfa FotoLook software. For further analysis of digital images a suitable macro built on Image Pro-Plus (Media Cybernetics) was used. Through this macro, background fluorescence was subtracted and rectangular non-overlapping areas of a defined magnitude (which was kept the same for every cell examined) were drawn within the cells so that to include as much of the cell as possible. Following background subtraction, all the fluorescing structures were automatically tracked and summed per cell. Since background was selected from tubulin polymer free areas, the sum of all grey levels above background represents the quantity of polymeric tubulin per cell. The results were then plotted as the average quantity of tubulin in the polymeric form per cell versus cell categories (interphase, mitotic, cytokinetic). Similar digital image analysis approaches have been used by other authors to quantify either Mts (Schwarzerová et al. 2002), or actin filaments (Cramer 1999). Acknowledgments We thank Dr M. Issidorides for allowing access facilities to the digital image analysis of the Cozzica Foundation. Thanks are also extended to Mr S. Yietos for developing the suitable macro for automation of tubulin polymer tracking and fluorescence intensity measurements. G. Komis was awarded a scholarship by the State Scholarship Foundation. 1 Corresponding author: E-mail, bgalatis@biol.uoa.gr; Fax, +003-010-7274702. View largeDownload slide Fig. 1 Dividing control root-tip cells after tubulin immunolabeling. Bar, 10 µm for all figures. (A) Cortical Mt array in an interphase cell. (B) Preprophase cell. The arrow marks the PPB. (C, D) Metaphase (C) and anaphase (D) spindle. (E) Phragmoplast. View largeDownload slide Fig. 1 Dividing control root-tip cells after tubulin immunolabeling. Bar, 10 µm for all figures. (A) Cortical Mt array in an interphase cell. (B) Preprophase cell. The arrow marks the PPB. (C, D) Metaphase (C) and anaphase (D) spindle. (E) Phragmoplast. View largeDownload slide Fig. 2 Living rhizodermal cells following 1 min (A) and 120 min (B) plasmolysis with 1 M mannitol, as they appear with DIC optics. The asterisks mark the periplasmic space. Bar, 10 µm. View largeDownload slide Fig. 2 Living rhizodermal cells following 1 min (A) and 120 min (B) plasmolysis with 1 M mannitol, as they appear with DIC optics. The asterisks mark the periplasmic space. Bar, 10 µm. View largeDownload slide Fig. 3 Electron micrographs of cells following plasmolysis with 1 M mannitol for 30 min. (A) The nucleus of a plasmolyzed interphase cell. Note the condensed chromatin. The arrows indicate the nuclear envelope. Bar, 1 µm. (B) Transverse section of a Hechtian strand (large arrow). The small arrows show the macrotubules, while the arrowheads point to the electron dense material of the periplasmic space. Bar, 0.2 µm. View largeDownload slide Fig. 3 Electron micrographs of cells following plasmolysis with 1 M mannitol for 30 min. (A) The nucleus of a plasmolyzed interphase cell. Note the condensed chromatin. The arrows indicate the nuclear envelope. Bar, 1 µm. (B) Transverse section of a Hechtian strand (large arrow). The small arrows show the macrotubules, while the arrowheads point to the electron dense material of the periplasmic space. Bar, 0.2 µm. View largeDownload slide Fig. 4 Tubulin immunolabeled root-tip cells following plasmolysis with 1 M mannitol for 30 min. Bar, 10 µm for all figures. (A) Cortical tubulin strands in an interphase meristematic cell. (B, C) Interphase meristematic cell in a cortical (B) and a median (C) plane. Numerous tubulin strands traverse the cortical (B) and perinuclear (C) cytoplasm. (D) Differentiating root-tip cell displaying numerous endoplasmic tubulin strands. N, nucleus. (E) Differentiating root-tip cell, in which endoplasmic tubulin strands form an impressive radial perinuclear system. N, nucleus. Inset. The nucleus after Hoechst 33258 staining. (F) Optical section through the cortex of differentiating root-tip cell. Note the convergence of tubulin strands on certain sites. View largeDownload slide Fig. 4 Tubulin immunolabeled root-tip cells following plasmolysis with 1 M mannitol for 30 min. Bar, 10 µm for all figures. (A) Cortical tubulin strands in an interphase meristematic cell. (B, C) Interphase meristematic cell in a cortical (B) and a median (C) plane. Numerous tubulin strands traverse the cortical (B) and perinuclear (C) cytoplasm. (D) Differentiating root-tip cell displaying numerous endoplasmic tubulin strands. N, nucleus. (E) Differentiating root-tip cell, in which endoplasmic tubulin strands form an impressive radial perinuclear system. N, nucleus. Inset. The nucleus after Hoechst 33258 staining. (F) Optical section through the cortex of differentiating root-tip cell. Note the convergence of tubulin strands on certain sites. View largeDownload slide Fig. 5 Histograms showing the quantity of tubulin in the polymeric form in control and plasmolyzed root-tip cells, as assessed by digital image analysis. View largeDownload slide Fig. 5 Histograms showing the quantity of tubulin in the polymeric form in control and plasmolyzed root-tip cells, as assessed by digital image analysis. View largeDownload slide Fig. 6 Tubulin immunolabeling in dividing root-tip cells following 30 min plasmolysis with 1 M mannitol. Bar, 10 µm for all figures. (A) Preprophase cell encompassing an atypical PPB (arrow) and numerous cortical tubulin strands outside the PPB region. (B–D) Atypical prophase (B) prometaphase (C) and metaphase (D) spindle. (E) The metaphase chromosomes of the cell shown in D after Hoechst 33258 staining. (F) Atypical anaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (G, H) Early cytokinetic cells possessing an atypical phragmoplast in the interzonal region. (I) Late cytokinetic cell displaying an atypical phragmoplast at the end of the cell plate. (J) The cytokinetic cell shown in I, as it appears with DIC optics. The arrow points to the cell plate. (K) Cytokinetic cell showing an atypical cell plate (arrow) as it appears with DIC optics. View largeDownload slide Fig. 6 Tubulin immunolabeling in dividing root-tip cells following 30 min plasmolysis with 1 M mannitol. Bar, 10 µm for all figures. (A) Preprophase cell encompassing an atypical PPB (arrow) and numerous cortical tubulin strands outside the PPB region. (B–D) Atypical prophase (B) prometaphase (C) and metaphase (D) spindle. (E) The metaphase chromosomes of the cell shown in D after Hoechst 33258 staining. (F) Atypical anaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (G, H) Early cytokinetic cells possessing an atypical phragmoplast in the interzonal region. (I) Late cytokinetic cell displaying an atypical phragmoplast at the end of the cell plate. (J) The cytokinetic cell shown in I, as it appears with DIC optics. The arrow points to the cell plate. (K) Cytokinetic cell showing an atypical cell plate (arrow) as it appears with DIC optics. View largeDownload slide Fig. 7 (A, B) Electron micrographs of macrotubules (arrows) in longitudinal (A) and transverse (B) sections. The arrowheads in B indicate the plasmalemma. P, periplasmic space. Bars, 0.2 µm. (C) Electron micrograph of a putative tubulin paracrystal. Bar, 0.2 µm. View largeDownload slide Fig. 7 (A, B) Electron micrographs of macrotubules (arrows) in longitudinal (A) and transverse (B) sections. The arrowheads in B indicate the plasmalemma. P, periplasmic space. Bars, 0.2 µm. (C) Electron micrograph of a putative tubulin paracrystal. Bar, 0.2 µm. View largeDownload slide Fig. 8 (A) Electron micrograph of a cytokinetic root-tip cell following 30 min plasmolysis with 1 M mannitol. The large arrows show an aberrant cell plate, the small ones the endoplasmic reticulum membrane arrays, while the arrowheads show macrotubule bundles of the atypical phragmoplast. N, nucleus. Bar, 2 µm. (B) Higher magnification of a macrotubule bundle of the atypical phragmoplast shown in A. Bar, 0.2 µm. View largeDownload slide Fig. 8 (A) Electron micrograph of a cytokinetic root-tip cell following 30 min plasmolysis with 1 M mannitol. The large arrows show an aberrant cell plate, the small ones the endoplasmic reticulum membrane arrays, while the arrowheads show macrotubule bundles of the atypical phragmoplast. N, nucleus. Bar, 2 µm. (B) Higher magnification of a macrotubule bundle of the atypical phragmoplast shown in A. Bar, 0.2 µm. View largeDownload slide Fig. 9 Electron (A, B) and light (C) micrographs of root-tip cells following 8 h plasmolysis with 1 M mannitol. (A) Interphase nuclei. The chromatin has resumed the normal organization (cf. with Fig. 3A). Bar, 2 µm. (B) Periplasmic space region in a higher magnification. Electron dense material occupies the whole periplasmic space. CW, cell wall. Bar, 2 µm. (C) Differentiating root-tip cells after PAS staining. The periplasmic spaces (arrows) are filled with material positive to PAS. Bar, 10 µm. View largeDownload slide Fig. 9 Electron (A, B) and light (C) micrographs of root-tip cells following 8 h plasmolysis with 1 M mannitol. (A) Interphase nuclei. The chromatin has resumed the normal organization (cf. with Fig. 3A). Bar, 2 µm. (B) Periplasmic space region in a higher magnification. Electron dense material occupies the whole periplasmic space. CW, cell wall. Bar, 2 µm. (C) Differentiating root-tip cells after PAS staining. The periplasmic spaces (arrows) are filled with material positive to PAS. Bar, 10 µm. View largeDownload slide Fig. 10 (A–D) Tubulin immunolabeling in root-tip cells treated with 1 M mannitol for 8 h. Bar, 10 µm for all figures. (A) Interphase cell showing cortical Mts. (B) Typical metaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (C) Anaphase spindle. (D) Phragmoplast. (E) Oryzalin-treated plasmolyzed root-tip cell. The tubulin strands have been disorganized. View largeDownload slide Fig. 10 (A–D) Tubulin immunolabeling in root-tip cells treated with 1 M mannitol for 8 h. Bar, 10 µm for all figures. (A) Interphase cell showing cortical Mts. (B) Typical metaphase spindle. Inset. The chromosomes after Hoechst 33258 staining. (C) Anaphase spindle. (D) Phragmoplast. (E) Oryzalin-treated plasmolyzed root-tip cell. The tubulin strands have been disorganized. View largeDownload slide Fig. 11 Oryzalin-treated living plasmolyzed rhizodermal cells observed with DIC optics. Bar, 10 µm for all figures. (A, B) The same rhizodermal cell after different times of plasmolysis with 1 M mannitol. The number in each figure shows the time of plasmolysis in min. The asterisks indicate the periplasmic space. Oryzalin pre-treatment 1 h. (C) Oryzalin-affected cell displaying numerous Hechtian strands (arrows) in the periplasmic space. Oryzalin pre-treatment 1 h. (D, E) Oryzalin-treated plasmolyzed cells. The number in each figure indicates the time of plasmolysis in min. Note the intense reduction of the protoplast volume. The arrow in D points to a protoplast region, completely detached from the cell wall, while the arrows in E point to subprotoplasts. Oryzalin pre-treatment 24 h. View largeDownload slide Fig. 11 Oryzalin-treated living plasmolyzed rhizodermal cells observed with DIC optics. Bar, 10 µm for all figures. (A, B) The same rhizodermal cell after different times of plasmolysis with 1 M mannitol. The number in each figure shows the time of plasmolysis in min. The asterisks indicate the periplasmic space. Oryzalin pre-treatment 1 h. (C) Oryzalin-affected cell displaying numerous Hechtian strands (arrows) in the periplasmic space. Oryzalin pre-treatment 1 h. (D, E) Oryzalin-treated plasmolyzed cells. The number in each figure indicates the time of plasmolysis in min. Note the intense reduction of the protoplast volume. The arrow in D points to a protoplast region, completely detached from the cell wall, while the arrows in E point to subprotoplasts. Oryzalin pre-treatment 24 h. View largeDownload slide Fig. 12 Histograms showing the volume of the plasmolyzed protoplast, as it has been estimated with Hoeffler’s method in untreated, oryzalin-treated and CB-treated rhizodermal cells, following 30 min plasmolysis with 1 M mannitol. The first three groups of histograms correspond to meristematic and differentiating cells, while the fourth corresponds to mature cells. View largeDownload slide Fig. 12 Histograms showing the volume of the plasmolyzed protoplast, as it has been estimated with Hoeffler’s method in untreated, oryzalin-treated and CB-treated rhizodermal cells, following 30 min plasmolysis with 1 M mannitol. The first three groups of histograms correspond to meristematic and differentiating cells, while the fourth corresponds to mature cells. 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Plant and Cell PhysiologyOxford University Press

Published: Aug 15, 2002

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