TY - JOUR
AU1 - Komis, G.
AU2 - Apostolakos, P.
AU3 - Galatis, B.
AB - Abstract Actin filament (AF) organization was studied during the plasmolytic cycle in leaf cells of Chlorophytoncomosum Thunb. In most cells the hyperosmotic treatment induced convex or concave plasmolysis and intense reorganization of the AF cytoskeleton. Thin cortical AFs disappeared and numerous cortical, subcortical and endoplasmic AFs arranged in thick and well‐organized bundles were formed. Plasmolysed cells displayed a significant increase in the overall AF content compared with the control cells. Cortical AF bundles were preferentially localized in the shrunken protoplast areas, lining the detached plasmalemma regions. The endoplasmic AF bundles were mainly found in the perinuclear cytoplasm and on the tonoplast surface. AFs also traversed some of the Hechtian strands. AF disorganization after cytochalasin B (CB) treatment induced dramatic changes in the pattern of plasmolysis, which lasted for a longer time and led to a greater decrease of the protoplast volume compared to the untreated cells. In many of the above cells the protoplasts assumed an ‘amoeboid’ form and were often subdivided into sub‐protoplasts. Soon after the removal of the plasmolytic solution both CB‐treated and untreated cells were deplasmolysed, while the AF cytoskeleton gradually reassumed the organization observed in the control cells. The findings of this study revealed for the first time in angiosperm cells that plasmolysis triggers an extensive reorganization of the AF cytoskeleton, which is involved in the regulation of protoplast shape and volume. The probable mechanism(s) leading to AF reorganization as well as the function(s) of the atypical AF arrays in plasmolysed cells are discussed. Key words: Actin filaments, Chlorophyton comosum, hyperosmotic stress, leaf cells, plasmolytic cycle. Received 3 January 2002; Accepted 4 April 2002 Introduction During plasmolysis of plant cells the plasmalemma is detached from the cell wall and the protoplast volume is significantly reduced, while the intracellular architecture experiences intense mechanical perturbations due to compaction (Oparka, 1994). In a previous study it was demonstrated that in plasmolysed dividing leaf cells of Chlorophyton comosum all microtubule arrays were disintegrated and free tubulin was incorporated into macrotubules and tubulin paracrystals. These atypical tubulin polymers are elongated, straight and rigid structures, which often appeared interconnected into complex networks (Komis et al., 2001). It has been suggested that those may offer mechanical support to the protoplast to resist forces exerted on it during plasmolysis (Komis et al., 2001). In the present study the effects of the hyperosmotic stress imposed by mannitol solutions on the organization of the actin filament (AF) cytoskeleton in the leaf cells of C. comosum were investigated. The disturbance of the AF organization in plasmolysed cells was expected, considering that, in plant cells, the osmotic stress induces (a) changes in cytosolic Ca2+ concentration (Busch, 1995; Knight et al., 1997, 1998; Cessna et al., 1998; Brownlee et al., 1999; Knight 2000); (b) activation of a mitogen‐activated protein kinase cascade (Hirt, 2000); and (c) rapid synthesis of polyphosphoinositides, which are part of the inositol signalling system (Munnik et al., 2000; Meijer et al., 2001). All these phenomena are immediately related to the mechanism(s) regulating AF organization (Janmey, 1994; Richelme et al., 2000; Sullivan et al., 2000). As far as is known the effect(s) of the hyperosmotic stress on AF organization in angiosperms have been studied only in epidermal cells of Alliumcepa (Lang‐Pauluzzi and Gunning, 2000) and Tradescantia virginiana (Cleary, 2001; see also Cleary cited in Gunning and Steer, 1996). In the plasmolysed epidermal cells of Allium cepa the cortical AFs were disorganized, while the other AF arrays were not affected detectably. Cleary (2001) studied the effect of plasmolysis mainly on asymmetrical divisions involved in stomata formation of Tradescantia virginiana. Moreover, AF rearrangement is involved in cell volume regulation in different eukaryotic cell systems that are often exposed to anisosmotic conditions. The AF cytoskeleton respond to osmotic stress in budding yeast (Chowdhury et al., 1992; Slaninova et al., 2000; Ooms et al., 2000), filamentus fungi (Bachewich and Heath, 1997), Dictyostelium discoideum (Aizawa et al., 1999; Zischka et al., 1999) and many different mammalian cell lines (Rizoli et al., 2000; Kapus et al., 2000). In particular, the role of AFs in many aspects of cell volume regulation in animal cells is well documented (Henson, 1999; Papakonstanti et al., 2000). Considering the above information, the probable involvement of AFs in volume regulation of the plasmolysed protoplasts of the leaf cells of C. comosum was also examined in the present work. This phenomenon has not been investigated so far. Materials and methods Plasmolysis Small leaf segments (approximately 2 mm in length) of C. comosum Thunb., corresponding to the leaf base, were excised and immersed in aqueous solutions of 0.3, 0.5 and 1 M mannitol for 30 min and were subsequently processed for AF localization. For examination of living material, the thinner leaf segments were chosen due to their transparency. Those were immersed into a drop of plasmolytic solution on a microscope slide, rapidly covered under a clean coverslip and monitored using the DIC optics of a Zeiss Axioplan microscope. In these specimens epidermal as well as subepidermal cells can be examined. Photomicrographs were taken on T‐MAX 400 film overrated at 1600 ASA. Cytochalasin‐B treatment Leaf segments were incubated in an aqueous solution of 100 µM cytochalasin B (CB; Sigma) for 30 min prior to plasmolysis. Subsequently, they were plasmolysed with 1 M aqueous mannitol solution further supplemented with 100 µM CB. Some of the leaf pieces were processed for AF labelling whereas some others were directly observed by DIC optics and photographed as described above. Additionally, after plasmolysis some leaf pieces were briefly fixed in 1% OsO4 in 1 M mannitol to allow long‐term observations on protoplast morphology. Deplasmolysis Following plasmolysis in 1 M mannitol, both CB‐treated and untreated leaf segments were returned to distilled water. Subsequently, they were either processed for AF localization or were monitored as previously described. For the observation of living material, leaf segments were first allowed to adhere on acid‐washed, poly‐l‐lysine coated coverslips. Subsequently, they were placed on top of a drop of plasmolytic solution and were monitored for a period of 30 min. Afterwards, plain distilled water was allowed to diffuse from one edge of the coverslip, while the excess of liquid was removed from the opposite site by the aid of filter paper. The immobilization of leaf segments prevented their dislocation during liquid exchange, thus allowing the observation of the same cells during the plasmolytic cycle. To test the viability of the CB‐treated or untreated cells after a plasmolytic cycle, some of the leaf pieces were re‐exposed to hyperosmotic treatment of the same magnitude. AF localization For staining of AFs in control and plasmolysed leaf cells, a fluorophore‐conjugated phalloidin‐staining regime was employed. The staining protocols used were based on previously published ones with extensive modifications and thus will be described in detail. All chemicals were purchased from Sigma unless stated otherwise. Control and treated leaf segments were fixed for 45 min in formaldehyde freshly prepared from hydrolysis of 4% w/v paraformaldehyde in either PEMD (50 mM K‐PIPES, 5 mM EGTA, 5 mM MgSO4, 5% v/v DMSO) pH 6.8 or phosphate‐buffered saline (PBS) pH 7.4 both supplemented with 1 M glycerol and 5 IUs of AlexaFluor 568‐conjugated phalloidin (Molecular Probes) per ml of fixative. Alternatively, prior to fixation, leaf segments were pretreated with the bifunctional protein crosslinker m‐maleimidobenzol‐N‐hydroxysuccinimide ester (MBS; 500 µM in PEMD plus 0.05% v/v Triton X‐100) for 30 min (Panteris et al., 1992, and references therein). For control experiments, leaf pieces were immersed in distilled water for 30 min before AF localization. Afterwards, specimens were thoroughly washed in PEMD for 30 min and they where then subjected to enzymatic digestion of the cell wall material using a cocktail of 1% cellulase Onozuka R‐10, 1% macerozyme (both from Yakult Honsha) and 3% pectinase (Fluka) in PEMD, pH 6.8 for 2 h. Leaf segments were then washed in PEMD and were subsequently forced through a wide‐bore Pasteur pipette in order to release cells. The resulting cell suspension was then filtered through a 200 µm mesh to remove unmacerated tissue. The cell suspension was mildly centrifuged at 500 rpm for 10 min in a benchtop clinical centrifuge. The supernatant containing cell debris was discarded and the loose pellet was resuspended in PEM (PEMD without DMSO). This procedure was repeated twice. Then 20 µl aliquots of the cell suspension were spread onto poly‐l‐lysine coated coverslips and were allowed to settle for 5 min in a humid chamber to prevent cells from air‐drying. After cells had adhered, they were permeabilized with 3% v/v Triton X‐100 in PBS pH 7.4, supplemented with 5 IUs AlexaFluor 568‐conjugated phalloidin per coverslip for 60 min. Finally, chromatin was counterstained for 60 min with 5 µg ml–1 Hoechst 33258 in PBS with the addition of 5 IUs per coverslip AlexaFluor 568‐phalloidin and specimens were finally mounted in antifade solution (0.1 mg ml–1 of p‐phenylenediamine in PBS, pH 8.0 made in 90% v/v glycerol). For fluorescence microscopy and photomicrography a Zeiss Axioplan microscope equipped with standard epifluorescence filters and Neofluar objectives was used. Micrographs were taken on Kodak T‐MAX 400 film at 1600 ASA. For the examination of AFs in intact leaf segments the same protocol was applied with the exception of the enzyme treatment, which was omitted. In this case photomicrography was not applicable due to high background generated by the overlapping cell layers. Assessment of the AF content To assess changes in the AF content due to plasmolysis, image analysis of either photomicrographic negatives or directly of fluorophore labelled specimens were used. Although the following approaches cannot be used to quantify in absolute numbers the cellular AF content, they are reliable for a comparison of the AF content between control and plasmolysed cells. For image analysis photomicrographic negatives of 100 control and 100 plasmolysed cells were scanned through an Agfa Duoscan scanner and captured as ‘tiff’ files using Agfa FotoLook software. For digital image analysis the Image‐Pro Plus software (Media Cybernetics) was used. Following background subtraction, at least five areas of predefined magnitude were chosen and the number of AFs that were automatically tracked within were measured, averaged and expressed as the amount of fluorescence per unit area. Background was measured in cytoplasmic areas devoid of AFs. Average fluorescence intensity per unit area of either plasmolysed or control cells (arbitrary units) was plotted as a histogram using MS Excel software (Microsoft Corp.). Digital image analysis to assess differences in the AF number between different cell populations has previously been used (Hallows et al., 1996; Schindelholz and Reber, 1999). Differences in fluorescence intensity were also estimated by measuring the overall fluorescence of Alexa‐phalloidin labelled cells. At least 500 cells from plasmolysed and control cells were compared, using the photometer coupled to the Zeiss Axioplan microscope, which converts fluorescence intensity to exposure time for photomicrography. The values obtained though this approach were comparable to those obtained from digital image analysis of the photomicrographic negatives. Results General remarks The hyperosmotic treatment of leaf pieces of C. comosum with 1 M mannitol induced plasmolysis in every cell type. Most of the epidermal and mesophyll cells (EC and MCs) examined, displayed convex or concave plasmolysis (Figs 1A–D, 3K, L). In a few plasmolysed cells, interconnected sub‐protoplasts were formed. Many large cells formed an extensive Hechtian strand network linking the protoplast with the cell wall. Examination of living ECs and MCs showed that the protoplast responds instantly to the hypertonic solution and that plasmolysis is completed within 1–5 min after immersion of the leaf segments in the plasmolytic solution. After 5 min the protoplast volume and shape remain fairly constant (Fig. 1A–D). The same plasmolysis pattern was induced by 0.5 and 0.3 M mannitol solutions. However, the number of the plasmolysed cells in these solutions was smaller, while the course of plasmolysis was prolonged compared with those induced by 1 M mannitol. A comparative study of living and isolated plasmolysed cells with DIC optics revealed that fixation and separation methods used in this study do not affect the form of the plasmolysed protoplasts appreciably. The AF staining protocol followed in this study yielded reproducible results in plasmolysed dividing, differentiating and mature cells. In the AF staining protocol the MBS prestabilization of AFs was abandoned, since Triton X‐100 used to facilitate penetration of MBS, brought about severe morphological alterations to the plasmolysed protoplasts. Fixation in PBS produced slightly better results than fixation in PEMD, probably due to the stabilizing effects of inorganic phosphates on AF structure (Rickard and Sheterline, 1988). Finally, the addition of substoichiometric quantities of Alexa‐phalloidin in the fixative protected the labile AFs against severing from formaldehyde. The higher quantum yield of Alexa 568 fluorophore when compared to rhodamine (Haugland, 2001) provided the means to compensate for the high levels of autofluorescence, and thus to increase the signal‐to‐noise ratio. AF organization in control cells Control leaf cells display all the AF arrays described elsewhere (Panteris et al., 1992; Cleary, 1995; Cleary and Mathesius, 1996). In all cell types examined the cortical cytoplasm was traversed by well‐organized arrays of fine AFs or AF bundles (Fig. 2A, B, E, H). In the vast majority of the interphase meristematic cells, cortical AFs were arranged perpendicularly to the longitudinal cell axis (Fig. 2A). In the preprophase–prophase cells, AF preprophase bands (AF‐PPBs) of various degrees of organization were routinely observed (Fig. 2C), a feature commonly observed in other plant species (Mineyuki, 1999). Differentiating and mature cells exhibited cortical AFs usually arranged in various orientations (Fig. 2B, E, H). Thick, mostly longitudinal and sparsely arranged AF bundles traversed the sub‐cortical cytoplasm, which is the site of cytoplasmic streaming (Fig. 2F). This AF system was more prominent in differentiating and mature leaf cells than in the meristematic ones, in which the cortical AF arrays dominated. In all cell types examined AF bundles were found in the perinuclear cytoplasm (Fig. 2D, G), which in many meristematic cells formed distinct perinuclear AF cages (Fig. 2D). In differentiating and mature cells AF bundles emerging from the perinuclear cytoplasm entered the transvacuolar cytoplasmic strands, often reaching the cell cortex (Fig. 2G). AF organization in plasmolysed cells In plasmolysed cells, the AF cytoskeleton is significantly affected. The changes observed were identical among all the mannitol solutions used (0.3, 0.5 and 1 M) as well as among all the epidermal and mesophyll cell types examined. To avoid repetition of the results the AF organization in cells plasmolysed with 1 M mannitol will be described only. A consistent effect of the hyperosmotic stress on the AF cytoskeleton is the disappearance of the fine cortical AFs. This was most obvious in the interphase plasmolysed meristematic cells. New AF systems consisting of thick AF bundles ran through the cortical cytoplasm. Additionally, subcortical and endoplasmic AF arrays of different organization from those found in control cells were observed (Fig. 3A–E). Notably, the AF‐PPB resisted plasmolysis in most preprophase cells (Fig. 3F), a phenomenon also observed in the plasmolysed ECs of Tradescantia (Cleary, 2001). At the PPB site the plasmalemma is not detached from the cell wall (Kagawa et al., 1992; Cleary, 2001; Komis et al., 2001). The most prominent effect of the hyperosmotic stress is that it induced a significant increase in the overall AF cell content. AFs were more numerous in plasmolysed cells than in the control ones (Fig. 3A–E cf. Fig. 2B, E, F, H) and formed thick bundles (Figs 3B–D, 4A). The data obtained using the digital image analysis of the photomicrographic negatives showed that this increase in plasmolysed differentiating cells is about 120% in relation to non‐plasmolysed cells (Fig. 5A). Estimations made by the microscope photometer revealed that the increase of the AF content in differentiating plasmolysed cells is about 150%, while in meristematic cells about 50% compared to control cells (Fig. 5B). This phenomenon was confirmed in ECs and MCs with all the mannitol solutions used. In plasmolysed cells most of the cortical and subcortical AF bundles were localized at the areas of intense protoplast shrinkage, where they formed a network lining the detached plasmalemma regions (Fig. 3B, G–L). Generally, the AF bundles were significantly enriched within the most shrunk protoplast regions (Figs 3H, K, 4A, B). In cells, which had undergone concave plasmolysis, an intense actin staining was localized underneath the plasmalemma, at the borders of the detached protoplast regions (Fig. 3G, K, L), while almost no staining was observed in the non‐detached ones. In many cells, AFs were also detected within most of the Hechtian strands (Fig. 3D). The endoplasmic AF bundles ran through the cytoplasm in various directions (Fig. 3E, I). In the majority of plasmolysed meristematic cells the endoplasmic AF bundles were arranged in a cage encircling the nucleus (Fig. 3E). Differentiating and mature cells displayed AFs juxtaposed with the tonoplast (Figs 3I–L, 4B). In some cases AFs completely surrounded the vacuoles (Fig. 4B). The AF organization in plasmolysed cells in intact tissue was similar to that described above. In all plasmolysed ECs examined numerous AF bundles were localized in the shrunken protoplast areas, lining the detached plasmalemma regions. Therefore, the cell separation procedure does not affect the AF organization. CB‐treated plasmolysed cells In plasmolysed cells CB induced AF disorganization and dramatic changes in the pattern of plasmolysis. In AF depleted plasmolysed ECs and MCs, the course of plasmolysis lasted for a longer time (Fig. 6A–D) and led to a greater decrease of the protoplast volume compared with the untreated cells (Fig. 6A–D cf. Fig. 1A–D). Frequently, the protoplast was almost completely detached from the cell wall (Figs 6D, 7A). As mentioned above, in cells treated with 1 M mannitol, plasmolysis was completed within 1–5 min after immersion in the hyperosmotic solution, while in those subjected to CB treatment continued for 30–60 min (Fig. 6A–D cf. Fig. 1A–D). In many CB‐treated plasmolysed cells the protoplast assumed an irregular form, usually ‘amoeboid’ with prominent protrusions (Fig. 7B–D). Frequently, this was subdivided into numerous subprotoplasts, which were usually separated from the main protoplast (Fig. 7E, F). On the surface of many CB‐treated protoplasts fine and rigid protoplasmic extensions, shorter and thicker than the Hechtian strands were observed (Figs 6D, 7A). The CB‐treated plasmolysed cells displayed many ‘vesicular elements’ exhibiting a rather homogeneous content occupying the periplasmic area (Fig. 7C–E). Besides, OsO4 fixation of CB‐treated cells, as well as of the untreated ones, revealed the existence of large osmiophilic bodies in the cytoplasm and in the vacuoles (Fig. 7F) not found in control cells. The nature and origin of these bodies remains to be elucidated. Deplasmolysis Within seconds after the replacement of the plasmolytic solution with water the plasmolysed protoplasts rounded up and deplasmolysis was rapidly carried out (Fig. 1E, F). During the protoplast re‐expansion the plasmalemma re‐incorporated the Hechtian strands. Finally, the protoplast is re‐appressed on the cell wall. Deplasmolysis also takes place in the CB‐treated plasmolysed cells. During this process the protoplast becomes round and rapidly expands, while the protoplasmic extensions are reincorporated by the protoplast (Fig. 6E, F). The sub‐protoplasts and the ‘vesicular elements’ located in the periplasmic space were also fused with the enlarging protoplast. Notably, the severe plasmolysis, experienced by the cells in the presence of CB, does not affect their viability even 60 min after the onset of plasmolysis (Fig. 6A–F). In deplasmolysed cells the thick AF bundles were disorganized and fine AFs reappeared. They formed the AF arrays found in control cells, i.e. cortical arrays, endoplasmic AF bundles traversing the perinuclear cytoplasm as well as others meandering among the vacuoles (Fig. 8A–D). Cells that underwent a complete plasmolytic cycle could be successfully re‐plasmolysed suggesting that they still retain their viability between the two‐ anisosmotic extreme treatments. However, during the second plasmolytic cycle, the protoplast remained plasmolysed for a short time. Then this is slowly deplasmolysed in the presence of mannitol solution (Fig. 9A, B). This was not the case for CB‐treated cells though. As soon as the deplasmolysed protoplast re‐encounters plasmolytic solution containing CB it initially shrinks but within seconds ‘explodes’ releasing cytoplasmic material. Discussion The changes in organization of the AF cytoskeleton occurring during the plasmolytic cycle in leaf cells of C. comosum are summarized as (a) the hyperosmotic stress induces the disappearance of the fine cortical AFs and the formation of numerous well‐organized cortical, subcortical and endoplasmic AF bundles; (b) the cortical AF bundles are preferentially localized in the shrunk protoplast areas, lining the detached plasmalemma regions: the endoplasmic AF bundles are mainly located in the perinuclear cytoplasm and on the tonoplast surface; (c) the experimental AF disorganization prolongs plasmolysis and induces dramatic changes in shape and volume of the plasmolysed protoplast; and (d) during deplasmolysis, the normal AF cytoskeleton is reinstated quite rapidly. The findings of the present study show clearly that the AF cytoskeleton in the angiosperm C. comosum, is intimately involved in the mechanism by which the cells regulate the shape and the volume of the protoplast during plasmolysis. Some of the phenomena mentioned above will be discussed in the following sections. AF polymerization in plasmolysed cells The induction of AF polymerization in plasmolysed cells observed for the first time here in a plant species, has been described in other diverse biological systems exposed to hypertonic conditions (Hoffmann and Pedersen, 1998). For example, in Erlich ascites tumour cells, a brief exposure to a hypertonic challenge results in approximately a 25% elevation of the F‐actin content (Pedersen et al., 1999). Similarly, the overall actin content is elevated in Dictyostelium amoebae experiencing a hyperosmotic treatment (Zischka et al., 1999). Therefore, the induction of AF polymerization is a common response of animal, fungal and plant cells to hyperosmotic stress. In animal cells the excellent mechanical properties of the AF networks seem to relieve the forces imposed on the cell through volume reduction and compaction (Janmey, 1994). It is known that the hyperosmotic treatment triggers a signalling pathway that involves an increase in the cytosolic Ca2+ concentration (Pedersen et al., 1998; Reddy, 2001) and the generation of polyphosphoinositides (Munnik et al., 2000; Meijer et al., 2001; Munnik, 2001). In animal cells these pathways directly integrate the stress condition to the AF cytoskeleton by inducing AF polymerization and/or by affecting the AF physical properties (Janmey, 1998). The hyperosmotic stress in plants causes an elevation of cytosolic Ca2+ concentration by recruiting intra‐ and/or extracellular stores (Busch, 1995; Knight et al., 1997, 1998; Cessna et al., 1998; Brownlee et al., 1999; Knight, 2000). This Ca2+ mobilization in the plasmolysed C.comosum leaf cells, could induce AF formation directly by increasing the affinity for polymerization of G‐actin subunits (Strzelecka‐Golaszewska, 2001) or indirectly through a series of Ca2+‐dependent signal transduction events involving the regulation of actin dynamics (Janmey, 1994, 1998). AFs and plasmalemma protection During plasmolysis certain plasmalemma areas retain their attachments to the cell wall, while some others are retracted following protoplast shrinkage (Oparka, 1994; Lang‐Pauluzzi, 2000). As a result shearing forces are generated, which might injure the plasmalemma (Oparka, 1994). This work reveals the existence of a positional relationship between cortical AF bundles and the plasmalemma at those sites where the most intense shearing forces are produced. In cells exhibiting concave plasmolysis, AFs are significantly enriched underneath the plasmalemma, preferentially near the boundaries between the detached and non‐detached regions of the latter. Moreover, in plasmolysed cells displaying convex plasmolysis, AFs line the detached plasmalemma regions. The partial persistence of plasmalemma–cell wall attachments in plasmolysed cells may raise a challenge against plasmalemma integrity through stretching. It is thus necessary for the plasmolysed protoplast to develop a mechanism to compensate for the incoming injury (Oparka, 1994). In animal cells, being under mechanical stress, the cytoskeleton provides support to the plasmalemma by reinforcing the cortical framework preferentially at the sites of extensive force application (Ingber, 1997; Ko and McCulloch, 2000). A characteristic example is the endothelial cells that line the blood vessels, which are continuously under shear stress (Kano et al., 2000) imposed by the blood stream. To compensate the stress, these cells form an extensive cortical network of AF bundles, the ‘stress fibres’, which are strongly inducible upon force application (Galbraith et al., 1998). Often, they coalign with the direction of fluid flow under shear stress (Galbraith et al., 1998; Frame and Sarelius, 2000). Current work suggests a role for AF remodelling as a response to mechanical challenge (Ingber, 1997). Stress fibres are able to contract, serving as a compensatory mechanism against mechanical injury of the plasmalemma (Arora et al., 1999; Katoh et al., 1998). In plasmolysed leaf cells of C. comosum the AF network formed underneath the plasmalemma might protect against shearing possibly through an analogous mechanism like the contraction of stress fibres. A similar role has been also attributed to the cortical AF network formed in Dictyostelium hyperosmotically stressed cells (Aizawa et al., 1999). Alternatively, it could result in an increased rigidity of the plasmalemma at the sites of maximal tension. AF bundles found in close association with the vacuoles in plasmolysed cells of C. comosum may offer mechanical support to the tonoplast as well. In plasmolysed plant cells, numerous multilamellar vesicles cut off from the tonoplast which at high external salt concentration became condensed into osmiophilic bodies (Singh and Johnson‐Flanagan, 1987). These structures probably reserve membrane material, to be used for the tonoplast re‐expansion during deplasmolysis (Oparka, 1994). Osmiophilic bodies observed in the plasmolysed cells of C. comosum, but not in the control ones, probably represent surplus of membrane material. AFs and protoplast shape regulation The findings of this work support the hypothesis that in plasmolysed C. comosum leaf cells the AF network is somehow involved in the regulation of the protoplast shape. The effects of CB further support the above suggestion. AF depletion from the plasmolysed protoplasts results in anomalous ‘amoeboid’ forms, never observed in cells plasmolysed in the absence of CB. Protoplasmic extensions similar to those found in plasmolysed protoplasts isolated from cold acclimated leaves of rye (Dowgert and Steponkus, 1984; Gordon‐Kamm and Steponkus, 1984) were observed in this material after CB treatment. It should be noted here that the regulatory role of the AFs in the shape of the plasmolysed protoplasts of C. comosum is expressed in the absence of microtubules. The latter are disintegrated in hyperosmotically treated cells (Komis et al., 2001). The tubulin paracrystals and macrotubules, which are formed, may offer mechanical support to the protoplast (Komis et al., 2001). Whether the tubulin paracrystal network underlies the formation of ‘amoeboid’ protoplast protrusions in the CB‐treated plasmolysed cells remains to be seen. The sub‐protoplasts and ‘vesicular elements’ encountered in the CB‐treated plasmolysed cells examined were not always, if at all, linked to the main protoplast. By contrast, sub‐protoplasts were rarely observed in plasmolysed cells in the absence of CB and were always attached to the main protoplast through AF‐rich cytoplasmic bridges. AF disruption may result in the excision of sub‐protoplasts as a result of plasmalemma destabilization and blebbing. AFs and protoplast volume regulation Disruption of AFs in CB‐treated plasmolysed cells of C. comosum uncoupled the ability of the protoplast to maintain a constant volume under hyperosmotic conditions. Therefore, plasmolysis should trigger a protective mechanism, to enable the cell to attenuate the loss of water and the protoplast volume reduction (Lee‐Stadelmann and Stadelmann, 1989). This is achieved by the increase in the concentration of small organic and chemically inert osmolytes in the plasmolysed protoplast (Hare et al., 1998; Tabaeizadeh, 1998). Their synthesis, transport and accumulation is a relatively slow procedure (Lang et al., 1998; Tabaeizadeh, 1998). Therefore, osmotic and volume regulation is initially carried out by the transfer of ions from the apoplast into the protoplast through plasmalemma ion channels, which are inducible under hyperosmotic conditions (Lang et al., 1998). The activation of such a regulatory mechanism in the plasmolysed cells of C. comosum is indicated by the fact that when they undergo a second plasmolytic cycle they are almost completely deplasmolysed in the presence of the mannitol solution. In animal cells, cytoskeleton‐mediated cell volume regulation after exposure to anisosmotic conditions is mostly attributed to the capacity of cytoskeletal elements, mostly AFs, in controlling ion channels and transporter activities (Ko and McCulloch, 2000; Papakonstanti et al., 2000). There are many ways by which the cytoskeleton is implicated in ion channel function (Janmey, 1998; Khurana, 2000; Szaszi et al., 2000). In hyperosmotically treated animal cells the cell volume is reconstituted though a process termed ‘regulatory volume increase’ (RVI). As described above, exposure to hypertonic solutions generally induces AF polymerization. Excessive AFs are required, among others, for the activation of ion channels (Lang et al., 1998; Pedersen et al., 1999; Szaszi et al., 2000), which in turn trigger the RVI process. In hyperosmotically C. comosum treated cells, the volume regulatory response is expressed differentially during two successive plasmolytic cycles. At the first plasmolytic cycle, where no net volume increase is observed, it was documented as the ability of the plasmolysed protoplast to maintain a constant volume. This observation suggests the accumulation of osmolytes or ions in the protoplast to counterbalance the extracellular hypertonicity. During the second plasmolytic cycle the accumulation of the osmolytes should be high enough to drive a net volume increase. The ability of the protoplast to withstand continuous shrinkage during the first plasmolytic cycle and the capacity to undergo volume increase during the second, are strongly inhibited by CB. This implies that an AF‐dependent mechanism, probably controlling the plasmalemma ion channel and aquaporin regulation, might be responsible for protoplast volume maintenance. It should be noted that the implication of the AFs in plant cell volume regulation, is not well documented. Reversible changes in cortical AF organization related to changes of cell volume have been observed only in stomata of Vicia faba and Commelina communis. In open stomata, guard cells display well‐organized radial cortical AF arrays, which disintegrate when the stomata close (Kim et al., 1995; Eun and Lee, 1997), changes probably mediated by cytosolic Ca2+ levels and by protein kinase and phosphatase activities (Hwang and Lee, 2001). It has been suggested that AFs are involved in the regulation of the guard cell volume, by modulation of the activity of the plasmalemma ion channels (Hwang et al., 1997), as it probably happens in the case of plasmolysed cells of C. comosum. Acknowledgements 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 actin filament tracking and fluorescence intensity measurements. G Komis was awarded a scholarship by the State Scholarship Foundation. View largeDownload slide Fig. 1. Living ECs as they appear under DIC optics during the first plasmolytic cycle (A–D, plasmolysis; E, F, deplasmolysis). Numbers in (A) to (D) indicate time lapsed (in min) after immersion of the tissue in the plasmolytic solution. Numbers in (E) and (F) indicate time in seconds following substitution of the plasmolytic solution by water. Bar 10 µm. View largeDownload slide Fig. 1. Living ECs as they appear under DIC optics during the first plasmolytic cycle (A–D, plasmolysis; E, F, deplasmolysis). Numbers in (A) to (D) indicate time lapsed (in min) after immersion of the tissue in the plasmolytic solution. Numbers in (E) and (F) indicate time in seconds following substitution of the plasmolytic solution by water. Bar 10 µm. View largeDownload slide Fig. 2. AF organization in control cells. Bar 10 µm. (A, B) Cortical AFs in a meristematic (A) and in a differentiating (B) cell. (C) AF‐PPB (arrow). (D) Endoplasmic AFs in a meristematic cell. Those are mostly located around the nucleus. (E) Cortical AFs of a differentiating MC. (F) Subcortical AFs of a differentiating cell. (G) Endoplasmic AFs of a mature cell. (H) Cortical AFs of a mature cell. View largeDownload slide Fig. 2. AF organization in control cells. Bar 10 µm. (A, B) Cortical AFs in a meristematic (A) and in a differentiating (B) cell. (C) AF‐PPB (arrow). (D) Endoplasmic AFs in a meristematic cell. Those are mostly located around the nucleus. (E) Cortical AFs of a differentiating MC. (F) Subcortical AFs of a differentiating cell. (G) Endoplasmic AFs of a mature cell. (H) Cortical AFs of a mature cell. View largeDownload slide Fig. 3. AF organization in cells plasmolysed for 30 min in 1 M aqueous mannitol solution. Bar 10 µm. (A) Meristematic cell displaying convex plasmolysis. AF bundles underlying the plasmalemma. (B) Numerous well‐organized AF bundles underlying the plasmalemma in this differentiating plasmolysed cell. (C, D) Mature plasmolysed cells displaying a well‐organized network of cortical/subcortical AF bundles. Arrowheads in (D), point to Hechtian strands traversed by AFs. (E) Endoplasmic AF bundles in a meristematic plasmolysed cell. (F) AF‐PPB (arrow) in a plasmolysed cell. (G) Cell displaying concave plasmolysis. Intense actin staining can be observed under the detached regions of the plasmalemma (arrowheads). (H) Most of the AF bundles in this plasmolysed cell are located in the most shrunken region of the protoplast (arrow). (I) Endoplasmic AFs in a differentiating plasmolysed cell. Many AFs are localized on the surface of the nucleus (N) and the vacuoles (arrowheads). (J) The cell displayed in I under DIC optics. (K, L) Differentiating cells displaying concave plasmolysis. Many AF bundles underlie the detached regions of the plasmalemma (arrows). Arrowheads point to AF bundles in contact with vacuoles. View largeDownload slide Fig. 3. AF organization in cells plasmolysed for 30 min in 1 M aqueous mannitol solution. Bar 10 µm. (A) Meristematic cell displaying convex plasmolysis. AF bundles underlying the plasmalemma. (B) Numerous well‐organized AF bundles underlying the plasmalemma in this differentiating plasmolysed cell. (C, D) Mature plasmolysed cells displaying a well‐organized network of cortical/subcortical AF bundles. Arrowheads in (D), point to Hechtian strands traversed by AFs. (E) Endoplasmic AF bundles in a meristematic plasmolysed cell. (F) AF‐PPB (arrow) in a plasmolysed cell. (G) Cell displaying concave plasmolysis. Intense actin staining can be observed under the detached regions of the plasmalemma (arrowheads). (H) Most of the AF bundles in this plasmolysed cell are located in the most shrunken region of the protoplast (arrow). (I) Endoplasmic AFs in a differentiating plasmolysed cell. Many AFs are localized on the surface of the nucleus (N) and the vacuoles (arrowheads). (J) The cell displayed in I under DIC optics. (K, L) Differentiating cells displaying concave plasmolysis. Many AF bundles underlie the detached regions of the plasmalemma (arrows). Arrowheads point to AF bundles in contact with vacuoles. View largeDownload slide Fig. 4. (A) Almost all AFs in this plasmolysed cell are located within the protoplasmic bridge (arrow) interconnecting neighboring sub‐protoplasts. (B) Mature plasmolysed cell. Many AFs can be observed on the surface of the vacuole (V) as well as within the intensely shrunken region of the protoplast (arrow). Bar 10 µm. View largeDownload slide Fig. 4. (A) Almost all AFs in this plasmolysed cell are located within the protoplasmic bridge (arrow) interconnecting neighboring sub‐protoplasts. (B) Mature plasmolysed cell. Many AFs can be observed on the surface of the vacuole (V) as well as within the intensely shrunken region of the protoplast (arrow). Bar 10 µm. View largeDownload slide Fig. 5. Control cell (white bars), plasmolysed cell (grey bars). (A, B) Histograms displaying the average AF fluorescence intensity of control and plasmolysed leaf cells. (A) Histogram of the average AF fluorescence intensity measured through digital image analysis. Sample size: 100 differentiating control cells, 100 differentiating plasmolysed cells. (B) Histogram of the average AF fluorescence intensity measured through microscopic. Sample size 500 control cells, 500 plasmolysed cells (a, differentiating cells; b, meristematic cells). View largeDownload slide Fig. 5. Control cell (white bars), plasmolysed cell (grey bars). (A, B) Histograms displaying the average AF fluorescence intensity of control and plasmolysed leaf cells. (A) Histogram of the average AF fluorescence intensity measured through digital image analysis. Sample size: 100 differentiating control cells, 100 differentiating plasmolysed cells. (B) Histogram of the average AF fluorescence intensity measured through microscopic. Sample size 500 control cells, 500 plasmolysed cells (a, differentiating cells; b, meristematic cells). View largeDownload slide Fig. 6. Living CB‐treated ECs as they appear under DIC optics during the first plasmolytic cycle (A–D, plasmolysis; E, F, deplasmolysis). Arrowheads in (D) point to protoplasmic extensions. Numbers in (A) to (D) correspond to time in minutes after immersion of the tissue in 1 M mannitol solution supplemented with 100 µM CB. Numbers in (E) and (F) indicate time in seconds after replacement of the plasmolytic solution with water. Bar 10 µm. View largeDownload slide Fig. 6. Living CB‐treated ECs as they appear under DIC optics during the first plasmolytic cycle (A–D, plasmolysis; E, F, deplasmolysis). Arrowheads in (D) point to protoplasmic extensions. Numbers in (A) to (D) correspond to time in minutes after immersion of the tissue in 1 M mannitol solution supplemented with 100 µM CB. Numbers in (E) and (F) indicate time in seconds after replacement of the plasmolytic solution with water. Bar 10 µm. View largeDownload slide Fig. 7. CB‐treated plasmolysed ECs under DIC optics. Bars 10 µm. (A) In this cell the protoplast is completely detached from the cell wall. Arrow points to a protoplasmic extension. (B, C) In these cell groups, the plasmolysed protoplasts have assumed an ‘amoeboid’ form. Arrows in C mark ‘vesicular elements’. (D) ‘Amoeboid’ plasmolysed protoplast exhibiting discrete protoplasmic protrusions (arrows). Arrowhead shows a ‘vesicular element’. (E) In this plasmolysed cell the protoplast (asterisk) has suffered very intense shrinkage. In the space between the protoplast and the cell wall sub‐protoplasts (arrows) and ‘vesicular elements’ (arrowheads) can be observed. (F) Osmiophilic bodies in CB‐treated plasmolysed cells fixed with OsO4. Arrows show sub‐protoplasts. View largeDownload slide Fig. 7. CB‐treated plasmolysed ECs under DIC optics. Bars 10 µm. (A) In this cell the protoplast is completely detached from the cell wall. Arrow points to a protoplasmic extension. (B, C) In these cell groups, the plasmolysed protoplasts have assumed an ‘amoeboid’ form. Arrows in C mark ‘vesicular elements’. (D) ‘Amoeboid’ plasmolysed protoplast exhibiting discrete protoplasmic protrusions (arrows). Arrowhead shows a ‘vesicular element’. (E) In this plasmolysed cell the protoplast (asterisk) has suffered very intense shrinkage. In the space between the protoplast and the cell wall sub‐protoplasts (arrows) and ‘vesicular elements’ (arrowheads) can be observed. (F) Osmiophilic bodies in CB‐treated plasmolysed cells fixed with OsO4. Arrows show sub‐protoplasts. View largeDownload slide Fig. 8. AF organization in deplasmolysed cells. Bar 10 µm. (A) Cortical AFs in a meristematic cell. (B–D) Endoplasmic AFs in a meristematic (B), differentiating (C), and mature (D) cell. View largeDownload slide Fig. 8. AF organization in deplasmolysed cells. Bar 10 µm. (A) Cortical AFs in a meristematic cell. (B–D) Endoplasmic AFs in a meristematic (B), differentiating (C), and mature (D) cell. View largeDownload slide Fig. 9. Living ECs under DIC optics during the second plasmolytic cycle. It is clearly evident that the protoplast volume increases in the presence of the plasmolytic solution. Numbers indicate time in minutes after the initiation of the second plasmolytic cycle (starting immediately as cells completely deplasmolysed and water was replaced by the plasmolytic solution). Bar 10 µm. View largeDownload slide Fig. 9. Living ECs under DIC optics during the second plasmolytic cycle. It is clearly evident that the protoplast volume increases in the presence of the plasmolytic solution. Numbers indicate time in minutes after the initiation of the second plasmolytic cycle (starting immediately as cells completely deplasmolysed and water was replaced by the plasmolytic solution). Bar 10 µm. References AizawaH, Katadae M, Maruya M, Sameshima M, Murakami‐Murofushi K, Yahara I. 1999. Hyperosmotic stress‐induced reorganization of actin bundles in Dictyostelium cells over‐expressing cofilin. Genes to Cells  4, 311–324. Google Scholar AroraPD, Janmey PA, McCulloch CA. 1999. A role for gelsolin in stress fiber‐dependent cell contraction. Experimental Cell Research  250, 155–167. Google Scholar BachewichCL, Heath IB. 1997. Differential cytoplasm–plasma membrane–cell wall adhesion patterns and their relationships to hyphal tip growth and organelle motility. Protoplasma  200, 71–86. Google Scholar BrownleeC, Goddard H, Hetherington AM, Peake L‐A. 1999. Specificity and integration of responses: Ca2+ as a signal in polarity and osmotic regulation. Journal of Experimental Botany  50, 1001–1011. Google Scholar BuschDS. 1995. Calcium regulation in plant cells and its role in signaling. Annual Review of Plant Physiology and Plant Molecular Biology  46, 95–122. Google Scholar CessnaSG, Chandra S, Low PS. 1998. Hypo‐osmotic shock of tobacco cells stimulates Ca2+ fluxes deriving first from external and then internal Ca2+ stores. Journal of Biological Chemistry  273, 27286–27291. Google Scholar ChowdhuryS, Smith KW, Gustin MC. 1992. Osmotic stress and the yeast cytoskeleton: phenotype‐specific suppression of an actin mutation. Journal of Cell Biology  118, 561–571. Google Scholar ClearyAL. 1995. F‐actin redistributions at the division site in living Tradescantia stomatal complexes as revealed by microinjection of rhodamine‐phalloidin. Protoplasma  185, 152–165. Google Scholar ClearyAL. 2001. Plasma membrane–cell wall connections: roles in mitosis and cytokinesis revealed by plasmolysis of Tradescantia virginiana leaf epidermal cells. Protoplasma  215, 21–34. Google Scholar ClearyAL, Mathesius U. 1996. Rearrangements of F‐actin during stomatogenesis visualized by confocal microscopy in fixed and permeabilized Tradescantia leaf epidermis. Botanica Acta  109, 15–24. Google Scholar DowgertMF, Steponkus PL. 1984. Behavior of the plasma membrane of isolated protoplasts during a freeze–thaw cycle. Plant Physiology  75, 1139–1151. Google Scholar EunSO, Lee Y. 1997. Actin filaments of guard cells are reorganized in response to light and abscisic acid. Plant Physiology  115, 1491–1498. Google Scholar FrameMD, Sarelius IH. 2000. Flow‐induced cytoskeletal changes in endothelial cells growing on curved surfaces. Microcirculation  7, 419–427. Google Scholar GalbraithCG, Skalak R, Chien S. 1998. Shear stress induces spatial reorganization of the endothelial cell cytoskeleton. Cell  Motility and the Cytoskeleton  40, 317–330. Google Scholar Gordon‐KammWJ, Steponkus PL. 1984. The influence of cold acclimation on the behavior of the plasma membrane following osmotic contraction of isolated protoplasts. Protoplasma  123, 161–173. Google Scholar GunningBES, Steer MW. 1996. Plant cell biology: structure and function. Boston, MA: Jones and Bartlett. Google Scholar HallowsKR, Law F‐Y, Packman CH, Knauf PA. 1996. Changes in cytoskeletal actin content, F‐actin distribution, and surface morphology during HL‐60 cell volume regulation. Journal of Cell Physiolog y 167, 60–71. Google Scholar HarePD, Cress WA, Van Staden J. 1998. Dissecting the roles of osmolyte accumulation during stress. Plant, Cell and Environment  21, 535–553. Google Scholar HauglandRP. 2001. Handbook of fluorescent probes and research chemicals. Oregon: Molecular Probes. Google Scholar HensonJH. 1999. Relationships between the actin cytoskeleton and cell volume regulation. Microscopy Research and Techniques  47, 155–162. Google Scholar HirtH. 2000. MAP kinases in plant signal transduction. Results and Problems in Cell Differentiation  27, 1–9. Google Scholar HoffmannEK, Pedersen SF. 1998. Sensors and signal transduction in the activation of cell volume regulatory ion transport systems. Contributions in Nephrology  123, 50–78. Google Scholar HwangJ‐U, Lee Y. 2001. Abscisic acid‐induced actin reorganization in guard cells of dayflower is mediated by cytosolic calcium levels and by protein kinase and protein phosphatase activities. Plant Physiology  125, 2120–2128. Google Scholar HwangJ‐U, Suh S, Yi H, Kim J, Lee Y. 1997. Actin filaments modulate both stomatal opening and inward K+‐channel activities in guard cells of Vicia faba L. Plant Physiology  115, 335–342. Google Scholar IngberDE. 1997. Tensegrity: the architectural basis of mechanotransduction. Annual Review of Physiology  59, 575–599. Google Scholar JanmeyPA. 1994. Phosphoinositides and calcium as regulators of cellular actin. Annual Review of Physiology  56, 169–191. Google Scholar JanmeyPA. 1998. The cytoskeleton and cell signaling: component localization and mechanical coupling. Physiological Reviews  78, 763–781. Google Scholar KagawaT, Kadota A, Wada M. 1992. The junction between the plasma membrane and the cell wall in fern protonemal cells, as visualized after plasmolysis, and its dependence on arrays of cortical microtubules. Protoplasma  170, 186–190. Google Scholar KanoY, Katoh K, Fujiwara K. 2000. Lateral zone of cell–cell adhesion as the major fluid shear stress‐related signal transduction site. Circulation Research   86, 425–433. Google Scholar KapusA, Di Ciano C, Sun J, Zhan X, Kim L, Wong TW, Rotstein OD. 2000. Cell volume‐dependent phosphorylation of proteins of the cortical cytoskeleton and cell‐cell contact sites. The role of Fyn and FER kinases. Journal of Biological Chemistry   275, 32289–32298. Google Scholar KatohK, Kano Y, Masuda M, Onishi H, Fujiwara K. 1998. Isolation and contraction of the stress fiber. Molecular Biology of the Cell  9, 1919–1938. Google Scholar KhuranaS. 2000. Role of actin cytoskeleton in regulation of ion transport: examples from epithelial cells. Journal of Membrane Biology  178, 73–87. Google Scholar KimM, Hepler PK, Eun SO, Ha KS, Lee Y. 1995. Actin filaments in mature guard cells are radically distributed and involved in stomatal movement. Plant Physiology  109, 1077–1087. Google Scholar KnightH. 2000. Calcium signaling during abiotic stress in plants. International Review of Cytology  195, 269–324. Google Scholar KnightH, Trewavas AJ, Knight MR. 1997. Calcium signalling in Arabidopsis thaliana responding to drought and salinity. The Plant Journal  12, 1067–1078. Google Scholar KnightH, Brandt S, Knight MR. 1998. A history of stress alters drought calcium signalling pathways in Arabidopsis. The Plant Journal  16, 681–687. Google Scholar KoKS, McCulloch CA. 2000. Partners in protection: interdependence of cytoskeleton and plasma membrane in adaptations to applied forces. Journal of Membrane Biology  174, 85–95. Google Scholar KomisG, Apostolakos P, Galatis B. 2001. Altered patterns of tubulin polymerization in dividing leaf cells of Chlorophyton comosum after a hyperosmotic treatment. New Phytologist  149, 193–207. Google Scholar LangF, Busch GL, Ritter M, Volkl H, Waldegger S, Gulbins E, Haussinger D. 1998. Functional significance of cell volume regulatory mechanisms. Physiological Reviews  78, 247–306. Google Scholar Lang‐PauluzziI. 2000. The behaviour of the plasma membrane during plasmolysis: a study by UV microscopy. Journal of Microscopy   198, 188–198. Google Scholar Lang‐PauluzziI, Gunning BES. 2000. A plasmolytic cycle: the fate of cytoskeletal elements. Protoplasma  212, 174–185. Google Scholar Lee‐StadelmannOY, Stadelmann EJ. 1989. Plasmolysis and deplasmolysis. Methods in Enzymology  174, 225–247. Google Scholar MeijerHJ, Arisz SA, Van Himbergen JA, Musgrave A, Munnik T. 2001. Hyperosmotic stress rapidly generates lyso‐phosphatidic acid in Chlamydomonas. The Plant Journal  25, 541–548. Google Scholar MineyukiY. 1999. The preprophase band of microtubules: its function as a cytokinetic apparatus in higher plants. International Review of Cytology  187, 1–49. Google Scholar MunnikT. 2001. Phosphatidic acid: an emerging plant lipid second messenger. Trends in Plant Science  6, 227–233. Google Scholar MunnikT, Meijer HJ, Ter Riet B, Hirt H, Frank W, Bartels D, Musgrave A. 2000. Hyperosmotic stress stimulates phospholipase D activity and elevates the levels of phosphatidic acid and diacylglycerol pyrophosphate. The Plant Journal  22, 147–154. Google Scholar OomsLM, McColl BK, Wiradjaja F, Wijayaratnam AP, Gleeson P, Gething MJ, Sambrook J, Mitchell CA. 2000. The yeast inositol polyphosphate 5‐phosphatases inp52p and inp53p translocate to actin patches following hyperosmotic stress: mechanism for regulating phosphatidylinositol 4,5‐bisphosphate at plasma membrane invaginations. Molecular and Cellular Biology  20, 9376–9390. Google Scholar OparkaKJ. 1994. Plasmolysis: new insights into an old process. New Phytologist  126, 571–591. Google Scholar PanterisE, Apostolakos P, Galatis B. 1992. The organization of F‐actin in root tip cells of Adiantum capillus veneris throughout the cell cycle. A double label fluorescence microscopy study. Protoplasma  170, 128–137. Google Scholar PapakonstantiEA, Vardaki EA, StournarasC. 2000. Actin cytoskeleton: a signaling sensor in cell volume regulation. Cellular Physiology and Biochemistry  10, 257–264. Google Scholar PedersenSF, Jorgensen NK, Hoffmann EK. 1998. Dynamics of Ca2+i and pHi in Ehrlich ascites tumor cells after Ca2+‐mobilizing agonists or exposure to hypertonic solution. Pflugers Archives  436, 199–210. Google Scholar PedersenSF, Mills JW, Hoffmann EK. 1999. Role of the F‐actin cytoskeleton in the RVD and RVI processes in Ehrlich ascites tumor cells. Experimental Cell Research   252, 63–74. Google Scholar ReddyAS. 2001. Calcium: silver bullet in signaling. Plant Science  160, 381–404. Google Scholar RichelmeF, Benoliel AM, Bongrand P. 2000. Dynamic study of cell mechanical and structural responses to rapid changes of calcium level. Cell Motility and the Cytoskeleton  45, 93–105. Google Scholar RickardJE, Sheterline P. 1988. Effect of ATP removal and inorganic phosphate on length redistribution of sheared actin filament populations. Evidence for a mechanism of end‐to‐end annealing. Journal of Molecular Biology  201, 675–681. Google Scholar RizoliSB, Rotstein OD, Parodo J, Phillips MJ, Kapus A. 2000. Hypertonic inhibition of exocytosis in neutrophils: central role for osmotic actin skeleton remodeling. American Journal of Physiology   279, C 619–633. Google Scholar SchindelholzB, Reber BF. 1999. Quantitative estimation of F‐actin in single growth cones. Methods   18, 487–492. Google Scholar SinghJ, Johnson‐Flanagan AM. 1987. Membrane alterations in winter rye and Brassica napus cells during lethal freezing and plasmolysis. Plant, Cell and Environment  10, 163–168. Google Scholar SlaninovaI, Sestak S, Svoboda A, Farkas V. 2000. Cell wall and cytoskeleton reorganization as the response to hyperosmotic shock in Saccharomyces cerevisiae. Archives of Microbiology   173, 245–252. Google Scholar Strzelecka‐GolaszewskaH. 2001. Divalent cations, nucleotides, and actin structure. Results and Problems in Cell Differentiation  32, 23–41. Google Scholar SullivanR, Burnham M, Török K, Koffer A. 2000. Calmodulin regulates the disassembly of cortical F‐actin in mast cells but is not required for secretion. Cell  Calcium  28, 33–46. Google Scholar SzasziK, Grinstein S, Orlowski J, Kapus A. 2000. Regulation of the epithelial Na(+)/H(+) exchanger isoform by the cytoskeleton. Cellular Physiology and Biochemistry   10, 265–272. Google Scholar TabaeizadehZ. 1998. Drought‐induced responses in plant cells. International Review of Cytology  182, 193–247. Google Scholar ZischkaH, Oehme F, Pintsch T, Ott A, Keller H, Kellermann J, Schuster SC. 1999. Rearrangement of cortex proteins constitutes an osmoprotective mechanism in Dictyostelium. EMBO Journal  18, 4241–4249. Google Scholar
TI - Hyperosmotic stress‐induced actin filament reorganization in leaf cells of Chlorophyton comosum
JF - Journal of Experimental Botany
DO - 10.1093/jxb/erf018
DA - 2002-08-01
UR - https://www.deepdyve.com/lp/oxford-university-press/hyperosmotic-stress-induced-actin-filament-reorganization-in-leaf-AlpgnLPHCd
SP - 1699
EP - 1710
VL - 53
IS - 375
DP - DeepDyve
ER -