A plasmolytic cycle: The fate of cytoskeletal elements - Springer Link

Protoplasma (2000) 212:174-185

PROTOPLASMA 9 Springer-Verlag 2000 Printed in Austria

A plasmolytic cycle: the fate of cytoskeletal elements I. Lang-Pauluzzi t'* and B. E. S. Gunning 2 1 Institute of Plant Physiology, University of Vienna, Vienna and 2 Plant Cell Biology Group, Research School of Biological Sciences, Australian National University, Canberra, A.C.T. Received August 24, 1999 Accepted October 3, 1999 Dedicated to Professor Walter Gustav Url on the occasion of his 70th birthday

Summary. In most plant cells, transfer to hypertonic solutions causes osmotic loss of water from the vacuole and detachment of the living protoplast from the cell wall (plasmolysis). This process is reversible and after removal of the plasmolytie solution, protoplasts can re-expand to their original size (deplasmolysis). We have investigated this phenomenon with special reference to cytoskeletal elements in onion inner epidermal cells. The main processes of plasmolysis seem to be membrane dependent because destabilization of cytoskeletal elements had only minor effects on plasmolysis speed and form. In most cells, the array of cortical microtubules is similar to that found in nonplasmolyzed states except that longitudinal patterns seen in some control cells were never observed in plasmolyzed protoplasts of onion inner epidermis. As soon as deplasmolysis starts, cortical microtubules become disrupted and only slowly regenerate to form an oblique array, similar to most nontreated cells. Actin microfilaments responded rapidly to the plasmolysis-induced deformation of the protoplast and adapted to its new form without marked changes in organization and structure. Both actin microfilaments and microtubules can be present in Hechtian strands, which, in plasmolyzed cells, connect the cell wall to the protoplast. Anticytoskeletal drugs did not affect the formation of Hechtian strands. Keywords: Cytoskeleton; Hechtian strands; Latrunculin B; Onion inner epidermis; Oryzalin; Plasmolysis.

Abbreviations: DIC differential interference contrast; DiOC6(3) 3,3-dihexyloxacarbocyanine iodide

Introduction

Plasmolysis is a phenomenon of living plant cells. Its most obvious feature is the detachment of the living protoplast from the cell wall, induced artificially due

* Correspondence and reprints: Institute of Plant Physiology, University of Vienna, Althanstrasse 14, A-1091 Vienna, Austria.

to loss of water from the vacuole by hypertonic solutions. According to the plasmolytic solution used, different forms of plasmolysis have been described: concave, convex, cap plasmolysis, band plasmolysis, systrophe, and the formation of subprotoplasts (reviewed by Oparka 1994). Plasmolysis and deplasmolysis were extensively studied in onion epidermis by Stadelmann (1964). Many analyses of the dynamic behavior of plant cells during plasmolysis have been made by Url (e.g., Url 1960, 1971, 1974), including a considerable number of scientific films. In hypertonic sugar solutions, onion inner epidermal cells undergo convex plasmolysis. The protoplast, however, is not detached completely from the cell wall but forms thin, cytoplasmic threads which connect the plasma membrane with the cell wall. These strands have been named after Hecht (1912) and are refered to as Hechtian strands (Bachewich and Heath 1997, Oparka et al. 1996, Oparka 1994, Schnepf et al. 1986, Smith 1972, Sitte 1963). The reverse process of plasmolysis, the re-expansion of the protoplast due to osmotic water uptake, is called deplasmolysis. Together, plasmolysis and deplasmolysis are part of a plasmolytic cycle. Although often used in cell physiology classes to demonstrate semipermeability of membranes, the phenomenon of plasmolysis has only rarely been studied in respect of subcellular changes in the cytoplasm. Sitte (1963) first used electron microscopy to analyze ultrastructural changes of Elodea leaf cells in different plasmolytic solutions. However, many questions remained. In plasmolyzed cells, membrane

I. Lang-Pauluzzi and B. E. S. Gunning: Cytoskeletal elements in plasmolytic cycle

surface area is deleted into osmotically induced endocytotic vesicles (Oparka et al. 1990, Gordon-Kamm and Steponkus 1984), but a considerable amount of membrane material is also preserved in Hechtian strands (Oparka et al. 1994). In deplasmolysis, vesicles are not reused, whereas Hechtian strands are reincorporated into the plasma membrane. The structure of their attachment sites is a subject of recent investigations (Canut et al. 1998, Bachewich and Heath 1997, Reuzeau et al. 1997b, Pont-Lezica et al. 1993). Cytoskeletal elements have been shown to be part of Hechtian strands in fungal hyphae (Bachewich and Heath 1997), mosses (Schnepf et al. 1986), and Tradescantia epidermal cells (Cleary cited in Gunning and Steer 1996), and there is a growing body of evidence for the involvement of cytoskeletal elements in the attachment sites of Hechtian strands and wall-tomembrane linkers. These protein complexes are possible candidates for cell-cell communication and signal transduction routes in plant cells (Canut et al. 1998; Reuzeau et al. 1997a, b; Pont-Lezica et al. 1993). Cortical arrays of microtubules and actin normally lie just within the plasma membrane and follow its contours; they too must be adjusted during plasmolysis. In this study we have investigated their behavior in onion inner epidermis cells during a plasmolytic cycle in sucrose solutions. Furthermore, we analyzed the fate and form of plasmolysis in these cells after application of cytoskeleton-disrupting drugs.

Material and m e t h o d s Plant material Onion (Allium cepa L.) inner epidermal peels were prepared as described in Lichtscheidl and Url (1990). The single cell layer peels were allowed to float cuticle side up on distilled water for at least 30 rain to let the shock of the preparation subside. Best results for sucrose plasmolysis were achieved by floating the cells for 5 min on 0.8 M sucrose solution and another 30 min on 0.5 M sucrose solution. Deplasmolysis was carried out in 0.25 M sucrose solution (10 min) followed by a transfer to distilled water for complete reexpansion of the protoplast.

Cytoskeletal drugs For destruction of microtubules, cells were treated with 10 gM oryzalin (Riedel-de Hahn) for 10 min prior to plasmolysis. Actin microfilaments were depolymerized in 10 gM latrunculin B (CaP biochem) for 10 min prior to transfer to plasmolytic solutions. Organelle movement stopped immediately after application of latrunculin B. For immunofluorescence staining, epidermal peels were mounted cuticle side down on coverslips and secured with sticky tape. During the fixation and staining process, cells were stored in a wet chamber.

I75

Fixation and labellings Nonplasmolyzed ceils were fixed for 30 min in 4% paraformaldehyde or a mixture of 4% paraformaldehyde and 0.5% glutaraldehyde in PHEM buffer [60 mM piperazine-N,N'-bis(2-ethanesulfonic acid), 25 mM HEPES, 10 mM EGTA, 4 mM MgC12, pH 6.9]. This buffer was also used for rinsing the cells between the staining steps. Plasmolyzed cells were fixed in 0.5 M sucrose solution containing 4% paraformaldehyde. Pectinase (Sigma) treatment (0.05% in distilled water, 5 min) was followed by 40 rain of membrane permeation in 0.2% Triton X-100 (Sigma) in PHEM buffer. The primary antibody was applied overnight at room temperature. For microtubule staining, we used mouse anti-c~-tubulin (Sigma; clone B512), 1 : 1500 in PBS (phosphate-buffered saline). Cells were incubated with the secondary antibody, a sheep anti-mouse fluorescein isothiocyanate conjugate (1 : 30 in PBS; Sigma) for 1 h at 37 ~ Cells were mounted on a 9 : 1 mixture of Mowiol 40-88 (Sigma) and pphenylenediamine dihydrochloride (Sigma-Aldrich). Slides were stored at 4 ~ to allow the mounting medium to dry and observed the next day. For immunostaining of actin, we used a rabbit antiactin antibody (ICN; clone C4) in combination with an anti-rabbit fluorescein isothiocyanate conjugate (Sigma). For phalloidin staining, cells were treated with a permeabilization buffer (Traas et al. 1987) with 1% mannitol, 50 mM piperazine-N,N'-bis(2-ethanesulfonic acid, 5 mM EGTA, 2 mM MgSO4, 0.01% Nonidet P-40 (Sigma), 5% dimethyl sulfoxide. Actin microfilaments were then stained by directly applying Texas Red-phalloidin (Calbiochem; 1 : 60 in permeabilization buffer). Visualization of the plasma membrane, endoplasmic reticulum, and Hechtian strands was achieved by application of the membranespecific dye 3,3-dihexyloxacarbocyanine iodide [DiOC6(3)] (Molecular Probes). We used a concentration of 5 ~tg/ml in water or plasmolytic solution, respectively. The dye was applied directly to living cells.

Microscopy For immunofluorescence microscopy, we used a Bio-Rad MRC-600 confocal laser scanning system attached to a Zeiss Axiovert 10 inverted microscope. High-numerical-aperture oil immersion objectives x40 (numerical aperture, 1.3) and x60 (numerical aperture, 1.25) were used. Images were integrated by averaging 3-5 scans with a Kalman filter. Cells were optically sectioned in 1 gm increments. Reconstruction of the periclinal cell surface was carried out by maximum pixel projection of image stacks with Confocal Assistant version 4 (Bio-Rad). DIC (differential interference contrast) images were taken either on the confocal microscope using the transmission mode or on a Zeiss Axioplan with objectives x20 (numerical aperture, 0.5), x40 (numerical aperture, 1.0), and MOO (numerical aperture, 1.3). Photographs were taken on TechPan 25 film (Kodak). Alternatively, we used a Reichert Biovar microscope with objectives >425 (numerical aperture, 0.45) and x40 (numerical aperture, 0.65) and a 100 ASA film (Kodak T-Max).

Results

Plasmolysis Onion inner epidermis cells are shown in Fig. i a. In 0.5 M sucrose solutions, plasmolysis results in the formation of a round protoplast (Fig. 2 a), so-called

176


I. Lang-Pauluzziand B. E. S. Gunning: Cytoskeletal elements in plasmolyticcycle

177

convex plasmolysis (Oparka 1994). In comparison to plasmolytic salt solutions like CaC12 or KC1, which slowly penetrate into the protoplast and alter the structure of organelles as well as the size of the protoplast, sucrose has no such effects (Sitte 1963). Even prolonged treatment of cells in plasmolytic sucrose solution up to several hours did not result in re-expansion of the protoplast. Hechtian strands (Hecht 1912) maintain a connection between the plasmolyzed protoplast and the cell wall. In onion inner epidermal cells, Hechtian strands have a diameter of 0.5 to 1 ~tm. They can be observed with good D I C optics but are much easier to image after staining with DiOC6(3) dye (Fig. 4). Hechtian strands can branch (Fig. 3b) and sometimes contain organelles as well as endoplasmic reticulum and cytoskeletal elements (Figs. 3b and 5). At the cell cortex, Hechtian strands emerge from a membranous network which remains firmly attached to the wall. Light-microscopical studies show that this reticulum contains organelles and endoplasmic reticulum. In plasmolyzed cells, it forms bulbs at both termini of broken plasmodesmata (Fig. 3 a).

in cases where the microtubule fluorescence in Hechtian strands branches (Fig. 3 b). Microtubules in Hechtian strands can form looplike structures close to the cell wall (Fig. 3 b).

Microtubules

Destabilization of cytoskeletal elements

In nonplasmolyzed cells of onion inner epidermis, microtubules are located in the cortical layer of cytoplasm in either longitudinal or transverse or oblique arrays (Fig. l b ) . Plasmolysis causes a repositioning of cortical microtubules to fit the smaller protoplast, and thus their arrays must become slightly more dense. The organization and disposition of cortical microtubules remains similar to nonplasmolyzed cells except for longitudinal orientations, which could not be detected in plasmolyzed cells. Most cells showed oblique or transverse cortical microtubles after plasmolysis (Fig. 2 b). Their arrays are not grossly distorted from pattern s seen in controls, and there are regions where parallel microtubules, presumably relics of original parallel arrays, can Still be seen. A b o u t 5 to 10% of Hechtian strands contain microtubules (Fig. 3 b), and they must be present in bundles

Actin microfilaments Before plasmolysis, actin microfilaments are present in bundles throughout the cytoplasm, in transvacuolar cytoplasmic strands where they provide tracks for organelle movement, and in a fine cortical network (Cleary and Mathesius 1996, Sonobe and Shibaoka 1989, Traas et al. 1987). After plasmolysis, transvacuolar bundles of actin microfilaments are preserved within the smaller protoplast (Fig. 5). The array of actin is similar to control cells and approximately 1 or 2% of Hechtian strands are stained with Texas Redphalloidin. However, it is difficult to observe the fine cortical actin array in plasmolyzed cells. This fine structure could be disrupted during the detachment of the protoplast from the cell wall. Organelle movement continues during the whole process of plasmolysis, suggesting that actin micro filaments remain functional.

Oryzalin Cells were treated with 1.D gM oryzalin prior to plasmolysis (Fig. 6a). Distribution of cytoplasm and organelle movement were not affected by oryzalin. In sucrose solution, detachment of the protoplast is smooth; Hechtian strands are formed even after drug treatment and the rate of plasmolysis was not altered significantly (data not shown). Light microscopical studies did not show differences to nontreated plasmolyzed cells. Latrunculin B Destruction of actin microfilaments with 1 0 g M latrunculin B prior to plasmolysis stopped organelle movement immediately in onion epidermal cells. This

Fig. 1 a, b. Inner epidermal cells of Allium cepabefore plasmolysis,a DIC image, b Indirect immunofluorescencestaining of cortical microtubules. The array of parallel microtubules can be longitudinal,random, or transverse. Bars: 100 gm Fig. 2a, b. Convex plasmolysis of the cells in 0.5 M sucrose solution; the protoplast is detached from the cell wall. a DIC image, b Microtubule staining.After plasmolysis,the orientation of microtubules remains parallel. Microtubules are present in Hechtian strands (arrows). Bars: 50 gm

178


Fig. 3a, b. Detailed image of Hechtian strands, a DIC image.Thin threads connect the protoplast with the cell wall.The strands are straight and can contain cytoplasm, endoplasmic reticulum, and other organelles. At both sides of the cell wall, bulbs of cytoplasm (Hechtian reticulation) are visible (arrows). Hechtian strands are connected with that reticulation, h Microtubule staining, high magnification of Hechtian strands and reticulation. Microtubules in branching strands (arrowheads). At the cell wall, microtubules form a looplike structure (arrow). Bars: 20 gm

I. Lang-Pauluzzi and B. E. S. Gunning: Cytoskeletalelements in plasmolyticcycle suggests a rapid destabilization of actin microfilaments, as has been described previously in mammalian cells (Epstein et al. 1999) and fungal hyphae (Bachewich and Heath 1997, Gupta and Heath 1997). The cytoplasm however is not evenly distributed, and cytoplasmic strands are missing (Fig. 6b). Plasmolysis forms of onion cells are still similar to control cells without latrunculin treatment. Immediately after application of the plasmolytic solution, the protoplast starts to retract smoothly from the wall and Hechtian strands are formed. The speed of the process is comparable to untreated cells; it takes about 30 rain to reach the final plasmolyzed state. Addition of latrunculin B and oryzalin together prior to plasmolysis resulted in cessation of organelle movement and induction of a fixation-like state of the ceils. During plasmolysis, however, the protoplast retracted from the cell wall in convex plasmolysis forms (data not shown). Hechtian strands could not be detected and the cells died during deplasmolysis. Deplasmolysis Within seconds after addition of water to plasmolyzed cells the protoplast rounds up and Hechtian strands disintegrate into rows of droplets. During these very first stages of deplasmolysis, the protoplast always retracts a little further before increasing in size again due to osmotic water uptake. Re-expansion then proceeds rapidly, taking a few minutes to complete deplasmolysis. The plasma membrane reincorporates Hechtian strands and any larger portions of cytoplasm that lay along them (data not shown). The plasmolytic cycle is complete when the protoplast is appressed to the cell wall again. The whole process takes less than 10 rain in onion inner epidermal cells. Organelle movement is sustained throughout deplasmolysis. Endoplasmic reticulum remains clearly visible as tubular and vesicular structures, similar to nonplasmolyzed cells where the cortical endoplasmic reticulure is positioned close to the plasma membrane (Hepler et al. 1990). Its highly dynamic tubules can be interconnected to form a net, thin bundles or a reticulum of polygonal meshes of different sizes (Lichtscheidl 1995). These pictures of the cortical endoplasmic reticulum in control cells of onion inner epidermis show high similarities to the Hechtian network that remains attached to the cell wall in plasmolyzed cells.

179

Microtubules In contrast to plasmolysis, where arrays of cortical microtubules are not altered dramatically, re-expansion of the protoplast during deplasmolysis is accompanied by marked reorganization of microtubules. In the early stages of deplasmolysis, the array tends to break up into a random network and discrete short lengths (Fig. 7a). As deplasmolysis proceeds, the microtubules display more and more order, and when deplasmolysis is complete, they are once again arranged parallel to one another (Figs. 7 a-d). Reorganization of microtubules lags behind deplasmolysis: the protoplast has almost reached its original size when the array is still disrupted. In onion epidermal cells, the restoration of the microtubule array takes about 20-30 min longer than deplasmolysis. Actin microfilaments No dramatic changes to the cytoplasmic actin cytoskeleton occur during plasmolysis or deplasmolysis. Actin microfilaments are still visible as bundles within the cytoplasm as the protoplast expands after plasmolysis. Organelle movement is sustained. We were not able to detect a fine cortical actin array in plasmolyzed or deplasmolyzed cells. This could be due to the fixation procedure. However, it is possible that this fine cortical structure becomes disrupted during plasmolysis although light microscopical studies show movements of small cell organelles within the Hechtian reticulation in plasmolyzed cells (data not shown), suggesting the presence of actin within that structure. There is, however, no direct evidence that actin microfilaments within the Hechtian reticulation correspond to the cortical actin array seen in Tradescantia virginiana (Cleary and Mathesius 1996). Drug treatment Cells that were treated with oryzalin and latrunculin B showed normal deplasmolysis, and the rates of deplasmolysis were the same as in nontreated cells. Light microscopical studies of cells that were treated with latrunculin B showed vesicular endoplasmic reticulum and no organelle movement. However, the protoplast re-expanded after addition of hypotonic solution, indicating functional membranes and reincorporation of membrane material from Hechtian strands.

180

I. Lang-Pauluzzi and B. E. S; Gunning: Cytoskeletal elements in plasmoly~ic cycle


Discussion

In plasmolyzed onion inner epidermal cells, cortical microtubules retain the general conformation that they possessed in nonplasmolyzed cells, with the exception of longitudinal arrays, which occur in some unplasmolyzed cells but were never seen after plasmolysis. This indicates reorganizations of cortical microtubules during plasmolysis. Actin microfilaments are not subject to dramatic changes during plasmolysis. However, with the methods applied here, we could only detect cytoplasmic actin microfilaments but not the fine cortical array of actin microfilaments after plasmolysis. Actin microfilaments and microtubules can occur in Hechtian strands, perhaps contributing to contacts between the retracted protoplast and the cell wall. During deplasmolysis, the cortical microtubule array does not re-expand congruently with the plasma membrane but becomes disrupted into an irregular array of randomly oriented, short fragments. These gradually increase in length and are rearranged into parallel arrays; finally, a pattern similar to nonplasmolyzed cells is established. Actin microfilaments re-expand gradually with the protoplast during deplasmolysis; no breakdown comparable to that of the microtubule system was detected. In onion inner epidermal cells, the plasmolytic cycle was not significantly affected by disruption of cytoskeletal elements. Our results indicate that the osmotic force during plasmolysis must be stronger than any cytoskeletal stabilization of the protoplast. We could not observe significant differences in the speed of plasmolysis when cytoskeletal elements were destroyed by treatment with antitubulin or antiactin drugs. As proposed by Palta and Lee-Stadelmann (1983), isolated protoplasts act as ideal osmometers, and we see here that this applies irrespectively of their cytoskeletal state. At least the major processes of plasmolysis are membrane dependent, and any role that cytoskeletal elements may have in stabilizing the cytoplasm, protoplast

181

shape, cytoplasmic viscosity, or hydraulic conductivity, seems to be of relatively minor consequence. In plasmolyzed cells, the congruence between cortical microtubules and cellulose microfibrils in the cell wall is lost, but even in the absence of the normal contact with the plasma membrane and cell wall, a cortical array of microtubules remains stable over a period of several hours and even days, when cells are kept alive (Lang-Pauluzzi unpubl, data). Since the plasma membrane retains contact with the cell wall along the sides of the cells in most plasmolyzed onion cells and via Hechtian strands at the cell ends where the protoplast is detached from the wall, membrane proteins that are linked to elements in the cell wall, hence stabilizing the microtubule array in its original confguration, might not be affected very much by plasmolysis. In Boodlea protoplasts, the orientation of cortical microtubules has been shown to be dependent on the contact of the plasma membrane to the cell wall, and the array of the microtubules in plasmolyzed cells is stabilized by anchors towards the cell wall, especially to cellulose (Mizuta et al. 1995). The situation in plasmolyzed cells is similar to that in isolated protoplasts, but there a new cell wall is synthesized. Extensive studies on algal protoplasts by Galway and Hardham (1986) have shown that cortical microtubules are disordered after isolation, but within 3 to 4 h microtubules became reorganized into an ordered, symmetrical array, concomitant with the first detectable cell wall synthesis. In our plasmolyzed onion protoplasts there was insufficient time for any wall regeneration. Early studies by Eschrich (1957) showed local callose depositions within the cell wall of plasmolyzed onion epidermis cells, but the formation of a new wall layer was never observed. Moreover, the retention of the contact between the protoplast and the cell wall by way of Hechtian strands is without counterpart in isolated protoplasts and perhaps is a factor in the failure to commence wall regeneration in plasmolyzed onion epidermal cells. Most of the

Fig. 4. Staining of Hechtian strands with the membrane-specific dye DiOC6(3). The strands can branch and contain mitochondria (arrows) that are brightly stained. At the cell wall, the strands merge with the Hechtian reticulation. Bar: 50 gm Fig. 5. Actin labelling of plasmolyzed cells with Texas Red-phalloidin. Bundles of actin microfilaments are visible throughout the cytoplasm. Hechtian strands also contain actin (arrows). Bar: 50 gm Fig. 6a, b. Plasmolysis in presence of cytoskeletal drugs, a Treatment with 10 p.M oryzalin destroys microtubules; DIC image. Convexplasmolysis form, mid-focal plane. The cytoplasm is equally distributed and organelle motility is normal, b Treatment with 10 ~tM latrunculin B. Immediate stop of organelle movement, the cytoplasm is accumulating in spotlike areas. Plasmolysis form is convex. Bars: 50 gm

182



cortical microtubules remain in close contact with the plasma membrane of plasmolyzed protoplasts, and only a few microtubules are present within Hechtian strands. Orientations of microtubules at final stages of plasmolysis were not the same in all cells. With the methods used here, we cannot say whether the orientation of microtubules changes through time, as in case of the Mougeotia protoplasts studied by Galway and Hardham (1986). With one exception, considered below, it is likely that the orientation of microtubules that was present before plasmolysis was retained in our experiments until the plasmolyticum was changed to induce deplasmolysis. It should be possible to check these events directly in live cells now that transformants with green-fluorescent-protein-labelled microtubules are available (Ueda et al. 1999). Another approach would be to microinject labelled tubulin into living cells, but so far our attempts to do this failed due to methodical difficulties. As noted above, cortical microtubules still appear parallel, in oblique or transverse orientation after plasmolysis, reflecting the situation before plasmolysis, but there is an interesting exception. Although longitudinal arrays of cortical microtubules are common in nonplasmolyzed cells, we could not find corresponding longitudinal microtubule orientations in plasmolyzed cells, suggesting that longitudinal arrays are relatively unstable and become oblique. This reorganization of microtubules must occur during the process of plasmolysis. In Spirogyra cells, Iwata (1995) also observed alterations in orientation, depending on the concentration of plasmolyticum that was used. The reorientation must occur as quickly as plasmolysis itself. Rapid changes of microtubule arrays are known in a number of other situations, including developmental events (Zandomeni and Schopfer 1994, Wymer and Lloyd 1996, Hush and Overall 1996, Kropf et al. 1997). The rather stable pattern of cortical microtubules in plasmolyzed cells becomes rapidly disrupted as soon as deplasmolysis starts and only slowly regenerates to form an oblique array, similar to arrays in most nontreated cells (Fig. 7 a-d). Galway and Hardham (1989) showed similar results after oryzalin-induced disas-

183

sembly of microtubules in isolated protoplasts of the alga Mougeotia sp. immediately after the isolation process of protoplasts (Galway and Hardam 1986). During protoplast regeneration, microtubules first formed short fragments, then clusters and longer microtubules that are coaligned in localized areas, and finally highly organized cortical microtubule arrays that were indistinguishable from those in untreated protoplasts. Oryzalin treatment of onion epidermal cells gave similar time courses; however, the disruption that occurs when deplasmolysis starts differs in that there is no complete dissolution of the microtubule array but merely a transient fragmentation and disorganization. Although the reasons for the breakdown of microtubules during the initial stages of deplasmolyis remain unclear, it is possible that the rapid change in plasma membrane surface area that occurs at that moment breaks membrane-microtubule connections, resulting in a loss of stability. An other interesting effect of plasmolysis is the drastic alteration of membrane potential that has been shown in nonturgid cells of Chara australis (McCulloch and Beilby 1997). Conductance of the plasma membrane increases with time and, after plasmolysis, is irreversible. It remains to be seen how these findings relate to microtubule orientation or membrane expansion during deplasmolysis. However, a regulation of the orientation of cortical microtubules in response to plasmolytic solutions is possible, as shown in Spirogyra cells (Iwata 1995), where after depolymerization, cortical microtubules were allowed to recover in mannitol solutions of different concentrations (0.1-0.3 M). Low concentrations resulted in transverse orientations, in higher osmotic solutions, cortical microtubules were arranged obliquely or longitudinally. A transfer to water caused deplasmolysis and reorganization of cortical microtubules into transverse arrays. Once again, it should be possible to monitor these phenomena in cells with green-fluorescent-protein-labelled microtubules. Actin microfilaments proved to be very resilient and responded rapidly to plasmolysis-induced deformation of the protoplast and associated rearrangement

Fig. 7a--d. Microtubule staining at different stages of deplasmolysis, a At the beginning of deplasmolysis, microtubules are cut down into small rods and are randomly oriented, b After 10 rain in distilled water, microtubules start to rearrange in parallel arrays and increase in length, c A n o t h e r 10 min later, some cells show almost normal, parallel microtubule distribution (cell in the middle), others are still more plasmolyzed, and microtubules are still randomly oriented (cell at the left), d Complete deplasmolysis. The protoplast fills the cell wall again, long cortical microtubules are arranged parallelly. Bars: 50 gm

184

[. Lang-Pauluzzi and B. E. S, Gunning: Cytoskeletal elements in plasmolytic cycle

of cytoplasmic strands (see also Lichtscheidl and Url 1987, Cleary 1995, Balugka et al. 1997). We find that during plasmolysis, actin microfilaments can adapt to the smaller protoplast without m a r k e d change in organization and structure. Again, in deplasmolysis, cytoplasmic strands, organelle m o v e m e n t , and actin microfilaments are retained throughout the whole process. Most of the filamentous actin of a plasmolyzed cell is present in the protoplast in cytoplasmic strands, but with the methods we applied, we were not able to detect the fine cortical actin that occurs in unplasmolyzed onion epidermal cells and has also been shown in Tradescantia leaf epidermis (Cleary and Mathesius 1996). Using high-resolution light microscopy, however, organelle m o v e m e n t within the Hechtian reticulation can be detected, indicating the presence of cortical actin in plasmolyzed cells. We found both classes of cytoskeletal elements within Hechtian strands (Figs. 2 b, 3 b, and 5). They m a y thus participate in connecting the protoplast to the cell wall. The diameter of Hechtian strands varies between 80 and 250 nm (Sitte 1963, Schnepf et al. 1986) or up to 2 ~m in moss rhizoids (Smith 1972). The strands can branch ( O p a r k a 1994) and incorporate endoplasmic reticulum, mitochondria, and small elements of the cytoplasm. With a diameter of 24 nm (Gunning and Steer 1996) microtubules can easily be a c c o m m o d a t e d in Hechtian strands. In p r o t o n e m a cells of the moss Funaria hygrometrica, Schnepf et al. (1986) were able to demonstrate microtubules within Hechtian strands by transmission electron microscopy. The looplike tubulin structure next to the cell wall seen here in onion epidermal cells could help to stabilize the Hechtian network in this region of the cell. However, cells whose microtubules have been r e m o v e d by oryzalin treatment still develop Hechtian strands, so a role for microtubules in anchoring seems doubtful. Actin is a c o m p o n e n t of t r a n s m e m b r a n e linkers for cytoskeleton-plasma m e m b r a n e - e x t r a c e l l u l a r matrix attachments in animal cells (Sheetz et al. 1998, Burridge et al. 1997). In plants, actin microfilaments have been reported in Hechtian strands of plasmolyzed fungal hyphae (Bachewich and H e a t h 1997) and in Tradescantia epidermal cells (Cleary cited in Gunning and Steer 1996). In our studies of onion epidermal cells, actin micro filaments appear far less abundant in Hechtian strands than microtubules. This fact, however, could be due to the difficulties of preservation of actin microfilaments in Hechtian strands when the cell is in the final stage of plasmolysis. The diffi-

culties of preserving fine cortical actin have been documented by Cleary and Mathesius (1996). Our observation of actin microfilaments in Hechtian strands does not necessarily m e a n that they are components of attachment sites but they could be derived f r o m cortical actin (Cleary and Mathesius 1996, Wasteneys et al. 1996, Sonobe and Shibaoka 1989) and could also provide links to the cortical endoplasmic reticulum ( O p a r k a 1994, Pont-Lezica etal. 1993, H e p l e r et al. 1990, Lichtscheidl et al. 1990, Lichtscheidl and Url 1990). Destruction of actin microfilaments with latrunculin B resulted in the cessation of organelle m o v e m e n t and the loss of cytoplasmic strands in onion cells, as also shown in fungal hyphae (Bachewich and H e a t h 1997), but plasmolysis occurred with the normal speed, and neither the plasmolysis forms nor the Hechtian strands were altered significantly.

Acknowledgments Many thanks to Ann Cleary,Irene Lichtscheidl, and Owen Schwartz for technical assistance and helpful discussions. Parts of this work were carried out at the Research School of Biological Sciences, Australian National University,and were supported by a grant from the Austrian Ministry for Science, Research, and Arts (nr. 951487).

References Bachewich CL, 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 Balugka F,Vitha S, Barlow PW,Volkmann D (1997) Rearrangements of F-actin arrays in growing cells of intact maize root apex tissues: a major developmental switch occurs in the postmitotic transition region. Eur J Biol Chem 72:113-121 Burridge K, Chrzanowska-Wodnicka M, Zhong C (1997) Focal adhesion assembly.Trends Cell Biol 7:342-347 Canut H, Carrasco A, Galaud J-R Cassan C, Bouyssou H, Vita N, Ferrara R Pont-Lezica R (1998) High affinity RGD-binding sites at the plasma membrane of Arabidopsis thaliana links the cell wall. Plant J 16:63-71 Cleary A (1995) F-actin redistributions at the division site in living Tradescantia stomatal complexes as revealed by microinjection of rhodamine-phaltoidin. Protoplasma 185:152-165 - Mathesius U (1996) Rearrangements of F-actin during stomatogenesis visualized by confocal microscopy in fixed and permabilized Tradescantia leaf epidermis. Bot Acta 109:15-24 Epstein DL, Rowlette LL, Roberts BC (1999) Actomyosin drug effects and aqueous outflow function. Invest Ophthalmol Vis Sci 40:74-81 Erwee MG, Goodwin PB (1984) Characterisation of the Egeria densa leaf symplast:response to plasmolysis,deplasmolysisand to aromatic amino acids. Protoplasma 122:162-168 Eschrich W (1957) Kallosebildung in plasmolysierten Allium cepaEpidermen. Planta 48:578-586


-

Galway ME, Hardham A R (1986) Microtubule reorganisation, cell wall synthesis and establishment of the axis of elongation in regenerating protoplasts of the alga Mougeotia. Protoplasma 135: 130-143 - (1989) Oryzalin-induced microtubule disassembly and recovery in regenerating protoplasts of the alga Mougeotia. J Plant Physiol 135:337-345 Gordon-Kamm WJ, Steponkus PL (1984) The behavior of the plasma membrane following osmotic contraction of isolated protoplasts: implications in freezing injury. Protoplasma 123:83-94 Gunning BES, Steer MW (1996) Bildatlas zur Biologie der Pflanzenzelle: Struktur und Funktion, 4th edn. Gustav Fischer, Stuttgart Gupta GD, Heath IB (1997) Actin disruption by latrunculin B causes turgor-related changes in tip growth of Saprolegnia ferax hyphae. Fung Genet Biol 21:64-75 Hecht K (1912) Studien fiber den Vorgang der Plasmolyse. Beitr Biol Pflanzen 11:133-189 Hepler PK, Palevitz BA, Lancelle SA, McCauley MM, Lichtscheidl I (1990) Cortical endoplasmic reticulum in plants. J Cell Sci 96: 355-373 Hush JM, Overall RL (1996) Cortical microtubule reorientation in higher plants: dynamics and regulation. J Microsc 181:129-139 Iwata K (1995) Regulation of the orientation of cortical microtubules in Spirogyra cells. J Plant Res 108:531-534 Kjellbom R Snogerup L, StOhr C, Reuzeau C, McCabe PE Pennell RI (1997) Oxidative cross-linking of plasma membrane arabinogalaetan proteins. Plant J 12:1189-1196 Kropf DL, Williamson RE, Wasteneys GO (1997) Microtubule orientation and dynamics in elongating characean internodal cells following cytosolic acidification, induction of pH bands, or premature growth arrest. Protoplasma 197:188-198 Lichtscheidl IK (1995) Organization and dynamics of endoplasmic reticulum: Allium cepa L. inner epidermis. Wiss Film 47:95-109 - Url WG (1987) Investigation of the protoplasm of Alliurn cepa inner epidermal cells using ultraviolet microscopy. Eur J Cell Biol 43:93-97 - (1990) Organization and dynamics of cortical endoplasmic reticulum in inner epidermal cells of onion bulb scales. Protoplasma 157:203-215 - Laneelle SA, Hepler PK (1990) Actin-endoplasmic reticulum complexes in Drosera: their structural relationship with the plasmalemma, nucleus, and organelles in cells prepared by high pressure freezing. Protoplasma 155:116-126 McCulloch SR, Beilby MJ (1997) The electrophysiology pf plasmolysed cells of Chara austral&.J Exp Bot 312:1383-1392 Mizuta S, Tsuji T, Tsurumi S (1995) Role of plasma membrane proteins and cell wall in meridional arrangement of cortical microtubules in Boodlea coacta. Protoplasma 189:123--131 Oparka KJ (1994) Plasmolysis: new insights into an old process. New Phytol 126:571-591 Prior DAM, Harris N (1990) Osmotic induction of fluid-phase endocytosis in onion epidermal cells. Planta 180:555-561 - Crawford JW (1994) Behavior of plasma membrane, cortical endoplasmic reticulum and plasmodesmata during plasmolysis of onion epidermal cells. Plant Cell Environ 17:163-171 - - (1996) Membrane conservation during plasmolysis. In: Smallwood M, Knox JR Bowles DJ (eds) Membranes: specialized functions in plants. Bios Scientific Publishers, Oxford, pp 3%56 -

-

-

-

185

Palta JR Lee-Stadelmann OY (1983) Vacuolated plant cells as ideal osmometer: reversibility and limits of plasmolysis, and estimation of protoplasm volume in control and water-stress-tolerant cells. Plant Cell Environ 6:601-610 Pont-Lezica RF, McNally JG, Pickard BG (1993) Wall-to-membrane linkers in onion epidermis: some hypotheses. Plant Cell Environ 16:111-123 Porter JC, Hogg N (1998) Integrins take partners: cross-talk between integrins and other membrane receptors. Trends Cell Biol 8: 390-396 Reuzeau C, Doolittle KW, McNalty JG, Pickard BG (1997a) Covisualisation in living onion cells of putative integrin, putative spectrin, actin, putative intermediate filaments and other proteins at the cell membrane and in an endomembranous sheath. Protoplasma 199:173-197 - McNally JG, Pickard BG (1997b) The endomembrane sheath: a key structure for understanding the plant cell? Protoplasma 200: 1-9 Schnepf E, Deichgr~iber G, Bopp M (1986) Growth, cell wall formation and differentiation in the protonema of the moss, Funaria hygrometrica: effects of plasmolysis on the developmental program and its expression. Protoplasma 133:50-65 Sheetz MR Felsenfeld DR Galbraith CG (1998) Cell migration: regulation of force on extracellular matrix-integrin complexes. Trends Cell Biol 8:51-54 Sitte P (1963) Zellfeinbau bei Plasmolyse. Protoplasma 57: 304333 Smith DL (1972) Staining and osmotic properties of young gametophytes of Polypodium ulgare L. and their bearing on rhizoid function. Protoplasma 74:465-479 Sonobe S, Shibaoka H (1989) Cortical fine actin filaments in higher plant cells visualized by rhodamine-phalloidin after pretreatment with m-maleimidobenzoyl N-hydroxysuccinimide ester. Protoplasma 148:80-86 Stadelmann E (1964) Zu Plasmolyse und Deplasmolyse yon AlliumEpidermen. Protoplasma 59:14-68 Traas JA, Doonan JH, Rawlings D J, Shaw PJ, Watts J, Lloyd CW (1987) An actin network is present in the cytoplasm throughout the cell cycle of carrot cells and associates with the dividing nucleus. J Cell Biol 105:387-395 Ueda K, Matsuyama T, Hashimoto T (1999) Visualization of microtubules in living cells of transgenic Arabidopsis thaliana. Protoplasma 206:201-206 Url W (1960) Rosettensystrophe in Thioharnstoff-Zucker-Mischl6sungen. Protoplasma 52:260-273 - (1971) The site of penetration resistance to water in plant protoplasts. Protoplasma 72:427-447 - (1974) Plasmolyse und Zytorrhyse. Institute for Scientific Films, G6ttingen, Film C 1144, pp 3-14 Wasteneys GO, Collings DA, Gunning BES, Hepler PK, Menzel D (1996) Actin in living and fixed characean internodal cells: identification of a cortical array of fine actin strands and chloroplast actin rings. Protoplasma 190:25-38 Wymer C, Lloyd C (1996) Dynamic microtubules: implications for cell wall patterns. Trends Plant Sci 1:222-228 Zandomeni K, Schopfer P (1994) Mechanosensory microtubule reorientation in the epidermis of maize coleptiles subjected to bending stress. Protoplasma 182:96-101