Cytochalasin is known to inhibit cytoplasmic streaming rapidly in characean cells without disassembling their actin bundles. Lower cytochalasin concentrations than those needed for streaming inhibition are now shown to disrupt bundle assembly and, over longer periods, assembled bundles. After local wounding, cytochalasin limited bundle regeneration to the production of polygons and straight, discontinuous bundles that rarely connected to bundles outside the wound. The regenerated bundles supported only scattered organelle movements, whereas long, oriented bundles of control cells were connected to those outside the wound and supported bulk endoplasmic streaming. Unwounded Chara plants cultured for up to 2 weeks in 1 μM-cytochalasin maintained normal bundle orientation and rapid cytoplasmic streaming, but the mean number of bundles per file of chloroplasts fell from 5·2 in controls to 2·0 in growing cells and 3·4 in nongrowing cells. These structural effects seem more likely than the streaming inhibition to reflect cytochalasin’s in vitro effect of blocking extension at the barbed but not the pointed end of F-actin. In particular, cytochalasin inhibited the extension into the wound of bundles in which only the barbed ends of filaments would be exposed. However, short lengths of isolated bundles grew within the wound and bundle growth in the intact cell continued, albeit in modified form. It is suggested that these examples of continuing bundle growth involve cytochalasin-resistant mechanisms that are not wholly dependent on barbed-end filament growth.

Cytoplasmic streaming in characean algae involves subcortical bundles of unipolar actin filaments (Williamson, 1975; Palevitz & Hepler, 1975; Kersey et al. 1976). Groups of bundles lie beneath each chloroplast file, anchored at the boundary between the streaming endoplasm and the surrounding sleeve of stationary cortical cytoplasm. Streaming is inhibited by cytochalasin B (Wessels et al. 1971; Williamson, 1972, 1975; Bradley, 1973; Bostrom & Walker, 1976; Nagai & Kamiya, 1977; Kuroda & Kamiya, 1981; Nothnagel et al. 1981), a fungal metabolite inhibiting many actin-based processes.

Cytochalasin may affect actin in several ways, but at low concentrations it binds in vitro to the barbed ends of actin filaments where G-actin is added most rapidly (MacLean-Fletcher & Pollard, 1980; Pollard & Mooseker, 1981). At equilibrium in the presence of cytochalasin, short filaments coexist with an increased concentration of G-actin (Hartwig & Stossel, 1979; Tellam & Frieden, 1982). The actin cytoskeleton is usually severely disrupted when animal cells are treated with cytochalasin and the mechanism of its action is not always clear, although valuable attempts have been made to relate the concentration dependence of different in vivo effects to the concentration dependence of different in vitro effects on actin (Yahara et al. 1982). The uncertainty is even greater with the characean algae, where alterations to the actin bundles in the presence of cytochalasin were not detected by light or electron microscopy (Wessels et al. 1971; Bradley, 1973; Williamson, 1972, 1975). (Binding of a fluorescent phallotoxin was reduced in cells that had been internally perfused with cytochalasin but, since the effect occurred more slowly than streaming inhibition (Nothnagel et al. 1981), its relevance to that inhibition is doubtful.) Such stability to cytochalasin, however, is not general in plant cells (Blatt et al. 1980; Hoch & Staples, 1983 ; Parthasarathy, 1985 ; Perdue & Parthasarathy, 1985 ; Witztum & Parthasarathy, 1985). The actin cytoskeleton of the mature characean cell differs from that of most other cells in not undergoing major developmental-, environmental- or cell-cycle-related changes during the cell’s life of many months. Such stability might be associated with mechanisms restricting the exchange of G-actin at filament ends, mechanisms that might also offer some stability towards cytochalasin. We therefore studied cytochalasin’s effects on Chara over longer periods than previously used and in two situations where actin bundles would be changing. The first was the regrowth of actin bundles following their local destruction (Kamitsubo, 1972; Williamson et al. 1984) and the second was the growth of actin bundles required to maintain continuous, nearly longitudinal bundles in extending cells. We found that both existing and growing actin bundles are affected by concentrations of cytochalasin that do not inhibit streaming along assembled bundles.

Plant material

Chara corallina was grown in a glasshouse and in the laboratory with room and window light. Plastic bins (75 1) with a 3 cm layer of soil were used for glasshouse cultures; 201 glass aquaria with 1 cm of agar (Bacto-Agar, Difco Laboratories) were used in the laboratory. The nutrient solution (broadly similar to that used previously; Williamson, 1975) contained (mM): CaCl2, 0·1; MgSO2, 0·1; Na2CO3, 0·2; NH4CI, 0·04; NaCl, 0·5; KC1, 0-1; morpholinopropane sulphonic acid, 0·5; together with (μM):KH2PO4, 0·86; FeCh, 2·48; nitrilotriacetic acid, 10·5; ZnCl2, 0’74; MnCl2, 2·1×10−2; CoCl2, 1·55×10−2; CuCl2, 2·99×10−2; Na2O3, 1-98; Na2MoO4, 0·49; pH7·0. Cultures in soil were started with plants collected locally (mainly from Lake Ginnindera, Belconnen, ACT), cultures in agar with 20–30 apical cuttings from soil-cultured plants. The cutting comprised the growing point with (normally) one expanded internode (>20 mm long) that was pushed into agar cooled to just above its gelling temperature (≈30°C).

Experimental

To determine the effects of cytochalasin B (Sigma Chemical Co.) on the numbers of actin bundles in Chara internodal cells, a 10 mM solution in dimethyl sulphoxide (DMSO) was added to an agar culture to give a final cytochalasin concentration of 1 μM and a final DMSO concentration of 0·01 % (v/v). Controls received an equal volume of DMSO. The additions were made roughly 2 weeks after planting at a time of rapid growth. The length of each macroscopic internodal cell on every plant was measured with a ruler at intervals of one to several days. This gave the growth history of every cell until harvested for immunofluorescence.

The effects of cell length and the presence or absence of cell growth on actin bundle number were studied in similar experiments, except that no additions were made to the media and all cells were harvested 21 or 22 days after planting to minimize any changes due to culture age.

Chloroplast-free ‘windows’ were prepared (Wiliamson et al. 1984) in internodal cells (≈5 cm long) grown in agar-based cultures and cytochalasin B (1 or 10μM) or DMSO (0·01 or 0·1 %, v/v) was added shortly afterwards.

Immunofluorescence

The preparation techniques have been described (Williamson et al. 1984) and also the monoclonal antibodies CC2 and CC6 (Williamson et al. 1986) that bind to the subcortical actin bundles. The number of bundles per file of chloroplasts was counted for each clearly focused file on photographs taken at 6–12 sites along each cell. Counting was done without knowledge of the treatment received by the cells and areas around the neutral lines were avoided.

Microscopy

The techniques for fluorescence microscopy have been described (Williamson et al. 1984). Differential interference contrast microscopy used video recording (Suzaki & Williamson, 1985). A temporal filter (Arlunya TF 4000, Vermont, Victoria 3133, Australia) could enhance stationary cortical structures and filter out moving, endoplasmic structures.

Cell cultures

All cells with lengths exceeding the 10 mm needed for perfusion showed typical (Williamson et al. 1986) arrays of bundles at the cortex-endoplasm interface (Fig. 1A). They did not contain detectable bundle ends such as were readily visible around and within windows during the early stages of regeneration (see fig. 3, Williamson et al. 1984). The number of bundles per file of chloroplasts showed slight variations with cell length and between growing and older, non-growing cells (Fig. 2). Culturing cells in 1 μM-cytochalasin reduced the number of bundles per chloroplast file (Fig. 3). This effect, detectable within 24h, was more pronounced in growing than in non-growing cells (Fig. 1B,C; Fig. 3) and could be seen in vivo (Fig. ID), proving that it was not a perfusion artefact. There was no evidence of discontinuities or disorientation in the depleted bundle array. The apparent thickness of the remaining bundles was increased whether seen in vivo (Fig. ID), as enhanced brightness by immunofluorescence (Fig. 1C), or by scanning electron microscopy (not shown). The number of bundles per chloroplast file was slightly reduced by DMSO alone as the solvent for the cytochalasin (Fig. 3). Cells depleted of bundles by cytochalasin treatment still streamed actively.

Bundle regeneration in the presence of DMSO alone was similar to that described previously (Williamson et al. 1984). After 12–24 h of regeneration, immunofluorescence (Fig. 4) revealed long curving lengths of actin bundles that were particularly abundant towards the edges of a window where they curved round to join the oppositely polarized arrays formerly separated by the neutral line. The centres of such windows were occupied by varying amounts of less-ordered bundles. We have previously suggested explanations for the origin and orientation of these bundles (Williamson el al. 1984).

Immunofluorescence observations were supplemented by video microscopy of cells prior to their fixation. The general features of organelle movements in the windows of control cells allowed more than 12 h for regeneration were readily related to the immunofluorescence image: bulk streaming with a velocity comparable to that outside the window followed a U-turn at each side of the window where immunofluorescence showed long, curving actin bundles (see Fig. 4). Only intermittent movements of usually single organelles occurred in various directions around the central region, where actin bundles were fewer and less ordered. In vivo observations revealed few ôf the bundles that could be demonstrated by immunofluorescence.

Treatment of cells with either 1 μM or 10μM-cytochalasin markedly reduced the quantity of regenerated actin bundles and altered their arrangement. Whereas controls with DMSO alone had an extensive bundle system after 12 h of regeneration (Fig. 4), cytochalasin-treated cells had fewer bundles even after 21 h (Fig. 5A). The long, gently curving bundles seen in controls (Fig. 4) were absent. Short lengths of usually rather straight, thick bundles and variously shaped rings or polygons (Fig. 5A) were prominent to varying degrees in different cells. The regenerated bundles were rarely connected to bundles outside the window and lacked a predominant orientation. The rings and polygons always enclosed chloroplasts emitting red autofluorescence (Fig. 5B,C) but not all such chloroplasts were thus enclosed. Chloroplasts retaining red autofluorescence must have reached the window after its creation by irradiation, since this induces white chloroplast autofluorescence (Williamson et al. 1984). They could have originated from the endoplasm of characean algae, which contains a few chloroplasts that usually rotate as they are carried in the streaming cytoplasm (see Kamiya, 1962). Very occasionally such a chloroplast was indeed seen to become fixed to the cortical cytoplasm in the window, thus halting its rotational and translational movements.

Towards the sides of cytochalasin-treated windows, endoplasm moved en masse in a U-turn across the neutral line. The velocity was slow (≈5 ¼m s-1) compared to the velocity of streaming outside the window (=50μms-1) and comparable to velocities of the U-turn flow seen in control windows before they contained regenerated bundles. In the central regions, such mass flow of endoplasm was absent. Discontinuous movements of single organelles occurred throughout such windows in various directions and over limited distances. These movements were seen close to the cortex-endoplasm boundary but not deeper in the endoplasm. Towards the sides of the window, discontinuous organelle movements in various directions could overlie bulk flow of the deeper endoplasm that was following a U-turn pathway. As in controls, fibres were only occasionally seen where organelles were actively moving. To emphasize that severe inhibition of regeneration occurred without inhibition of streaming elsewhere in the cell, the velocity of streaming outside the window just before fixation was 55 μms-1 for the cell shown in Fig. 5A, 93 % of the velocity just before cytochalasin was applied.

The results will be discussed in two contexts: the action of cytochalasin on characean cells and the assembly of characean actin bundles.

Cytochalasin action

Some amendments are required to the theories of how cytochalasin affects characean cells. Earlier studies showed that relatively high cytochalasin concentrations (40–200μM) inhibited streaming in a few minutes with little if any disruption of the actin bundles (see Introduction). These conclusions remain valid but the present, longer-term experiments show that actin bundle assembly and organization are sensitive to cytochalasin concentrations that are insufficient to inhibit streaming. The one instance of severe inhibition by 1 μM-cytochalasin occurs in the window. It may reasonably be attributed to the scarcity of regenerated bundles rather than to any inhibition of bundle function, since previously assembled bundles continue to support rapid streaming outside the window.

Cytochalasin’s known effects on actin polymerization can more readily explain the effects on bundle assembly than the inhibition of streaming. Similar concentrations produce the in vivo bundle assembly effect (⪖ 1 μM) and the in vitro polymerization effect (2μM; MacLean-Fletcher & Pollard, 1980; Pollard & Mooseker, 1981), whereas streaming along pre-existing bundles is virtually insensitive to such low concentrations (Bradley, 1973; this study). As will be discussed, the effects of cytochalasin on bundle assembly in various situations differ in ways that may relate to its in vitro effects on actin polymerization.

There remain, however, strong grounds for believing that higher cytochalasin concentrations inhibit streaming by acting directly on the force-generating actin system, even if this does not involve depolymerization. Cytochalasin inhibits motive force production by affecting the stationary, cortical cytoplasm rather than the streaming endoplasm (Nagai & Kamiya, 1977; Kuroda & Kamiya, 1981) and, most persuasively, inhibition is rapid and complete in demembranated models where ATP, pH, Ca and other variables are controlled (Williamson, 1975; NothnageleZ al. 1981; Shimmen & Tazawa, 1983). Moreover, cytochalasin rapidly makes the actin bundles insoluble in low salt solutions (Williamson, 1978; Williamson et al. 1985) and, more slowly, decreases their binding of phallotoxin (Nothnagel et al. 1981). Such an idea of two or more in vivo effects with different concentration dependencies accords well with the ideas of Yahara et al. (1982). They postulated for mammalian cells that several in vivo effects with different concentration dependencies reflected multiple in vitro effects on actin with similarly disparate concentration dependencies.

Bundle assembly

Following the earlier immunofluorescence studies, two origins were proposed for bundles regenerating in windows (Williamson et al. 1984): by growth as extensions of the ends of upstream bundles (i.e. those delivering endoplasm to the window), and by growth following independent initiation within the windows. The mechanism of assembly in normally growing cells might be different again since intact cells lack the visible bundle ends suggested to support growth in the window. Bundle assembly itself probably involves filament growth, cross-linking and anchorage to the cortex, but many variations can be imagined in the order, subcellular location and detailed mechanism by which these basic steps could generate a bundle.

The effects of cytochalasin further support the existence of more than one mechanism for bundle assembly in characean cells. The extension of the upstream bundles into windows is very efficiently blocked by cytochalasin. This inhibition may depend on cytochalasin’s ability to cap the barbed ends of actin filaments that would be exposed where the upstream bundles were severed. However, other bundles do regenerate in the windows of both control and cytochalasin-treated cells without connections to the bundle endings at the window edge. With cytochalasin, they form either chloroplast-associated polygons or fibres that generally seem thicker, straighter and less branched than those regenerated in controls. This indicates that a second mechanism(s) of assembly is operating that is less sensitive to cytochalasin than that which extends the upstream bundles. The continuation of some bundle growth is reasonable since nucleation (Tellam & Frieden, 1982), filament annealing (MacLean-Fletcher & Pollard, 1980) and pointed-end extension (Pollard & Mooseker, 1981) of F-actin all continue in vitro even when cytochalasin blocks barbed-end monomer addition.

The loops and polygons in cytochalasin-treated windows surround chloroplasts whose red autofluorescence indicates their arrival after the irradiation that created the window. Settlement of chloroplasts travelling in the streaming endoplasm was directly observed in a few cases. Specific binding molecules perhaps link the chloroplast envelope to the specialized layer of cortical cytoplasm beneath the plasma membrane (Williamson, 1985). Since chloroplasts with red autofluorescence in control windows are not surrounded by loops or polygons when processed for immunofluorescence, cytochalasin may promote their formation. However, chloroplasts travelling in the endoplasm rotate (see Kamiya, 1962), so that they may arrive in the window already carrying an actin loop to propel rotation. Cytochalasin might therefore prevent the disassembly of such loops, perhaps by its ability to block the removal as well as the addition of monomer at the barbed end of actin filaments (Pollard & Mooseker, 1981).

Bundles regenerating in cytochalasin-treated windows are not aligned like those in control windows (compare Fig. 4 with 5A). This is surprising since the postulated orienting agent (passive endoplasmic flow; Williamson étal. 1984) is still present. The composition of the bundles assembled in the presence of cytochalasin may be altered so that they are less easily deformed and can therefore resist the forces experienced when aligned at an angle to the direction of flow. Alternatively, flow may be sufficiently rapid only immediately adjacent to the upstream bundle ends so that the mechanism fails when cytochalasin blocks their extension.

In unwounded cells, the average number of actin bundles per chloroplast file remains nearly constant as a cell grows from ≈ 10 mm to 50 mm. Interruptions in the actin bundles were never seen. While cytochalasin reduced the number of bundles associated with each file of chloroplasts, neither bundle continuity nor orientation was disrupted in the way recorded in the windows. These bundles must also grow by a mechanism that lacks the obligatory dependence on barbed-end extension that makes the extension of upstream bundles into the window highly sensitive to cytochalasin. It seems more likely that the actin bundles are stretched and aligned by cell extension, presumably recruiting more actin filaments along their length to maintain their thickness approximately. (Such stretching and alignment is well documented for the wall and for the chloroplasts of the cortical cytoplasm (Taizet al. 1981; Green, 1964).) Since cytochalasin reduces the number of bundles in nongrowing cells, even the structure of these apparently stable bundles is cytochalasin-sensitive. It remains to be determined whether cytochalasin displaces the G:F equilibrium towards unpolymerized actin (Tellam & Frieden, 1982) and what causes the thickening of individual bundles.

Thus we consider that the present results strengthen our previous conclusion (Williamson et al. 1984) that actin bundle assembly in Chara is unlikely to proceed through a single mechanism. The minimum requirement seems to be for two mechanisms. First, a mechanism to extend the upstream bundles from the window edge that is highly cytochalasin-sensitive, most probably as a result of an obligate dependence on the extension of barbed filament ends. Second, one or more mechanisms operating in the window and in growing cells, that assemble thickened but functional bundles in the presence of cytochalasin. An assembly mechanism would be cytochalasin-resistant if it lacked an obligate requirement for barbed-end extension of F-actin. These results suggest that low concentrations of cytochalasin affect actin polymerization in characean cells, while earlier studies suggest that higher concentrations rapidly inhibit streaming by abolishing force generation by assembled actin bundles.

We thank Jean Perkin and Peter Jablonski for their helpful comments.

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