Planta 9 Springer-Verlag1988

Planta (1988)173:442 446

Microfilament bundles of F-actin in Spirogyra observed by fluorescence microscopy Yasue Goto and Katsumi Ueda* Biological Laboratory, Nara Women's University, Nara 630, Japan

Abstract. Microfilament bundles (MFBs) of F-actin were observed by fluorescence microscopy in cells of Spirogyra treated with rhodamine-phalloidin. Four types of M F B s could be recognized on the basis of locality and appearance: those dispersed in the cytoplasm near the cell surface; those beneath the plasma membrane running parallel to each other; those at the edges of the chloroplast; and those surrounding the nucleus. Each type exhibited a unique behavior during the cell cycle. Microfilament bundles dispersed in the cytoplasm came together at the middle of the cell to form a fibril ring at the mitotic prophase. The fibril ring decreased in diameter, causing the development of a furrow in the protoplast that progressed from the outside to the inside. After the completion of furrowing, the MFBs in the fibril ring dispersed beneath the plasma membrane. Microfilament bundles surrounding the nucleus formed a net-like cage which became invisible at the mitotic anaphase, while MFBs seen at the chloroplast edges persisted there during the cell cycle without changing their position. Parallel MFBs running perpendicular to the long axis of the cell were seen at all stages in the cell cycle. Key words: Actin filaments - Cell cycle Cell division - Chlorophyta Microfilament bundles Spirogyra

Introduction Microfilaments (MFs) in plant cells have been less studied than those in animal cells, mainly because of the difficulty in preservation during cell preparation for microscopy. Microfilaments were first observed by electron microscopy in Nitella cells and * To whom correspondence should be addressed Abbreviations." MF = microfilament; MFB microfilament bundle; MT = microtubule

assumed to provide the driving force for cytoplasmic streaming (Nagai and Rebhun 1966). Since then, several papers have appeared on the ultrastructure of MFs in plants in relation to cytoplasmic streaming (Pickett-Heaps 1967; Franke et al. 1972; Palevitz and Hepler 1975; Nagai and H a y a m a 1979; Dazy et al. 1981; Yamaguchi and Nagai 1981). That these M F s are composed of Factin has been well demonstrated by experiments using heavy meromyosin (Dazy et al. 1981 ; K o o p 1981) and cytochalasin B (Williamson 1972; Bradley 1973). More recently, specific binding between phalloidin and F-actin has been demonstrated (Barak et al. 1980), and fluorescence-labeled phalloidin has been used for detecting the microfilament bundles (MFBs) of F-actin in plant cells (Barak et al. 1980; Pesacreta etal. 1982; Gunning and Wick 1985). This method has demonstrated the occurrence of MFBs of F-actin in several kinds of cells, including conifer root cells (Pesacreta et al. 1982), angiosperm root-hairs (Lloyd and Wells 1985), Cobaea seed hairs (Quader et al. 1986), meristematic cells of Alliurn (Clayton and Lloyd 1985), epidermal cells of barley and oats (Parthasarathy 1985), and pollen-tube cells (Perdue and Parthasarathy 1985). In the present investigation, rhodamine-phalloidin was used to observe the behavior of F-actin MFBs in Spirogyra cells at various stages in the cell cycle. The functions they may perform during the cell cycle are discussed. Material and methods The Spirogyra used in the present investigation was isolated from a pond in Nara by one of the authors (K.U.) and cultured in the medium of Waris (1950) supplemented by 5 ml soil extract in 100 ml culture medium. The species name could not be identified because the alga formed zygospores neither in the culture medium nor in the natural habitat. Microfilament bundles of F-actin were detected by fluorescence microscopy (Olympus, Tokyo, Japan; type BH2 RFA,

Y. Goto and K. Ueda: Microfilament bundles of F-actin in Spirogyra with a green exciter filter) in cells mounted in a solution prepared by dissolving 5"10 7 M rhodamine-phalloidin (Molecular Probes, Junction City, Ore., USA) in I ml of 50 mM 1,4-piperazineethanesulfonic acid (Pipes) buffer (pH 7.3) (Wako Chemicals, Osaka, Japan) containing 2.5 mM ethylene glycol bis(fl-aminoethyl ether)-N,N,N',NMetraacetic acid (EGTA) (Wako Chemicals), 0.5 mM MgCI2, 0.05 mM phenylmethylsulphonyl fluoride (Wako Chemicals), 0.5% Triton X-100 (Wako Chemicals), 0.0025% leupeptin (Peptide Institute, Minoh City, Japan), and 0.05% n-propyl gallate. Some cells were treated with a nonfluorescent phalloidin solution for 20 min before the rhodamine-phalloidin treatment. The nonfluorescent phalloidin solution was prepared by replacing 5.10-~M rhodamine-phalloidin in the staining solution with 5 mM phalloidin (Sigma Chemical Co., St. Louis, Mo., USA). For detection of microtubules by fluorescence microscopy, cells were fixed with 3.7% paraformaldehyde (Wako Chemicals) in 100 mM Pipes buffer (pH 6.9) containing 3 mM EGTA. After washing with Pipes buffer for 1 min, the cells were airdried on a cover slide and were cut with a razor blade at their terminal regions. They were immersed for 1 h in Pipes buffer containing 5 mM EGTA, 5 mM MgC12, 1% Triton X-100, 0.4M mannitol, and 0.0025% leupeptin. These cells were treated with a mouse monoclonal antitubulin (Amersham International, Amersham, Bucks., UK), followed by fluorescein isothiocyanate (FITC)-labeled antimouse immunoglobulin G from sheep (Amersham International). A blue exciter filter was used for observation of FITC-fluorescence.

Results

Cells of Spirogyra are cylindrical, each containing a single nucleus in its center and in the species used in our investigations two or three chloroplast bands coiled in spirals directly beneath the plasma membrane of the cell (Fig. 1). Fluorescent filaments can be seen in the cytoplasm throughout the cell cycle when cells are treated by rhodamine-phalloidin. These filaments become invisible by a pretreatment with nonfluorescent phalloidin which specifically binds to F-actin. Accordingly, fluorescent filaments combined with rhodamine-phalloidin are assumed to be cytoskeletal MFBs consisting of F-actin. The MFBs are found in the cells in four different locations. Firstly, MFBs are dispersed in the cytoplasm near the surface of the cylindrical cells, most of them running nearly parallel with the long axis of the cell (Fig. 2). Figure 3 is a light photomicrograph of the same cell area as Fig. 2. These MFBs vary in thickness, possibly because of differences in the number of F-actin molecules bundled together to make a visible filament. Secondly, MFBs are seen near the spiral chloroplasts. Some are situated longitudinally along the edges of the chloroplast bands (Figs. 4, 5, arrows), while others project laterally from the edges of the chloroplast bands and connect to the edges of neighboring chloroplasts (Fig. 4). A third type of MFBs tightly

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surround the interphase nucleus (Figs. 6, 7) forming a finely interwoven net-like cage as apparent in the surface view (Fig. 7). The thickness of the MFBs at this location is less than that at other sites. Finally, the fourth type of MFBs is detectable beneath the cell surface. These MFBs are thin and run parallel to each other and are perpendicular to the long axis of the cell (Fig. 8). At a similar position in the cell, abundant microtubules (MTs) are also present, running in the same direction (Fig. 20). Microtubules with a distribution pattern similar to that of the MFBs near the nuclei or near the chloroplast bands were not observed. In cells in early mitotic prophase, some of the dispersed MFBs come together at the middle of cylindrical cells to form a fibril ring situated just beneath the plasma membrane (Fig. 9). Many wavy MFBs are connected to the fibril ring, implying that they may eventually be incorporated into it. Figure 10 shows the fibril ring in a mid-prophase cell. The MFBs are packed more tightly than those at the previous stages. The net-like cage of MFBs surrounding the nucleus is most conspicuous at this stage. In cells in the mitotic metaphase, the fibril rings are similar in appearance to those in mid-prophase cells. The net-like cage of MFBs does not cover the entire metaphase spindle, but only its equator, leaving large uncovered areas at both spindle poles (Fig. 11). The MFBs situated at the chloroplast edges can be seen in all cells during mitosis, but no MFBs are detected in metaphase or anaphase spindles. Net-like cages of MFBs surrounding individual chromosome groups cannot be recognized from early anaphase to mid-telophase. The same cell area with a fibril ring and two early telophase nuclei is shown from Fig. 12 to Fig. 14. A faint trace of protoplast-furrowing is visible surrounding the cell (Fig. 13, arrows). The fluorescent fibril ring is situated just on the furrow (Fig. 12). However, fluorescence is not detectable at the positions of the nuclei (compare Figs. 12 and 14). The netlike cage of MFBs can be observed around nuclei at mid-telophase (Figs. 15, 16). The furrowing of the protoplast has advanced slightly towards the cell center in the cell of Fig. 15 where the fibril ring is in tight contact with the furrow. The diameter of the fibril ring continuously decreases with the development of the furrow, until the cell is finally divided into two daughter cells (Figs. 1719). After the completion of cell division, the MFBs in the ring become dispersed in the cytoplasm over the entire surface areas of the new cells. The MFBs loosened and partially dispersed from the fibril ring are shown in Fig. 19.

Fig. 1. A Spirogyra cell with three chloroplast bands and a nucleus (N). x 250. Fig. 2. Microfilament bundles in the cytoplasm near the cell surface, x 600. Fig. 3. The same cell area as in Fig. 2 observed with a light microscope, x 600. Fig. 4. Microfilament bundles between the chloroplast bands, x 600. Fig. 5. Microfilament bundles along the edges of chloroplast bands (arrows). x 600. Fig. 6. Microfilament bundles tightly surrounding the nucleus, which appear as a bright circle in the optical cross section. x 900. Fig. 7. Network of MFBs surrounding the nucleus from the surface view. x 900. Fig. 8. Microfilament bundles beneath the plasma membrane running perpendicular to the long cell axis. • 1200. Fig. 9. Microfilament bundles in the early prophase cell, which have come together at the middle of the cell. x 900. Fig. 10. Fibril ring in the mid-prophase cell. x 900. Fig. 11. Net-like cage of MFBs at the equator of the metaphase spindle, x 900. Fig. 12. Fibril ring at the early telophase cell. x 600. Fig. 13. The same cell area as in Fig. 12 observed with a light microscope. The trace of the furrowing can be recognized (arrows). x 600. Fig. 14. The same cell area as in Fig. 13 focused at the nuclei (N). x 600. Bars represent I0 gm

Y. Goto and K. Ueda : Microfilament bundles of F-actin in Spirogyra

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Fig. 15. Fibril ring in the mid-telophase cell. The arrow points to the net-like cage around a new daughter nucleus, x 600. Fig. 16. The same cell area as in Fig. 15 observed with a light microscope. N, Nucleus. x 600. Fig. 17. Fibril ring with the diameter about half the cell diameter, x 600. Fig. 18. The same cell area as in Fig. 17. N, Nucleus. x 600. Fig. 19. Fibril ring with the minimized diameter. • 900. Fig. 20. Microtubules running parallel to each other beneath the plasma membrane, x 1300. Bars represent 10 gm

Discussion

Four distinct types of MFBs could be recognized on the basis of location and appearance in Spirogyra; those dispersed in the cytoplasm near the cell surface, those beneath the plasma membrane running parallel to each other, those at the chloroplast edges, and those surrounding the nucleus. Each type exhibits its own typical behavior during the cell cycle. The MFBs in the cytoplasm near the cell surface show conspicuous dislocation, ring formation, and dispersion during the cell cycle. The time when dispersed MFBs come together at the middle of the cylindrical cell to form a fibril ring coincides with the start of protoplast furrowing. Furrow development is always accompanied by a decrease in ring diameter. After the completion of protoplast furrowing, the MFBs disperse again. These findings indicate that the MFBs may be closely related to cytokinesis by furrowing. A configura-

tional change in F-actin probably takes place in the fibril ring, resulting in the diminish of the diameter of the ring and causing the development of protoplast furrowing from the outside towards the inside. In higher plants, cytokinesis starts from the cell center toward the cell periphery in the phragmoplast. Microfilaments in the phragmoplast which run perpendicular to the cell plate have been reported in the root tip of Alliurn (Clayton and Lloyd 1985) and in stamen hair cells of Tradescantia (Gunning and Wick 1985). In these cases, MFs are not directly involved in cell-plate formation, although indirect participation of MFs in cytokinesis such as the acceleration of transport of material needed for cytokinesis has been proposed (Clayton and Lloyd 1985). Close association between MFs and MTs has been reported in fungal cells (Hoch and Staples 1983, 1985). In Spirogyra, MFBs running parallel to each other beneath the cell surface may combine

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Y. Goto and K. Ueda: Microfilament bundles of F-actin in Spirogyra

to form a rigid cytoskeleton with MTs running parallel to the same direction. It seems likely, however, that such an association of MFs with MTs is limited to the MFs running parallel to each other. Other MFBs, distributed beneath the plasma membrane and at the chloroplast edges, are not associated with MTs detectable by immunofluorescence microscopy. Similarly, the MTs in the metaphase and anaphase spindles in Spirogyra also have no associated MFs as evident by the absence of rhodamine fluorescence. In animal cells, MFs in metaphase spindles have been reported to be absent, on the basis of immunofluorescence (Aubin et al. 1979). Microfilament bundles at the chloroplast edges persist in Spirogyra cells without recognizable morphological changes during the cell cycle. Chloroplasts in Spirogyra are bands running spirally beneath the plasma membrane of the cylindrical cells. They are not flat but concave, with the edges close to the plasma membrane and the centers separated away from it. The MFBs seen at the chloroplast edges seem to be located within the very narrow spaces between the plasma membranes and the chloroplasts, and serve probably to connect chloroplasts with the plasma membrane and attach them to it. Microfilament bundles connecting neighboring chloroplast bands may act as a cytoskeleton to maintain a constant distance between such adjacent chloroplast bands. That both types of chloroplast-associated MFBs continue to be present in cells during mitosis is quite in accord with the fact that the position of the spiral chloroplast bands remains unchanged in the cell during mitosis. A network or a net-like cage of MFBs surrounding the nucleus is observed at most stages in the cell cycle, except at the mitotic anaphase. The net-like cage of MFBs may have the function of maintaining the position of the nuclei in cells of Spirogyra. This possibility is supported by observations on nuclear positioning in Funaria (Schmiedel and Schnepf 1980) and in Lepidium sativum (Hensel 1985). Nuclei in both these plants have been noted to be moved to abnormal positions upon treatment of the tissue with cytochalasin B. References Aubin, J.E., Weber, K., Osborn, M. (1979) Analysis of actin and microfilament-associated proteins in the mitotic spindle and cleavage furrow of PtK2 cells by immunofluorescence microscopy. Exp. Cell Res. 124, 93-109 Barak, L.S., Yocum, R.R., Nothnagel, E.A., Webb, W.W. (1980) Fluorescence staining of the actin cytoskeleton in

living cells with 7-nitrobenz-2-oxa-l,3-diazole-phallacidin. Proc. Natl. Acad. Sci. USA 77, 980-984 Bradley, M.O. (1973) Microfilaments and cytoplasmic streaming: Inhibition of streaming by cytochalasin. J. Cell Sci. 12, 327-334 Clayton, L., Lloyd, C.W. (1985) Actin organization during the cell cycle in meristematic plant cells. Actin is present in the cytokinetic phragmoplast. Exp. Cell Res. 156, 231 238 Dazy, A.C., Hoursiangou-Neubrun, D., Sauron, M.E. (1981) Evidence for actin in the marine alga Acetabularia mediterranea. Biol. Cell 41,235-238 Franke, W.W., Herth, W., van der Woude, W.J., Morre, D.J. (1972) Tubular and filamentous structures in pollen tubes: Possible involvement as guide elements in protoplasmic streaming and vectorial migration of secretory vesicles. Planta 105, 317-341 Gunning, B.E.S., Wick, S.M. (1985) Preprophase bands, phragmoplasts, and spatial control of cytokinesis. J. Cell Sci., Suppl. No. 2, 157-179 Hensel, W. (1985) Cytochalasin B affects the structural polarity of statocytes from cress roots (Lepidium sativum L.). Protoplasma 129, 178 187 Hoch, H.C., Staples, R.C. (1983) Visualization of actin in situ by rhodamine-conjugated phalloin in the fungus Uromyces phaseoli. Eur. J. Cell Biol. 32, 5~58 Hoch, H.C., Staples, R.C. (1985) The microtubule cytoskeleton in hyphae of Uromyces phaseoli germlings : Its relationship to the region of nucleation and to the F-actin cytoskeleton. Protoplasma 124, 11~122 Koop, H.U. (1981) Protoplasmic streaming in Acetabularia. Protoplasma 109, 143-157 Lloyd, C.W., Wells, B. (1985) Microtubules are at the tips of root hairs and form helical patterns corresponding to inner wall fibrils. J. Cell Sci. 75, 225-238 Nagai, R., Rebhun, L.I. (1966) Cytoplasmic microfilaments in streaming Nitella cells. J. Ultrastruct. Res. 14, 571-589 Nagai, R., Hayama, T. (1979) Ultrastructure of the endoplasmic factor responsible for cytoplasmic streaming in Chara internodal cells. J. Cell Sci. 36, 121-136 Palevitz, B.A., Hepler, P.K. (1975) Identification of actin in situ at the ectoplasm-endoplasm interface of Nitella. J. Cell Biol. 65, 29-38 Parthasarathy, M.V. (1985) F-actin architecture in coleoptile epidermal cells. Eur. J. Cell Biol. 39, 1-12 Perdue, T.D., Parthasarathy, M.V. (1985) In situ localization of F-actin in pollen tubes. Eur. J. Cell Biol. 39, 13-20 Pesacreta, T.C., Carley, W.W., Webb, W.W., Parthasarathy, M.V. (1982) F-actin in conifer roots. Proc. Natl. Acad. Sci. USA 79, 2898-2901 Pickett-Heaps, J.D. (1967) Ultrastructure and differentiation in Chara sp. I. Vegetative cells. Aust. J. Biol. Sci. 20, 539 551 Quader, H., Deichgraber, G., Schnepf, E. (1986) The cytoskeleton of Cobaea seed hairs : Patterning during cell-wall differentiation. Planta 168, 1-10 Schmiedei, G., Schnepf, E. (1980) Polarity and growth of caulonema tip cells of the moss Funaria hygrometrica. Planta 147, 405-413 Waris, H. (1950) Cytophysiological study on Micrasterias. I. Nuclear and cell division. Physiol. Plant. 3, 1-16 Williamson, R.E. (1972) A light-microscope study of the action of cytochalasin B on the cells and isolated cytoplasm of the Characeae. J. Cell Sci. 10, 811 819 Yamaguchi, Y., Nagai, R. (1981) Motile apparatus in Vallisneria leaf cells. I. Organization of microfilaments. J. Cell Sci. 48, 193-205 Received 26 June; accepted 5 October 1987