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Plant Physiol. (1997) 11 5: 1491-1 498 Actin Filaments of Guard Cells Are Reorganized in Response to Light and Abscisic Acid’ Soon-Ok Eun and Youngsook Lee* Department of Life Science (S.-O.E., Y.L.), School of Environmental Engineering (Y.L.), Pohang University of Science and Technology, Pohang, 790-784, Korea; and lnstitute of Molecular Biology, Academia Sinica, Nankang, Taipei, Taiwan 11 529, Republic of China (Y.L.) actin monomers and tubulin dimers, respectively, and reassemble at spatially defined sites of the cell. Traditionally, they have been known to participate in diverse processes such as mitosis, cytokinesis, cytoplasmic streaming, intraand intercellular transport, and cell shaping by providing a framework in animal cells and by directing wall deposition in plant cells. Recently, it has become evident that they also function as a signal transducer; upon the perception of extracellular stimuli the cytoskeleton is rapidly reorganized, and these structural changes in turn affect activities of other signal-mediating molecules. This has been well demonstrated in animal cells (Cantiello et al., 1991; Schwiebert et al., 1994; Carraway and Carraway, 1995; Diakonova et al., 1995; Prat et al., 1996). Actin in yeasts serves a similar function in the relay of pheromonestimulated responses (Leeuw et al., 1995). In plants actin filaments provide a matrix for components of the phosphatidylinositol cycle, a major pathway in transducing extracellular signals across the plasma membrane. For example, phosphatidylinositol 4-kinase (Xu et al., 1992), phosphoinositide-specific phospholipase C (Huang et al., 1997), and diacylglycerol kinase (Tan and Boss, 1992) are associated with the detergent-extracted cytoskeleton. The phospholipase C activity of broad bean (Viciafaba L.) leaves is inhibited by an actin-binding protein, profilin (Drabak et al., 1994). Thus, the state of actin polymerization in plant cells can be critica1 in activating and recruiting signal molecules to a site where they interact, as in other cells. We previously showed the presence of actin filaments in mature guard cells and demonstrated that application of actin antagonists perturbed stomatal behavior (Kim et al., 1995). From these observations we suggested that the dynamic feature of actin filaments may be important in guard cell signaling . Since the organization of actin filaments typically changes when they are involved in cell signaling, we investigated the possibility that the distribution of actin filaments in guard cells actually changes during normal stomatal movements. We report that actin filaments of illuminated guard cells are radially organized at the cell cortex, and that the radial arrays are rapidly disassembled when stomatal closing is induced by ABA. Likewise, the radial pattern of cortical actin filaments are abolished in guard cells of dark-closed stomata. We recently showed that treatment with actin antagonists perturbed stomatal behavior in Commelina communis 1. leaf epidermis and therefore suggested that dynamic changes in actin are necessary for signal responses in guard cells (M. Kim, P.K. Hepler, S.O. Eun, K . 4 . Ha, Y. Lee [1995]Plant Physiol 109: 1077-1084). Here we show that actin filaments of guard cells, visualized by immunofluorescence microscopy, change their distribution in response to physiological stimuli. When stomata were open under white-light illumination, actin filaments were localized in the cortex of guard cells, arranged in a pattern that radiates from the stomatal pore. In marked contrast, for guard cells of stomata closed by darkness or by abscisic acid, the actin organization was characterized by short fragments randomly oriented and diffusely labeled along the pore site. Upon abscisic acid treatment, the radial pattern of actin arrays in the illuminated guard cells began to disintegrate within a few minutes and was completely disintegrated in the majority of labeled guard cells by 60 min. Unlike actin filaments, microtubules of guard cells retained an unaltered organization under all conditions tested. These results further support the involvement of actin filaments in signal transduction pathways of guard cells. A set of guard cells surrounding stomata of terrestrial plants function much like sliding doors in a building, opening to allow the CO, uptake required for photosynthesis and closing to reduce water loss during periods of water deficit. Such regulation is initiated by sensing environmental and interna1 stimuli such as light, humidity, CO,, and the plant-stress hormone ABA, and is accomplished by osmotic volume changes of the cells. Previous studies have implicated heterotrimeric G-proteins, the H+ pump, and the movement of various ions regulated by ion channels in these processes (for review, see Assmann, 1993). Thus, guard cells provide an ideal system in which to examine whether other molecules, including cytoskeletal elements, take part in plant signaling and, if so, how they interact with better-characterized ones. Actin filaments and microtubules are dynamic cellular components; they disassemble into their building units, This research was supported by the Basic Science Research Fund of Pohang University of Science and Technology and the Science and Engineering Foundation of Korea (grant no. 61-0507056-2 awarded to Y.L.) and by a postdoctoral fellowship from Korea Science and Engineering Foundation awarded to 5-O.E. * Corresponding author; e-mail [email protected]; fax 82-562- Abbreviations: FITC, fluorescein isothiocyanate; TRITC, tetramethyl rhodamine isothiocyanate. 279-2199. 1491 Eun and Lee 1492 MATERIALS A N D METHODS Plant Material Commelina communis L. plants were grown in a greenhouse at the controlled temperature of 18 to 22°C. The light period was 13 to 16 h with the maximum light intensity of 1000 pmol m-'s-l at noon. We used the youngest fully expanded leaves of 4- to 5-week-old plants. Conditions for Opening or Closing of Stomata To compare the organization of cytoskeletal elements in guard cells of open and closed stomata, plant materials were treated under various conditions before fixation. Guard cells of open stomata were obtained under two different conditions: 2 to 3 h before the beginning of the light period from well-watered plants in water-saturated (100% RH) air and 3 h after the beginning of the light period under 300 to 400 pmol m-' s-l of white light. Guard cells of closed stomata were obtained in three different ways: 2 to 3 h before the beginning of the light period in drier air (RH <70%), in darkness 3 h after the beginning of the normal light period, and ABA treatment of leaf epidermal peels approximately 3 h after the beginning of the light period under white light of about 300 pmol m-' s-'. In each experiment the open or closed state of the stomatal aperture was confirmed microscopically before fixation. For analysis of the effect of ABA on the state of actin over time, epidermal peels were floated on a buffer containing 10 mM K+-Mes (pH 6.1), 0.5 mM EGTA, and 50 mM KC1. When stomata were wide open, ABA was added to a final concentration of 10 p ~Multiple . samples were taken at 3, 6, 9, 12, 30, and 60 min after the onset of ABA treatment. Severa1 samples from each time were examined for the stomatal size, and the rest were fixed immediately for immunolocalization. Visualization of Actin Filaments and Microtubules Actin filaments were immunolocalized as described previously (Kim et al., 1995) with the following modifications. The abaxial epidermis was peeled from mature leaves and fixed at room temperature or 37°C for 1 h with mmaleimidobenzoyl N-hydroxysuccinimide ester dissolved to 200 p~ in 50 miv phosphate buffer (pH 6.8) containing 5 mM EGTA, 0.05% (v/v) Triton X-100, 0.3 mM phenylmethylsulfonic acid, 1.5 p~ aprotinin, 2 p~ pepstatin, and 20 piv leupeptin. The epidermal peels were frozen in liquid nitrogen and then cracked into small pieces with prechilled metal blocks. They were further fractured if necessary by tapping gently with the tips of forceps. They were extracted for 2 h with 1%(v/v) Triton X-100 in the buffer used for fixation and then rinsed with PBS. The separate blocking step was omitted. The small epidermal fragments were sequentially incubated with a 1:20 dilution of monoclonal actin antibody, a 1 : l O O dilution of biotinylated goat anti-mouse IgM, and a 1:40 dilution of streptavidin-FITC. Antibodies (Amersham) were diluted with PBS containing Plant Physiol. Vol. 11 5, 1997 1%BSA and 0.05% (v/v) Triton X-100. A11 incubations were carried out at 37°C. The duration was 45 min for each of the primary and the secondary antibodies and 30 min with streptavidin-FITC. The labeled samples were mounted with the medium containing 90% glycerin, 10% PBS (pH 8.9 adjusted with NaHCO,), and 0.1% p-phenylenediamine. They were observed and recorded under an epifluorescence microscope equipped with Automatic-2 exposure system (Zeiss) using T-Max 400 film (Kodak). The sample-handling procedures for labeling microtubules of leaf epidermis were very similar to those given for localization of actin filaments except for some differences during fixation and incubation with antibodies: epidermal tissue was fixed at room temperature with 3.7% paraformaldehyde, 0.2% picric acid, 0.05% (v/v) Triton X-100, and 5 mM EGTA, prepared in 50 miv phosphate buffer (pH 6.8); anti-/3-tubulin (Amersham) and FITC-conjugated antimouse IgG (Sigma) were diluted to 1:50 and 1:200 and used as primary and secondary antibodies, respectively. For double labeling of actin filaments and microtubules in the same cells, leaf epidermis was processed as described above for the preservation of actin filaments until they were incubated with biotinylated secondary antibodies. They were further incubated with anti-p-tubulin and then with a mixture of 1:40 streptavidin-FITC and 1:80 TRITC-anti-mouse IgG (Sigma). Fluorescence from FITC and TRITC was observed using a fluorescence microscope (Optiphot-2, Nikon) equipped with filter blocks of a narrow band pass (for FITC: excitation 465-495 nm, barrier BA515-555; for TRITC: excitation 540/25 nm, barrier BA605/55). Images were recorded on T-Max 400 film using a Microflex UFX-DX photographic attachment (Nikon). Since the aim of this study was to examine cytoskeletal distributions in guard cells in relation to stomatal regulation, it would be ideal to use a fixative that maintains both stomatal size and the integrity of cytoskeletal structure. To our knowledge, a fixation procedure that can retain actin filaments in fully open or completely closed stomata has not yet been reported. In our hands, m-maleimidobenzoyl N-hydroxysuccinimide ester, a protein cross-linker that requires detergents to get into the cell, is the best fixative for preservation of actin filaments in guard cells, as in other types of plant cells. Fixing the cells in detergent-containing solutions inevitably destroys the intactness of the plasma membrane, and therefore it entails the collapse of the stomata1 aperture. However, the clear differences that are observed between labeled guard cells of open versus closed stomata suggest that the fixation procedures themselves do not alter the integrity of the cytoskeletal structure. RESULTS Actin Filaments in Cuard Cells of Open Stomata When stomata were opened by illumination, actin in guard cells was in its filamentous form and localized at the cortex in a radial pattern: the filaments were closely spaced at the center of the ventral side, the side of guard cells near the stomatal pore, and spread out toward the opposite side, the dorsal side (Fig. 1). Some of the filaments were Stimulus-Dependent Actin Reorganization in Guard Cells 1493 cortical near the dorsal side of the cell but became partially cytoplasmic near the nucleus, which was often located close to the stomatal pore side and also stained brightly with actin antibodies. Stomatal opening is typically promoted by high plant water potential and high ambient humidity (Assmann, 1993; Kearns and Assmann, 1993). Stomata of C. communis leaves under these conditions were sometimes wide open, even in the absence of light, approximately 2 to 3 h before the beginning of the light period. In this case, actin was also localized in a radial pattern indistinguishable from that observed in the guard cells swollen by illumination (data not shown). Actin Filaments in Guard Cells of Dark-Closed Stomata Figure I. Actin filaments in guard cells of stomata open under light. The original stomatal aperture was not maintained during the fixation. Fine actin filaments are localized at the cortex of the guard cells. Some microfilaments appear to be branched (arrows). The radial pattern of actin filaments are apparent in the guard cells, where they are in focus. Diffuse staining from the nucleus (n) is seen at the center of the guard cells near the stomatal pore. The microfilaments in some parts of the cell on the right side are out of focus because of the convexity of guard cells in the epidermal surface. Bar represents 10 /Jim. branched in the direction of the dorsal side. Although the continuity of the actin filaments along the paradermal surface of guard cells was evident when focusing through the cell depth, the cortical actin filaments in guard cells could rarely all be focused in a single plane. This observation indicates that the radial actin arrays are located very close to the plasma membrane, which is highly convex because of the pronounced three-dimensional shape of guard cells. In some guard cells a few actin filaments appeared to be connected to the nuclear envelope; therefore, they were When plants were kept under dark conditions with moderate humidity, stomata most often remained completely closed 2 to 3 h before and after the onset of the usual light period. There was no difference in actin labeling in the guard cells of dark-closed stomata at these two different times of the day. Furthermore, actin labeling in these cells was entirely distinct from that observed in guard cells of open stomata: fluorescence was either diffuse (Fig. 2, A and B) or shown as randomly distributed fragments (Fig. 2C). The diffuse staining was seen through the depth of the cells and more concentrated near the ventral side of the guard cells (Fig. 2, A and B). On rare occasions, long filaments running parallel to the long axis of the guard cell were labeled in the subcortical cytoplasm (Fig. 2D). ABA Effects on Actin Filaments To determine whether actin organization in the guard cells of stomata closed by ABA is distinct from that observed in the guard cells of dark-closed stomata and to examine dynamics of actin reorganization in guard cells, we performed time-scale experiments with ABA. Different Figure 2. Actin filaments in guard cells of dark-closed stomata. Various patterns of actin labeling are shown in three different sets of guard cells. Right-side cell of each pair in A to C and both cells in D are labeled. Diffuse staining shown on the ventral side of the guard cells throughout the cell depth (focal plane was at the cortex in A and the nucleus in B), and randomly oriented short cortical fragments (C) were the most common patterns. Relatively long filaments along the length of the cell (D) were occasionally observed. Bar represents 10 /j.m. 1494 Plant Physiol. Vol. 115, 1997 Eun and Lee 100 0 10 20 30 40 50 60 v Time in ABA (min) Figure 3. Left, Cortical actin patterns in permeabilized ABA-treated guard cells. A, Long radial filaments, spanning most of the cell width. B, Shorter filaments but still in the radial pattern. C, Fragments in a random orientation or diffuse labeling. Right, Time plot of stomatal size (A) and the percentage of guard cells with the radial actin filaments (A and B patterns, •) after the onset of ABA treatment at time 0. Bar in A represents 10 /xm. from samples that had been kept for hours under either stomatal opening or closing conditions, epidermal pieces taken in the process of ABA treatment showed large variations in stomatal size and actin filament patterns. Thus, we measured stomatal size and categorized cortical actin patterns into three groups: (a) long radial filaments spanning most of the cell width, (b) shorter filaments still in the radial direction, and (c) fragments in a random orientation or diffuse labeling (Fig. 3; Table I). Before ABA treatment the dominant pattern was the radial arrays; guard cells showing long or short radial filaments consisted of 90% of the total guard cells labeled. After 3 min in ABA, although the actin pattern in the majority of guard cells was still the radial one, the population of guard cells with this organized pattern was substantially reduced compared to that before ABA treatment. At the same time, the stomatal size was also reduced. During the extended period of ABA treatment, diffuse or spotty labeling lacking any radial arrays of actin, the pattern similar to that observed in the guard cells of dark-closed stomata, became more prevalent, reaching 85 and 98% after 30 and 60 min, respectively. The gradual but steady disassembly of actin filaments during ABA-induced stomatal closing was observed in all timecourse experiments we performed (n = 4). A similar trend of actin depolymerization with a faster time course was observed under the conditions of rapid stomatal closure, addition of 10 /XM ABA in 30 mM KC1 and 10 mM K+-Mes (pH 6.1), without EGTA (data not shown). Whereas actin filaments at the cortex disappeared, in the subcortical area, where not many actin filaments were localized in open guard cells, long filaments similar to those shown in Figure 2D were observed frequently in guard cells treated with ABA for 60 min. Microtubules in Guard Cells Actin filaments and microtubules in plant cells are often intimately associated in the cell cortex, and their stability is interdependent. We localized microtubules to investigate whether microtubules in guard cells are also redistributed in response to stimuli that alter actin distribution and whether changes in one cytoskeletal element influence the polymerization state of the other. Cortical microtubules were arranged in a radial pattern similar to that of actin filaments in guard cells of open stomata (Fig. 4). However, compared with actin filaments, microtubules appeared denser, with smaller angles between individual arrays. In addition, microtubule arrays from one side of a guard cell reached the other side without branching (Fig. 4). Another difference was that there was little labeling of tubulin in the subcortical cytoplasm and around the nuclear envelope. Thus, most microtubules in Table I. Time course of ABA effects on actin filaments in guard cells and stomatal aperture Pattern of Actin Filaments as Shown in Figure 3 Time in ABA A min 0 3 6 9 12 30 60 B Labeled Cell No. Stomatal Size ± SE (n = 40) C j*m % of total labeled cells 67 53 19 12 23 24 21 23 23 60 10 65 118 99 62 93 18 2 0 27 13 2 55 85 98 33 97 81 12.9 11.8 11.2 10.5 8.1 5.8 ± ± ± ± ± ± 0.3 0.3 0.2 0.3 0.4 0.3 3.2 ± 0.2 Stimulus-Dependent Actin Reorganization in Guard Cells 1495 Figure 4. Microtubules in guard cells. Microlubules are distributed in a radial pattern in the illuminated guard cells (A) and in the guard cells treated with ABA for 3 min (B) and 30 min (C and D). C and D show intact microtubules in the two peridermal regions of the same pair of guard cells. Bar represents 10 urn. guard cells appeared to be localized in the cortex of the cells. Double labeling of actin filaments and microtubules in the same guard cells confirmed these differences in the organization of the two cytoskeletal elements (Fig. 5, A-D). However, the most striking difference between microtubules and actin filaments was their response to stimuli. The circumferential organization of microtubules in illuminated guard cells (Fig. 4A) was not affected by incubation of epidermis with ABA (Fig. 4, B-D) or under darkness (Fig. 5D), whereas these conditions caused depolymerization of actin filaments (Figs. 2, A-C and 5B). DISCUSSION We previously suggested that actin is an essential component in signal transduction pathways of guard cells based on the data that the actin antagonists cytochalasin D and phalloidin, which resulted in a net decrease and increase in actin filaments in guard cells, respectively, interfered with stomatal movements (Kim et al., 1995). The results presented in this paper further support the hypothesis by clearly demonstrating fast reorganization of actin in guard cells in response to physiological stimuli. During the last decade accumulating evidence has shown that actin filaments, microtubules, and the proteins that bind to the cytoskeletal elements are functionally associated with signal transducers. In this regard, the degree and the location of cytoskeletal assembly are critical for inducing proper responses of the cell. In animal cells actin filaments are readily reorganized by chemotactic stimuli (Howard and Meyer, 1984; Condeelis, 1993), growth factors (Rijken et al., 1991; Ridley et al., 1992; Nobes et al., 1995), and extracellular matrix (Hartwig, 1992). Furthermore, when cytoskeletal reorganization experiences interference, cellular responses to the stimuli are interrupted (Ridley and Hall, 1992; Tominaga et al., 1993; Peppelenbosch et al., 1995; Takaishi et al., 1995). Actin responses to growth substances in plant cells are not well documented. However, actin filaments in plant cells do change their organization; for example, during the cell cycle (Seagull et al., 1987; Cleary et al., 1992; Zhang et al., 1993), differentiation (Cho and Wick, 1990; Cleary, 1995), interaction with fungus (Kobayashi et al., 1994), and phototropic responses (Meske and Hartmann, 1995; Mineyuki et al., 1995). Although the turnover rate of actin filaments in plant cells has not been determined, interphase microtubule dynamics in plant cells was demonstrated to be faster than that in animal cells (Hush et al., 1994). The response of actin filaments to ABA we observed in guard cells is faster than any hormonal reorganization of cytoskeleton demonstrated to date in plant cells. Another particular characteristic of actin filaments in guard cells is that the changes in the structure are reversible in response to cyclic changes of environmental conditions. These qualities make actin in guard cells suitable for a signal-mediating component, like its counterpart in animal cells. Guard cells of stomata that opened under two different conditions showed the same radial pattern of actin organization. Similarly, both darkness and ABA caused stomatal closure, resulted in the disappearance of cortical filaments, and led to almost identical diffuse/spotty patterns of actin. These results imply that structural changes in actin are not limited to a particular signal but are common to physiological signals that open or close stomata. Our temporal studies with ABA demonstrated that disintegration of cortical actin filaments in guard cells occurs in parallel with closing of stomata; both changes were apparent within 3 min after the onset of ABA treatment and progressed further along with time. More advanced techniques that allow ABA treatment of guard cells that have been loaded with labeled phalloidin or tagged actin may clarify whether actin reorganization precedes stomatal closing. Therefore, does actin depolymerization always favor closing of stomata? Our previous data do not support this idea, since cytochalasin D, which abolished the radial actin filaments in guard cells, enhanced stomatal opening. Fusicoccin, another fungal toxin that enhances stomatal opening, also caused actin depolymerization under light (S.-O. Eun and Y. Lee, unpublished data). Moreover, both opening and closing movements are inhibited by phalloidin, which increases the number of the filamentous form of actin filaments in guard cells (Kim et al., 1995). Therefore, 1496 Eun and Lee Figure 5. Organization of actin and microtubules in guard cells. Double labeling of actin (A and B) and microtubules (C and D) in guard cells of open (A and C) and closed (B and D) stomata under light and darkness, respectively. E and F, Schematic drawing of cytoskeletal organization in guard cells (actin in E and microtubules in F). Bar represents 10 /xm. we propose that depolymerization of actin allows changes in stomatal aperture but does not determine the direction of change. For better understanding of roles of actin cytoskeleton in guard cells, it is necessary to identify molecules that are Plant Physiol. Vol. 115, 1997 affected by dynamic changes in actin structure. The physical state of actin in animal cells affects activities of lipidhydrolyzing enzymes, receptor- and nonreceptor-protein kinases, G-proteins, and their modulators (Carraway and Carraway, 1995), as well as ion channels (Cantiello et al., 1991; Schwiebert et al., 1994; Prat et al., 1996). Our patchclamping data showed that activities of K + channels in guard cells are certainly influenced by application of actin antagonists (Hwang et al., 1997). We are currently investigating other possible target molecules of actin in guard cells. Equally important is elucidating the signaling pathways that reside upstream of actin in guard cells. Actin polymerization is modulated by numerous actin-binding proteins. However, few of them have been characterized in guard cells to date. Thus, an understanding of the regulation of polymerization and depolymerization of actin in guard cells awaits more detailed information concerning characteristics of these actin-binding proteins. Intracellular Ca 2+ levels are also known to affect actin filaments, although these effects may be indirect by activating a group of actin-severing proteins such as gelsolin, as has been suggested in mammalian cells. It is interesting that ABA provokes both a cytosolic [Ca 2+ ] increase (Irving et al., 1992; McAinsh et al., 1992) and actin depolymerization (Table I) within several minutes in guard cells of C. communis. It would be informative to understand how these two are related to each other in guard cell signaling. In a different manner, a subfamily of small GTP-binding protein, Rho, functions as a molecular switch in the regulation of actin in animal and fungal systems. A full-length Rho cDNA isolated from pea seedlings (Yang and Watson, 1993) and partially characterized genes in several different plants (Lee and Lee, 1996) suggest its universal presence in higher plants. We have preliminary results implicating the existence of Rho proteins in C. communis guard cells and its participation in stomatal movements. These investigations will likely unravel important new aspects of signal transduction in plant cells. It has been shown that actin filaments and microtubules in plant cells are often closely aligned, and alteration in one results in reorganization of the other. Disruption of actin filaments with cytochalasin D interfered with reorganization and/or stability of microtubules in onion mitotic cells (Eleftheriou and Palevitz, 1992) and during the development of cotton fibers (Seagull, 1990) and wheat mesophyll cells (Wernicke and Jung, 1992). However, we do not consider the role of actin filaments in guard cells in such a connection because the changing state of actin filaments did not affect the well-organized microtubules in guard cells. In fact, the radial pattern of microtubules remained intact in guard cells regardless of the size of their stomata. Whether microtubules change their distributions and play a role in stomatal movements is equivocal. Our studies showed no changes in microtubules in guard cells of C. communis in response to darkness or ABA, and treatment of C. communis leaf epidermis with the microtubule antagonists taxol or oryzalin did not affect stomatal aperture (data not shown), suggesting that microtubules are not involved in stomatal movements. In broad bean guard cells as well, pharmacological disruption of microtubules did not inter- Stimulus-Dependent Actin Reorganization in Guard Cells fere w i t h stomatal behavior (Jiang e t al., 1996). However, stomatal opening in Tradescantia virginiana was inhibited by colchicine, a microtubule-depolymerizing alkaloid (Couot-Gastelier and Louguet, 1992), and microtubules in broad bean g u a r d cells of open stomata became fragmented when stomata were closed by ABA (Jiang e t al., 1996). Further investigations a r e necessary t o clearly understand whether t h e differences between these reports and w h a t we observed a r e due to differences i n t h e plant materials or in t h e techniques used. In conclusion, actin cytoskeleton of guard cells undergoes changes i n their organization i n response t o changes i n physiologically important stimuli. These results are consistent w i t h our earlier pharmacological d a t a and provide a foundation t o claim actin as a component i n g u a r d cell signal transduction pathways. ACKNOWLEDCMENTS We thank Drs. Virginia S. Berg and Richard C. Crain for their helpful comments concerning the manuscript, Mr. Shi-In Kim for the management of plants, and Ms. Jae-Ung Hwang for assistance with the time-course experiments and many helpful discussions. Received May 19, 1997; accepted September 7, 1997. Copyright Clearance Center: 0032-0889/97/ 115/1491/OS. LITERATURE CITED Assmann SM (1993) Signal transduction in guard cells. 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