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Journal of Experimental Botany, Vol. 54, No. 380, Plant Reproductive Biology Special Issue, pp. 103±113, January 2003 DOI: 10.1093/jxb/erg003 The actin cytoskeleton is a target of the self-incompatibility response in Papaver rhoeas C. J. Staiger1 and V. E. Franklin-Tong2,3 1 Department of Biological Sciences, Purdue University, West Lafayette, IN 47907-1392, USA 2 School of Biosciences, University of Birmingham, Edgbaston B15 2TT, UK Received 26 April 2002; Accepted 11 July 2002 Abstract The integration of signals received by a cell, and their transduction to targets, is essential for all cellular responses. The cytoskeleton has been identi®ed as a major target of signalling cascades in both animal and plant cells. Self-incompatibility (SI) in Papaver rhoeas involves an allele-speci®c recognition between stigmatic S-proteins and pollen, resulting in the inhibition of incompatible pollen. This highly speci®c response triggers a Ca2+-dependent signalling cascade in incompatible pollen when a stigmatic S-protein interacts with it. It has been demonstrated recently that SI induces dramatic alterations in the organization of the pollen actin cytoskeleton. This implicates the actin cytoskeleton as a key target for the SI-stimulated signals. The cytological alterations to the actin cytoskeleton that are triggered in response to SI are described here and there seem to be several stages that are distinguishable temporally. Evidence was obtained that Factin depolymerization is also stimulated. The current understanding that the actin cytoskeleton is a target for the signals triggered by the SI response is discussed. It is suggested that these F-actin alterations may be Ca2+-mediated and that this could be a mechanism whereby SI-induced tip growth inhibition is achieved. The potential for actin-binding proteins to act as key mediators of this response is discussed and the mechanisms that may be responsible for effecting these changes are described. In particular, the parallels between sustained actin rearrangements during SI and in apoptosis of animal cells are considered. 3 Key words: Actin binding proteins, actin cytoskeleton, Ca2+ signalling, pollen, self-incompatibility, signal transduction. Introduction The cytoskeleton is a major target of signalling events in plant cells. Both the microtubule and the actin ®lament cytoskeleton in plants are known to rearrange when numerous, diverse external stimuli are applied (Nick, 1999; Staiger, 2000). A major focus of plant cytoskeleton research is the identi®cation of signals that regulate actin dynamics (Staiger et al., 2000). In recent years there have been several studies in plant systems that describe actin rearrangements in response to speci®c, de®ned physiological stimuli. These include responses of stomatal guard cells to abscisic acid or light; root hairs responding to Nod factors from Rhizobium bacteria; and pollen tubes responding to self-incompatibility proteins (Eun and Lee, 1997; CaÂrdenas et al., 1998; Miller et al., 1999; Geitmann et al., 2000). Although many of the components responsible for regulating actin dynamics are well-de®ned in animal cells and yeast, considerably less is known for plants. Several actin-binding proteins (ABPs) have been identi®ed in plants, but their role in vivo is not yet well-de®ned. However, only a few ABPs which could regulate ®lament dynamics in pollen tubes have been identi®ed and characterized, namely, pro®lin, ADF/co®lin and 135ABP (Yokota and Shimmen, 2000; Hepler et al., 2001; McCurdy et al., 2001). Furthermore, although the ABPs have been implicated in mediating certain cytological changes, their involvement in speci®c physiologically relevant situations has not been demonstrated. No study on To whom correspondence should be addressed. Fax: +44 (0)121 414 5925. E-mail: [email protected] ã Society for Experimental Biology 2003 104 Staiger and Franklin-Tong plants has attempted to characterize the effect that signalling stimuli have on actin dynamics or, more speci®cally, the state of actin polymerization. Tip growing cells provide a good system whereby the signals responsible for mediating changes to the actin cytoskeleton may be studied. In plants, pollen tubes represent a well-characterized cell where tip growth has been studied in detail. Tip growth involves targeted vesicle secretion and several signalling components have been identi®ed as being associated with, or necessary for, pollen tube growth. These include a tip-based cytosolic free Ca2+ ([Ca2+]i) gradient, Ca2+-dependent protein kinase, Rho-like small GTP-binding proteins, and cAMP (Estruch et al., 1994; Pierson et al., 1996; Messerli and Robinson, 1997; Zheng and Yang, 2000; Moutinho et al., 2001). Although it is thought that the control of pollen tube growth involves a complex interplay between the cytoskeleton and signalling cascades, there is currently little direct evidence for this. It is well established that an intact actin cytoskeleton is essential for pollen germination and tip growth. Furthermore, it has been demonstrated that both actin organization and ®lamentous (F-)actin levels are involved/ important for pollen tube tip growth (Gibbon et al., 1999; Geitmann and Emons, 2000; Fu et al., 2001; Vidali et al., 2001). Despite this wealth of knowledge relating to these individual components, direct causal links between these components have not yet been established. The self-incompatibility (SI) response in the ®eld poppy, Papaver rhoeas L., provides a good example of a speci®c and genetically controlled system that employs a signal-mediated inhibition of tip growth. SI involves highly speci®c `self' recognition of pollen to prevent self-fertilization and consequent inbreeding. In this species, SI is determined by a single, multi-allelic S gene that is gametophytically controlled (Lawrence et al., 1978). S allele-speci®c self-incompatibility (S) proteins are secreted by the stigma and interact with pollen, resulting in the rapid inhibition of growth of `self' or incompatible, but not compatible pollen. It is proposed that the stigmatic Sproteins act as signal molecules that interact with a pollen S-speci®c receptor that has yet to be identi®ed. The identi®cation and cloning of the stigmatic S gene (Foote et al., 1994; Walker et al., 1996; Kurup et al., 1998) and the availability of an in vitro system for SI (Franklin-Tong et al., 1988) has allowed the dissection of the signalling cascade triggered by the SI response. Possibly the ®rst events are S-speci®c increases in [Ca2+]i (Franklin-Tong et al., 1993, 1997), which involves in¯ux (Franklin-Tong et al., 2002; Rudd and Franklin-Tong, 2003). This is thought to trigger the Ca2+-dependent phosphorylation of a pollen protein, p26 (Rudd et al., 1996) and other signalling events, including a putative programmed cell death signalling cascade (Rudd and Franklin-Tong, 2003). More recently, it has been demonstrated that the SI response induces rapid alterations in actin cytoarchitecture in incompatible pollen tubes (Geitmann et al., 2000; Snowman et al., 2000b). Recent advances in the understanding of the actin cytoskeleton as a target for the SI signalling cascade(s) are reviewed here, together with a discussion of potential ABPs that may be involved in alterations induced by SI. Actin in normally growing pollen tubes The F-actin cytoskeleton has been visualized in ®xed pollen tubes using rhodamine-phalloidin and either ¯uorescence or confocal microscopy (see Geitmann et al., 2000, for full details of methodology). m-maleimidobenzoyl N-hydroxysuccinimide (MBS) ester has been used, which gives good quality actin con®gurations which are not dissimilar in quality to those obtained using rapid freeze ®xation (Doris and Steer, 1996; Miller et al., 1999). Figure 1a shows an image of a typical poppy pollen tube grown in vitro and subsequently ®xed and visualized using this method. Typically, there are three main `zones' of Factin in the pollen tube: long, even arrays of longitudinal actin ®lament bundles in the shank of the tube; a subapical meshwork of F-actin in a `basket-like' con®guration and, at the tip, an actin-depleted zone. This is very similar to the actin con®guration which has been described for both ®xed and living pollen tubes using both phalloidin and a GFPmouse::talin fusion protein (Miller et al., 1996; Kost et al., 1998; Fu et al., 2001). SI induces S-speci®c alterations to the F-actin cytoskeleton Several dramatic alterations to the F-actin cytoskeleton are triggered by the induction of the SI response, which was achieved by the addition of S-proteins to incompatible pollen growing in vitro (Geitmann et al., 2000; Snowman et al., 2000a, b). These are illustrated in Fig. 1. The ®rst detectable changes were extremely rapid, being detected at between 30 s and 2 min. Figure 1b shows that the distinctive subapical `basket-like' con®guration had lost its organization and actin bundles appeared to be displaced into the apical region. This very rapidly gave the appearance of a big `blob' of F-actin in the apex of the pollen tube, as illustrated in Fig. 1c. This tip-localized actin is a well-recognized feature of tip growing cells that have stopped elongating (Lancelle et al., 1997; de Ruijter et al., 1999; Miller et al., 1999). The appearance of this marker corresponds to the time-period when growth is arrested (A Geitmann, AMC Emons, VE Franklin-Tong, unpublished data). Very early in the SI response, pronounced phalloidin labelling of F-actin adjacent to the plasma membrane was detected (Fig. 1b, c). Optical sectioning, using confocal microscopy, revealed this more clearly, as shown in Fig. 1d. Whether this represents an increase in cortical Self-incompatibility response in poppy 105 Fig. 1. Alterations in the actin cytoskeleton induced by the SI response. All images show rhodamine-phalloidin staining of F-actin using wide®eld ¯uorescence microscopy, except (d), which shows a confocal optical section. The scale bar in (i) is 10 mm. (a) Untreated pollen tube showing normal F-actin con®guration. (b) 2 min SI showing actin at tip and evidence of F-actin at the plasma membrane. (c) 5 min SI showing a large `blob' of F-actin at the tip and further evidence of cortical F-actin. (d) 2 min SI, single optical section obtained using confocal microscopy shows both the F-actin at the plasma membrane and loss of F-actin in the cytoplasm. (e) 5 min SI shows loss of actin ®laments in the tip region, with a speckled F-actin appearance. (f) 10 min SI shows further breakdown of F-actin bundles, and punctate actin structures in the cytosol and at the cortex. (g) 60 min SI shows larger punctate actin structures. (h) Compatible SI reactions at 20 min and (i) 60 min show no alterations in the Factin organization. Since identical biologically active S-proteins were used in incompatible reactions, this demonstrates S-speci®city. actin (i.e. movement of actin to this region or new polymerization), or is simply remaining F-actin that is more apparent due to the loss of signal in the lumen of the pollen tube is not known. However, the impression is that there is an increase in actin in this cortical region. This needs further investigation. It is of interest that a predominance of cortical actin localization has also been observed in animal cells undergoing apoptosis (Palladini et al., 1996; GueÂnal et al., 1997). There is preliminary evidence that programmed cell death (PCD) is likely to be induced in the SI response (Jordan et al., 2000; Rudd and FranklinTong, 2003). However, this potential link is, at present, speculative, and needs to be explored further. Concomitant with this, the overall intensity of phalloidin staining was reduced substantially within 2±5 min after challenge (see Fig. 1d), indicating loss of F-actin. By 5±10 min after SI challenge, the distinctive longitudinal F-actin bundles were no longer the prominent feature in the pollen tube shank, and very few actin ®lament bundles were visible in the cytoplasm, particularly in the region within ~50 mm of the tip. The phalloidin labelling of F-actin gave a ®ne, speckled appearance, as shown in Fig. 1e suggesting that severing or depolymerization might have occurred. These ®ne fragments of actin were detected throughout the cytoplasm, but a distinct distribution was visualized at the cortex. Further changes were detected at later time points, and between 10±20 min the F-actin appear to be organized into larger aggregates, as shown in Fig. 1f. These larger, punctate foci of actin were visualized throughout the pollen tube cytoplasm, although they also displayed a prominent cortical localization. They appeared to increase in size over time and became more discernible as distinct structures, as illustrated in Fig. 1g. Nucleation is not implied by the use of the word foci, but is considered to be the best word to describe these aggregates of actin. Studies indicate that these Factin structures persist for at least 3 h. Cortical actin patches are a typical feature in yeast and, in recent years, the ABPs and protein kinases involved in the formation and dynamics of these structures have been studied intensely (Pruyne and Bretscher, 2000; Pelham and Chang, 2001). However, there are very few reports of actin patch-like structures in other eukaryotic cells. It is not known whether the punctate foci of F-actin are structurally or functionally equivalent to these actin patches. With the exception of the large focal adhesions that occur in pollen tubes growing in vivo (Pierson et al., 1986; Lord, 1994), actin aggregates have not been reported previously in pollen tubes. Furthermore, the few reports of punctate actin structures in higher plants have a contentious status as they are not readily reproducible. For example, they were visualized in bean root hairs undergoing nodulation factor 106 Staiger and Franklin-Tong Fig. 2. Quanti®cation of F-actin changes induced by SI. The histogram shows the percentages of pollen tubes displaying normal distribution of F-actin ®laments (class I), membrane-localized F-actin ®laments (class II) or the membrane-localized fragmented F-actin `punctate foci' (class III) following an incompatible challenge. `Others' indicate morphologies that do not ®t these classes. [ã Kluwer Academic Publishers. From: Snowman et al. (2000). Actin rearrangements in pollen tubes are stimulated by the selfincompatibility (SI) response in Papaver rhoeas L. In: Staiger CJ, Baluska F, Volkmann D, Barlow P, eds. Actin: a dynamic framework for multiple plant cell functions, Dordrecht, The Netherlands: Kluwer Academic Publishers.] signalling (CaÂrdenas et al., 1998), but were not detected in other studies of Nod factor stimulation (Miller et al., 1999). In lower plants, although a cortical actin patch is detected during the establishment of polarity in algae (Braun and Wasteneys, 1998; Alessa and Kropf, 1999), this appears very different to the numerous actin foci that have been observed here. The nature of the actin patches stimulated by the SI response, and identi®cation of the components that interact with them, clearly need to be investigated further in order to obtain some understanding of their functional signi®cance. The speci®city of the alterations in actin con®guration was established by the use of several controls. The same biologically active S-proteins were used to challenge compatible pollen, thereby demonstrating the absolute Sspeci®city of the actin response because the only difference between the responses was the S genotype of the pollen. Addition of germination medium and heatdenatured S-proteins that were biologically inactive to incompatible pollen tubes acted as further controls. No inhibition of growth and no alterations in the actin cytoskeleton were observed in any of these situations (Geitmann et al., 2000; Snowman et al., 2000b). This is illustrated in Fig. 1h, i, which shows that 20 min and 60 min after a compatible challenge, the actin con®guration appears normal. These controls demonstrate clearly that the actin alterations observed are triggered speci®cally by an incompatible SI response. Quantitation of the alterations in F-actin organization during the SI response at different time intervals has helped to de®ne the timing of SI-induced events. This is Fig. 3. SI-induced F-actin alterations also occur in pollen grains. All images show rhodamine-phalloidin staining of F-actin. Images were obtained using confocal microscopy and are full projections in (a), (c) and (d), and single optical sections in (b), (d) and (f). Scale bar in (a) is 10 mm. (a) A normal pollen grain at 20 min has germinated and shows organized F-actin bundles. (b) An optical section through the pollen grain in (a) shows discernible cytosolic F-actin. (c) A pollen grain at 20 min post-SI has retained F-actin organization at the cortex, but (d) an optical section shows that virtually all cytosolic F-actin has been lost. (e, f) 60 min post-SI F-actin is organized into punctate foci that is distributed throughout the cytosol and cortex. shown in Fig. 2. Pollen tubes were scored according to the organization of their actin cytoskeleton following SI challenge. The initial movement of actin bundles at the tip was not scored, since virtually all of the incompatible pollen tubes showed this within 1±2 min. Three main classes of F-actin organization were scored. Class I was that found in normally growing pollen tubes (as described earlier). Class II were pollen tubes with a predominance of F-actin adjacent to the plasma membrane; while class III were those with membrane-localized punctate foci of actin. The histogram in Fig. 2 shows that after SI induction, an increase in class II pollen tubes is detected concomitant with a decrease in the number of class I pollen tubes. Subsequently, the number of class II pollen tubes decreases, and a proportional increase in pollen tubes of class III is detected. After a compatible challenge 80% of the pollen tubes counted were in the class I category (data not shown; Geitmann et al., 2000). These data not only Self-incompatibility response in poppy 107 show the initial speed of the response, but also establish that the cortical predominance of F-actin occurs early and prior to the formation of punctate foci of actin at the cell cortex. Although the SI response can be reproduced in pollen tubes that have been grown for several hours, this is not what generally happens in vivo, where inhibition normally occurs in pollen grains, either before or soon after germination. This being the case, F-actin con®gurations have also been examined semi-vivo, by incorporating Sproteins into an agarose-based germination medium, and in vivo (Geitmann et al., 2000). The incompatible pollen tubes displayed very similar alterations to the F-actin con®guration as those challenged in vitro. It was also examined whether the SI actin response occurs in ungerminated pollen grains. These SI inductions in pollen grains gave a similar, but delayed, actin response to those seen in pollen tubes (Snowman et al., 2002). At 20 min, normal pollen tubes had germinated, and had a wellorganized F-actin cytoarchitecture, as illustrated in Fig. 3a and b. By contrast, at 20 min post-SI, much of the cytoplasmic F-actin was lost, although F-actin staining adjacent to the plasma membrane was retained, as illustrated in Fig. 3c and d. Punctate foci of actin were detectable at 60 min (as shown in Fig. 3e, f) and continued to develop over several hours. This demonstrates that the SI-stimulated actin changes can occur in ungerminated pollen grains. SI in this species, although often occurring before germination, almost always occurs after polarity has been established, since deposition of callose is at a single colpal aperture, rather than at all three. The 20 min delay in the pollen grain actin response, which corresponds to the time taken to germinate, suggests that polarity perhaps needs to be established before the actin cytoskeleton can be targeted. This suggests that it is polarized tip growth that is targeted in this response. Further studies are required to test this hypothesis. Nevertheless, these data provide convincing evidence that the pollen actin cytoskeleton is targeted by the SI-induced signalling cascades. Cessation of growth is not responsible for the actin alterations In order to establish whether the SI-induced alterations in actin con®guration observed were a primary effect stimulated by incompatible S-proteins or were merely a secondary effect of the consequent arrest of growth, pollen tubes were subjected to several treatments that caused inhibition of pollen tube growth. Gd3+, caffeine and EGTA, in appropriate concentrations can effectively arrest pollen tube growth (Pierson et al., 1994, 1996). Although Gd3+ and caffeine caused a displacement of the subapical basket of actin, and EGTA caused only a very slight change to the subapical mesh, the general organization of the long arrays of actin bundles in the pollen tube shank Fig. 4. Latrunculin B induces changes to the F-actin cytoskeleton in pollen tubes. Effects of latrunculin B (10 min treatment) on F-actin in pollen tubes of P. rhoeas. Pollen tubes were treated, ®xed and stained with rhodamine-phalloidin. Images are single optical sections obtained using confocal microscopy. Scale bar is 10 mm. (a, b) 100 nM LatB. were intact and unchanged after these treatments (Geitmann et al., 2000). Since this non-speci®c inhibition of growth does not stimulate the gross changes in actin con®guration observed in the SI response, these data demonstrate that the SI-induced events are not merely secondary consequences of inhibition of growth, but are likely to be stimulated by the SI signalling cascade(s). Could actin depolymerization be responsible for some of the response? These observations suggested that actin depolymerization might be involved in mediating some of the cytoskeletal changes stimulated in the SI response. In order to test this possibility, the actin depolymerizing drugs latrunculin A (LatA) and latrunculin B (LatB) were used, which have been shown to bind to monomeric G-actin, thereby reducing the pool of available actin subunits for polymerization (Spector et al., 1989). This results in actin ®lament depolymerization; apparently LatB can completely deplete cellular F-actin (Pendleton and Koffer, 2001). A few minutes of treatment of pollen tubes with 50 nM LatA resulted in the characteristic actin con®guration of the pollen tube F-actin being replaced by a network of diffuse actin arrays. However, phalloidin staining adjacent to the plasma membrane remained strong. Treatment of pollen tubes with 100 nM LatB for 10 min resulted in a signi®cant loss of strongly-staining actin bundles. Short, weakly stained actin ®lament bundles were detected throughout the pollen tube, with a distinct signal adjacent to the plasma membrane, as shown in Fig. 4a and b. This suggests that there is a population of F-actin that is perhaps more resistant to depolymerization most likely because of interactions with unknown ABPs in this region. Treatment with 1 mM LatB resulted in further apparent fragmentation of the F-actin cytoskeleton, giving a speckled appearance. 108 Staiger and Franklin-Tong These studies indicate that though the effect of LatB has some similarity to the intermediate stages detected in incompatible pollen tubes, the alterations to F-actin induced by lantrunculin depolymerization are subtly different to the changes in organization induced by SI. This suggests that, although depolymerization is likely to play a role in the SI response, other factors are, not surprisingly, likely to be involved. Since the imaging data for the SI-induced F-actin changes suggested that there was likely to be a decrease in F-actin in incompatible pollen tubes as, qualitatively, the phalloidin signal appears to be reduced upon S-protein challenge, it was decided to investigate this possibility further. Quanti®cation of F-actin levels in pollen In order to investigate the alterations to actin in incompatible pollen further, a quantitative approach has recently been used. Although a number of studies have provided microscopic analysis of alterations to the actin cytoskeleton in response to a variety of stimuli in plant cells, there are few examples of quanti®cation and biochemical analysis of the changes. Indeed, currently, there exists only one published report of quanti®ed changes to the Factin concentration in a plant system, and this is in maize pollen (Gibbon et al., 1999). This study used an adaptation of a biochemical method based on the number of ¯uorescent-phalloidin binding sites which has been used to measure F-actin content in activated neutrophils and yeast (Howard and Oresajo, 1985b; Lillie and Brown, 1994). This assay was used here to determine the concentration of actin in polymeric form in pollen (Snowman et al., 2002). The levels of F-actin in pollen throughout hydration, germination and growth for several hours were surprisingly constant. The mean concentration of actin in polymeric form was 15.4 mM (SE 62.7; n=22). A test was then done to see if changes in F-actin concentration could be detected by adding LatB, which will depolymerize F-actin (Gibbon et al., 1999). Pollen challenged for 10 min with 100 nM and 1 mM LatB revealed signi®cant decreases in F-actin levels of ~60% and 45%, respectively (Snowman et al., 2002). This demonstrated that the phalloidin-binding assay was appropriate for measuring rapid changes in F-actin levels in P. rhoeas pollen. The SI response involves F-actin depolymerization Since the F-actin concentration in pollen could be effectively measured, it was examined whether an incompatible SI challenge resulted in alterations to this. The previous observations had suggested that depolymerization might be stimulated, since there was a loss of phalloidin staining in the cytoplasm and the F-actin bundles were lost, being replaced by a speckled array of ®ne F-actin fragments. However, this could potentially suggest fragmentation or cleavage rather than depolymerization per se. It was therefore important to establish what type of actin alterations are being evoked. Use of the phalloidin-binding assay has revealed a rapid, large and sustained decrease in F-actin levels as a consequence of an incompatible response. Even within 1 min, which was the earliest time-point taken, there was a signi®cant reduction in F-actin compared to untreated pollen. By 5 min after SI induction, F-actin levels were less than 50% that of the controls. F-actin levels decreased further during the 60 min time period over which these studies were made (Snowman et al., 2002). These data clearly demonstrate rapid and sustained F-actin depolymerization that was S-speci®c, since the relevant controls of compatible pollen treated with the same S-proteins and also incompatible pollen challenged with germination medium or inactive S-proteins did not show changes in Factin levels (data not shown). This very large reduction in F-actin was observed in both incompatible pollen grains and pollen tubes. The overall F-actin reduction as a consequence of SI challenge was 74% in tubes and 56% in grains, compared with the GM controls at 60 min. One interesting difference between the response in pollen grains and pollen tubes was that the response in pollen grains was delayed by ~20 min, which corresponds to the time taken for polarity to be established. This might suggest that the SI-induced actin depolymerization acts on tip growth. This idea is substantiated by the studies with maize pollen, which demonstrate that differences in sensitivity to LatB are not due to ungerminated grains having a more stable actin cytoskeleton than actively growing pollen tubes (Gibbon et al., 1999). Increases in cytosolic calcium result in changes in F-actin How the changes in the actin cytoskeleton described here relate to the other SI induced signalling events already identi®ed is an important question. It is well established that Ca2+ acts as a second messenger in the SI response. Increases in [Ca2+]i are probably the ®rst events in this signalling cascade, and are detectable within a few seconds of the SI-interaction (Franklin-Tong et al., 1997, 2002). The ®rst changes to the F-actin cytoskeleton were detected at the earliest time point possible for ®xed cells, 30 s. Whether Ca2+ is involved, either directly or indirectly, in these changes is an important question to answer. Therefore an investigation as to whether F-actin may be a target for a Ca2+-dependent signalling cascade has begun. Treatments known to induce increases in [Ca2+]i in pollen tubes were used. The Ca2+ ionophore, A23187 and mastoparan (Franklin-Tong and Franklin, 1993; FranklinTong et al., 1996) were used to see whether arti®cial increases in [Ca2+]i will stimulate changes to the status of Self-incompatibility response in poppy 109 F-actin in pollen tubes. Both of these drug treatments resulted in changes to the actin cytoskeleton that had some broad similarities to those stimulated by SI. Features included fragmented and bundled F-actin, small punctate foci of F-actin, and later, larger punctate foci of F-actin (data not shown; Snowman et al., 2002). considerable decreases in F-actin, indicating depolymerization, stimulated by these drugs were also measured (data not shown; Snowman et al., 2002). Since increases in [Ca2+] can stimulate actin depolymerization, it suggests that the SIinduced Ca2+ increases target the actin cytoskeleton. Early studies show that increasing [Ca2+]i arti®cially can disrupt actin ®laments in pollen tubes (Kohno and Shimmen, 1987, 1988). However, although a gelsolin-like severing activity was considered, this is rather speculative, as there is no ®rm evidence for this suggestion. Discussion Changes to the actin cytoskeleton cytoarchitecture stimulated by the SI response are described here, together with preliminary studies that begin to quantify some of the biochemical changes to the actin cytoskeleton. Several alterations to the pollen F-actin cytoskeleton stimulated by the SI response in P. rhoeas have been demonstrated. Furthermore, a rapid and considerable F-actin depolymerization in incompatible pollen tubes has recently been measured. Quantitative measurements of changes to Factin levels in a plant cell by a de®ned biological stimulus have not previously been reported, although Gibbon et al. (1999) provided the ®rst such measurements in pollen from maize. Although there are several studies describing changes to the plant actin cytoskeleton in response to physiological stimuli, they are all descriptive (Eun and Lee, 1997; Kobayashi et al., 1997; CaÂrdenas et al., 1998; Miller et al., 1999). It is, therefore, not known if these alterations involve reorganization or changes in the polymerization status of actin. Depolymerization of actin during SI There are surprisingly few examples of actin depolymerization stimulated in response to a biologically-relevant ligand, as extracellular stimuli generally appear to lead to the polymerization of actin ®laments (Carlsson et al., 1979; Howard and Oresajo, 1985a; Hall et al., 1988; Greenberg et al., 1991; Downey et al., 1992; Becker and Hart, 1999). The few examples of measurements of actin depolymerization include the stress response of yeast (Lillie and Brown, 1994; Yeh and Haarer, 1996), the collapse of growth cones in response to chemo-repulsive axonal guidance molecules (Aizawa et al., 2001), and vasopressin stimulation of toad bladder epithelial cells (Ding et al., 1991). However, it is striking that depolymerization is generally transient. For example, during the heat shock response of yeast a 25±50% reduction in F-actin within 30 s is observed, but levels return to normal within 3 min (Lillie and Brown, 1994). By contrast, SI-induced pollen depolymerization continues for at least 60 min. This suggests that the formation of actin punctae is due to actin reorganization rather than re-polymerization. Since these studies clearly demonstrate that alterations to the actin cytoskeleton also continue well after the inhibition of tip growth, this suggests that SI-induced signalling events also occur after the inhibition of growth. Other SI studies also indicate this (Rudd and Franklin-Tong, 2003). It is proposed that some of these events may function to make this arrest of growth irreversible. Actin-binding proteins implicated in the pollen SI response The dramatic and rapid changes of the actin cytoskeleton after S-protein challenge raise the question of how these alterations are mediated. Actin-binding proteins (ABPs) are thought to be responsible for modulating changes in actin organization and dynamics. Depolymerization could result from a loss of side-binding proteins, capping of ®lament ends, increased severing activity, or stimulation of a sequestering protein. However, only a few ABPs have been identi®ed and characterized in pollen tubes. These are pro®lin, ADF/co®lin and 135-ABP (Hepler et al., 2001; McCurdy et al., 2001) Although ADF/co®lin can cause actin depolymerization and is regulated by Ca2+ indirectly, increases in [Ca2+]i are expected to inhibit, rather than activate F-actin depolymerization by ADF (Smertenko et al., 1998; Allwood et al., 2001). 135-ABP is a villin-like protein with Ca2+/calmodulin-regulated actin-bundling activity (Yokota et al., 2000). However, no demonstration of an actin capping, severing or depolymerizing activity for 135-ABP has been made yet. Since a villin-like protein, Drosophila QUAIL, lacks detectable severing and depolymerizing activity (Matova et al., 1999) it is unwise to assume biochemical properties on the basis of sequence homology. Furthermore, as there are no data on cellular [135-ABP] in pollen or accurate measurements of its af®nity for F-actin, its potential function in SI is currently impossible to assess. Pollen pro®lin has recently been shown to have increased actin-sequestering activity at elevated, but physiological calcium concentrations (Kovar et al., 2000). This ABP seems a likely contributor to actin depolymerization during SI, but probably does not act alone. Detailed analysis of cellular concentrations and binding constants for these and other yet to be identi®ed pollen ABPs will be necessary before the actin remodelling that occurs during SI is fully understood. A potential link between actin and PCD? There is good evidence that apoptosis in some animal cell types (e.g. HL-60 cells) can trigger F-actin depolymeriza- 110 Staiger and Franklin-Tong tion; moreover, this status is maintained for several hours (Levee et al., 1996; Korichneva and HaÈmmerling, 1999; Rao et al., 1999). Thus, triggering of apoptosis is virtually the only current system where long-term actin depolymerization has been measured. The large actin depolymerization that is sustained for at least 1 h in incompatible pollen is highly reminiscent of events triggered by apoptosis. This suggests that a programmed cell death (PCD) signalling pathway might be triggered by SI in incompatible pollen. Since there is preliminary evidence that PCD is activated in pollen by SI (Rudd and Franklin-Tong, 2003), this is not pure speculation, though further investigations to establish a ®rm link need to be made. Activated caspase-3, a key effector of apoptosis, has been shown to have the capacity to cleave several cellular proteins in animal cells. Both actin and ABPs are implicated as targets for activated caspases. Proteolytic cleavage of actin has been observed in some animal cell types (Brown et al., 1997; GueÂnal et al., 1997; Korichneva and HaÈmmerling, 1999; Paddenberg et al., 2001), although this is not a universal phenomenon (Levee et al., 1996; Bursch et al., 2000). Furthermore, there is evidence that several ABPs are also targets for caspase activity (Kothakota et al., 1997; Janmey, 1998; Umeda et al., 2001), including gelsolin. The gelsolin cleavage product can disrupt F-actin in the absence of Ca2+ and may preferentially sever F-actin rather than bind monomeric actin (Kothakota et al., 1997). Furthermore, this gelsolin cleavage product can trigger apoptosis (Kothakota et al., 1997). Recent results from studies on animal cells indicate that the regulation of actin polymerization may be an important apoptotic morphological effector. There is convincing evidence that F-actin depolymerization may act as a effector of apoptosis in some cell lines (Janmey, 1998; Korichneva and HaÈmmerling, 1999; Rao et al., 1999). It has recently been demonstrated that depolymerization of actin can activate caspases and cytochrome c release from mitochondria, thereby stimulating apoptosis (Paul et al., 2002). However, how depolymerization of the actin cytoskeleton is achieved and maintained during apoptosis is currently not known. Whether gelsolin is involved in these cases is not yet known. The existence of PCD in plant cells is now widely accepted (Greenberg, 1997; del Pozo and Lam, 1998; Lam et al., 2001). Although there are no clear caspase orthologues in the Arabidopsis genome, a number of studies indicate that caspase-like activities are triggered by PCD in plant cells (D'Silva et al., 1998; del Pozo and Lam, 1998; Korthout et al., 2000; Richael et al., 2001). Since there is evidence that PCD is stimulated in the SI response (Jordan et al., 2000), with preliminary evidence for a caspase-like activity stimulated in the SI response (Rudd and Franklin-Tong, 2003), this might indicate a potential reason why actin depolymerization is sustained. Thus, it was postulated that a PCD signalling cascade is triggered by the SI response and actin depolymerization may potentially play an active role in the SI response by stimulating caspase activity. This is, at present, the only explanation (based on the available literature) for sustained actin depolymerization observed during SI. Further studies to explore the evidence for a PCD signalling cascade and to provide ®rm links between this and the sustained depolymerization of F-actin are required. In summary, changes to the actin cytoskeleton during the SI response have been described. There is evidence for a rapid, massive and sustained depolymerization of Factin, and that this is induced via increased [Ca2+]. However, surprisingly, the most obvious (and at present, the only) candidate for this Ca2+-dependent depolymerization, namely, pro®lin, is unlikely to be solely responsible for the extent of depolymerization. The sustained nature of the depolymerization is highly unusual and points to a possible link with a PCD signalling cascade. Acknowledgements The work described here was carried out by Benjamin Snowman funded by a BBSRC Biochemistry and Cell Biology Committee studentship and by Anja Geitmann, in collaboration with Anne-Mie Emons, funded by a Mare Curie Fellowship. This work was funded by USDA-NRICGP support to CJS and BBSRC support to VEFT. Thanks are due to BBSRC for a special travel grant that allowed collaborative work between CJS and VEFT. References Aizawa H, Wakatsuki S, Ishii A et al. 2001. 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