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Journal of Experimental Botany, Vol. 60, No. 2, pp. 495–508, 2009 doi:10.1093/jxb/ern321 Advance Access publication 26 December, 2008 REVIEW PAPER Organelle motility in the pollen tube: a tale of 20 years Giampiero Cai* and Mauro Cresti Dipartimento Scienze Ambientali, Università di Siena, via Mattioli 4, I-53100 Siena, Italy Received 10 September 2008; Revised 18 November 2008; Accepted 18 November 2008 Abstract Organelle movement is an evident feature of pollen tubes and is essential for the process of tube growth because it enables the proper distribution of organelles and the accumulation of secretory vesicles in the tube apex. Organelles move along the actin filaments through dynamic interactions with myosin but other proteins are probably responsible for control of this activity. The role of microtubules and microtubule-based motors is less clear and somewhat enigmatic. Nevertheless, the pollen tube is an excellent cell model in which to study and analyse the molecular mechanisms that drive and control organelle motility in relation to plant cell expansion. Current knowledge and the main scientific discoveries in this field of research over the last 20 years are summarized here. Future prospects in the study of the molecular mechanisms that mediate organelle transport and vesicle accumulation during pollen tube elongation are also discussed. Key words: Actin filaments, cell growth, cytoplasmic movement, kinesin, microtubules, myosin, organelles, pollen tube. Introduction: the importance of actin filaments Sexual reproduction in seed plants (gymnosperms and angiosperms) shows a critical evolutionary step consisting of the use of a specialized male gametophyte (pollen) and its developmental product (the pollen tube). The advent of the pollen grain enabled male gametes of seed plants to be transported over long distances and to resist adverse environmental conditions. Development of pollen and pollen tubes, coupled with the evolution of specialized organs (flowers) in angiosperms, allowed plants progressively to do without water for fertilization and to invade dry land (Cresti et al., 1992). The pollen tube is a tubular cell that allows the male gametes to navigate from male to female flower organs and to do without specialized motile devices, like flagella, that are required for sperm movement in lower plants (Tiezzi et al., 1991). The growth of pollen tubes is a specialized type of cell expansion (apical growth) characterized by the fusion of Golgi-derived secretory vesicles in a restricted cellular domain (the tube tip). Three main domains of pollen tubes have been proposed: the apical region is the growth domain, the subapical region is a transition domain, while the base domain is more or less similar to other plant cells and contains the typical repertoire of plant cell organelles. As in other plant cells, organelles and vesicles move rapidly back and forth along the main axis of the pollen tube: this activity (‘cytoplasmic streaming’) is required to maintain the distribution of membraneous structures and to focus Golgi vesicles in the growth domain of the pollen tube. Movement of pollen tube organelles is persistent but single organelles behave independently of each other and move at different speeds (Fig. 1A). Trafficking of membrane structures occurs toward the tip, mainly along the tube flanks; in the subapex, more or less in alignment with a specific actin array (the actin fringe), organelles reverse their direction of movement and turn back along the central actin cables. Golgi-derived secretory vesicles accumulate in the tip domain by moving along peripheral actin cables; vesicles are probably finally delivered to the tip region by the actin fringe while recycling and endocytotic vesicles are then driven back along the central actin filaments (Fig. 1B). Cytoplasmic streaming is a key feature of plant cells, whose large size obliged them to set up an intense circulation of organelles in order to facilitate intracellular transport and the distribution of resources. The pollen tube is, for many reasons, an excellent cell model in which to study the molecular events underlying * To whom correspondence should be addressed: E-mail: [email protected] ª The Author [2008]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved. For Permissions, please e-mail: [email protected] 496 | Cai and Cresti Fig. 1. (A) Movement of organelles in tobacco pollen tubes as shown by DIC video-microscopy. A single pollen tube is shown after 3 h of growth. Specific organelles are indicated as representative of different patterns of motility. The organelle indicated by an arrowhead moves very fast (1.5 lm s 1), while the two organelles indicated by arrows move very slowly; in particular, the organelle marked by a black arrow is apparently immobile. The organelle labelled with a white line moves slowly (0.5 lm s 1). Numbers at the bottom left are time in seconds. Bar: 5 lm. (B) Schematic drawing of organelle and vesicle trafficking in a pollen tube. Both structures move toward the apex along the flanks of the pollen tube; in the subapical region, organelles reverse direction and turn back in the centre of the pollen tubes. Vesicles are directed toward the apex where they fuse with the plasma membrane; vesicles that do not penetrate the apex are recycled together with endocytotic vesicles. organelle transport. It is a single haploid cell, the ultrastructure of which is well known in different species, and can be manipulated easily. Although most growth events occur in a spatially-limited region (the tip), the cell shows many features typical of plant cells, such as interactions with other cells (the female stigma cells), polarized growth, and time-dependent cell wall construction. Pollen tubes can be analysed by different techniques, such as immunocytology, biochemistry, genetic transformation, and live cell imaging. This is strictly possible in all species, but a proper combination of different pollen species (Nicotiana tabacum, Lilium longiflorum, Arabidopsis thaliana, etc) provides a valuable plant cell model. The aim of this review is to summarize scientific discoveries in the field of organelle movement and pollen tube growth in the last 20 years. We start by reviewing preliminary observations on the critical role of actin filaments and end by describing the final fate of secretory vesicles and the interplay between vesicles and actin filaments in endo/exocytosis. The choice of this span of time was intentional. In 1988, the 10th Symposium on Sexual Reproduction in Higher Plants (co-organized by M Cresti), was held in Siena (Italy). The opening lecture by Professor Jack Heslop-Harrison (1920–1998) was focused on organelle transport in pollen tubes (‘The pollen tube: motility and the cytoskeleton’) and on the use of videoenhanced microscopy to correlate the movement of pollen tube organelles with the organization of actin filaments and microtubules (Heslop-Harrison and Heslop-Harrison, 1988). In their meticulous pioneering work, the HeslopHarrisons showed the motility of organelles in older segments of pollen tubes and a putative correlation between organelle transport and actin cytoskeleton. In 1988, two other key manuscripts on organelle movement in pollen tubes were published. Heslop-Harrison et al. (1988) showed Organelle movement in pollen tubes | 497 that treatment with colchicine did not affect pollen tube growth and organelle transport but may influence shaping of the generative cell during its course through the tube. Work by Kohno and Shimmen (1988) revealed that isolated pollen organelles moved along Characeae actin bundles in an ATP-dependent manner and that motility was influenced by Ca2+. The manuscripts indicated that actin filaments were the main tracks along which pollen tube organelles moved and indirectly suggested that myosinbased organelle movement was regulated by variations in Ca2+ concentrations. Organization of the cytoskeleton in angiosperm pollen tubes was already partly known. Condeelis (1974) identified F-actin in pollen tubes of Amaryllis belladonna, later related to the movement of pollen nuclei (Cresti et al., 1976). Many years later, the availability of specific probes enabled researchers to show that actin filaments and microtubules are arranged along the longitudinal axis of pollen tubes, indicating the extent of overlapping as well as several differences in the apical (growth) region (Pierson et al., 1986). It was immediately clear that cytoskeletal organization in the tip domain of pollen tubes was hard to decipher because of fixation artefacts due to the dynamics of the tip cytoskeleton. Using different chemical procedures, Pierson (1988) showed differential organization of actin filaments in the tip domain of pollen tubes and presumed ‘actin foci’ in the pollen tube flanks; since similar structures are involved in cell–cell interaction in animal cells, the pollen ‘actin foci’ were postulated to have a role in the process of communication between pollen tube and style. Soon after, the actinbased motor, myosin was discovered. Using antibodies to animal myosins, Tang et al. (1989a) identified a myosin heavy-chain polypeptide of 175 kDa in Nicotiana pollen tubes. Based on its localization, myosin was deduced to be involved in the movement of vesicles, organelles, and the male germ unit in pollen tubes. Similar results were obtained with different types of myosin antibodies (HeslopHarrison and Heslop-Harrison, 1989b), sustaining the model that actin and myosin were the main protein mechanism guiding organelle transport. In order to understand the function of myosins, more information was also needed on the distribution of actin filaments. Anti-actin antibodies revealed axially oriented strands of actin filaments distributed alongside the pollen tube except at the very tip. Conventional staining with rhodamine–phalloidin provided similar results (Tang et al., 1989b). Actin filaments apparently organized as bundles along the pollen tube length but showed a distinct pattern in the growing domain, suggesting that differential organization of actin was required for tube elongation. Similar results were obtained using microwave-accelerated DMSOpermeabilisation and TRITC-phalloidin staining (HeslopHarrison and Heslop-Harrison, 1991). How do the distribution of myosins and the organization of actin filaments correlate with organelle transport and zonation in pollen tubes? Zonation is a way of describing the different domains of pollen tubes based on their membrane content: the vesicular, organelle, nuclear, and vacuole regions (Cresti et al., 1977). However, these areas are not separated by defined boundaries and exchange of materials among them is a prerequisite for growth. Thus, zonation is the result of dynamic activities that, for example, constrain vesicles in the tip region and prevent large-sized organelles from entering. Correlations between actin filaments and organelle distribution were analysed by Heslop-Harrison and Heslop-Harrison (1989a), who found that cytochalasins B and D affected pollen tube growth and rapidly stopped organelle circulation and movement of vegetative nuclei and generative cells. A striking characteristic of organelle circulation in pollen tubes is their tipward movement along the tube flanks and backward transport in the centre (the so-called ‘reverse fountain’). The reversal of organelle flow (which occurs in the apical 5–15 lm of growing pollen tubes) prevented large-sized organelles (like lipid globuli and amyloplasts) from penetrating the apical region but allowed them to arrest and reverse their movement in the subapical domain (Heslop-Harrison and Heslop-Harrison, 1990). These findings suggested that the subapical region contained a sort of filter that discriminated the different organelles. Based on cytological observations, the main candidate for filtering activity was the actin cytoskeleton, described as a network of cross-linked actin filaments in the tip and ordered, orientated bundles in the older parts. The inability of large-size organelles to enter the apex was also presumed to depend on the dense mass of tiplocalized vesicles (Pierson et al., 1990). Vacuoles in the central zone were observed to be very motile and the cytoplasmic strands in the vacuolar region contained elements of the endoplasmic reticulum along which organelles moved in a saltatory manner (Pierson et al., 1990). These findings suggested that the endoplasmic reticulum was associated with actin filaments, which, in turn, supported the movement of organelles. This speculative hypothesis would later be confirmed using GFP-tagged Golgi bodies (Nebenführ and Staehelin, 2001). The relationships between endoplasmic reticulum and Golgi bodies were also investigated. Pollen tubes treated with Brefeldin A showed dense cytoplasm containing tubular reticulum in the tip domain and rapidly dissociating and disappearing Golgi bodies, which fused with the endoplasmic reticulum (Rutten and Knuiman, 1993). The indecipherable role of microtubules Most of the evidence discussed above indicated that actin filaments were the main pathways along which organelles and vesicles moved in the pollen tube. The basic evidence that colchicine did not affect pollen tube growth was indirect proof that microtubules were not critical for organelle transport and tube elongation. However, a number of researchers were investigating the distribution and putative function of microtubules in pollen. Aström (1992) showed that most pollen tube microtubules contain tyrosinated a-tubulin whereas acetylated a-tubulin was detected in the generative cell, kinetochore fibres, and cell plate of the phragmoplast; acetylation of a-tubulin was thought to 498 | Cai and Cresti be correlated with more stable microtubules. A breakthrough in the pollen microtubule world was the immunological identification of a 105 kDa kinesin-related protein in Nicotiana tabacum (tobacco) pollen tubes (Tiezzi et al., 1992). The kinesin-related homologue of pollen tubes localized in the apical and subapical regions of pollen tubes, probably in association with vesicular material. The protein also co-localized with pollen tube microtubules (Cai et al., 1993). Further confirmation of the presence of kinesinrelated proteins came from studies in Corylus avellana pollen, which contained a 100 kDa kinesin-related protein associated with Golgi vesicles (Liu et al., 1994). The presence of kinesin-related proteins in pollen tubes cast doubts on the role of microtubules in organelle transport, however, detailed observations on the distribution of microtubules in tobacco pollen tubes did not clarify the presence of microtubules in the tip domain and their involvement in vesicle transport (Del Casino et al., 1993). In addition, the absence of microtubule-based motility data weakened the preliminary indications on the role of microtubules in vesicle transport. The microtubule-motor story became a little more complicated a couple of years later when dynein-related polypeptides were identified in tobacco pollen tubes (Moscatelli et al., 1995) associated with pollen tube organelles (Moscatelli et al., 1998), indicating that the microtubule motor repertoire of pollen tubes may comprise different motor classes. Although the presence of dynein in plants is still debated, dyneins seem to have been lost from higher plants (Wickstead and Gull, 2007) and their role seems to have been replaced by the large kinesin family. Although the microtubule story was highly promising, the indisputable lack of colchicine- and oryzalin-related effects on tube elongation was a crucial obstacle for these preliminary discoveries. To be more precise, oryzalin was reported to affect the pulsatory growth of pollen tubes (Geitmann et al., 1995) and generative cell movements (Aström et al., 1995), but these findings were hardly related to organelle movement and motors. Multiple myosins? Although actin–myosin interactions were considered to play a major role in organelle transport in pollen tubes, the only myosin identified was a 175 kDa polypeptide detected by heterologous antibodies (Tang et al., 1989a). Like other cell systems, it seemed possible that pollen tubes contained different classes of myosins, putatively involved in different functions. Using antibodies to different myosin classes, Miller et al. (1995) observed that myosin-I seemed to move the generative cell and vegetative nucleus along the pollen tube, while myosin-V moved small organelles (vesicles) and myosins-I and II moved large organelles in two directions. A different antibody against heavy meromyosin detected a 174 kDa polypeptide associated with cytoplasmic organelles of tobacco pollen tubes, most fluorescent spots being concentrated in the tip region below the plasma membrane. This finding was also confirmed by immunoelectron microscopy that revealed gold particles around vesicle-like structures (Tirlapur et al., 1995). To sum up, data obtained in the middle 1990s indicated the presence of various myosins, whose different staining patterns were probably due to the use of heterologous antibodies and the cross-reactivity of antibodies with different myosin classes. A unified model was missing. Immunological detection was not the only way to reveal myosin-like proteins in pollen tubes. Kohno et al. (1990) had already described the main biochemical properties of one pollen myosin; the same research group was able partially to purify one myosin polypeptide (Kohno et al., 1992) and isolate the complete 170 kDa motor protein (Yokota and Shimmen, 1994). The myosin of Lilium longiflorum (lily) pollen tubes showed properties typical of the myosin family and was localized as spots in the tip domain of Lilium longiflorum pollen tubes and throughout the pollen tubes of tobacco (Yokota et al., 1995). These results were the first evidence of a pollen-specific myosin and suggested that the protein might be associated with, and involved in trafficking of different organelle classes. It was evident that angiosperm pollen tubes contained at least one myosin of 170 kDa; proofs for other myosins essentially depended on the antibody used for detection. The presence of different myosins in pollen tubes was also suggested again more recently (Lovy-Wheeler et al., 2007) after evidence that different organelle classes of pollen tubes move differentially. Since the genome of Arabidopsis thaliana contains 17 myosins belonging to families VIII and XI (Lee and Liu, 2004), pollen tubes presumably contain different members of the myosin XI family, each involved in the movement of different organelles and vesicles. Clearly, interpretation of the function of myosin(s) in pollen tubes is limited by the absence of genetic data. The presence of genes coding for myosin VIII and XI in the plant genome (Jiang and Ramachandran, 2004) suggests that the previous localization of myosin-I, II, and V should be re-examined in a new light. For example, the previously identified 175 kDa (Tang et al., 1989a) and 174 kDa myosins (Tirlapur et al., 1995) could correspond to the 170/175 kDa myosins identified later (Yokota et al., 1995), assuming that polyclonal antibodies to myosin-II cross-react with plant myosin-XI. Since there is functional homology between animal myosin-V and plant myosin-XI (Li and Nebenführ, 2007), antibodies to myosin-V may indirectly indicate that plant myosin-XI moves organelles and vesicles in the pollen tube (Miller et al., 1995). Myosin-VIII is yet to be discovered and may be responsible for the endocytotic process in the pollen tube apex (Golomb et al., 2008). The binding of different myosins with specific organelles is a further fascinating research field; myosins probably associate with the membrane of plant organelles by means of Rab proteins (Hashimoto et al., 2008), which regulate the binding and, consequently, the activity of motor proteins. In any case, to learn more about myosin functions, cDNAs encoding myosins expressed in pollen tubes need to be cloned and used for functional studies. Organelle movement in pollen tubes | 499 Fast progress on actin dynamics through gene cloning and GFP technology In the second half of the 1990s, advances in molecular biology techniques opened new research perspectives in the field of organelle movement. Immunological techniques had been extremely important in uncovering the distribution of cytoskeletal proteins, however, alternative procedures were needed to obtain new information. Techniques of genetic engineering and DNA cloning allowed researchers to move from basic cytoskeletal proteins to the discovery of many different accessory proteins involved in actin dynamics. Analysis of organelle movement made progress through the observation of single organelles after labelling with specific dyes, exploiting specific computer software. Researchers inferred that movements of membraneous structures were random in the pollen tube tip but were more directed in older cell regions (de Win et al., 1998). These findings indicated indirectly that the organization of the actin cytoskeleton changed from the tip to the base of the pollen tubes. The differential distribution of actin in living pollen tubes was also revealed by genetic transformation with GFP-labelled actin-binding proteins (reviewed in Wilsen et al., 2006). However, a characteristic pattern of labelling was obtained with certain probes. High expression levels of probes also caused structural rearrangements of actin filaments and inhibited growth. Although these markers are important technological advances, their use does not yet allow complete visualization of the actin cytoskeleton in growing pollen tubes and may cause artefacts and prevent pollen tube growth. At the same time, interest in pollen tubes was captured by the study of mechanisms regulating growth and transduction of external signals into internal processes. In this context, analysis of ion influx and dynamics provides insights into the relationships between ion concentrations and actin filaments. Dissipation of the tip-focused Ca2+ gradient enhanced entry of large organelles into the tube tip (Pierson et al., 1994), suggesting that Ca2+ controlled the organization of actin filaments, as proposed by Kohno and Shimmen (1987). Another paper described the presence of an alkaline band delimiting the apical domain and defining the acidic tip of pollen tubes (Feijo et al., 1999). It was suggested that the apical acidic domain correlated with the growth process, probably facilitating vesicle movements and exocytosis. The alkaline band probably also represented the area where organelles reversed their motion (‘reverse fountain’) by regulating the assembly of actin filaments through pH-sensitive actin-binding proteins. Reversal of organelle motility in the subapical region presumably required appropriate local organization of actin filaments. In this context, Li et al. (2001) described the presence of circular F-actin bundles in pollen tubes, missing in the initial 10-20 lm of the tubes and well correlated with cytoplasmic streaming of organelles. Although the circular F-actin bundles were not further described, they could speculatively represent the track for bidirectional cytoplasmic streaming in pollen tubes. The so-called former ‘actin foci’ represented an attempt to associate actin dynamics with extracellular signals as in animal cells. To understand the dynamics of actin filaments in relationships with external cues it was necessary to discover proteins that mediate the response of actin to extracellular signals. Rop (‘Rho-of-plants’) GTPases were found in the growth sites of pollen tubes and were assumed to control polar growth (Zheng and Yang, 2000). RopGTPases influence the dynamics of tip-localized actin in pollen tubes by transforming short actin bundles of the apical domain into a filamentous network; overexpression of Rop-GTPases also induces the formation of transverse actin bands behind the tip (Fu et al., 2001). Speculatively, external signals are converted by tip-localized Rop-GTPases into messengers that modulate the organization of actin filaments and thus the motility of organelles and the growth of pollen tubes. The Rop-mediated equilibrium between G- and F-actin requires the activity of accessory intermediate proteins. Villin was identified in lily pollen tubes and localized in association with actin bundles (Vidali et al., 1999). Villins are Ca2+-dependent, severing, capping, and nucleating proteins and may participate in F-actin fragmentation and nucleation in the pollen tube apex following activation by the Ca2+ gradient. Like villin, LdABP41 is a Ca2+dependent protein that cuts actin filaments into short filaments and may operate in combination with the tip-focused Ca2+ gradient (Fan et al., 2004). Villin also stimulates the formation of unipolar bundles of actin filaments. Pollen tubes combine villin with gelsolin (Huang et al., 2004; Xiang et al., 2007) and profilin (Vidali and Hepler, 1997) to organize actin filaments into two different and partly autonomous arrays (the actin cables in pollen tube shanks and shorter filaments in the apical and subapical regions), required for the processes of organelle transport and tube growth. Although tip growth is more sensitive to actin inhibitors than cytoplasmic streaming, the two activities are related because tip growth requires a continuous supply of secretory vesicles, indicating that pollen tube growth requires the assembly of different subsets of actin filaments, which have different sensitivity to actin drugs (Vidali et al., 2001). Consequently, pollen tube growth is the result of cytoplasmic streaming, accumulation of secretory vesicles in the tip domain (which depends on longitudinal actin filaments) and the rate of vesicle fusion (which may require a different actin array). The number of actin-modulating proteins of pollen tubes discovered increased rapidly. Actin-depolymerizing factors (ADF) were identified as proteins showing pH-sensitive actin-binding and depolymerizing activity (Allwood et al., 2002). The alkaline band in pollen tubes was therefore postulated to enhance the activity of ADFs and shift actin toward the monomeric form. Thus the tip-focused Ca2+ gradient and the alkaline band are critical markers of reverse streaming activity (Cardenas et al., 2005). The organization of actin filaments was also found to depend on the activity of formin, a group of actin-nucleating proteins that enhanced the formation of actin cables in the pollen 500 | Cai and Cresti tube (Cheung and Wu, 2004) and promoted reversal of streaming in the apical domain. The entire process of pollen tube growth (comprising organelle/vesicle transport and endo/exocytosis) necessarily required equilibrium to be maintained between G- and F-actin. Recognizing the number and role of the molecular actors, which ultimately regulate actin dynamics, and their relationships with differential ion concentrations is a crucial forthcoming challenge. Dynamics of specific organelle classes Analysis of actin dynamics was accompanied by the visualization of individual membrane classes in living pollen tubes using specific markers. For example, FM4-64, a marker of vesicles in the tube apex, highlighted the organization of secretory vesicles in the ‘apical clear zone’ and revealed distinct pathways of vesicle movement toward and away from the apex (Parton et al., 2001). Vesicle distribution was found to fluctuate in the apical domain of pollen tubes, suggesting that accumulation of secretory vesicles in the pollen tube apex is not continuous but shows a periodicity that depends on the actin cytoskeleton but not on the secretion rate (Parton et al., 2003). Again, these findings suggested that organelle and vesicle motility is a process unrelated to secretion and growth. Visualization of GFP-Rab11b-tagged vesicles in the inverted conical region of tobacco pollen tubes after Latrunculin treatment again indicated that accumulation of vesicles in the tip is closely linked to the actin cytoskeleton (de Graaf et al., 2005). Other cell compartments (specifically, endoplasmic reticulum, mitochondria, and vacuoles) were monitored during pollen tube growth and their movement was shown to depend on actin filaments but apparently not on microtubules. Movement of endoplasmic reticulum followed the direction of the main cytoplasmic streaming in pollen tubes, traveling apically along the pollen tube flanks and reversing its motion in the ‘actin fringe’ domain. Mitochondria were found predominantly in the subapical region and showed a movement similar to that of the endoplasmic reticulum. In contrast, the vacuole was formed by thin longitudinal tubules undergoing active motion but never extending into the apical domain (Lovy-Wheeler et al., 2007). Although cytoplasmic streaming generated the circular motion of membraneous structures throughout the pollen tube, the above findings showed that organelle distribution was neither uniform nor accidental but precisely controlled. A key question now is whether different myosins or different regulation of the same myosin may be responsible for differential organelle distribution. Organelles and vesicles also move along microtubules Apart from preliminary observations on putative kinesinrelated protein in pollen tubes, many findings suggested that a role of microtubules during in vitro pollen tube growth was unnecessary. Evidence that colchicine and oryzalin treatment caused few or no effects on the elongation rate of pollen tubes led to the consideration that actin filaments were the only motor system supporting organelle transport. More consistent evidence that microtubules may take part in organelle movement came with the discovery of pollen tube proteins showing microtubule-gliding ability. A 90 kDa kinesin-like protein from tobacco pollen tubes was identified after screening of proteins co-pelleting with microtubules and reacting with a peptide antibody against a highly conserved region in the motor domain of the kinesin superfamily (Cai et al., 2000); the purified 90 kDa protein induced microtubules to glide in motility assays. This finding was followed by evidence that organelles from tobacco pollen tubes were capable of moving with different velocities along microtubules in vitro (Romagnoli et al., 2003). A 105 kDa organelle-associated motor was identified as functionally, biochemically, and immunologically related to kinesin. This indicated that the movement of pollen tube organelles was not exclusively actin-based but also depended on microtubule-based motors. Further support for this finding came from evidence that isolated pollen tube mitochondria moved along microtubules and actin filaments in vitro (Romagnoli et al., 2007). Movement was slow and continuous along microtubules but very fast and irregular along actin filaments. Results also indicated that pollen vesicles and mitochondria bound to identical myosins but specific kinesins, because the 90 kDa kinesin was found in association with mitochondria but not with Golgi vesicles. The sum of these elements suggested that microtubules and kinesins contributed to the disposition of mitochondria in pollen tubes. Similar results were also found in cells of Arabidopsis thaliana (Ni et al., 2005). The presence of an additional kinesin-like protein associated with Golgi bodies of Nicotiana pollen tubes (Wei et al., 2005) further sustained the model that organelles also move along microtubules and that pollen tubes may contain different microtubule-based motors. Golgi-associated kinesins are not peculiar to pollen tubes, also being found in Arabidopsis trichomes (Lu et al., 2005). Although the contribution of microtubules and microtubulebased motors may be superfluous during pollen tube growth in vitro, it is hypothesized that the role of microtubules in organelle motility may be more essential in vivo, when pollen tube growth is fastest (Joos et al., 1995). Actin-microtubule and related proteins The presence of an additional microtubule-dependent transport system suggested that actin filaments and microtubules may co-operate in the motility of organelles and vesicles during pollen tube growth. The information that the velocity of membranes along the two filamentous systems was drastically different indicated that the specific role of actin- and microtubule-motors was different. The combined use of myosin and kinesin to achieve the proper final delivery of organelles (so-called ‘functional co-operation’) Organelle movement in pollen tubes | 501 suggested (albeit not necessarily) that actin filaments and microtubules communicated in some way. While the organization of actin filaments has been described in great detail and the cortical actin fringe is acknowledged as a typical feature of growing pollen tubes (Lovy-Wheeler et al., 2005), detailed information on the distribution of microtubules in living pollen tubes is still lacking or under progress (Cheung et al., 2008). Microtubules and actin filaments are organized differently in pollen tubes but also show some similarities. Both are disposed in bundles along the main axis, with actin filaments also being present in the cytoplasm while microtubules are mostly found in the cell cortex. In the cortex, single actin filaments are usually observed to be aligned with microtubules and elements of the endoplasmic reticulum; microtubules appear to be cross-linked to actin filaments and to the plasma membrane (Lancelle et al., 1987). Close association of microtubules with actin filaments has also been shown by immunogold labelling, indicating that microtubules may act as guide elements for actin filaments or vice versa (Lancelle and Hepler, 1991). Co-localization of microtubules and actin filaments in the pollen tube cortex has also been revealed by double labelling with fluorescent probes, suggesting functional relationships with organelle movement and the organization of the cell cortex (Pierson et al., 1989). A schematic drawing of actin filaments and microtubules (based on visualization with GFP-fluorescent probes) and their hypothetical relationships in pollen tubes is shown in Fig. 2A. Although a kind of interplay between microtubules and actin filaments probably exists, its implications are enigmatic and controversial. Microtubules are probably not involved in the capacity of pollen tubes to overcome mechanical obstacles, whereas actin filaments seem very important. Conversely, maintenance of the direction of growth in vitro was susceptible to microtubule degradation whereas actin filaments were less or non critical (Gossot and Geitmann, 2007). In addition, the organization of microtubules probably depended on actin filaments, while actin filaments seemed to be independent of microtubules. These findings suggested that microtubules and actin filaments were physically connected in some way. In this context, a microtubule-associated protein, SB401, was identified in the pollen of Solanum berthaultii and shown to bind to actin filaments (Huang et al., 2007). The possible interplay between actin filaments and microtubules and the concept of ‘functional co-operation’ possibly implied the need for proteins mediating interactions between actin filaments and microtubules and their effects of their mutually dependent architecture on organelle movement. Inadequate data on microtubule-associated proteins (MAPs) in pollen tubes makes any type of realistic hypothesis even more difficult, although a search of the literature and BLAST comparison of proteins brought forth several MAP-homologous proteins expressed in Arabidopsis thaliana and putatively involved in different functions (Gardiner and Marc, 2003). MAPs have still to be demonstrated in pollen tubes and it is not clear whether they are required for organelle movement. Results of in vitro motility assays indicated that pollen tube organelles move constantly along single microtubules, suggesting that microtubule bundles may not be necessary (Romagnoli et al., 2007). The same ambiguity also concerns post-translational modification of tubulin. In addition to microtubule stabilization, acetylated and tyrosinated a-tubulins (Aström, 1992; Del Casino et al., 1993) may possibly be correlated with the activity of motor proteins and the regulation of microtubule-mediated trafficking of organelles, as proposed by Dunn et al., (2008). The effects of MAP-mediated differential density of microtubules on organelle movement is an open question. Actin, vesicles and the regulation of exocytosis/endocytosis The final fate of Golgi-derived secretory vesicles in pollen tubes is fusion with the apical plasma membrane. Relationships between endoplasmic reticulum, Golgi bodies, and vesicles have been studied using Brefeldin A, which induced subapical membrane accumulation derived from the backflow of apical vesicles and from Golgi-derived membranes. The Brefeldin-induced membrane aggregation showed periodic movements, which were not related to periodic fluctuations of Ca2+. This evidence suggests that the Golgi system is a central element in the production and recycling of vesicular materials; these activities depend on the actin cytoskeleton (Parton et al., 2003). Thus, the growth of pollen tubes is essentially based on focusing Golgi-derived vesicles that accumulate through the motor activity of the actin–myosin complex and fuse in specific sites of the pollen tube apex (polarized exocytosis). From the early work of Steer and Steer (1989), it emerged that the number of secretory vesicles in the pollen tube apex largely exceeded the number effectively required for growth, suggesting that excess plasma membrane must be recovered by other mechanisms (endocytosis). The relationship between vesicle trafficking and endo/ exocytosis is a critical aspect of pollen tube growth. Vesicle streaming and tube growth can be unplugged by Latrunculin B (Cardenas et al., 2005), suggesting that vesicle movement and fusion are separated by an undefined boundary. Thus, current discussion about vesicle motility (which is based on cytoskeleton-membrane interactions) also needs to consider the subsequent phase of vesicle fusion (which includes regulation and the dynamics of this process). Recent research indicated that exocytosis is mediated by small (400 nm) vesicles and occurs in a region 3–10 lm from the tube apex (Zonia and Munnik, 2008). This model of subapex-delimited vesicular secretion has also been suggested by other authors using spatiotemporal correlation spectroscopy and fluorescence recovery after photobleaching; secretory vesicles were shown to move towards the apex in the pollen tube cortex and accumulate close to the tip, returning through the tube centre (Bove et al., 2008). The velocity of secretory vesicles matched the speed of Golgi vesicles as measured in vitro along actin filaments and microtubules (Romagnoli et al., 2007). These findings imply 502 | Cai and Cresti Fig. 2. (A) Schematic distribution of actin filaments and microtubules in a pollen tube as shown by GFP-tagged proteins. Actin filaments are found in the cortical and central regions of the tube. They presumably arise as very short filaments from formin-containing membraneassociated proteins (only shown at the apex/subapex border); the actin fringe may represent a transition domain and a growth regiondelimiting domain; actin cables in pollen tube shanks are probably formed by bundle-forming proteins (such as villin). Microtubules exist prevalently in bundles (formed by uncharacterized proteins) in the cortical cytoplasm and their organization in the subapex is still unclear; presence of a microtubule fringe has been proposed (Lovy-Wheeler et al., 2007), but its organization and spatial relationships with the actin fringe are unknown. Microtubules probably associate with the plasma membrane through p161 (Cai et al., 2005) and, hypothetically, with actin filaments through uncharacterized proteins. Sites of origin and polarity of microtubules are not known. Monomers of actin and tubulin are probably present in the apical region. Objects are not drawn to scale. (B) Regulation of cytoplasmic streaming. This picture is derived from (A) but is essentially focused on the relationships between the signal transduction pathway and the pollen tube cytoskeleton. Ca2+ is considered to be a main effector in the regulation of organelle movement. It enters the pollen tube in the tip region (1) and generates a local increase in concentration (2), which also depends on the concerted activity of phospholipase C (PLC), phosphoinositol biphosphate (PIP2), inositol triphosphate (IP3) (3), and of Rop GTPases activity (4). The oscillatory increase in Ca2+ concentration activates the villin/gelsolin/profilin group, shifting the equilibrium of actin toward the monomeric form (5). This protein group works in combination with ADFs (not reported) and the alkaline band, the latter produced by active efflux of H+ (6). The increase in Ca2+ concentration affects the Organelle movement in pollen tubes | 503 relationships between the zone of active secretion and the actin cytoskeleton. Vesicles are known to move along the actin filaments, but the precise role of actin in the secretion mechanism is still unclear. Identification of the actin fringe in pollen tubes (Lovy-Wheeler et al., 2005) suggested the hypothesis of its role in vesicle trafficking and fusion. Since the actin fringe starts about 1–5 lm behind the apex and extends for 5–10 lm, the region of the actin fringe approximately coincides with the region of vesicle secretion. Speculatively, exocytic vesicles may traffic directly to the site of exocytosis through the actin fringe or the actin fringe may somehow label or delimit the site of exocytosis. A putative model may therefore be that accumulation of secretory vesicles in the tube apex depends on longitudinal actin filaments (Cardenas et al., 2008) while the actin fringe probably acts as a final track for conveying vesicles specifically to the site of growth. The apex and subapex of pollen tubes may also possibly be the region of actin assembly. New actin filaments probably arise from membraneassociated protein complexes containing formin. Overexpression of formin in pollen tubes induces the formation of actin filaments, arising from the plasma membrane, in the cytoplasm. Formin-dependent actin polymerization is therefore important for polar growth of pollen tubes (Cheung and Wu, 2004). Actin filaments almost certainly arise from the plasma membrane as very short filaments and then organize into the actin fringe, which is therefore a transition domain between growing actin filaments in the apex/ subapex and longitudinal actin cables. The actin fringe may also act as a functional domain to delimit the region of growth. Little is known about the organization of microtubules in the pollen tube apex. Despite evidences that microtubules are absent from the apical domain, GFPlabelled EB1 indicated that microtubules may lie closer to the pollen tube tip than previously thought (Cheung et al., 2008). The presence of microtubules in close proximity to the tip may also depend on the bending features of pollen tubes (Foissner et al., 2002) suggesting a role of microtubules in the guiding process. The presence of a microtubule fringe in the subapex of pollen tubes was suggested by microtubule labelling with fluorescent taxol (Lovy-Wheeler et al., 2007) but the implications for growth and relationships with the actin fringe are unclear. The exocytotic event is closely related to the opposite process of endocytosis, which most likely occurs in two distinct regions of the pollen tube, along the apex, and in the distal region (Zonia and Munnik, 2008; Moscatelli et al., 2007). In this context, hydrodynamic flux was assumed to regulate exocytosis and endocytosis because hypotonic treatment (and thus cell swelling) stimulated exocytosis and reduced endocytosis, while hypertonic treatment (and cell shrinking) stimulated endocytosis but decreased exocytosis. The growth rate of pollen tubes is, consequently, the result of fine-tuning of the apical volume of pollen tubes, which influences the activity of endocytosis (at the apex and along the distal tube) and exocytosis (in the subapical region) and thus defines the boundary of the growth zone. Visualization of endocytosis by fluorescent phospholipids (Lisboa et al., 2008) indicated that the plasma membrane retrieved was re-integrated into the internal membrane pool and redistributed at the base and apex of the pollen tube. Depolymerization of actin filaments abolished plasma membrane retrieval, whereas oryzalin had no effect, suggesting that actin filaments but not microtubules may be responsible for the endocytotic process. Exocytotic/endocytotic activity is closely related to the process of pollen tube growth, which is, in turn, regulated by several external cues and intracellular messengers. The latter presumably mediate the effects of growth regulators by affecting the rate of vesicle fusion and the delivery of vesicles to the apical domain through modulation of the actin cytoskeleton. Among the different molecules that establish signal transduction, Ca2+ was suggested to play a key role in vesicle-dependent secretion of cell wall components, while Rop-GTPases may play a crucial role in the fusion of secretory vesicles and endocytosis (Camacho and Malho, 2003). Other authors have reported that Rop-GTPase activity influences F-actin dynamics in the pollen tube tip (Fu et al., 2001). Such activity probably depends on two counteracting downstream pathways involving Rop-interactive CRIB-containing proteins (RICs), which contain a binding motif to Cdc42/Rac (Wu et al., 2001). Specifically, it was found that RIC4 (overexpression of which promoted actin assembly) determined the accumulation of secretory vesicles but inhibited exocytosis; on the other hand, disassembly of actin by RIC3 was required for exocytosis (Lee et al., 2008). This again showed that the process of vesicle transport in the tube tip and exocytosis depend on distinct mechanisms and that fine-tuning of the equilibrium between F- and G-actin is essential for tube growth. The activity of Rop-GTPases is probably mediated by protein kinase activity. The presence of type 2A protein phosphatase inhibitors had dramatic effects on the cytoskeleton (randomly oriented actin filaments and microtubules, alteration of actin bundle alignment, and disassembly of microtubules) (Foissner et al., 2002), which were similar to those observed after overexpression of Rop1 (Fu et al., 2001). Phosphatidylinositol 4,5-bisphosphate, inositol 1,4,5-trisphosphate, and phosphatidic acid have also been suggested to participate in the control of vesicle fusion/retrieval and to modulate the motor activity of myosin (7) through the associated calmodulin light chain (not shown). Regulation of actin filaments and inactivation of myosin prevent large-sized organelles from entering the apical domain (8). Secretory vesicles also move along actin filaments using myosin activity (9) but it is not known whether this myosin is affected by variations in Ca2+ concentrations. Vesicles enter and accumulate in the apical domain (10) and fuse with the plasma membrane (11) or turn back toward the grain (12). Nothing is known about the regulation of microtubules, but they could be regulated by phospholipase D (not shown). The polymerization state of actin filaments may also regulate microtubule dynamics (13). Objects are not drawn to scale. 504 | Cai and Cresti organization of actin filaments (Monteiro et al., 2005a). Expression of a phospholipase C (PLC) in pollen tubes of Petunia inflata raised the apical Ca2+ gradient, which disorganized the actin filaments (Dowd et al., 2006). This finding suggested that PLC hydrolysed phosphatidylinositol 4,5-bisphosphate to the signalling molecules inositol 1,4,5trisphosphate and diacyl glycerol (DAG). Inositol 1,4,5trisphosphate is thought to regulate intracellular Ca2+ concentrations, which are known to affect actin organization. The negative effects of expression of PLC were reversed by treatment with low concentrations of the actin drug Latrunculin B, suggesting that PLC alters actin structure in growing tube tips and restricts growth to the very tip of pollen tubes. Overexpressed PLC also inhibited tube growth and accumulated laterally at the pollen tube tip plasma membrane mimicking the distribution of phosphatidylinositol 4,5bisphosphate. Diacylglycerol (DAG) also showed a localization similar to that of phosphatidylinositol 4,5-bisphosphate (Helling et al., 2006). Phospholipase D (PLD) and its product phosphatidic acid (PA) also play a role in the polarization of pollen tubes and their activity can be blocked by primary alcohols, like 1-butanol. When inhibited by 1butanol, pollen tube growth can be restored by taxol treatment, indicating that microtubules may be one target of PLD activity. This hypothesis is supported by the finding that PLD is linked to microtubule dynamics (Dhonukshe et al., 2003). The role of microtubules (if any) in vesicle targeting and fusion is unclear. Although kinesin-related proteins have been localized in apical vesicles (Tiezzi et al., 1992), disorganization of microtubules does not influence pollen tube growth. Nevertheless, the evidence that oryzalin treatment affected pulsatory growth (Geitmann et al., 1995) and directional growth of pollen tubes (Gossot and Geitmann, 2007) indirectly suggests that microtubules may interfere with the growth mechanism. In addition, the finding that plant phospholipase D associates with membranes and microtubules (Gardiner et al., 2001), coupled with the effects of protein phosphatase inhibitors on pollen tube microtubules (Foissner et al., 2002), indicates interplay between the signal transduction pathway and the microtubule cytoskeleton. On the basis of the above information, it is possible to postulate relationships between the actin cytoskeleton and the endo/exocytotic process (Fig. 2B). Ca2+ is presumed to enter pollen tubes at the very tip (1) increasing its local concentrations (2) (Messerli and Robinson, 1997). Variation in Ca2+ concentrations is also thought to increase after stimulation of the phospholipase C (PLC), phosphoinositolbiphosphate (PIP2) and inositol-triphosphate (IP3) pathway (3) (Monteiro et al., 2005b). Rop-GTPases are also believed to regulate Ca2+ concentrations (4) (Hwang et al., 2005). Local oscillatory increases in Ca2+ concentration are considered to activate the protein group consisting of villin, gelsolin, and profilin, shifting the equilibrium towards Gactin and preventing polymerization/elongation of actin filaments (5) (Yokota et al., 2005). This protein group probably works in combination with ADFs (not reported in the figure) and the alkaline band, which is probably produced by active discharge of H+ (6) (Chen et al., 2002). An increase in Ca2+ concentrations also affects the motor activity of myosin (7) through the associated calmodulin light chain (not shown) (Yokota et al., 1999). Regulated elongation of actin filaments along with the inactivation of myosin probably prevents large-sized organelles from entering the apical domain (8) but allows them to turn back. Secretory vesicles also move along actin filaments using the motor activity of myosin (9). It is not known whether vesicle-associated myosin is affected by increases in Ca2+ concentrations; in any case, vesicles penetrate and accumulate in the apical domain (10) where they fuse with the plasma membrane (11). Vesicles that do not fuse may return to the subapical region, together with endocytic vesicles (12); this tip-to-subapex movement may rely on actin and myosin. Nothing is known about the regulation of microtubule activities; speculatively, they may be regulated by phospholipase D (not shown), or the polymerization state of actin filaments may regulate microtubule dynamics (13) (Gossot and Geitmann, 2007; Poulter et al., 2008). Conclusions and future prospects Advances in the last 20 years of research have offered important new insights into the functions of actin filaments and microtubules in organelle transport inside pollen tubes. Since the early evidence of organelle movement along actin filaments, current research is now focusing on the relationships between actin/tubulin dynamics, organelle/vesicle transport, and pollen tube growth. A great amount of data indicates that actin filaments are the main tracks of organelle and vesicle transport; actin filaments support cytoplasmic streaming in the basal domain and accumulation of vesicles in the tip domain. External signals appear to be critical for redirecting tip growth when the pollen tube has to follow signals from female tissues; external signals presumably modulate the activity of motor proteins and organization of actin filaments. The role of microtubules is still unclear, although their function could be more prominent for the growth of pollen tubes in vivo. Whether actin and the microtubule cytoskeleton co-operate in motor activity is still unclear and there are major challenges to be solved in the near future. Some of these questions are listed below. (1) Do organelles of the same class move independently? The motility pattern of mitochondria, Golgi bodies, endoplasmic reticulum, and vesicles needs to be clarified. We do not know whether single organelles move independently of the others or ignore whether or not the trafficking of each organelle class depends on that of other classes. Do specific motor receptors exist? Does this imply involvement of multiple myosins? (2) Is organelle transport regulated by functional cooperation? It is current opinion that organelle transport in eukaryotic cells is achieved by regulating the activities Organelle movement in pollen tubes | 505 Fig. 3. Hypothetical interactions of pollen tube organelles with both actin filaments and microtubules. Speculatively, organelles move rapidly along actin filaments using myosin as motor protein; organelles may be stopped in specific sites of the pollen tube for finer positioning through dynamic interaction of kinesin with microtubules. Microtubule-actin interacting proteins are hypothetical; they may help organize cytoskeletal architecture but their role in organelle transport is hard to decipher. The different sized arrows indicate the approximate intensity of motility along actin and microtubules. Objects are not drawn to scale. of motor proteins of different families (Goode et al., 2000). This model has been the subject of relatively little study in plant cells, although there is evidences that myosin and kinesin may ultimately be required for organelle trafficking (Wei et al., 2005). Since organelles interact dynamically with actin filaments and microtubules, the pollen tube may be an excellent model for studying this process; unfortunately, we do not know how many myosins and kinesins are present in pollen tubes, and how motor proteins co-operate in organelle and vesicle movement. A speculative basic model is reported in Fig. 3. (3) What is the relationship between external signals, motor-mediated organelle/vesicle delivery, and pollen tube growth? Delivery of vesicles to the apical region supports pollen tube growth, which is under the control of external signals from female tissues (Herrero, 2001). Although indications suggest that myosin activity in pollen tubes is controlled by Ca2+ concentrations (Yokota et al., 1999), no direct evidence as yet links the signal transduction pathway to motor activity. Does control of motor activity simply rely on the regulation of cytoskeletal filaments? (4) What is the exact role of microtubule motors in organelle/vesicle transport? There is evidences that pollen tube organelles and vesicles move along microtubules (Romagnoli et al., 2007) and that microtubules may affect the pulsed directional growth of pollen tubes (Gossot and Geitmann, 2007; Geitmann et al., 1995); however, more information is needed on the role of microtubule motors in these processes. Gene sequences of kinesins and inhibition of expression by iRNA may provide some answers. (5) What is the effect of localized vesicle delivery on cell wall construction in pollen tubes? In order to appreciate the contribution of the cytoskeleton to the pollen tube shaping process, it is essential to determine the secretion pattern of cell wall-synthesizing enzymes, their activation scheme, and dependence on exocytosis and actin filaments or microtubules. Is proper organization of the cytoskeleton required for pollen tube cell wall construction? How do these activities depend on molecular motors? (6) What is the relationship between the actin fringe, the alkaline band, and secretion of vesicles? This feature merits considerable attention because the actin fringe (presumably under the control of the alkaline band) is a key element in vesicle secretion. Does the actin fringe define the secretion zone or does it drive secretory vesicles to the secretion site? 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