Pollen tube growth: a delicate equilibrium between secretory and. Membrane recycling in pollen tube growth

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1 1 Title: Pollen tube growth: a delicate equilibrium between secretory and 2 endocytic pathways Running title: Membrane recycling in pollen tube growth Authors: Alessandra Moscatelli 1 * and Aurora Irene Idilli 1 1 Dipartimento di Biologia L. Gorini, Università degli Studi di Milano, Via Celoria 26, 20133, Milano (Italy) *Author for correspondance. Tel: ; Fax: : ; alessandra.moscatelli@unimi.it. Supported by the FIRST Research Program of the University of Milano 1

2 32 33 ABSTRACT Although pollen tube growth is a prerequisite for higher plant fertilization and seed production, the processes leading to pollen tube emission and elongation are crucial for understanding basic mechanisms of tip growth. It was generally accepted that pollen tube elongation occurs by accumulation and fusion of Golgi-derived secretory vesicles (SVs) in the apical region, or clear zone, where they were thought to fuse with a restricted area of the apical plasma membrane (PM), defining the apical growth domain. Fusion of SVs at the tip reverses outside cell wall material and provides new segments of PM. However, electron microscopy studies have clearly shown that the PM incorporated at the tip greatly exceeds elongation and a mechanism of PM retrieval was already postulated in the mid of nineteenth century. Recent studies on endocytosis during pollen tube growth showed that different endocytic pathways occurred in distinct zones of the tube, including the apex, and led to a new hypothesis to explain vesicle accumulation at the tip, namely that endocytic vesicles contribute substantially to V- shaped vesicle accumulation in addition to SVs and that exocytosis does not involve the entire apical domain. New insights suggested the intriguing hypothesis that modulation between exo- and endocytosis in the apex contributes to maintain PM polarity in terms of lipid/protein composition and showed distinct degradation pathways that could have different functions in the physiology of the cell. Pollen tube growth in vivo is closely regulated by interaction with style molecules. The study of endocytosis and membrane recycling in pollen tubes opens new perspectives to studying pollen tube-style interactions in vivo Key words: pollen tube, endocytosis, exocytosis, membrane recycling, polarized growth

3 The Angiosperm male gametophyte is characterized by a pulsate growth Sexual reproduction is a crucial step in the life cycle of plants. In Angiosperms, the process starts with pollination or deposition of the male gametophyte on the stigma surface. After the pollen-stigma interaction, pollen grains hydrate and germinate, producing a cell protrusion known as pollen tube (Taylor and Hepler 1997). Pollen tubes grow in the style and function as biological channels to convey generative/sperm cells to the embryo sacs for fertilization. While the correct mechanism of pollen tube growth is a prerequisite for sperm cell transport, pollen tubes are model cells for understanding basic mechanisms regulating tip growth (Taylor and Hepler 1997; Hepler et al. 2001; Krichevsky et al. 2007). Pollen tube growth results from extreme secretory activity, inducing growth rates that in Nicotiana tabacum (L.) may reach nm/sec (Hepler et al. 2001). Much data supports the idea that Golgi-derived SVs fuse with a restricted area of the pollen PM, establishing a PM growth domain where the pollen tube emerges. Ultrastructural studies have shown that many organelles are not uniformly distributed in the pollen tube cytoplasm. The tip region shows distinct zoning: an apical region or clear zone (5-10 μm from the tip) hosts typically a V-shaped vesicle accumulation, a subapical organelle-rich zone, a nuclear zone and a vacuolated zone that may extend toward the grain (Li et al. 1997; Taylor and Hepler 1997; Hepler et al. 2001; Cheung and Wu 2008). As the pollen tube grows, callose deposition occurs in the subapical regions and progressively isolates portions close to the grain from the cytoplasmic regions, containing the biosynthetic machinery and the male germ unit, in the apical regions. Pollen tube may have an oscillating or pulsate pattern of growth (Li et al. 1997). Oscillating growth, typical of fungal hyphae and pollen tubes of Angiosperm species such as Lilium longiflorum (L.), is characterized by fables changes in growth rate. On the other hand, pulsate growth, typical of Nicotiana tabacum (L.), is characterized by slow phases of growth followed by sudden steps (pulses) (Picton and Steer 1982). Studies on cell wall deposition suggest that slow periods of growth coincide with 3

4 increases in cell wall thickness due to deposition of pectins and arabinagalactan proteins (AGPs), while during pulses a thinner cell wall is formed (Li et al. 1994; Li et al. 1995; Li et al. 1997). While in Lilium pollen tubes, wall components are uniformly secreted along the cell, in tobacco pollen tubes the pulsate growth induces the formation of rings containing AGPs, the period of which corresponds to that of the pulses (Li et al. 1995). The mechanisms enhabling the pulsate growth are not completely clear. It has been claimed that the balance between turgor pressure and cell wall rigidity of pollen tubes plays an important role in regulating this process (Messerli and Robinson 2003). However, since measurement of turgor showed that it does not change in time (Benkert et al. 1997) it has been proposed that the mechanism of oscillatory growth could be primarily a consequence of the mechanical properties of the cell wall and periodic ion fluxes in the cytosol (Holdaway-Clarke and Hepler 2003). The importance of pectin secretion in regulating the cell wall plasticity of pollen tubes required for extension in the apical region has been demonstrated. During the growth process pectins elaborated in the Golgi apparatus are secreted in their methyl-esterified form by fusion of SVs in specific regions of the tip (Li et al. 2002; Krichevsky et al. 2007). Homogalcturan, the linear polymer of α-1,4-d-galacturonic acid, the principal component of pectins, is progressively de-esterified by pectin methyl esterases (PMEs), secreted, together with pectins, by the same SVs (Li et al. 2002). In the pollen tube, some PMEs have been observed to be co-expressed with several PME inhibitors (PMEIs), suggesting that the concerted action of PMEs and PMEIs plays an important role in regulating cell wall plasticity of the tube (Bosch and Hepler 2005). Recent studies showed a polarized localization of PMEI isoforms in the tip PM of tobacco pollen tubes and suggested that the endocytic process is involved in removing these proteins from the subapical regions and in recycling them to the tip as the pollen tube elongates, thus maintaining cell wall plasticity at the apex (Rockel et al. 2008). Pectin de-esterification 116 exposes carboxylic groups and favours their binding with Ca 2+ ions, leading to an increase in cell wall rigidity. The function of PMEs during cell wall formation in plants requires a close regulation of enzyme activity; in fact PME activity is modulated by 4

5 chemical characteristics of the environment, especially changes in ph. A localized reduction in ph, due to de-esterification, may activate enzymes that modify the cell wall plasticity such as pectate lyase and polygalacturonidase (Catoire et al. 1998). The essential function of PMEs in cell wall regulation is especially explicit in vivo when changes in the direction of tube growth require drastic changes in cell wall plasticity. Pollen tubes of Arabidopsis mutants, with a 20% reduction in PME activity, show inhibited growth efficiency and altered the cell wall mechanical properties (Jiang et al. 2005). Ion fluxes across the PM and could also help explain the pulsate growth of pollen tubes (Holdaway-Clarke and Hepler 2003). Critical constraints for pollen tube growth are mechanisms that maintain Ca 2+ and H + ion gradients in the tip region of the tube (Hepler et al. 2001; Robinson and Messerli 2002; Holdaway-Clarke and Hepler 2003; Feijo et al. 2004). In lily pollen tubes, the Ca 2+ concentrations in the apex (up to 5 μm from the apical PM) can reach values of 10 μm and as low as 200 nm just behind the tip (20 μm from the apical PM) (Pierson et al. 1994; Malhò et al. 2000). Recent studies showed that the dissipation of the Ca 2+ gradient inhibits pollen tube growth. On the other hand, reorientation of the Ca 2+ gradient in the tip region leads to a parallel reorientation of the direction of growth (Pierson et al. 1994; Malhò et al. 2000; Holdaway-Clarke and Hepler 2003). The entrance of Ca 2+ ions seems to occur by stretch-activated Ca 2+ channels in the apical PM, during the extension phase of the growth process (Dutta and Robinson 2004). A significant contribution to apical Ca 2+ homeostasis is also played by subapical Ca 2+ pumps that lower cytosolic Ca 2+ immediately behind the tip, contributing to maintain the steep Ca 2+ gradient in the apex. A pharmacological approach inhibiting 142 Ca 2+ channels demonstrated a direct relationship between ion concentration and oscillatory growth (Geitman and Cresti 1998). Some observations also showed that the Ca 2+ gradient at the tip determines where SVs fuse with the PM: a high concentration of Ca 2+ helps maintain actin filaments (AFs) dynamic in this area and inhibits cytoplasmic streaming of SVs in the tip, allowing them to fuse with the apical PM (Taylor and Hepler 1997; Hepler et al. 2001; Camacho and Malhò 2003; Cardenas et al. 2008). When the 5

6 Ca 2+ gradient is dissipated, AFs extend to the apical PM and acquire the ability to transport organelles in the clear zone, leading to growth inhibition (Pierson et al. 1994). Interestingly, recent evidence on the dynamics of SV accumulation in the tip region indicated that the density of SVs in the tip region oscillates with the same period as pulsate growth (Parton et al. 2001). This means that peaks of growth coincide with decline in accumulation of V-shaped fluorescence in the apex, which suggests massive periodic fusion of SVs with the apical PM during rapid growth. Moreover, in addition to 155 exchange with the extracellular matrix, Ca 2+ release from internal membranous compartments could also contribute to the dynamics of this ion, in view of the abundance of smooth endoplasmic reticulum (ER) in the clear zone and in subapical regions of the tube (Franklin-Tong et al. 1996). Based on recent data on cell wall deposition and periodic cytosolic Ca 2+ influx, a new model for pulsate growth, combining different factors such as turgor pressure, regulation of cell wall plasticity, Ca 2+ dependent exocytosis and endocytic membrane recycling was recently developed (Kroeger et al. 2008). Other ion transporters play important roles in pollen tube growth. The pollen tube apex appears to be slightly acidic, since cytosolic ph is generally one unit lower than that of the alkaline zone, situated immediately behind the clear zone (Feijo et al. 1999). Substantial proton influx in the apical region and parallel proton efflux in the subapical regions have been demonstrated and presumably help to maintain the H + gradient in the tip (Feijo et al. 1999). Obviously, localized ion fluxes imply restricted localized presence of proteins involved in their transport. Pollen tubes expressing GFP-H + - ATPase revealed that this pump is mainly localized in the flanks of the tip, supporting the idea that a proton influx occurs in the tip while a proton efflux is maintained in the subapical regions (Certal et al. 2008). An influx of K + ions at the tip has also been observed, and analysis of Arabidopsis mutants sustained the hypothesis that this ion could play a role in pollen tube growth and in the efficiency of male gamete transport (Fan et al. 2001). Analogously, Cl - ion 6

7 dynamics is reputed to regulate cytoplasmic hydrodynamics, critical for tube growth (Zonia et al. 2002). It therefore seems that coordinated cell wall deposition and tube elongation require the concerted action of ion fluxes, AFs organization, exo-endocytosis and GTPaseregulated signalling pathways (Lin et al. 1996; Kost et al. 1999; Fu et al. 2001; Zonia et al. 2002; Robinson and Messerli 2002; Kost 2008). Most of these factors regulating pollen tube growth help maintain a polarized distribution of proteins/lipids in the PM as the pollen tube elongates, supporting the idea that an equilibrium between the insertion and retrieval of proteins/lipids plays an important role during pollen tube growth (Kost et al. 1999; Potocky et al. 2003; Helling et al. 2006; Dowd et al. 2006). Polarized protein/lipid composition of the PM implies selective removal/recycling of molecules regulating the polarity of cells and plant development, such as growth regulators and ion transporters between different PM domains (Baluska et al. 2003; Murphy et al. 2005; Geldner and Jurgens 2006). This process is especially significant for highly specialized cells such as pollen tubes, which only grow at the apex by an extreme form of polar growth, known as tip growth (Hepler et al. 2001). During pollen tube growth, the high rate of SV fusion at the tip largely exceeds PM extension; this observation led to the hypothesis that an internalization mechanism could be involved in regulating membrane internal economy (Steer and Steer 1989; Derksen et al. 1995). On the other hand, recent evidence showed that targeted exocytosis and endocytosis are also regulated by phosphoinosides (IP 3 and PIP 2 ) and phosphatidic acid (PA) through modulation of AF organization and SV fusion Pollen tube growth depends on polarized exocytosis Pollen tubes follow a precise, regulated tip growth pattern based on transport and accumulation of post-golgi SVs at the extreme tip, where on fusion with a restricted region of the PM, they secrete cell-wall material and provide new segments of PM (Taylor and Hepler 1997; Hepler et al. 2001; Krichevsky et al. 2007). As the pollen tube grows, actomyosin-dependent reverse fountain-like cytoplasmic streaming drives SVs 7

8 forward to the apex where cytoplasmic streaming does not occur (Hepler et al. 2001). Reverse fountain-like cytoplasmic streaming is thought to be responsible for V-shaped vesicle accumulation in the tip region, where SVs fuse with the apical PM (Fig. 1, orange arrows). Electron microscope observations after rapid freeze fixation and substitution of Lilium pollen tubes showed that morphologically different vesicles were present in the tip: a pool of small vesicles, considered to be SVs, were localized at the very tip, close to the PM while vesicles of greater diameter were observed in the clear zone, distal to the apical PM (Lancelle and Hepler 1992). Two-directional movements of individual vesicles were recently analysed using computer-assisted video microscopy (Wang et al. 2005). Movement of vesicles toward the apex occurs in cortical regions of the tube and proceeds up to 7-10 μm from the tip, where some of the vesicles invert their trajectory and are transported backward the grain, by streaming in the central zone of the tube cytoplasm (Hepler et al. 2001; Parton et al. 2001; Wang et al. 2005) (Fig. 1, green arrows). However, a substantial portion of vesicles are captured and maintained in the apical region, leading to the typical V-shaped accumulation or clear zone (up to 10 μm from the tip) (Fig. 1) where it is generally accepted that only Brownian movement of vesicles is observed (de Win et al. 1999). Corrected SV fusion with the PM, cell wall deposition and tube elongation require coordination of GTPase-regulated signalling pathways, exo-endocytosis, cytoskeleton organization and ion fluxes. Small Rho-GTPases have been identified in all eukaryotic cells and participate in a wide variety of processes as different as AFs organization and membrane trafficking (de Curtis 2008). The best characterized function of Rho- GTPases and the Rho homologues Rac and Cdc 42 is the regulation of AF organization in different cell types (Hall 1998), and it has also been shown that rearrangement of cortical AFs regulates secretion in animal cells (Aunis 1998). On the other hand, Cdc42 is thought to control the AF dynamics at the cell surface, allowing polarized secretion and bud growth in yeast (Hirata et al. 1998). Although this protein superfamily comprises Rho, Rac e Cdc42 isoforms (Takai et al. 2001), plant Rho-GTPases are represented by a small group of proteins that seems to be related to Rac of animals. We 8

9 therefore refer to plant RhoGTPases as Rac/Rop-GTPases (Winge et al. 1997). In physiological conditions Rac/Rop-GTPases in pollen tubes are localized in a restricted area of the apical PM, a region which has long been considered the whole growth domain, and are key regulators of polar cell expansion. (Kost et al. 1999; Zheng and Yang 2000; Fu et al. 2001; Gu et al. 2004). As in other systems, pollen tube Rac/Rop appears to coordinate AF organization, membrane traffic and Ca 2+ signalling, and to controls lipid-based signal transduction pathways (Kost et al. 1999; Fu et al. 2001; Gu et al. 2003; Gu et al. 2004; Kost 2008). In physiological conditions, the actin cytoskeleton consists of thick longitudinally oriented actin cables in the shank of the tube and fine filaments close to the tip (Hepler et al. 2001, Cheung and Whu 2008). The absence of actin bundles in the apex has been considered a prerequisite for fusion of SVs with the apical PM domain, since drugs upsetting AF dynamics at the tip, and the consequent formation of long AFs determine the growth inhibition and loss of cytoplasmic zoning (Pierson et al. 1994). Different models have been proposed to explain participation of the actin cytoskeleton in polarized growth. A model proposed that AFs and cell wall help maintain the turgor pressure by a Ca 2+ -dependent mechanism. It has also been proposed that the force generated by actin polymerization could promote the cell extension phase (Picton and Steer 1982). Another model proposed that fine AFs in the tip region function as a filter to capture SVs that will form the typical V-shaped accumulation before their fusion with the PM (Heslop-Harrison and Heslop-Harrison 1990). Although more recent data showed that AFs in the tip region are dynamic and that different actin structures can be observed during pulsate growth in time lapse experiments, actin bundles were never observed in the apex of normally growing pollen tubes (Fu et al. 2001). In tobacco, an actin ring has been reported in the subapical region of pollen tubes expressing GFP-talin (Kost et al. 1998). A collar of F-actin, 5 μm behind the tip, and a dense meshwork of AFs in the collar region was observed in chemically fixed pollen tubes of different species (Gibbon et al. 1999). The subapical actin structures (5 μm from the tip) seem to be composed of fine AFs distinct from the 9

10 actin cables associated with cytoplasmic streaming (Hepler et al. 2001). The shifting of actin cables, typical of the shank of the tube, and the AFs arrays in the apical-subapical regions of the tube seem to involve the function of the alkaline band. The alkaline band in pollen tubes was postulated to enhance the activity of actin depolymerization factors and shift actin toward its monomeric form. The steep-focused Ca 2+ gradient and the alkaline band are therefore critical determinants of AF polymerization state and reverse streaming (Cardenas et al. 2008). The observation that low concentrations of latrunculin B disrupt the subapical fine AFs without affecting tube growth and cytoplasmic streaming suggests that subapical actin arrays either target SVs to the tip or maintain cytoplasmic organization (Gibbon et al. 1999). Studies using pollen specific Arabidopsis thaliana Rac/Rop1 in tobacco pollen tubes showed that it localized in the apical dome of the PM where it acted as central switch in the control of tip growth. Blocking Rop signalling by expression of dominant negative Rac/Rop 1 mutant caused inhibition of pollen tube growth, whereas constitutively active Rop1 mutant induced isotropic growth (Kost et al. 1999). Analogously, expression of a constitutively active form of Arabidopsis thaliana Rac/Rop5 in tobacco pollen tubes apparently induced actin polimerization and actin bundle formation in the tip and caused pollen tubes to form balloons, suggesting complete loss of polarity (Kost et al. 1999). By contrast, AFs in pollen tubes expressing a dominant negative form of Rac/Rop5 were finer and less organized and pollen tube growth was strongly inhibited (Kost et al. 1999). Further studies using the GFP-tagged actin binding domain of mouse talin in tobacco pollen tubes showed a dynamic form of tip-localized F-actin, called short actin bundles (SABs), and provided the first direct evidence linking Rho-GTPases to actin organization in the control of cell polarity in plants. The dynamic of SABs during polar growth is regulated by Arabidopsis thaliana Rac/Rop1. When overexpressed, Rac/Rop1 induces transformation of SABs into a network of fine filaments and formation of transverse actin bundles behind the tip, leading to depolarized growth (Fu et al. 2001). As a consequence, SVs fuse with a wider zone of PM, determining swelling of the apical region (Fu et al. 2001). To 10

11 contrary, pollen tube elongation was strongly inhibited by expression of a null mutant of Rac/Rop1-GTPase, that seemed to block apical secretion (Fu et al. 2001). These changes were due to ectopic localization of Rac/Rop1 in the apical region of the PM and were suppressed by overexpression of a guanine dissociation inhibitor which removed ectopically localized Rac/Rop1 (Fu et al. 2001). Phosphatidylinositol monophosphate kinases which synthesize phosphatidyl inositol 4,5-bisphosphate (PIP 2 ) have been identified as Rac/Rho effectors in animal cells (Hartwig 1995). Synthesis of PIP 2 allows Rho-GTPases to control targeted exocytosis by regulating membrane trafficking and to modify the AF organization by interacting with acting-binding proteins (Van Aelst and D Souza-Schorey 1997; de Curtis 2008). Hay and coworkers (1995) suggested that PIP 2 regulated organization of AFs and polarized exocytosis by altering the lipid composition of membrane microdomains and/or by recruiting proteins that mediate membrane fusion. Therefore, localized synthesis of PIP 2 and other membrane lipids was proposed as a key step in the organization of signalling (Irvine 1998) and as observed in animal cells, PIP 2 may be the effector of Rac/Rop in pollen tube (Kost et al. 1999). In addition to acting as effector, PIP 2 may also function as a precursor for the generation of other signalling molecules at the tip. Hydrolysis of PIP 2 by phospholipase C (PLC), which results in the formation of 1,4,5-trisphosphate (IP 3 ) and diacyl glycerol (DAG), followed by IP 3 -dependent Ca 2+ release from internal stores, plays a crucial role in regulating Ca 2+ homeostasis (Yang and Kazanietz 2003). This pathway has been shown to have an essential function in poppy pollen tube elongation: 313 PLC-induced hydrolysis of tip-localized PIP 2 and IP 3 -mediated Ca 2+ release in the cytoplasm may be involved in establishing the Ca 2+ gradient (Franklin-Tong et al. 1996). Recent studies showed that PIP 2 and PLCs maintain a differential position in the PM as the pollen tube grows and that endocytosis and membrane recycling play a central role in regulating the targeted localization. A specific isoform of PLC (Nt PLC3) has been identified in tobacco pollen tubes, where it appears to be constantly targeted in the flanks of the tip and never accumulates at the extreme apex (Helling et al. 2006). PLC3 was hypothesized to limit the spread of PIP 2 to the subapical regions of the tip while 11

12 generating DAG laterally at the pollen tube tip PM. DAG has been observed to be continuously retrieved from subapical parts of the PM and reinserted in the apical domain by endocytic membrane recycling (Helling at al. 2006). In a parallel study the dynamics and function of another isoform of PLC (PLC1) were revealed: it also localizes in regions limiting the tip domain, in a pattern complementary to that of PIP 2 (Dowd et al. 2006). It therefore appears that the polarized targeting and endocytic cycling of PLCs plays a key role in maintaining the proper lipid composition of the tip growth domain and in promoting polarized exocytosis. PA has also been shown to participate in regulation of exo-endocytosis. PA is a product of phospholipase D activity and can also arise as the end product of PIP 2 hydrolysis. PA promotes membrane curvature and formation of SVs and has a crucial role in the structural integrity of the Golgi apparatus (Sweeny et al. 2002) and in cytoskeletal reorganization (O Luanaigh et al. 2002). In pollen tubes, decreasing the availability of 334 PA (with butan-1-ol) disrupts the Ca 2+ gradient and transport and accumulation of vesicles in the apical region (Monteiro et al. 2005). PIP 2, IP 3 and PA therefore seem to exercise a concerted action modulating endo- and exocytosis and actin organization playing a key role in the establishment and maintenance of polarity. For a long time it was accepted that the clear zone was entirely composed of SVs and that the whole apical PM was the site of exocytosis (Taylor and Hepler 1997; Hepler et al. 2001). Recently, FM-dyes (as FM4-64 and FM1-43) made it possible to dissect vesicle trafficking in plant cells and to follow exocytic and endocytic pathways in vivo (Bolte et al. 2004). This approach revealed that secretory and endocytic vesicles both contribute V-shaped apical accumulation (Moscatelli et al. 2007; Zonia and Munnick 2008; Bove et al. 2008) (Fig. 1) and that endocytosis could be important both for maintaining membrane homeostasis in the cell and for the polarized distribution of proteins and lipids on the PM

13 Endocytosis and membrane recycling Endocytosis is involved in many cell processes in eukaryotic cells, such as nutrient uptake, down-regulation of PM receptors, and PM recycling and signalling. Mechanisms of endocytosis involving clathrin-dependent (CD) or clathrin independent (CI) internalization have been characterized and shown to coexist in animal cells (Baba et al. 2001; Nichols and Lippincot-Schwartz 2001; Kirkham et al. 2005) and plants, where receptor-mediated internalization (Holstein 2002; Ortiz-Zapater et al. 2006), phagocytosis (Son et al. 2003) and fluid phase endocytosis (Baluska et al. 2004) have been observed. Moreover, plant structural sterols and lipid rafts (Mongrand et al. 2004; Lefebvre et al. 2007) suggest that endocytic processes, similar to fluid phase or caveolae/lipid raft-mediated endocytosis in animals, may also be active in plants. In animals, vesicles derived from the PM during receptor-mediated (CD) endocytosis are delivered to a first compartment called early endosome (EE), a tubulovesicular structure that was characterized by specific sets of Rab-GTPases (Gruenberg 2001; Zerial and McBride 2001). This compartment is regarded as a sorting station for internalized molecules. Understanding of the mechanisms regulating endocytosis in plants lags far behind that of animals and is mostly concentrated on CD internalization. Studies of plant endocytosis using cationized ferritin as probe have revealed the ultrastructure of some membraneous compartments involved in internalization and trafficking during receptormediated endocytosis (Tanchak et al. 1988; Samuels and Bisalputra 1990; Fowke et al. 1991). In plants, the ultrastructure of EEs is still matter of debate, although an intense recycling pathway to the PM has been detected using styryl FM dyes and charged nanogold as probes (Ueda et al. 2001; Onelli et al. 2008). Although plants have the common basic molecular machinery that regulates membrane trafficking in other eukaryotes (Holstein 2002; Otegui and Spitzer 2008), they have evolved specialized molecular and structural mechanisms. A good example of functional divergence is the endosomal RabGTPase family. Rab proteins coordinate important stages of membrane trafficking (Molendijk et al. 2004). In mammals, Rab11 and Rab4 GTPases are associated with recycling endosomes. While Rab11 orthologs constitute the largest 13

14 subgroup of plant RabGTPases (Ueda and Nakano 2002; Vernoud et al. 2003) and was shown to play a role in targeting of transport vesicles at the tip of tobacco pollen tube (de Graaf et al. 2005), no Rab4 orthologs have been identified in plants (Rutherfod and Moore 2002). The Rab5 members regulate early endosomal functions in animal cells. The Arabidopsis genome encodes three Rab5-like proteins (ARA7) but their localization is not on EEs as expected. Although ARA7 was suggested to be present on EEs involved in recycling auxin efflux PIN1 to the PM (Ueda et al. 2004; Geldner 2004), there is convincing evidence that ARA7 localizes on late endosomes (LEs) in tobacco leaf epidermal cells, instead of in EEs, and is involved in vacuolar trafficking, because mutations that impair the function of ARA7 cause delocalization of vacuolar markers (Kotzer et al. 2004). Alternatively, colocalization of SNARE SYP41 and vacuolar H + ATPase subunit 1a, but not ARA7, in compartments of the trans Golgi network (TGN) suggest that the TGN can be regarded as an EE in plants (Dettmer et al. 2006; Lam et al. 2007). Further evidence supporting TGN as an early compartment in the plant endocytic pathway comes from ultrastructural studies tracking cationized ferritin in protoplasts. In these studies, ferritin was first observed in Golgi stacks, TGN and a compartment called partially coated reticulum, that probably corresponds to TGN membranes (Pesacreta and Lucas 1985; Tanchak et al. 1988). However, recently a tubulo-vesicular EE similar to that described in animal cells was observed in tobacco protoplasts as the first compartment receiving recently endocytosed, positively charged nanogold from the PM (Onelli et al. 2008). The classical view of endosomes as stable membranous compartments has been changed by new data showing them to be a continuum of maturing compartments (Perret et al. 2005; Rink et al. 2005). In animals and plants, the degradation pathway seems to involve production of multivesicular bodies (MVBs) by EEs. In animals, MVBs are thought to originate from the vesicular portion of EEs and to fuse or mature into LEs and finally to fuse with lysosomes. In plants, the degradative pathway seems to involve the maturation of early into late endosomes (Ueda et al. 2001) which have been identified as MVBs or prevacuolar compartments (PVCs). Direct fusion of PVCs with 14

15 vacuoles without involvement of vesicle transfer has been hypothesized, as suggested for lysosomes in animals (Luzio et al. 2000; Gruenberg and Stenmark 2004). The endocytic pathway is known to intersect secretory and biosynthetic processes during recycling to the PM and protein transport to lysosomes or vacuoles (Henkel et al. 1996, Ang et al. 2004). MVBs/PVCs have been suggested to be involved in both the secretory and endocytic pathways (Tse et al. 2004): plant PVCs are considered an intermediate for delivering proteins to storage vacuoles and for sorting materials to lytic compartments (Jiang and Rogers 1998). Cross-talk between endocytic and secretory pathways was recently established in root cells. During maturation of EEs into LEs through MVBs, sorting of cargo proteins toward various destinations is conducted by sorting endosomes, multivesicular compartments that redirect proteins to various destinations such as vacuoles, TGN and the PM for recycling, suggesting that sorting endosomes could be a conserved organelle in eukaryotes (Jaillais et al. 2008). Constitutive cycling mediates membrane traffic between endosomes and the PM, allowing rapid changes in PM composition (Royle and Murrel-Lagnado 2003). In animal cells, this recycling process responds to hormonal stimuli that control the rate of protein recycling and modulate PM protein concentrations, including those of ion channels and receptors. Hormones have also been shown to affect plant endocytosis and constitutive recycling. The phytohormone auxin has a general inhibitory effect on endocytosis (Paciorek et al. 2005) whereas abscisic acid upregulates internalization of KAT1 K + channel, which play an important role in stomatal opening (Sutter et al. 2007). Other proteins cycling between endosomes and PM are water channel PM intrinsic protein 2, PM H + -ATPase and auxin carrier efflux PIN1 (Dhonukshe et al. 2007). Evidence showing that the high level of fusion of SVs exceeds PM extension suggest that a mechanism of PM retrieval and recycling regulates the membrane economy of growing pollen tubes (Steer and Steer 1989). More recently, accurate electron microscope analysis of serial sections of pollen tubes revealed the presence of coated vesicles in regions just behind the clear zone (Derksen et al. 1995). This observation, together with data proving the existence of sequences coding for a clathrin heavy 15

16 chain-like polypeptide (Blackbourn and Jackson 1996), supported the idea that CD endocytosis functions during pollen tube growth in recycling excess PM secreted at the tip (Derksen et al. 1995). The endocytic process in pollen tubes of several species was recently analyzed (Parton et al. 2001) using FM4-64 (Bolte et al. 2004). This approach showed that most internalized PM of pollen tubes accumulated at the tip, in which a typical V-shaped fluorescent region was observed, suggesting that most of the internalized PM was redirected into the secretory pathway (Parton et al. 2001; Moscatelli et al. 2007). However, because this probe inserts homogeneously into the PM, it shows the endocytic process as a whole but does not provide information about different endocytic modes that might occur in pollen tubes. A dissection of the endocytic process in pollen tubes was done in yeast using charged nanogold as probe (Prescianotto- Baschong and Riezmann 1998). Positively and negatively charged nanogold particles bind negatively and positively charged residues on the PM, respectively, showing the fate of internalized PM segments in different regions at ultrastructural level. Use of positively and negatively charged nanogold together with a specific type of fluorochrome showed different internalization events taking place in specific PM regions, such as the subapical zones and, suprisingly, also the tip domain (Fig. 1, greenish-yellow and orange vesicles, respectively) (Moscatelli et al. 2007). The distribution pattern of positively charged nanogold involved organelles such as cis, medial and trans elements of Golgi bodies, smooth ER and vacuoles, whereas negatively charged nanogold was only found in a limited number of vesicles in the tip region and in vacuoles. Sometimes negatively charged gold particles were observed in vesicular-tubular elements related to the trans face of Golgi bodies, but not in cis or medial cisternae (Fig. 1 shows internalization model). Retrieval of excess PM secreted during exocytosis has been shown in animal and plant cells with high secretory activity. In neurons, internalized PM is trapped in synaptic vesicles at the tip and reused by the secretory pathway (Henkel et al. 1996); similarly, in pollen tubes, use of FM4-64 showed endocytosis of PM that then concentrated in the tip region (Fig. 2C), presumably only to be used again for exocytosis 16

17 and to maintain membrane homeostasis in the cell (Parton et al. 2001). Positively and negatively charged nanogold showed spatially distinct internalization patterns, because the former was internalized in the sides of tubes, presumably in the organelle-rich zone (up to 25 μm from the tip). This internalization pattern is in line with previous studies that showed coated vesicles μm from the tip (Derksen et al. 1995) and suggested that a CD pathway occurs in this region (see below). In animal cells, internalized vesicles were first delivered to EEs, a sorting station of tubulovesicular structure characterized by specific sets of Rab-GTPases (Gruenberg 2001; Zerial and McBride 2001). In plant cells, a large number of Rab and SNARE homologues have been found in the Arabidopsis genome, but data on their localization and interactions is controversial (Geldner and Jurgens 2006). Compartments having a morphology similar to that described in animal EEs were not observed in pollen tubes; positively charged nanogold was seen immediately below the PM in vesicles that could be newly invaginated or EEs; in the latter case, they did not have tubulo-vesicular structure. Interestingly, the transport of positively charged nanogold involved Golgi bodies. Positively charged nanogold was observed in vesicles associated with the trans face of Golgi apparatus, which is in line with the hypothesis that elements of the TGN could function as early endosomal compartments in plants (Dettmer et al. 2006; Geldner and Jurgens, 2006). Stained vesicles associated with the rims of cis and medial cisternae of Golgi bodies and studies using FM dyes together with a specific marker for Golgi bodies, such as BodiPy TR ceramide, revealed that this organelle also functions as a station for delivering internalized PM to the secretory pathway (Fig. 2 B-E). Positively charged nanogold was also observed in tubules of smooth ER dispersed in the apical region. In pollen tubes, apical smooth ER has been postulated to play a main role in homeostasis of Ca 2+, because it functions as an internal Ca 2+ store (Hepler et al. 2001) from which Ca 2+ ions may be mobilized to the cytosol by a IP 3 -regulated mechanism (Kost 2008); delivery of internalized membranes to smooth ER could play a role in maintaining the integrity of this compartment. Positively charged nanogold has been observed in internalizing vesicles containing fibrillar material, similar to cell wall components outside 17

18 the protoplast, suggesting that removing a portion of cell wall could help regulate wall plasticity during elongation, cooperating with cell wall loosening enzymes such as polygalacturonases and pectate lyases (Wen et al. 1999; Ren and Kermode 2000). Internalization of pectins and AGPs is known in somatic plant cells, after which cell wall molecules are delivered to brefeldin A (BFA)-induced compartments, suggesting that they could also be recycled into the secretory pathway to limit de novo synthesis by the cell wall (Baluska et al. 2002; Baluska et al. 2005). On the contrary, negatively charged nanogold was internalized in the tip region of the tube, in which it appeared inside a few vesicles (Fig. 1, pink and orange vesicles). Organelles such as ER or cis, medial and trans cisternae of Golgi bodies were not seen to participate in negatively charged gold transport, suggesting that this probe is not recycled into the secretory pathway through the Golgi apparatus. However, sometimes gold particles were observed in vesicles closely associated with the trans face of Golgi bodies, from which vesicles seemed to continue and to fuse with a compartment probably involved in degradation. These observations again supported the idea that the TGN could be a real meeting point between degradative and biosynthetic pathways in plants, and that negatively charged nanogold is basically sent to degradative compartments, although pulse-chase experiments suggest the possibility of limited recycling in the tip region. One hypothesis is that negatively charged nanogold reveals a second exocytotic pathway involved in maintaining the necessary PM polarity/remodelling in the apical subapical PM during tip growth. A mechanism of endocytosis at the tip was recently shown to be involved in recycling DAG and PLC in the apical/subapical regions of growing tobacco pollen tubes (Helling et al. 2006). Observing the movements of FM4-64-stained vesicles in the tip region, Parton and colleagues (2001) observed an apparent cycling of bright spots 10 μm from the apex back to the tip that could be related to internalization-recycling of negatively charged nanogold in the apex (Moscatelli et al. 2007). 521 Endocytosis has also been implicated in removing K + ion channels in order to accommodate changes in surface area of guard cells following osmotic changes (Hurst et al. 2004; Meckel et al. 2004). Tip growing cells, such as pollen tubes, have a series of 18

19 ion transporters that function in specific regions of the PM. Among these, Ca 2+ channels and H + proton pumps in the tip (Hepler et al. 2001) and Ca 2+ /K + pumps in the subapical regions regulate ion homestasis necessary for organization of Afs and exocytosis (Lovy- Wheeler et al. 2005; Kost 2008). The polarized localization of these complexes in different regions of the PM could require internalization/repositioning, explaining cycling of endocytic vesicles limited to the tip region. Recently, Dhonukshe et al. (2007) reported data on constitutive CD endocytosis of PIN efflux carriers in Arabidopsis protoplasts and suggested that CD internalization could be the predominant system in plants. However, the occurrence of CI mechanisms in specialized cells such as pollen tubes could not be excluded. Data reported by Derksen et al. (1995) showed that SVs deliver 430 μm 2 membrane/min, of which only 50 μm 2 is used for expansion, leaving an excess of 380 μm 2. The endocytic activity behind the tip, taken at its maximum of 225 μm 2 /min, cannot retrieve a membrane surface of this size. Because only the area delimited by coated pits or coated vesicles was considered in this case, part of the internalization could actually occur by CI mechanisms. Analysis of pollen tubes treated with ikarugamycin (Ika), an inhibitor of CD endocytosis (Hasumi et al. 1992; Luo et al. 2001), showed that CD endocytosis could be involved in internalization of PM to be recycled to the secretory pathway, because most tubes did not show the canonical V-shaped fluorescence in the tip region. This was in line with previous observations suggesting that CD endocytosis was involved in the removal of excess PM in growing root hairs and developing cell plates (Emons and Traas 1986; Otegui and Stahelin 2000; Baluska et al. 2005; Tahara et al. 2007). Uptake of negatively charged nanogold at the tip seemed to be severely impaired, because vacuoles never contained gold particles. However, because the degradative pathway conveying positively charged nanogold to vacuoles was intact, it is clear that distinct degradative pathways can function and it might also be worthwhile studying CI internalization modes in pollen tubes. Bright fluorescence of the PM behind the tip often suggested that the subapical region could be a point of substantial CD endocytosis. Statistical analysis of fluorescence with Ika showed a significant inhibition of internalization in the flanks of the 19

20 tube, indicating the importance of recycling for tube elongation: in general, tube growth reduced but did not cease in the presence of Ika and it was quite common to observe a reduction of fluorescence in the tip PM during FM4-64 internalization in time-lapse experiments, which suggested that tip growth proceeds at a reduced rate by de novo synthesis by the cell wall and PM (Moscatelli et al. 2007). The evidence of internalization processes at the tip led us to consider a new hypothesis to explain vesicle accumulation at the apex, namely that endocytic vesicles might contribute substantially to V-shaped vesicle accumulation in addition to SVs (see model in Fig. 1). Use of different FM dyes such as FM4-64 and FM1-43 in the same pollen tubes allowed us to identify different vesicle pools and to map the sites of exo- and endocytosis in the clear zone of tobacco pollen tubes. Pulse chase experiments showed that while exocytosis occurred at the edges of the apical PM domain (3-10 μm from the tip), endocytic activity was localized in the central part of the apical dome (Zonia and Munnik 2008) (Fig. 1). Membrane retrieval from the apical PM led to generation of small endocytic vesicles that moved to the endosomal compartment by reverse transport (Zonia and Munnik 2008). Accurate analysis of vesicle dynamics confirmed the presence of a constitutive endocytic pathway in the central domain of the apical PM, and sustained a new model to explain tip growth (Bove et al. 2008). SVs, transported to the apex by forward cortical cytoplasmic streaming, can fuse with specific regions of the PM flanking the central apical dome (3-10 μm), and not with the entire apical PM. FRAP experiments in the exo- and endocytosis sites of the apex revealed that not all of the SVs conveyed to the tip region fused with PM and some of them took more that one passage to the apex before they fused with the PM (Bove et al. 2008). Endocytosis for in vivo growth regulation Pollen tube growth in vivo is closely regulated by its interaction with style molecules. The study of endocytosis in pollen tubes opens new perspectives for pollen tube-style interactions in vivo. Particularly in the self-incompatibity process which characterizes N. alata (McClure and Franklin-Tong 2006), the compatibility/incompatibility of pollen tubes 20

21 growing in the style seems to be determined by sequestration in vacuole-like compartments of S-RNases which are internalized in both kinds of tubes (Goldraij et al. 2006). However, in the case of incompatible pollen, S-RNases escape from vacuolelike vesicles and act as degradation enzymes. After compatible pollination, S-RNases remain sequestered within vacuoles and pollen tubes can grow through the style (Goldraij et al. 2006). It is not yet known whether internalisation of S-RNases follows different mechanisms in compatible and incompatible pollination. However, the mechanism of internalisation (CD or CI) could determine the destiny of internalised materials into different transport pathways and it would be of great interest to study the mechanism by which S-RNases escape their vacuole-like compartments and become toxic for the tube after incompatible pollination. An RNase-binding 120 Kd glycoprotein is also taken up by Nicotiana alata (L.) pollen tubes growing in the style and has been observed in vacuoles together with RNases (Lind et al. 1996; Goldraij et al. 2006). Other pistil proteins are internalized in pollen tubes by endocytosis in Lilium longiflorum (L.) (Kim et al. 2006). However, the endocytic mechanisms responsible for their internalization and transport through the endomembrane system still need to be characterized

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34 Forward and backward cytoplasmic streaming are shown by orange and green arrows in the cortex and central area of the cytoplasm, respectively Figure 2. Colocalization of FM4-64 with BODIPY-labelled membranes. (A) Bright field of pollen tube chosen for the analysis. (B) BODIPY TR-Ceramide-stained Golgi bodies are distributed in the cytoplasm and concentrate in the tip region to form a collar-like structure (arrows) just behind the extreme tip. (C) FM4-64 staining pattern 30 minutes after addition of the fluorochrome. FM4-64 stains the PM and labelled spots show typical V-shaped accumulation at the apex. (D) Colocalization analysis 3 minutes after addition of FM4-64 showing probe staining overlapping in the collar-like structure and tip (white spots). (D) Cytofluorograms enable the intensity distribution of green and red channels to be visualized in a 2D scatter plot 30 minutes after FM4-64 addition. The Region of Interest (ROI, in yellow) select colocalized pixels. Bar, μ10 m

35 Figure V G - - V - G Negatively charged nanogold + Positively charged nanogold Secretory Vesicles (SVs) Anterograde cytoplasmic streaming and PM recycling Retrograde cytoplasmic streaming PM recycling clathrin uncoated lateral endocytic vesicles clathrin coated lateral endocytic vesicles clathrin coated apical endocytic vesicles clathrin uncoated apical endocytic vesicles Sites of exocytosis

36 Figure 2 BF BP TR Cer FM4-64 A BP TR Cer and FM4-64 B C D E