Spread the news: systemic dissemination and local impact of Ca2+ signals along the phloem pathway

Journal of Experimental Botany, Apr 2014

We explored the idea of whether electropotential waves (EPWs) primarily act as vehicles for systemic spread of Ca2+ signals. EPW-associated Ca2+ influx may trigger generation and amplification of countless long-distance signals along the phloem pathway given the fact that gating of Ca2+-permeable channels is a universal response to biotic and abiotic challenges. Despite fundamental differences, both action and variation potentials are associated with a sudden Ca2+ influx. Both EPWs probably disperse in the lateral direction, which could be of essential functional significance. A vast set of Ca2+-permeable channels, some of which have been localized, is required for Ca2+-modulated events in sieve elements. There, Ca2+-permeable channels are clustered and create so-called Ca2+ hotspots, which play a pivotal role in sieve element occlusion. Occlusion mechanisms play a central part in the interaction between plants and phytopathogens (e.g. aphids or phytoplasmas) and in transient re-organization of the vascular symplasm. It is argued that Ca2+-triggered systemic signalling occurs in partly overlapping waves. The forefront of EPWs may be accompanied by a burst of free Ca2+ ions and Ca2+-binding proteins in the sieve tube sap, with a far-reaching impact on target cells. Lateral dispersion of EPWs may induce diverse Ca2+ influx and handling patterns (Ca2+ signatures) in various cell types lining the sieve tubes. As a result, a variety of cascades may trigger the fabrication of signals such as phytohormones, proteins, or RNA species released into the sap stream after product-related lag times. Moreover, transient reorganization of the vascular symplasm could modify cascades in disjunct vascular cells.

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Spread the news: systemic dissemination and local impact of Ca2+ signals along the phloem pathway

Journal of Experimental Botany Spread the news: systemic dissemination and local impact of Ca2+ signals along the phloem pathway Aart J. E. van Bel 1 2 Alexandra C. U. Furch 1 Torsten Will 1 Stefanie V. Buxa 1 Rita Musetti 0 Jens B. Hafke 3 0 Department of Agricultural and Environmental Sciences, University of Udine , Via delle Scienze 208, 33100 Udine , Italy 1 Institute of Phytopathology and Applied Zoology, Centre for BioSystems, Land Use and Nutrition, Justus-Liebig-University , Heinrich- Buff-Ring 26-32, D-35392 Giessen , Germany 2 Institute of General Botany, Justus-Liebig University , Senckenbergstrasse 17, D-35390 Giessen , Germany 3 Institute of Plant Physiology, Justus-Liebig University , Senckenbergstrasse 3, D-35390 Giessen , Germany We explored the idea of whether electropotential waves (EPWs) primarily act as vehicles for systemic spread of Ca2+ signals. EPW-associated Ca2+ influx may trigger generation and amplification of countless long-distance signals along the phloem pathway given the fact that gating of Ca2+-permeable channels is a universal response to biotic and abiotic challenges. Despite fundamental differences, both action and variation potentials are associated with a sudden Ca2+ influx. Both EPWs probably disperse in the lateral direction, which could be of essential functional significance. A vast set of Ca2+-permeable channels, some of which have been localized, is required for Ca2+-modulated events in sieve elements. There, Ca2+-permeable channels are clustered and create so-called Ca2+ hotspots, which play a pivotal role in sieve element occlusion. Occlusion mechanisms play a central part in the interaction between plants and phytopathogens (e.g. aphids or phytoplasmas) and in transient re-organization of the vascular symplasm. It is argued that Ca2+-triggered systemic signalling occurs in partly overlapping waves. The forefront of EPWs may be accompanied by a burst of free Ca2+ ions and Ca2+-binding proteins in the sieve tube sap, with a far-reaching impact on target cells. Lateral dispersion of EPWs may induce diverse Ca2+ influx and handling patterns (Ca2+ signatures) in various cell types lining the sieve tubes. As a result, a variety of cascades may trigger the fabrication of signals such as phytohormones, proteins, or RNA species released into the sap stream after product-related lag times. Moreover, transient reorganization of the vascular symplasm could modify cascades in disjunct vascular cells. Calcium hotspots; calcium signatures; eletropotential waves; long-distance signalling; phloem pathway; sieve element cytoskeleton; sieve elements; sieve tube occlusion Introduction In their natural habitat, plants are permanently exposed to countless abiotic and biotic changes imposing a permanent stress. The majority of environmental challenges are communicated via the extracellular microenvironment (i.e. the apoplasmic space) and, from there, via the plasma membrane to the intracellular space. The external stimuli are monitored by a vast battery of sensors which transform the external information into signals triggering adequate cell reactions. One of the initial events in sensing is a Ca2+ influx modulated by Ca2+-permeable channels at the plasma membrane. Since no Ca2+-selective channels have been identified with certainty in the plasma membrane of plant cells thus far (Kudla et  al., 2010), we will refer to these channels as ‘Ca2+-permeable’ (e.g. Sanders et al., 2002). Ca2+ influx elevates the cytosolic Ca2+ level according to stimulus-specific, spatio-temporal, and potentially cell-specific patterns designated Ca2+ signatures © The Author 2014. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: 10 s up to 30 min (Stahlberg and Cosgrove, 1997; Stahlberg et al., 2006). The propagation velocities are 5–10 times slower than those of APs (Stahlberg and Cosgrove, 1997) and the amplitude drops along the transmission path (Davies, 2004; Stahlberg et  al., 2005, 2006). As a result, VP amplitudes decrease with increasing distance from the stimulus site and finally extinguish (van Sambeek and Pickard, 1976). The slow repolarization of VPs might result from the shutdown of proton pumps as indicated by pH-dependent fluorochromes and the ineffectiveness of ion channel blockers (Stahlberg and Cosgrove, 1992, 1996). The inhibition of proton pump activity is not fully understood, but may be due to elevated Ca2+ levels (Kinoshita et  al., 1995; Hafke et  al., 2013). Proton pump activity may be equally suppressed by cytosolic Ca2+ in APs, but the reduced pump activity may be hardly detectable due to the lower Ca2+ influx during APs (see ‘Creation of Ca2+ hotspots in sieve elements’). Generation and propagation of VPs have only been observed in intact plants, whilst APs can propagate in isolated organs (Stahlberg et  al., 2006). Therefore, relaxation of the negative hydrostatic pressure in the xylem vessels is the likely source of VP generation (Stahlberg and Cosgrove, 1997). That VPs originate from events in xylem vessels (Fig. 1) was demonstrated by the fact that VPs, in contrast to APs, are able to traverse dead or poisoned areas (Stahlberg et al., 2006). Essential differences and functional similarities between APs and VPs The preceding sections disclose a few essential differences between the APs and VPs (Fig. 1). (i) APs and VPs are of a dissimilar nature (i.e. they are of an electrical or mechanistic origin) and, by implication, the Ca2+-permeable channels responsible for the initial depolarization are either voltagedependent or mechano-sensitive channels, respectively (Fig.  1). (ii) APs are generated in non-vascular or vascular cells, move longitudinally along the sieve tubes, and may disperse in the lateral direction to surrounding vascular cells (Fig.  1). In contrast, VPs are generated by vascular (xylem parenchyma) cells and move laterally across several cell layers to the sieve tubes, so that VPs reflect arrival of successive single depolarizations at the sieve element plasma membrane mimicking electrical propagation along the sieve tubes (Fig. 1; Malone, 1996; Pyatygin et al., 2008; van Bel et al., 2011a). These conclusions call for further exploration of the following questions: (i) which Ca2+-permeable channels reside in the vascular cells and where are they located (Fig. 2); and (ii) how does the symplasmic organization of the phloem strands enable combined longitudinal propagation and lateral dispersion of electrical information (Fig. 3)? Despite their profound differences, EPWs have one essential, functional feature in common. They are all associated with an initial elevation of cytosolic Ca2+ (Trebacz et al., 2006; Davies and Stankovic, 2006; Demidchik and Maathuis, 2007; McAinsh and Pittman, 2009), regardless of the involvement of voltage-dependent, mechano-sensitive, or ligand-activated Ca2+-permeable channels. Hence, the elevation of Ca2+ levels in sieve elements, the involvement of the sieve element cytoskeleton in Ca2+ influx mechanisms, and the impact of Ca2+ influx on sieve element biology are major issues in this frame (Figs. 4–6). Phloem-associated Ca2+-permeable channels and other channels involved in EPW propagation Types of Ca2+-permeable channels localized to the plasma membrane in plants In animals, highly selective Ca2+ channels are responsible for Ca2+ fluxes at the plasma membrane (Tsien et al., 1987; Tsien and Tsien, 1990; McAinsh and Pittman, 2009), whereas non-selective cation channels (NSCCs) or Ca2+-permeable channels seem to enable Ca2+ fluxes in plants to give rise to stimulus-specific Ca2+ signatures (Demidchik and Maathuis, 2007; McAinsh and Pittman, 2009). Ca2+ influx across the plasma membrane can be mediated by the following types of non-specific cation channels (Fig.  2A; Demidchik and Maathuis, 2007; McAinsh and Pittman, 2009): (i) HACCs: hyperpolarization-activated Ca2+-permeable channels, which are gated by an increase in membrane voltage, reactive oxygen species (ROS), and changes in the cytoplasmic Ca2+ level; (ii) DACCs: depolarization-activated Ca2+-permeable channels activated by a decrease in membrane voltage; (iii) MSCs: mechano-sensitive channels the gating of which is modulated by tensile forces exerted on membranes; (iv) CNCGs: cyclic nucleotide-gated channels activated by binding of cyclic nucleotides (e.g. cAMP, cGMP); and (v) GLRs: glutamate receptor-like channels activated by binding of amino acids. Regarding HACCs (in Arabidopsis root cells), the resting value of the membrane potential is more positive than their activation voltage, which, however, can shift to more positive membrane potentials brought about by increased Ca2+ levels (Demidchik et al., 2002; Demidchik and Maathuis, 2007; Miedema et al., 2008). It is likely that DACCs are engaged in cold-induced Ca2+ influx (White, 2009). A  member of the DACCs, named the maxi cation channel, was postulated to be responsible for the creation of complex temperature-dependent Ca2+ signatures (White and Ridout, 1999; White, 2004, 2009). Apart from their gating response to changing tensile forces (Demidchik and Maathuis, 2007), MSCs may act as primary temperature sensors (Minorsky and Spanswick, 1989; Monroy and Dhindsa, 1995; Plieth et  al., 1999), as demonstrated by the gradually increasing activity of MSCs at temperatures dropping below 20 °C (Ding and Pickard, 1993). Putative Ca2+-permeable channels lining the sieve elements Among all possible Ca2+-permeable channels, MSCs have been identified with certainty in the sieve element plasma membrane thus far. Forisome reactions in intact sieve elements (Knoblauch et  al., 2001) and in sieve element protoplasts (Hafke et al., 2007) evidenced Ca2+ influx in response to vigorous turgor changes. MSCs may also be crucial players in the activation of HACCs that may catalyse a long-lasting Ca2+ influx into sieve elements during the prolonged EPW phase after remote burning (Furch et  al., 2009). Recently, GLRs have been discovered in the phloem (Vincill et  al., 2013), but their cellular location is uncertain. Only circumstantial evidence has been obtained for other Ca2+-permeable channels in sieve tubes. Cold-shock induced Ca2+ influx into sieve elements (Thorpe et al., 2010; Hafke et al., 2013) could have been mediated by MSCs or DACCs (Ding and Pickard, 1993; Plieth, 1999; Plieth et al., 1999; White and Ridout, 1999; White, 2009). The wealth of potential ligands associated with VP generation (see ‘Presumptive significance of plasmodesmal connectivity for lateral VP dispersion’) renders the presence of ligandactivated channels on the sieve element plasma membrane highly plausible. Location of the Ca2+-permeable channels in sieve elements Early studies using BODIPY-DHP and antibodies localized voltage-dependent Ca2+-permeable channels to the sieve element (Volk and Franceschi, 2000). A more detailed approach using fluorochrome mixtures (Furch et al., 2009) and reaching a higher spatial resolution visualized Ca2+-permeable channels located in the plasma membrane and the endoplasmic reticulum (ER) stacks of sieve elements (Fig. 2B). Ca2+-permeable channels are unevenly localized to the sieve element plasma membrane. They are mostly aggregated in the vicinity of sieve plates and unilaterally branched plasmodesmata (pore–plasmodesm units; PPUs) towards the companion cells (Furch et al., 2009); hence, at the sieve element side facing the companion cell (Fig. 2B). A false-colour presentation of the ratio between BODIPY-DHP and RH-414 fluorescence confirmed preferential aggregation of Ca2+-permeable channels near sieve plates and PPU orifices (Fig. 2C). Furthermore, distribution of Ca2+-permeable channel clusters closely matched that of sieve element ER distribution in sieve elements (Fig. 2D), as was documented by double-label experiments (Fig. 2E). Cl– channels is entirely lacking, phloem-localized K+ channels of the AKT2/3 type were electrophysiologically characterized and linked to AP depolarization (Marten et al., 1999; Bauer et al., 2000; Lacombe et al., 2000; Deeken et al., 2002). Weak inward rectifying currents matching the features of AKT2/3 channels were recorded in sieve element protoplasts (Hafke et  al., 2007, 2013). The increasing permeability of AKT2/3 channels at more alkaline pH values (Marten et al., 1999) and the extracellular alkalinization during transmission of SPs (Zimmermann et  al., 2009) indicate that AKT2/3 channels are involved in membrane repolarization. As regards proton pumps, immunological approaches localized H+-ATPases to sieve elements and companion cells (Langhans et al., 2001). Symplasmic organization of phloem strands Deployment of other channels and pumps involved in EPW propagation Ultrastructure and plasmodesmal connectivity of sieve elements Although this review does not focus on other channels and pumps involved in EPWs, we will pay some marginal attention to the few facts known. Whilst information on sieve element In sieve elements, cellular substructure is reduced to a plasma membrane envelope lined with a thin margin of gelatinous cytoplasm (mictoplasm) containing a limited number of organelles (e.g. van Bel, 2003). Originally, the mictoplasm was defined as the mixture of cytoplasmic contents with the sieve element fluid (Engleman, 1965). This layer has been re-defined as mictoplasm for practical reasons (van Bel, 2003): Ca2+ concentrations vary greatly in this space during EPWs (Furch et al., 2009). The fact that the mictoplasmic layer is in open contact with the sap stream in the sieve element lumen is the consequence of tonoplast disintegration during sieve element ontogeny (Esau, 1969). Several organelles such as the nucleus, ribosomes, and Golgi apparatus are degraded during sieve element development (Behnke and Sjolund, 1990). The ER that may originate from the cortical ER (Hepler et al., 1990) survives the partial programmed cell death and is aggregated in regular stacks that are often oriented perpendicularly to the plasma membrane (Sjolund and Shih, 1983; Ehlers et al., 2000). The ER stacks are tethered to the plasma membrane and to each other by anchors of unknown nature (Ehlers et al., 2000) to prevent dragging by mass flow and resultant sieve pore occlusion. For the same reason, a special type of plastids of unknown function—considerably smaller than chloroplasts (Behnke and Sjolund, 1990)—are tethered to the plasma membrane (Ehlers et al., 2000). Microscopically visible clusters of phloem-specific structural proteins are located at the margins of the sieve element (Behnke and Sjolund, 1990; Knoblauch and van Bel, 1998) or even in the sieve-tube lumen (Froelich et al., 2011; Knoblauch and Oparka, 2012). In addition, there is a wealth of soluble proteinaceous components in sieve elements (Barnes et  al., 2004; Walz et  al., 2004; Giavalisco et  al., 2006; Aki et  al., 2008; Furch et al., 2010; Dinant and Lucas, 2013). It has been excluded for a long time that a complete cytoskeleton exists in sieve elements (Parthasaraty and Pesacreta, 1980; Thorsch and Esau, 1981; Evert, 1990), although circumstantial structural (Chaffey and Barlow, 2002) and chemical (Kulikova and Puryaseva, 2002; Barnes et  al., 2004; Walz et  al., 2004; Giavalisco et al., 2006; Aki et al., 2008) evidence favoured the opposite view. Recent confocal laser scanning microscopic, immunological, and physiological studies have probably ended the dispute by identification of a complete, parietally located actin network in sieve elements (Hafke et al., 2013). As inferred from dye coupling experiments, the sieve element precursor divides longitudinally and becomes transiently isolated from its neighbouring cells, which has been regarded as an instrument for developmental specialization (van Bel and van Rijen, 1994). The other daughter cell develops into 1–4 companion cells flanking each sieve element (Esau, 1969). Given its limited cellular equipment, a sieve element relies almost completely on its companion cell(s) for its survival, which makes communication between the two of paramount importance (van Bel 2003). Towards the end of the temporary symplasmic seclusion, so-called PPUs (van Bel and Kempers, 1997) arise that have the capacity to traffic a vast spectrum of substances including macromolecules between sieve element and companion cell(s) (Imlau et  al., 1999; Lucas et al., 2001, 2009). Plasmodesmata between companion cells and phloem parenchyma are sparse (Kempers et  al., 1998), which seems to present a symplasmic bottleneck. These plasmodesmata have never been studied in detail, but may be of special nature, since their opening state is related to source–sink relationships (Patrick and Offler, 1996; Hafke et  al., 2005). Moreover, the phloem-specific clostero- and luteoviruses are unable to pass this symplasmic border and, hence, are contained inside the sieve element–companion cell complexes (Stewart et al., 2013). Presumptive significance of plasmodesmal connectivity for lateral AP dispersion The electrical conductivity of the sieve element plasma membrane, the longevity of the sieve elements, and the high electrical conductance of sieve pores make sieve tubes ideal conduits for long-distance electrical signalling (van Bel and Ehlers, 2005). The restriction of longitudinal AP propagation to the sieve tubes indicates a high degree of electrical resistance in the plasmodesmal pathway from sieve elements to other cells (Fig. 3). The scarcity of plasmodesmata between companion cells and phloem parenchyma cells in transport phloem (Kempers et al., 1998), which are closed under sourcelimiting conditions, would fulfil the requirements for electrical insulation (Patrick and Offler, 1996; Hafke et al., 2005). On the other hand, it should be borne in mind that electrical currents are expected to pass plasmodesmata with extremely low molecular exclusion limits or even move along the membranes crossing the cytoplasmic sleeve. Moreover, permanent and full electrical insulation of sieve element–companion cell complexes is unlikely, as inferred from symplasmic unloading of excess photoassimilates under sink-limiting conditions (Patrick and Offler, 1996), to fill axial storage compartments along the phloem pathway rapidly. ‘Electrical leakiness’ (Fig. 3A) is indicated by small depolarizations of phloem parenchyma cells coincident with the passage of EPWs (Rhodes et  al., 1996). All in all, there is a good chance that the electrical insulation of sieve tubes is incomplete. Voltage-dependent Ca2+ channels in sieve elements would be the initiators of longitudinal AP propagation that is diverted by electrotonic leakage bringing about Ca2+ influx into vascular parenchyma cells. The concept of functional current leakage is supported by events in the excitable plant Mimosa pudica, in which long distances are covered by APs owing to an insulating sclerenchyma sheath around the sieve element–companion cell complexes (Fleurat-Lessard and Roblin, 1982). This shield is interrupted in the pulvini, where numerous plasmodesmata provide ample symplasmic access to flexor parenchyma cells (Fleurat-Lessard and Bonnemain, 1978) with inherent facilitation of current leakage. The flexor cells react to Ca2+ influx by instantaneous loss of osmotic substances giving rise to leaf and leaflet bending (Fleurat-Lessard and Bonnemain, 1978). These phenomena may exemplify less prominent events in non-excitable plants with lower rates of current leakage and less eye-catching reactions by the flanking parenchyma cells. Altogether, it appears that incomplete insulation of sieve tubes is not a defect, but highly functional in lateral dispersion of Ca2+ waves and Ca2+-mediated information. Presumptive significance of plasmodesmal connectivity for lateral VP dispersion While the lateral events accompanying APs allow a straightforward assessment, there is more room for speculation regarding VPs. Disturbance of the hydraulic equilibrium in xylem vessels leads to water intake by the adjacent parenchyma cells which causes membrane depolarization due to increased turgor (Malone and Stankovic, 1991; Stahlberg and Cosgrove, 1992, 1997; Mancuso, 1999; Davies, 2006). Therefore, receptor potentials are probably triggered here by mechano-sensitive Ca2+-permeable channels (probably MSCs), but the mode of subsequent lateral electrical transmission to sieve tubes is a matter of debate. As a first possibility (Fig.  3B), pressure-induced receptor potentials activate voltage-dependent Ca2+-permeable channels (perhaps DACCs) that generate EPW propagation towards the sieve tubes. The second and, at first sight, most likely option (Fig. 3C) is that the turgor of all vascular cells including the sieve tubes rises by intake of water after vessel damage, as argued for the mechanisms of cucurbit phloem exudation (Zimmermann et al., 2013). According to this scenario, VP propagation results from the action of mechano-sensitive channels which perceive local turgor changes in each vascular cell. This concept explains the attenuation of VPs with distance, provided that the relaxation in the vessels is increasingly dampened further away from the site of damage. Nevertheless, a few essential problems remain with this concept. Why is the VP generation not almost equally rapid along the vascular pathway, because pressure loss must propagate very quickly. In other words, why does it take so much longer for the VP to be expressed far away from the site of wounding, and why is the reaction to crushing so much more vigorous than to cutting, although the number of vessels damaged is approximately identical? Therefore, it has been postulated as a third alternative (Fig.  3D) that MSC-mediated Ca2+ influx triggers cascades that produce chemical signals (Ricca, 1916; van Sambeek and Pickard, 1976; van Sambeek et al., 1976; Boari and Malone, 1993; Malone, 1996; Stahlberg and Cosgrove, 1997; Mancuso, 1999; Pyatygin et  al., 2008). Oligosaccharides as well as the peptide systemin in solanacean species (Narvaez-Vasquez and Ryan, 2004) are potential messengers triggering VPs (Thain et al., 1995; Moyen and Johannes, 1996) after docking to ligand-activated Ca2+-permeable channels (Fig.  3D). The period to accumulate sufficient second messengers which may be correlated with the degree of relaxation would explain the increasing lag time between wounding and VP generation along the pathway. In view of the co-occurrence of diverse Ca2+-permeable channels in plasma membranes (Kudla et  al., 2010), combinations of the above scenarios are likely to occur. Irrespective of the mode of lateral EPW transmission, open plasmodesmata are compulsory (Fig.  3B, D; van Bel et  al., 2011a), unless information is transferred by lateral pressure An elevated mictoplasmic Ca2+ level may boost its own concentration by Ca2+-stimulated Ca2+ efflux from ER stacks in analogy to Ca2+-induced Ca2+ release at the tonoplast (CICRs, or calcium-induced calcium release channels; Bewell et al., 1999; Sanders et al., 2002). Similarly, Ca2+ would trigger presumptive Ca2+-dependent Ca2+ channels on the ER membranes (Fig. 4F; Furch et al., 2009; Hafke et al., 2009). Ca2+ recruitment from internal stores is an established event during cold shocks (Knight et  al., 1996; Gong et  al., 1998; White and Broadley, 2003). Further evidence (Furch et  al., 2009; Thorpe et al., 2010; van Bel, 2011a) also points to the ER as an important Ca2+ store which seems a major reason why ER stacks have been retained during sieve element evolution (Sjolund and Shih, 1983; van Bel, 2003). All in all, Ca2+ hotspots are probably created where high densities of Ca2+-permeable channels in the plasma membrane and an abundance of ER stacks meet (Fig. 4J; Hafke et al., 2009). In line with putative Ca2+ accumulation at these sites, the reactivity of forisomes increases when their tips are located in the vicinity of Ca2+ hotspots in sieve elements (Furch et  al., 2009). Further functional support for Ca2+ hotspots is provided by the fact that the forisome tips, being positioned between the ER stacks, are the only forisome parts that disperse as a reaction to weaker stimuli (Fig.  4J). The frequently perpendicular orientation of the ER stacks (Ehlers et al., 2000) facilitates insertion of the tips into a space (Furch et al., 2009), where Ca2+ levels may reach the threshold value needed for forisome dispersion. The interstices of the ER offer an undisturbed microenvironment for creation of Ca2+ hotspots (Furch et al., 2009; Hafke et al., 2009). Correlation between the Ca2+ concentration in hotspots and forisome responses As argued above, forisomes can be regarded as innate indicators for the Ca2+ thresholds and the approximate Ca2+ concentration in hotspots. APs seem to generate low-concentrated hotspots, since APs seldom lead to forisome responses (Fig. 4J) or, if they do so, lead to a slight wiggling of the forisome tails or a partial dispersion of the tips. Forisome dispersion coincident with prolonged EPW profiles indicates strong accumulation of Ca2+ at sieve element hotspots in response to VPs (Fig. 4J). Violent stimuli (burning, crushing) trigger APs and VPs in parallel that will collaborate in generating Ca2+ influx, the more so as Ca2+ potentiates it own hotspot concentration via Ca2+ liberation from ER stacks (Fig. 4J; Hafke et al., 2009). Ca2+ hotspots could also be meaningful for callose synthesis since Ca2+ concentrations required for this reaction greatly exceed those in the sieve tube sap (Hafke et  al., 2009; Furch et al., 2009), at least in vitro (Colombani et al., 2004). Involvement of the sieve element cytoskeleton in EPW propagation It has been known for a long time that cold shocks induce transient blockage of sieve tubes (Pickard and Minchin, 1990), which has been related to Ca2+ channel reactivity (Thorpe et  al., 2010). Cold shocks [>0.5  °C s–1 (Thorpe et  al., 2010) or 4.2  °C in less than a second (Hafke et  al., 2013)] induced sieve element depolarization followed by forisome dispersion (Fig.  5A). The depolarization was strongly reduced by the Ca2+ channel blocker La3+, and forisome dispersion also failed to occur. The apparent cold-triggered Ca2+ influx was originally ascribed to gating of mechano-sensitive Ca2+-permeable channels (Fig. 5A; Thorpe et al., 2010) due to a change of the tensile force exerted on the plasma membrane. Involvement of the cytoskeleton, however, was not excluded given the resemblance between cold-induced Ca2+ influx into the mictoplasm and other cell types (Knight et al., 1996; Plieth et al., 1999; White, 2009). The latter has become more plausible after the recent discovery of a complete, dense actin network in sieve elements (Fig. 5E; Hafke et al., 2013). The actin disruptor latrunculin A (Lat A) has similar inhibitory effects on the cold-induced events (depolarization and forisome dispersion) in the presence or absence of La3+ (Fig. 5B; Hafke et al., 2013). Their equal impact indicates that LatA and La3+ target the same Ca2+ influx mechanism that is linked in some way to actin action. All in all, the presumptive interaction between Ca2+permeable channels and actin (Hafke et  al., 2013) predicts that the cytoskeleton plays a pivotal role in EPW propagation. Interaction between Ca2+ channels and the cytoskeleton in sieve elements is further supported by an intimate connection between the plasma membrane and the actin meshwork as indicated by dense anti-actin immunochemical labelling of the face of the plasma membrane (Fig. 5C; Hafke et al., 2013). Forisomes probably must be kept in position for optimal sensing of Ca2+ changes in hotspots, although no compelling evidence for anchoring has been obtained thus far. The virtual absence of actin on dispersed forisomes (Fig. 5D; Hafke et al., 2013) seems to exclude that forisomes are linked to actin, unless actin filaments are torn apart during the fixation procedure due to forisome swelling. Other modes of linkage could be provided by protein filaments of unknown nature that anchor sieve element organelles to the plasma membrane (Fig. 5F; Ehlers et al., 2000) or tubulin. As expected, tubulin occurs in sieve elements (JBH, unpublished results) based on preliminary experiments using the tubulin disruptor oryzalin. Actin and tubulin may be coupled to different Ca2+ channels, since the activity of depolarization-activated (Mazars et al., 1997; Thion et al., 1998) and mechano-sensitive (Wang et al., 2004; Zhang et al., 2007) Ca2+-permeable channels was modulated by microtubules and microfilaments, respectively, in other cell types. The interaction between cytoskeleton elements and Ca2+ channels and the inherent cytoskeleton involvement in shaping Ca2+ signatures and triggering intracellular signal cascades (Mazars et  al., 1997; Trewavas and Malho, 1997; Drøbak et  al., 2004; Davies and Stankovic, 2006) may be of paramount significance for EPW propagation. The question now arises as to how actin and Ca2+-permeable channels are linked (Fig.  5F). In general, cytoskeleton disruptors that destabilize either F-actin (Liu and Luan, 1998; Wang et  al., 2004; Zhang et al., 2007) or microtubules (Thion et al., 1998) affect the action of ion channels. Protein complexes designated as ‘transducons’, which consist of an aggregate of receptors, Ca2+-permeable channels, bound calmodulin, protein kinases, and phosphatases, have been invoked to explain the intimate interaction between Ca2+ and the cytoskeleton (Trewavas and Malho, 1997) via members of the NETWORKED superfamily (Deeks et al., 2012). Transducons have been proposed to be tethered by integrins (Trewavas and Malho, 1997; Knepper et al., 2011) to the plasma membrane and cell wall. Ca2+-induced sieve element occlusion mechanisms: a safety design? Full sieve element occlusion achieved by forisomes had been a matter of debate (Peters et al., 2006) until in vitro experiments demonstrated that the swelling capacity was more than sufficient (Knoblauch et al., 2012). In intact V. faba plants, forisomes dispersed within seconds after EPW passage induced by burning and recontracted after 10–20 min (Fig.  6A–C; Furch et al., 2007, 2009). Forisome dispersion turned out to be quicker than callose production (Furch et al., 2007, 2009). By the time that a forisome had recontracted, probably due to active Ca2+ removal (e.g. Huda et al., 2013), callose build-up reached its maximum followed by a slower degradation up to 3 h (Fig. 6D; Furch et al., 2007, 2008, 2010). Both modes of occlusion are under the control of Ca2+ ions (Fig. 6E, F), the difference being that protein-mediated occlusion may have a lower Ca2+ threshold (50  μM; Furch et  al., 2009; Hafke et  al., 2009) than callose synthesis. In vitro callose synthesis required a concentration of 8 mM Ca2+ (Colombani et  al., 2004). Alternatively, the time lag of maximal callose deposition under the control of the Cal7 gene (Barratt et al., 2011; Xie et al., 2011) is due to the relative slowness of the complex de novo callose synthesis (Chen and Kim, 2009) with a Vmax of 45.5 nmol min–1 mg–1 (Li and Brown, 1993). A dual sieve plate occlusion mechanism was also found in Cucurbita maxima (Furch et al., 2010). Rapid, apparent coagulation of the phloem proteins, PP1 and PP2, several centimetres away from the site of burning preceded callose deposition (Furch et al., 2010). As shown for various species, callose deposition reaches its maximum after 10–30 min and is gradually degraded thereafter (Furch et al., 2008). Commensurate with the amount of callose deposited, PPUs reopen before the sieve pores do (Furch et al., 2007, 2008, 2010). Its occurrence in systematically distant families suggests that dual occlusion is widespread and functions to safeguard sieve tube contents. In this safety design, protein occlusion guarantees quick sieve plate sealing, which bridges the time until callose deposition is completed (van Bel et al., 2011a). Although evidence in favour of a dual occlusion mechanisms is growing, numerous questions have to be addressed, in particular concerning the diversity of occlusion mechanisms. (i) It is unclear if the Ca2+ thresholds for protein reactivity and callose synthesis are different (Fig.  6E, F). There might be a vast spectrum of Ca2+ thresholds needed for diverse occlusion mechanisms (Furch et  al., 2007, 2008, 2009, 2010), which in particular pertains to VPs which are positively related to the stimulus strength (Stahlberg and Cosgrove, 1997; Stahlberg et  al., 2006). (ii) Most probably, not every protein clogging event in sieve tubes is Ca2+ dependent. Forisomes comprise SEO proteins (Pélissier et al., 2008), a widespread family among dicotyledons (Rüping et al., 2010; Anstead et  al., 2012; Ernst et  al., 2012; Jekat et  al., 2012). SEO proteins are claimed to be Ca2+ binding in general (Ernst et al., 2012), although firm direct evidence seems to be lacking. Furthermore, PPs in cucurbit sieve tubes (Cronshaw and Sabnis, 1990; Dinant et al., 2003) do not belong to the SEO family (Ernst et al., 2012) and may react to (reactive) oxygen (species) or interact due to oxidation (Alosi et al., 1988). (iii) Since structural phloem-specific proteins are virtually absent in grasses (Eleftheriou, 1990), protein occlusion seems less important there (van Bel, 2003). However, emergence of protein plugs in gramineous sieve tubes indicates the presence of soluble proteins that are able to coagulate in response to injury (Will et  al., 2009). (iv) The capacity to remove Ca2+ from sieve elements may be decisive for the reversibility of occlusion and achieved by a battery of Ca2+ efflux facilitators (Kudla et al., 2010; Huda et al., 2013). Their activities bear strongly on the mechanisms of Ca2+ homeostasis in sieve elements. Ca2+ efflux facilitators such as Ca2+ ATPases at the plasma membrane and the endomembranes, as well as Ca2+ exchangers (McAinsh and Pittman, 2009; Kudla et al., 2010) could modulate Ca2+ signatures and are responsible for cytoplasmic Ca2+ homeostasis. It has been speculated that soluble Ca2+-binding proteins fine-tune and shape Ca2+ transients during signalling (McAinsh and Pittman, 2009). Given the wealth of soluble proteins in sieve tube sap (e.g. Nakamura et al., 1993; Lin et al., 2009; Gaupels et al., 2012; Dinant and Lucas, 2013), this type of Ca2+ sequestration may be of paramount importance for Ca2+ buffering in the sieve element. Ca2+-binding proteins associated with the cytoskeleton could also act as modulators of Ca2+ signatures (Malho et al., 1998). Relationships between Ca2+-mediated sieve element occlusion and pathogenic attacks Apart from the involvement of Ca2+-permeable channels in the long-distance signalling of pathogenic infections and the implementation of defence mechanisms (e.g. Lecourieux et al., 2006; Cheval et al., 2013), Ca2+ channels are also locally and directly involved in putting up anti-pathogenic barriers. As reported below, penetration of aphid stylets and the presence of phytoplasmas elicit sieve tube occlusion mechanisms related to Ca2+ influx. Aphid infestation Sieve elements in Lupinus albus and V. faba occlude instantaneously by virtue of forisome dispersion in response to micropipette impalement (tip diameter 1 μm) due to Ca2+ influx (van Bel and van Rijen, 1994; Knoblauch and van Bel, 1998). After impalement, cell wall Ca2+ will diffuse into the sieve element via the wound edges created by the micropipette (Fig.  7A; Will and van Bel, 2006). Concomitantly, sieve element turgor is dissipated by the large micropipette volume, which may affect the gating of mechano-sensitive Ca2+-permeable channels (Fig. 7A). Remarkably, aphid stylets which have a similar tip diameter do not cause forisome dispersion (Walker and Medina-Ortega, 2012). Pressure loss into the stylet is minimal due to the minute volume and sealing of the wound edges by gel saliva (Miles, 1999; Tjallingii, 2006; Will et  al., 2013) so that passive Ca2+ influx and activation of Ca2+-permeable channels are constrained (Fig. 7A; Will and van Bel, 2006). When sieve element occlusion is triggered by remote burning, feeding aphids react within a few seconds (Will et al., 2007; Furch et al., 2010). Several aphid species switch from ingestion to secretion of watery saliva probably to counteract sieve tube occlusion by Ca2+ binding (Fig. 7B–D; Will et al., 2007, 2009). In vitro studies using forisomes (Will et al., 2007), biochemical techniques (Will et  al., 2007), and proteomics (Carolan et  al., 2009; Rao et  al., 2013)  confirmed that watery saliva contains Ca2+-binding proteins. In vitro, dispersion of Ca2+-treated forisomes was reversed by addition of the Ca2+ chelator EDTA or watery saliva concentrate from the aphid species Megoura viciae (Fig. 7E–I; Will et al., 2007). As a supplementary function, Ca2+-binding proteins may interfere with Ca2+-mediated defence and signalling mechanisms (Will and van Bel, 2008). Up-regulation of genes encoding calmodulin, calmodulin-like proteins, calcium-dependent protein kinases, and calcium-binding reticulin in response to infestation (Coppola et  al., 2013) suggests a major role for Ca2+ in plant defence against aphids. The fact that Ca2+-binding proteins have been identified in the phloem-feeding green rice leafhopper (Hattori et  al., 2012) indicates that comparable strategies for suppression of plant defence may exist in diverse hemipteran families. Thus far, however, evidence in favour of in vivo suppression of Ca2+-induced sieve element occlusion by aphid saliva is lacking. On the contrary, forisome reversibility after leaf tip burning was found to be similar in distant sieve tubes with or without aphid stylet penetration in intact broadbean plants (Medina-Ortega and Walker, 2013). Infection by phytoplasms Phytoplasmas are frequently transmitted to plants by phloem-feeding leafhoppers and distributed via the sieve tubes (Christensen et  al., 2005; McLean and Hogenhout, 2013) which become occluded in response to the infection (Braun and Sinclair, 1978; Kartte and Seemüller, 1991; Musetti and Favali, 1999). Phytoplasma-infected sieve tubes in V.  faba contain consistently dispersed forisomes (Fig. 7J–M) hinting at Ca2+ levels appreciably higher than in healthy sieve tubes (Musetti et al., 2013). The Ca2+ concentration is indeed higher in infected sieve tubes (Fig. 7N, O). Moreover, the sieve tubes are sealed with thick deposits of callose (Fig. 7P, Q; Musetti et al., 2013). Undoubtedly, phytoplasmas induce Ca2+ influx with inherent consequences for forisome dispersion and callose synthesis. Thus far, it is unclear if sieve element occlusion is part of the plant’s strategy against phytoplasma spread or if phytoplasmas induce and explore symplasmic isolation for undisturbed multiplication. Physiological and genetic impact of EPWs A diversity of physiological and genetic remote responses to EPWs have been reported, many of which are likely to be due to Ca2+ influx (Kudla et al., 2010). EPWs induce the expression of the proteinase inhibitor gene (pin2; Wildon et  al., 1992; Pena-Cortes et  al., 1995; Stankovic and Davies, 1997) and other genes (Davies, 2004) in distant parts of tomato plants. EPW propagation and gene expression are linked by the fact that Ca2+ influx and a consequent, transient increase in cytosolic Ca2+ is required for pin2 gene expression (Fisahn et al., 2004). Interestingly, the level of IP3, a second messenger potentially responsible for Ca2+ liberation from ER stacks (Gilroy et al., 1990; Krol et al., 2003, 2004), abruptly increases after EPW passage (Davies, 2004). Furthermore, touch-triggered EPWs evoke an arsenal of transcriptional downstream responses pertinent to 2.5% of the genes (Braam, 2005) including the enhanced expression of Ca2+-binding proteins (Lee et  al., 2005). These data strongly suggests a relationship between EPWs, Ca2+ influx, and remote effects on gene expression. A range of physiological responses to EPWs have been documented (Retivin et al., 1997; Fromm and Lautner, 2012). A  Ca2+-controlled shutdown of proton pumps (Kinoshita et  al., 1995; Hafke et  al., 2013) during and after passage of a VP triggered by heating led to a transient decrease in the cytosolic pH from 7.0 to 6.4 and a concomitant increase of the apoplasmic pH from 4.5 to 5.2 (Grams et al., 2009). The lowered cytosolic pH, in turn, may bring about depressed CO2 uptake rates and reduced photosynthetic quantum yields (Koziolek et  al., 2004; Lautner et  al., 2005; Grams et  al., 2009). In keeping with these observations, occasional sudden changes in osmotic potential of sieve tubes (Knoblauch et al., 2001) or pressure waves may cause VPs involved in the restoration of the source–sink balance. Remarkably, APs triggered by sudden flooding of drought-stressed roots produced the opposite effect: they induced an increase in stomatal conductance and photosynthesis (Grams et al., 2007) in the absence of appreciable pH changes. Responses of distant cells to EPWs may depend on two major factors: the Ca2+ signatures in question and the equipment of the recipient cells. Ca2+ signatures (McAinsh and Pittman, 2009) will depend on the Ca2+-permeable channels involved, their cellular location, their Ca2+-mediated interaction, the available Ca2+ binding components, and the final Ca2+ compartmentation. There is a wealth of data on Ca2+ receptors and downstream signalling cascades (e.g. Dodd et al., 2010), and it seems that Ca2+ influx is of crucial importance for initiation of numerous cascades (Sanders et  al., 2002; Kudla et al., 2010). The diverse Ca2+ reactivity of various cell types has not yet been studied in detail, but there are obvious differences. Differential Ca2+ responsiveness of diverse cell types along the phloem pathway is exemplified by: sieve plate occlusion in sieve elements (Furch et  al., 2007, 2009), production of NO in companion cells (Gaupels et al., 2008), systemin production in phloem parenchyma cells (Narvaez-Vazquez and Ryan, 2004), and massive water release in pulvinar flexor cells (Fleurat-Lessard and Bonnemain, 1978). It will be fascinating to explore further the impact of EPWs on gene expression, metabolism, and physiology of distant conductive elements and adjoining cells. Speculations on whole-plant effects of EPW-modulated Ca2+ waves Symplasmic organization of phloem strands changes in response to Ca2+ fluxes Ca2+ influx during APs mostly is insufficient to induce forisome dispersion and sieve element occlusion (Fig. 4J). After passage of EPWs of sufficient strength, namely VPs or, in particular, a combination of VPs and APs (Fig. 4J), Ca2+ levels exceed an activation threshold. The resultant occlusion of the intercellular corridors may impose a transient symplasmic reorganization of the sieve tube tracks (Fig.  8). Apart from the proven occlusion of sieve plates and PPUs (Figs. 4,6), elevated Ca2+ levels in the adjoining parenchyma cells may induce the deposition of callose collars around their plasmodesmata, as demonstrated for several tissues (e.g. Tucker, 1990; Kauss and Jeblick, 1991; Radford et al., 1998; Holdaway-Clarke et  al., 2000; Sivaguru et  al., 2000, 2005; Michard et  al., 2011). Formation of callose deposits blocking photoassimilate loading by sieve tubes in response to APs (Fromm et al., 2013) is in agreement with this concept. Without the usual interaction with their neighbours, occlusion of symplasmic contacts would render vascular cells temporarily autonomous units (Figs. 8,9). Under these conditions, vascular cells may be able to switch to other cascades implementing more discrete metabolic or genetic programmes. The effects of (transient) plasmodesmal closure on cell autonomy were demonstrated for the differentiation of the stomatal apparatus (Palevitz and Hepler, 1985), the divergent development of the sieve element and companion cell (van Bel and van Rijen, 1994), the formation of symplasmic domains (Ehlers et al., 1999), the synchronization of metabolic activity (Ehlers and Kollmann, 2000), and the explosive elongation of cotton hair cells (Ruan et  al., 2001). As soon as the lifelines (PPUs) between companion cells and sieve elements have been restored, Ca2+-induced products can be released into the sieve elements and translocated to target cells when the sieve pores become re-opened (van Bel et al., 2011a). Ca2+-triggered systemic signalling occurs in partly overlapping waves Lateral transfer of EPWs, either focused in the pulvini (Fleurat-Lessard and Bonnemain, 1978) or distributed along the entire pathway (Rhodes et  al., 1996), may reflect a fundamental difference between EPWs in animals and plants. Instead of the minor ion displacements occurring in animals, gating of ion channels causes massive ion displacement in plants (Pyatygin et  al., 2008). Apart from the regulation of Ca2+ influx, ion displacement in plants may strongly contribute to ion homeostasis (e.g. Mummert and Gradmann, 1991; Trebacz et al., 1994; Zimmermann and Felle, 2009). Dissemination of electrical signalling implies that both cells along the phloem pathway and those at the termini of the phloem track are targets for EPWs. The multitude of potential combinations of Ca2+ influx and its differential effects on diverse cell types potentiate the complexity of the responses and provide an endless wealth of possibilities (Kudla et  al., 2010; Dempsey and Klessig, 2012) exemplified by the ‘myriad plant responses’ to herbivores (Walling, 2000). We explore the possibility—as advanced before in a less elaborate way (van Bel and Ehlers, 2005)—that phloem-borne signalling passes through partly overlapping waves which are distinct in time scale, site of origin, and nature (Fig.  9). At the forefront of EPWs, Ca2+ ions are released into sieve elements which may readily attach to constitutive Ca2+-binding proteins in the sieve tube sap such as Ca2+-dependent protein kinases (Nakamura et  al., 1993; Yoo et  al., 2002; Gaupels et al., 2012). Thus, the first wave of signals (time scale: seconds to minutes to arrive in target cells) may include free Ca2+ ions accompanied by Ca2+-activated or Ca2+-binding proteins. As a result, Ca2+ signatures induce proactive responses to imminent changes. The signatures will depend on the stimulus; that is, disparate signatures are obtained from diverse Ca2+permeable channels which are functionally linked with different cytoskeleton components (Mazars et al., 1997; Thion et al., 1998; Wang et al., 2004; Zhang et al., 2007). A second wave of signals (time scale: minutes to hours) may comprise compounds from the vascular parenchyma that are readily manufactured under the control of Ca2+ influx. If the EPW is accompanied by symplasmic reorganization, the longer residence time could make the stagnant contents of sieve elements into reaction vessels for Ca2+ binding to constitutive sieve element proteins, and vascular cells may follow alternative signalling cascades as argued above. Thus, without sieve element occlusion, the compounds released into the sieve tubes for further translocation may differ from those released after relief of symplasmic re-organization (Fig.  9). During this second stage, various parallel cascades may be initiated by Ca2+ influx. For instance, calmodulin-like and calmodulin (McCormack and Braam, 2003; Lee et al., 2005; McCormack et  al., 2005), as well as other specific Ca2+binding proteins (White and Broadley, 2003; Kudla et  al., 2010) are attached to the cytoskeleton (Malho et al., 1998). In this way, information conferred by Ca2+ signatures is decoded and transformed into protein–protein interactions, resulting in Ca2+-dependent phosphorylation cascades like transcriptional responses that lead to downstream reactions (Luan et al., 2002; Sanders et al., 2002; Kudla et al., 2010). Whether Ca2+ is directly related to the synthesis of jasmonic acid (Fisahn et al., 2004) and/or salicylic acid is uncertain, but there seems little doubt that Ca2+ ions are engaged in the action of jasmonic acid (Munemasa et al., 2011) and salicylic acid (Du et al., 2009; Boursiac et al., 2010). In addition, cytosolic Ca2+ elevation is linked to downstream nitric oxide production, as shown for companion cells (Gaupels et al., 2008) via the intervention of calmodulin-(like) proteins (Ma et al., 2008). The third wave (time scale: hours) would encompass longterm implementation of Ca2+ effects exemplified by the production of various types of RNA (Kehr and Buhtz, 2013), proteins (Lin et  al., 2009; Dinant and Lucas, 2013), and even lipidic substances (Guelette et al., 2012) present in the sieve tube sap. For the impact of Ca2+ signals on the production of macromolecules, the reader is referred to an excellent review (Kudla et al., 2010), but a few examples are given here. Ca2+ signals are converted into transcriptional responses for a fair number of genes (Lee et al., 2005; Kaplan et al., 2006) which may comprise ~3% of the protein-coding genes in Arabidopsis (Kudla et al., 2010). Many of these expression responses depend on Ca2+ regulation of the transcription factors (e.g. Finkler et  al., 2007). As an interesting note in the present context, one of these transcription factors interacts with the promoter of AtEDS1, a regulator of salicylic acid synthesis (Du et al., 2009). Macromolecules produced in the vascular cells and released into the sieve tube sap via PPUs (Lucas et al., 2001; Chen and Kim, 2006; Lough and Lucas, 2006; Ding and Itaya, 2007; Lin et  al., 2009) might find their way to target cells by molecular tagging (zip codes) so that compounds required for local and remote use can be distinguished (van Bel et  al., 2011b). In this way, macromolecules are recognized to remain within the sieve element into which they had been released or move either to companion cells along the pathway (Fisher et al., 1992; Golecki et al., 1999) or to sink cells. Interactions on the interface between ER stacks and the sieve element cytoskeleton may play a crucial part in the distribution of macromolecules inside the sieve element and delivery of macromolecules into the sieve tube sap. Presumably, some of the macromolecules are back-trafficked into companion cells by the aid of non-cell autonomous agents (Schulz, 1999; Itaya et al., 2000; Lucas et al., 2009). This ‘molecular hopping’ (van Bel et al., 2011b) may provide a complex basis for amplification or attenuation of systemic signals. Macromolecules enter sink cells via permanently widened plasmodesmata (Fisher and Cash-Clarke, 2000), each of which may demand specific entrance codes (Foster et al., 2002). Concluding remarks This review is a plea for further research on the link between EPWs and chemical systemic signalling. It appears to be worth investigating if and to what extent EPWs provide a common basis for the rapid distribution of Ca2+ signals. A  limited number of studies demonstrate the immense and remote effects of EPWs on the genetics and physiology of plants. There may be a few prime targets for investigation. (i) Which Ca2+-permeable channels are involved in the propagation of EPWs and the processing of electrical information in vascular cells? (ii) Is there an impact of a temporary symplasmic organiza tion on the production of signalling substances? (iii) Do the Ca2+ signatures and the resultant cascades depend on the nature of electrical signalling; for example, is there a difference between Ca2+ signatures induced by VPs or APs? (iv) Do the Ca2+ signatures and the resultant cascades depend on the receptor or target cell types; that is, are there cellspecific responses to identical Ca2+ signatures? (v) What is the relationship between Ca2+ influx and the production of various mobile signals along the pathway such as phytohormones, reactive oxygen species, lipidic substances, proteins, and RNA species? Aki T, Shigyo M, Nakano R, Yoneyama T, Yanagisawa S. 2008. Nano scale proteomics revealed the presence of regulatory proteins including three FT-like proteins in phloem and xylem saps from rice. Plant and Cell Physiology 49, 767–790. Alarcon JJ, Malone M. 1994. Substantial hydraulic signals are triggered by leaf-biting insects in tomato. Journal of Experimental Botany 45, 953–957. Alosi MC, Melroy DL, Park RB. 1988. The regulation of gelation of phloem exudate from Cucurbita fruit by dilution, glutathione, and glutathione reductase. Plant Physiology 86, 1089–1094. Anstead JA, Froehlich DR, Knoblauch M, Thompson GA. 2012. Arabidopsis P-protein filament formation requires both AtSEO1 and AtSEO2. Plant and Cell Physiology 53, 1033–1042. Barnes A, Bale J, Constantinidou C, Ashton P, Jones A, Pritchard J. 2004. Determining protein identity from sieve element sap in Ricinus communis L. by quadrupole time of flight (Q-TOF) mass spectrometry. Journal of Experimental Botany 55, 1473–1481. Barratt DHP, Kolling K, Graf A, Pike M, Calder G, Findlay K, Zeeman SC, Smith AM. 2011. Callose synthase GSL7 is necessary for normal phloem transport and inflorescence growth in Arabidopsis. Plant Physiology 155, 328–341. Bauer CS, Hoth S, Haga K, Phillipar K, Aoki N, Hedrich R. 2000. Differential expression and regulation of K+ channels in the maize coleoptile: molecular and biophysical analysis of cells isolated from cortex and vasculature. The Plant Journal 24, 139–145. Behnke HD, Sjolund RD. 1990. Sieve elements. Comparative structure, induction and development. Berlin: Springer. Benolken RM, Jacobson SM. 1970. Response properties of a sensory hair excised from Venus’s flytrap. Journal of General Physiology 56, 64–82. Bewell MA, Maathuis FJM, Allen GJ, Sanders D. 1999. Calciuminduced calcium release mediated by voltage activated cation channel in vacuolar vesicles from red beet. FEBS Letters 458, 41–44. Boari F, Malone M. 1993. Rapid and systemic hydraulic signals are induced by localized wounding in a wide range of species. Journal of Experimental Botany 44, 741–746. Bolsover S, Silver RA. 1991. Artifacts in calcium measurements: recognition and remedies. Trends in Cell Biology 1, 71–74. Boursiac Y, Lee SM, Romanowsky S, Blank R, Sladek C, Chung WS, Harper JF. 2010. Disruption of the vacuolar calcium-ATPases in Arabidopsis results in the activation of a salicylic acid-dependent programmed cell death pathway. Plant Physiology 154, 1158–1171. Braam J. 2005. In touch: plant responses to mechanical stimuli. New Phytologist 165, 373–389. Brauer M, Zhong W-J, Schobert C, Sanders D, Komor E. 1998. Free calcium ion concentration in the sieve-tube sap of Ricinus communis L. seedling. Planta 206, 103–107. Braun EJ, Sinclair WA. 1978. Translocation in phloem necrosisdiseased American elm seedlings. Pythopathology 68, 1733–1737. Brinckmann E, Lüttge U. 1974. Lichtabhängige Membranpotentialschwankungen und deren interzelluläre Weiterleitung bei panaschierten Photosynthesemutanten von Oenothera. Planta 119, 45–57. Buchen B, Hensel D, Sievers A. 1983. Polarity in mechanoreceptor cells of trigger hairs of Dionaea muscipula Ellis. Planta 158, 458–468. Carolan JC, Fitzroy CIJ, Ashton PD, Douglas AE, Wilkinson TL. 2009. The secreted salivary proteome of the pea aphid Acyrthosiphon pisum characterised by mass spectrometry. Proteomics 9, 2457–2467. Chaffey N, Barlow P. 2002. Myosin, microtubules, and microfilaments: co-operation between cytoskeletal components during cambial cell division and vascular differentiation in trees. Planta 214, 526–536. Chen X-Y, Kim J-Y. 2006. Transport of macromolecules through plasmodesmata and the phloem. Physiologia Plantarum 126, 560–571. Chen X-Y, Kim J-Y. 2009. Callose synthesis in higher plants. Plant Signaling and Behavior 4, 489–492. Cheval C, Aldon D, Galaud JB, Ranty B. 2013. Calcium– calmodulin-mediated regulation of plant immunity. Biochimica et Biophysica Acta 1833, 1766–1771 Christensen NM, Axelsen KB, Nicolaisen M, Schulz A. 2005. Phytoplasmas and their interactions with hosts. Trends in Plant Science 10, 526–535. Colombani A, Djerbi S, Bessueille L, Blomqvist K, Ohlsson A, Berglund T, Teeri TT, Bulone V. 2004. In vitro synthesis of (1→3)-β-d -glucan (callose) and cellulose by detergent extracts of membranes from cell suspension cultures of hybrid aspen. Cellulose 11, 313–327. Coppola V, Coppola M, Rocco M, et al. 2013. Transcriptomics and proteomic analysis of a compatible plant–aphid interaction reveals a predominant salicylic-acid plant response. BMC Genomics 14, 515. Cronshaw J, Sabnis DD. 1990. Phloem proteins. In: Behnke H-D, Sjolund RD, eds. Sieve elements. Comparative structure, induction and development. Berlin: Springer, 257–283. Davies E. 2004. New functions for electrical signals in plants. New Phytologist 161, 607–610. Davies E. 2006. Electrical signals in plants: facts and hypotheses. In: Volkov AG, ed. Plant electrophysiology. Berlin: Springer, 407–422. Davies E, Stankovic B. 2006. Electrical signals, the cytoskeleton and gene expression: a hypothesis on the coherence of the cellular responses to environmental insult. In: Baluska F, Mancuso S, Volkmann D, eds. Communication in plants. Berlin: Springer, 309–320. Davies E, Zawadzki T, Witters D. 1991. Electrical activity and signal transmission in plants: how do plants know? In: Penel C, Grepin H, eds. Plant signalling, plasma membrane and change of state. University of Geneva, 119–137. Deeken R, Geiger D, Fromm J, Koroleva O, Ache P, LangenfeldHeyser R, Sauer N, May ST, Hedrich R. 2002. Loss of AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216, 334–344. Deeks MJ, Calcutt JR, Ingle EKS, et al. 2012. A superfamily of actin-binding proteins at the actin-membrane nexus of higher plants. Current Biology 22, 1595–1600. Demidchik V, Bowen HC, Maathuis FJM, Shabala SN, Tester MA, White PJ, Davies JM. 2002. Arabidopsis thaliana root nonselective cation channels mediate calcium uptake and are involved in growth. The Plant Journal 32, 799–808. Demidchik V, Maathuis FJM. 2007. Physiological roles of nonselective cation channels in plants: from salt stress to signaling and development. New Phytologist 175, 387–404. Dempsey DA, Klessig D. 2012 SOS—too many signals for systemic acquired resistance. Trends in Plant Science 17, 538–535. Demuro A, Parker I. 2006. Imaging single-channel calcium microdomains. Cell Calcium 40, 413–422. Dinant S, Clark AM, Zhu Y, Vilaine F, Palauqui JC, Kusiak C, Thompson GA. 2003. Diversity of the superfamily of phloem lectins (phloem protein 2) in angiosperms. Plant Physiology 131, 114–128. Dinant S, Lucas WJ. 2013. Sieve elements: puzzling activities deciphered through proteomics studies. In: Thompson GA, van Bel AJE, eds. Phloem. Molecular cell biology, systemic communication, biotic interactions. Chichester, UK: Wiley-Blackwell, 157–185. Ding B, Itaya A. 2007. Control of directional macromolecular trafficking across specific cellular boundaries: a key to integrative plant biology. Journal of Integrative Plant Biology 49, 1227–1234. Ding JP, Pickard BG. 1993. Modulation of mechanosensitive calcium-selective cation channels by temperature. The Plant Journal 3, 713–720. Dodd AN, Kudla J, Sanders D. 2010. The language of calcium signaling. Annual Review of Plant Biology 61, 593–620. Drøbak BK, Franklin-Tong VE, Staiger CJ. 2004. The role of the actin cytoskeleton in plant cell signaling. New Phytologist 163, 13–30. Du L, Ali GS, Simons KA, Hou J, Yang T, Reddy AS, Poovaiah BW. 2009. Ca2+/calmodulin regulates salicylic acid-mediated plant immunity. Nature 437, 741–745. Dziubinska H, Trebacz K, Zawadzki T. 2001. Transmission route for action potentials and variation potentials in Helianthus annuus L. Journal of Plant Physiology 158, 1167–1172. Ehlers K, Binding H, Kollmann R. 1999. The formation of symplasmic domains by plugging of plasmodesmata: a general event in plant morphogenesis. Protoplasma 209, 181–192. Ehlers K, Knoblauch M, van Bel AJE. 2000. Ultrastructural features of well-preserved and injured sieve elements: minute clamps keep the phloem transport conduits free for mass flow. Protoplasma 214, 80–92. Ehlers K, Kollmann R. 2000. Synchronization of mitotic activity in protoplast-derived Solanum nigrum L. microcalluses is correlated with plasmodesmal connectivity. Planta 210, 269–278. Eleftheriou EP. 1990. Monocotyledons. In: Behnke H-D, Sjolund RD, eds. Sieve elements. Comparative structure, induction and development. Berlin: Springer, 139–159. Engleman E. 1965. Sieve elements of Impatiens sultanii. II. Developmental aspects. Annals of Botany 29, 103–104. Ernst AM, Jekat SB, Zielonka S, Müller B, Neumann U, Rüping B, Twyman RM, Krzyzanek V, Prüfer D, Noll G. 2012. Sieve element occlusion (SEO) genes encode structural phloem proteins involved in wound sealing of phloem. Proceedings of the National Academy of Sciences, USA 109, E1980–E1989. Esau K. 1969. The phloem. Encyclopedia of plant anatomy , Vol. 5. Berlin: Bornträger. Evert RF. 1990. Dicotyledons. In: Behnke H-D, Sjolund RD, eds. Sieve elements. Comparative structure, induction and development. Berlin: Springer, 103–137. Felle HH, Zimmermann MR. 2007. Systemic signalling in barley through action potentials. Planta 226, 203–214. Finkler A, Ashery-Padan R, Fromm H. 2007. CAMTAs; calmodulinbinding transcription factors from plants to human. FEBS Letters 581, 3893–3898. Fisahn J, Herde O, Willmitzer L, Pena-Cortes H. 2004. Analysis of a transient increase in cytosolic Ca2+ during the action potential of higher plants with high temporal resolution: requirement of Ca2+ transients for induction of jasmonic acid synthesis and PINII gene expression. Plant and Cell Physiology 45, 456–459. Fisher DB, Cash-Clarke CE. 2000. Sieve tube unloading and postphloem transport of fluorescent tracers and proteins injected into sieve tubes via severed aphid stylets. Plant Physiology 123, 125–137. Fisher DB, Wu K, Ku MSB. 1992. Turnover of soluble proteins in wheat sieve tube. Plant Physiology 100, 587–592. Fleurat-Lessard P, Bonnemain J-L. 1978. Structural and ultrastructural characteristics of the vascular apparatus of the sensitive plant Mimosa pudica L. Protoplasma 94, 127–143. Fleurat-Lessard P, Roblin G. 1982. Comparative histocytology of the petiole and the main pulvinus in Mimosa pudica L. Annals of Botany 50, 83–92. Foster TM, Lough TJ, Emerson SJ, Lee RH, Bowman JL, Foster RLS, Lucas WJ. 2002. A surveillance system regulates selective entry of RNA into the shoot apices. The Plant Cell 14, 1497–1508. Froelich DF, Mullendore DL, Jensen KH, Ross-Elliott TJ, Anstead JA, Thompson GA, Pelissier HC, Knoblauch M. 2011. Phloem ultrastructure and pressure flow; SEO protein agglomerations do not affect translocation. The Plant Cell 23, 4428–4445. Fromm J. 1991. Control of phloem unloading by action potentials in Mimosa. Physiologia Plantarum 83, 529–533. Fromm J, Bauer T. 1994. Action potentials in maize sieve tubes change phloem translocation. Journal of Experimental Botany 45, 463–469. Fromm J, Fei H. 1998. Electrical signaling and gas exchange in maize plants of drying soil. Plant Science 132, 203–213. Fromm J, Hajirezaei M-R, Becker VK, Lautner S. 2013. Electrical signalling along the phloem and its physiological response in the maize leaf. Frontiers in Plant Science 4, 239. Fromm J, Hajirezaei M, Wilke I. 1995. The biochemical response of electrical signaling in the reproductive system of Hibiscus plants. Plant Physiology 109, 375–384. Fromm J, Lautner S. 2006. Characteristics and functions of phloemtransmitted electrical signals in higher plants. In: Baluska F, Mancuso S, Volkmann D, eds. Communication in plants. Berlin: Springer, 321–332. Fromm J, Lautner S. 2012. Generation, transmission, and physiological effects of electrical signals in plants. In: Volkov AG, ed. Plant electrophysiology. Berlin: Springer, 207–232. Fromm J, Spanswick R. 1993. Characteristics of action potential in willow (Salix viminalis L). Journal of Experimental Botany 44, 1119–1125. Furch ACU, Hafke JB, Schulz A, van Bel AJE. 2007. Ca2+mediated remote control of reversible sieve tube occlusion in Vicia faba. Journal of Experimental Botany 58, 2827–2838. Furch ACU, Hafke JB, van Bel AJE. 2008. Plant- and stimulusspecific variations in remote-controlled sieve-tube occlusion. Plant Signaling and Behavior 3, 858–861. Furch ACU, van Bel AJE, Fricker MD, Felle HH, Fuchs M, Hafke JB. 2009. Sieve element Ca2+ channels as relay stations between remote stimuli and sieve tube occlusion. The Plant Cell 21, 2118–2132. Furch ACU, Zimmermann MR, Will T, Hafke JB, van Bel AJE. 2010. Remote-controlled stop of mass flow by biphasic occlusion in Cucurbita maxima. Journal of Experimental Botany 61, 3697–3708. Gaupels F, Furch ACU, Will T, Muir K, Kogel K-H, van Bel AJE. 2008. Nitric oxide generation in Vicia faba phloem cells reveals them to be sensitive detectors as well as possible systemic transducers of stress signals. New Phytologist 178, 634–646. Gaupels F, Sarioglu H, Beckman M, Hause B, Spannagl M, Draper J, Lindermayr C, Durner J. 2012. Deciphering systemic wound response of the pumpkin extrafascicular phloem by metabolomics and stable-isotope protein labeling. Plant Physiology 160, 2285–2299. Giavalisco P, Kapitza K, Kolasa A, Buhtz A, Kehr J. 2006. Towards the proteome of Brassica napus phloem sap. Proteomics 6, 896–909. Gilroy S, Read ND, Trewavas AJ. 1990. Elevation of cytoplasmic calcium by caged calcium or caged inositol triphosphate initiates stomatal closure. Nature 346, 769–771. Golecki B, Schulz A, Thompson GA. 1999. Translocation of structural P-proteins in the phloem. The Plant Cell 11, 127–140. Gong M, van der Luit AH, Knight MR, Trewavas AJ. 1998. Heatshock induced changes in intracellular Ca2+ level in tobacco seedlings in relation to thermotolerance. Plant Physiology 116, 429–437. Grams TEE, Koziolek C, Lautner S, Matyssek R, Fromm J. 2007. Distinct roles of electric and hydraulic signals on the reaction of leaf gas exchange upon re-irrigation in Zea mays. Plant, Cell and Environment 30, 79–84. Grams TEE, Lautner S, Felle HH, Matyssek R, Fromm J. 2009. Heat-induced electrical signals affect cytoplasmic and apoplastic pH as well as photosynthesis during propagation through the maize leaf. Plant, Cell and Environment 32, 319–326. Guelette BS, Benning UF, Hoffmann-Benning S. 2012. Identification of lipids and lipid-binding proteins in phloem exudates from Arabidopsis. Journal of Experimental Botany 63, 3603–3616. Hafke JB, Furch ACU, Fricker MD, van Bel AJE. 2009. Forisome dispersion in Vicia faba is triggered by Ca2+ hotspots created by concerted action of diverse Ca2+ channels in sieve elements. Plant Signaling and Behavior 4, 968–972. Hafke JB, van Amerongen JK, Kelling F, Furch ACU, Gaupels F, van Bel AJE. 2005. Thermodynamic battle for photosynthate acquisition between sieve tubes and adjoining parenchyma in transport phloem. Plant Physiology 138, 1527–1537. Imlau A, Truernit E, Sauer N. 1999. Cell-to-cell and long-distance trafficking of the green fluorescent protein in the phloem and symplastic unloading of the protein into the sinks. The Plant Cell 11, 309–322. Itaya A, Liang G, Woo Y-M, Nelson RS, Ding B. 2000. Nonspecific intercellular protein trafficking probed by green-fluorescent proteins in plants. Protoplasma 213, 165–175. Li L, Brown MR. 1993. β-Glucan synthesis in the cotton fiber. II. Regulation and kinetic properties of β-glucan synthases. Plant Physiology 101, 1143–1148. Liu K, Luan S. 1998. Voltage-dependent K+ channels as targets of osmosensing in guard cells. The Plant Cell 10, 1957–1970. Lin MK, Lee YJ, Lough TJ, Phinney BS, Lucas WJ. 2009. Analysis of the pumpkin phloem proteome provides insights into angiosperm sieve tube function. Molecular and Cellular Proteomics 8, 343–356. Llinàs R, Sugimori M, Silver RB. 1992. Microdomains of high calcium concentration in a presynaptic terminal. Science 256, 677–679. Llinàs R, Sugimori M, Silver RB. 1995. The concept of calcium concentration microdomains in synaptic transmission. Neuropharmacology 34, 1443–1451. Lough T, Lucas WJ. 2006. Integrative plant biology. Role of phloem long-distance macromolecular trafficking. Annual Review of Plant Biology 57, 203–232. Luan S, Kudla J, Rodriguez-Concepcion M, Yalovsky S, Gruissem W. 2002. Calmodulins and calcineurin B-like proteins: calcium sensors for specific signal response coupling in plants. The Plant Cell 14, 389–400. Lucas WJ, Ham B-K, Kim J-Y. 2009. Plasmodesmata—bridging the gap between neighbouring plant cells. Trends in Cell Biology 19, 495–503. Lucas WJ, Yoo B-C, Kragler F. 2001. RNA as a long-distance information macromolecule in plants. Nature Reviews of Molecular and Cellular Biology 2, 849–857. Lunevsky VZ, Zherelova OM, Vostrikov IY, Berestovsky GN. 1983. Excitation of Characeae cell membranes as a result of activation of calcium and chloride channels. Journal of Membrane Biology 72, 43–58. Ma W, Smigel A, Tsai YC, Braam J, Berkowitz GA. 2008. Innate immunity: cytosolic Ca2+ elevation is linked to downstream nitric oxide generation through the action of calmodulin or calmodulin-like proteins. Plant Physiology 148, 818–828. Malho R, Moutinho A, van der Luit A, Trewavas AJ. 1998. Spatial characteristics of calcium signalling: the calcium wave as a basic unit in plant cell calcium signalling. Philosophical Transactions of the Royal Society B: Biological Sciences 353, 1463–1473. Malone M. 1996. Rapid, long-distance signal transmission in higher plants. Advances in Botanical Research 22, 163–228. Malone M, Stankovic B. 1991. Surface potentials and hydraulic signals in wheat leaves following localized wounding by heat. Plant, Cell and Environment 14, 431–436. Mancuso S. 1999. Hydraulic and electrical transmission of woundinduced signals in Vitis vinifera. Australian Journal of Plant Physiology 26, 55–61. Marten I, Hoth S, Deeken R, Ketchum KA, Hoshi T, Hedrich R. 1999. AKT3, a phloem-localised K+ channel is blocked by protons. Proceedings of the National Academy of Sciences, USA 96, 7581–7586. controls the changes in cytosolic calcium of cold-shocked Nicotiana plumbaginifolia protoplasts. Cell Calcium 22, 413–420. Nakamura S, Hayashi H, Mori S, Chino M. 1993. Protein phosphorylation in the sieve tubes of rice plants. Plant and Cell Physiology 34, 927–933. Narvaez-Vazquez J, Ryan CA. 2004. The cellular localization of prosystemin: a functional role for phloem parenchyma in systemic wound signalling. Planta 218, 360–369. Ng CK, McAinsh MR. 2003. Encoding specificity in plant calcium signalling: hot-spotting the ups and downs and waves. Annals of Botany 92, 477–485. Okihara K, Ohkawa T, Tsutsui I, Kasai M. 1991. A Ca2+- and voltage-dependent Cl–-sensitive anion channel in the Chara plasmalemma: a patch clamp study. Plant and Cell Physiology 32, 593–601. Opritov VA, Pyatygin SS, Vodeneev VA. 2002. Direct coupling of action potential generation in cells of a higher plant (Cucurbita pepo) with the operation of an electrogenic pump. Russian Journal of Plant Physiology 49, 142–147. Overall R, Gunning BES. 1982. Intercellular communication in Azolla roots. I. Electrical coupling. Protoplasma 111, 151–160. Palevitz BA, Hepler PK. 1985. Changes in dye coupling of stomatal cells of Allium and Commelina demonstrated by microinjection of Lucifer yellow. Planta 164, 473–479. Parthasaraty MV, Pesacreta TC. 1980. Microfilaments in plant vascular cells. Canadian Journal of Botany 58, 807–815. Patrick JW, Offler CE. 1996. Post-sieve element transport of photoassimilates in sink regions. Journal of Experimental Botany 47, 1165–117. Pélissier HC, Peters WS, Collier R, van Bel AJE, Knoblauch M. 2008. GFP tagging of sieve element occlusion (SEO) proteins in green fluorescent forisomes. Plant and Cell Physiology 49, 1699–1710. Pena Cortes H, Fisahn J, Willmitzer L. 1995. Signals involved in wound-induced proteinase inhibitor II gene expression in tomato and potato plants. Proceedings of the National Academy of Sciences, USA 92, 4106–4113. Peters WS, Haffer D, Hanakam CB, van Bel AJE, Knoblauch M. 2010. Legume phylogeny and the evolution of a unique contractile apparatus that regulates phloem transport. American Journal of Botany 97, 797–808. Peters WS, van Bel AJE, Knoblauch M. 2006. The geometry of the forisome–sieve element–sieve plate complex in the phloem of Vicia faba L. leaflets. Journal of Experimental Botany 57, 3091–3098. Pickard B. 1973. Action potentials in plants. Botanical Reviews 39, 172–201. Pickard WF, Minchin PEH. 1990. The transient inhibition of phloem translocation in Phaseolus vulgaris by abrupt temperature drops, vibration, and electric shock. Journal of Experimental Botany 41, 1361–1369. Plieth C. 1999. Temperature sensing by plants: calcium-permeable channels as primary sensors—a model. Journal of Membrane Biology 172, 121–127. Pyatygin SS, Opritov VA, Vodeneev VA. 2008. Signaling role of action potential in higher plants. Russian Journal of Plant Physiology 55, 285–291. Radford JE, Vesk M, Overall RL. 1998. Callose deposition at plasmodesmata. Protoplasma 201, 30–37. Rao SAK, Carolan JC, Wilkinson TL. 2013. Proteomic profiling of cereal aphid saliva reveals both ubiquitous and adaptive secreted proteins. PLoS One 8, e57413. Retivin VG, Opritov VA, Fedulina SB. 1997. Generation of action potentials induces preadaptation of Cucurbita pepo L. stem tissues to freezing injury. Russian Journal of Plant Physiology 44, 432–442. Rhodes JD, Thain JF, Wildon DC. 1996. The pathway for systemic electrical signal conduction in the wounded tomato plant. Planta 200, 50–57. Ricca U. 1916. Soluzione d’un problema di fisiologia: la propagazione di stimulo nella Mimosa. Nuovo Giornale di Botanico Italiano 23, 51–170. Roblin G. 1985. Analysis of the variation potential induced by wounding in plants. Plant and Cell Physiology 26, 455–461. Roblin G, Bonnemain J-L. 1985. Propagation in Vicia faba stem of a potential variation by wounding. Plant and Cell Physiology 26, 1273–1283. Ruan Y-L, Llewellyn, Furbank RT. 2001. The control of singlecelled cotton fiber elongation by developmentally reversible gating of plasmodesmata and coordinate expression of sucrose and K+ transport genes and expansin. The Plant Cell 13, 47–60. Rüping B, Ernst AM, Jekat SB, Nordzieke S, Reineke AR, Müller B, Bornberg-Bauer E, Prüfer D, Noll GA. 2010. Molecular and phylogenetic characterization of the sieve element occlusion gene family in Fabaceae and non-Fabaceae plants. BMC Plant Biology 10, 219. Samejima M, Sibaoka T. 1983. Identification of the excitable cells in the petiole of Mimosa pudica by intracellular injection of procion yellow. Plant and Cell Physiology 24, 33–39. Sanders D, Pelloux J, Brownlee C, Harper JF. 2002. Calcium at the crossroads of signaling. The Plant Cell 14, 401–417. Schwan S, Menzel M, Fritzsche M, Heilmann A, Spohn U. 2009. Micromechanical measurements of P-protein aggregates (forisomes) from Vicia faba plants. Biophysics and Biochemistry 139, 99–105. Schulz A. 1999. Physiological control of plasmodesmatal gating. In: van Bel AJE, van Kesteren WJP, eds. Plasmodesmata. Structure, function, role in cell communication. Berlin: Springer, 173–204. Sibaoka T. 1966. Action potentials in plant organs. Symposia of the Society of Experimental Biology 20, 49–74. Sibaoka T. 1969. Physiology of rapid movements in plants. Annual Review of Plant Physiology 20, 165–184. Sibaoka T. 1991. Rapid plant movements triggered by action potentials. Journal of Plant Research 104, 73–95. Sinyukin AM, Britikov EA. 1967. Action potentials in the reproductive system of plants. Nature 215, 1278–1280. through plasmodesmata. A new mechanism of aluminium toxicity in plants. Plant Physiology 124, 991–1005. Sivaguru M, Yamamoto Y, Rengel Z, Ahn S-J, Matsumoto M. 2005. Early events responsible for aluminium toxicity symptoms in suspension-cultured tobacco cells. New Phytologist 165, 99–109. Sjolund RD, Shih CY. 1983. Freeze-fracture analysis of phloem structure in plant tissue cultures. I. The sieve element reticulum. Journal of Ultrastructure Research 82, 111–121. Stahlberg R, Cleland RE, Van Volkenburgh E. 2005. Decrement and amplification of slow wave potentials during their propagation in Helianthus annuus L. shoots. Planta 220, 550–558. Stahlberg R, Cleland RE, Van Volkenburgh E. 2006. Slow wave potentials—a propagating electrical signal unique to higher plants. In: Baluska F, Mancuso S, Volkmann D, eds. Communication in plants. Berlin: Springer, 291–308. Stahlberg R, Cosgrove DJ. 1992. Rapid alterations in growth rate and electrical potentials upon stem excision in pea seedlings. Planta 187, 523–531. Stahlberg R, Cosgrove DJ. 1996. Induction and ionic basis of slow wave potentials in seedlings of Pisum sativum L. Planta 200, 416–425. Stahlberg R, Cosgrove DJ. 1997. The propagation of slow wave potentials in pea epicotyls. Plant Physiology 113, 209–217. Stankovic B, Davies E. 1997. Intercellular communication in plants: electrical stimulation of proteinase inhibitor gene expression in tomato. Planta 202, 402–406. Stankovic B, Witters DL, Zawadzki T, Davies E. 1998. Action and variation potentials in sunflower: an analysis of their relationship and distinguishing characteristics. Physiologia Plantarum 103, 51–58. Stankovic B, Zawadzki T, Davies E. 1997. Characterization of the variation potential in sunflower. Plant Physiology 115, 1083–1088. Stewart LR, Ding B, Falk BW. 2013. Viroids and phloem-linited viruses: unique molecular probes of phloem biology. In: Thompson GA, van Bel AJE, eds. Phloem. Molecular cell biology, systemic communication, biotic interactions. Chichester: Wiley-Blackwell, 271–292. Thain JF, Gubb IR, Wildon DC. 1995 Depolarization of tomato leaf cells by oligogalacturonide elicitors. Plant, Cell and Environment 18, 211–214. Thiel G, Homann U, Plieth C. 1997. Ion channel activity during the action potential in Chara: a new insight with new techniques. Journal of Experimental Botany 48, 609–622. Thion L, Mazars C, Nacry P, Bouchez D, Moreau M, Ranjeva R, Thuleau P. 1998. Plasma membrane depolarization-activated calcium channels stimulated by microtubule-depolymerizing drugs in wild-type Arabidopsis thaliana protoplasts, display constitutively large activities and a longer half-life in ton 2 mutant cells affected in the organization of the cortical microtubules. The Plant Journal 13, 603–610. Thorpe MR, Furch ACU, Minchin PEH, Föller J, van Bel AJE, Hafke JB. 2010. Rapid cooling triggers forisome dispersion just before phloem transport stops. Plant, Cell and Environment 33, 259–271. Thorsch J, Esau K. 1981. Nuclear degeneration and the association of endoplasmic reticulum with the nuclear envelope and microtubules in maturing sieve elements of Gossypium hirsutum. Journal of Ultrastructure Research 74, 195–204. Tjallingii WF. 2006. Salivary secretions by aphids interacting with proteins of phloem wound responses. Journal of Experimental Botany 57, 739–745. Trebacz K, Dziubinska H, Krol E. 2006. Electrical signals in longdistance communication in plants. In: Baluska F, Mancuso S, Volkmann D, eds. Communication in plants. Berlin: Springer, 277–290. Trebacz K, Simonis W, Schönknecht G. 1994. Cytoplasmic Ca2+, K+, Cl– and NO3– activities in the liverwort Conocephalum conicum L. at rest and during action potentials. Plant Physiology 106, 1073–1084. Trewavas A. 1999. Le calcium, c’est la vie: calcium makes waves. Plant Physiology 120, 1–6. Trewavas AJ, Malho R. 1997. Signal perception and transduction: the origin of the phenotype. The Plant Cell 9, 1181–1195. Tsien RW, Hess P, Mc Cleskey EW, Rosenberg RL. 1987. Calcium channels: mechanisms of selectivity, permeation, and block. Annual Review of Biophysical Chemistry 16, 265–790. Tsien RW, Tsien RY. 1990. Calcium channels, stores and oscillations. Annual Review of Cell Biology 6, 715–760. Tsutsui I, Ohkawa T, Nagai R, Kishimoto U. 1986. Inhibition of Cl– channel activation in Chara corallina membrane by lanthanum ion. Plant and Cell Physiology 27, 1197–1200. Tucker EB. 1990. Calcium-loaded 1,2-bis(2-aminophenoxy) ethane-N,N,N′,N′-tetraacetic acid blocks cell-to-cell diffusion of carboxyfluorescein in staminal hairs of Setcreasea purpurea. Planta 182, 34–38. Tuteja N, Umate P, van Bel AJE. 2010. Forisomes: calciumpowered protein complexes with potential as ‘smart’ biomaterials. Trends in Biotechnology 28, 102–110. van Bel AJE. 2003. The phloem, a miracle of ingenuity. Plant, Cell and Environment 26, 125–149. van Bel AJE, Ehlers K. 2005. Electrical signalling via plasmodesmata. In: Oparka KJ, ed. Plasmodesmata. Oxford: Blackwell, 263–278. van Bel AJE, Kempers R. 1997. The pore/plasmodesm unit; key element in the interplay between sieve element and companion cell. Progress in Botany 58, 278–291. van Bel AJE, Knoblauch M, Furch ACU, Hafke JB. 2011a. (Questions)n on phloem biology. 1. Electropotential waves, Ca2+ fluxes and cellular cascades along the propagation pathway. Plant Science 181, 210–218. van Bel AJE, Furch ACU, Hafke JB, Knoblauch M, Patrick JW. 2011b. (Questions)n on phloem biology. 2. Mass flow, molecular hopping, distribution patterns and macromolecular signalling. Plant Science 181, 325–330. van Bel AJE, van Rijen HVM. 1994. Microelectrode-recorded development of the symplasmic autonomy of the sieve element/ companion cell complex in the stem phloem of Lupinus luteus L. Planta 192, 165–175. Vincill ED, Clarin AE, Molenda JN, Spalding EP. 2013. Interacting receptor-like proteins in phloem regulate lateral root initiation in Arabidopsis. The Plant Cell 25, 1304–1313. Fromm J , Lautner S. 2007 . Electrical signals and their physiological significance in plants . Plant, Cell and Environment 30 , 249 - 257 . 2013. Involvement of the sieve-element cytoskeleton in electrical responses to cold shocks . Plant Physiology 162 , 707 - 714 . Hafke JB , Furch ACU , Reitz MU , van Bel AJE . 2007 . Functional sieve element protoplasts . Plant Physiology 145 , 703 - 711 . Hafke JB , van Bel AJE . 2013 . Cellular basis of electrical potential waves along the phloem and impact of coincident Ca2+ fluxes . In: Thompson GA, van Bel AJE, eds. Phloem. Molecular cell biology, systemic communication, biotic interactions . Chichester: WileyBlackwell , 122 - 140 . Hattori M , Nakamura M , Komatsu S , Tsuchihara K , Tamura Y , Hasegawa T. 2012 . Molecular cloning of a novel calcium-binding protein in the secreted saliva of the green leafhopper Nephotettix cincticeps . Insect Biochemistry and Molecular Biology 42 , 1 - 9 . Hepler PK , Palevitz BA , Lancelle SA , McCauley MM , Lichtscheidl L. 1990 . Cortical endoplasmic reticulum in plants. Journal of Cell Science 96 , 355 - 373 . Hlavackova V , Krchnak P , Naus J , Novak O , Spundova M , Strnad M. 2006 . Electrical and chemical signals involved in shortterm systemic photosynthetic responses of tobacco plants to local burning . Planta 225 , 235 - 244 . Hodick D , Sievers A. 1988 . The action potential of Dionaea muscipula Ellis . Planta 174 , 8 - 18 . Hodick D , Sievers A. 1989 . On the mechanism of trap closure of Venus flytrap (Dionaea muscipula Ellis) . Planta 179 , 32 - 42 . 2000. Physiological elevations in cytoplasmic free calcium by cold or microinjection result in transient closure of higher plant plasmodesmata . Planta 210 , 329 - 335 . Holdaway-Clarke TL , Walker NA , Overall R . 1996 . Measurement of the electrical resistance of plasmodesmata and membranes of corn suspension cells . Planta 199 , 537 - 544 . Homann U , Thiel G. 1994 . Cl- and K+ channel currents during the action potential in Chara: simultaneous recording of membrane voltage and patch currents . Journal of Membrane Biology 141 , 297 - 309 . Houwink AL . 1935 . The conduction of excitation in Mimosa pudica . Recueil des Travaux Botaniques Neerlandais 32 , 51 - 91 . Huda KMK , Banu MSA , Tuteja R , Tuteja N. 2013 . Global calcium transducer P-type Ca2+-ATPases open new avenues for agriculture by regulating stress signalling . Journal of Experimental Botany 64 , 3099 - 3109 . Iijima T , Sibaoka T. 1981 . Action potentials in the trap-lobes of Aldrovanda vesiculosa . Plant and Cell Physiology 22 , 1595 - 1601 . Iijima T , Sibaoka T. 1982 . Propagation of action potentials over the trap-lobes of Aldrovanda vesiculosa . Plant and Cell Physiology 23 , 679 - 688 . Iijima T , Sibaoka T. 1985 . Membrane potentials in excitable cells of Aldrovanda vesiculosa trap-lobes . Plant and Cell Physiology 26 , 1 - 13 . Jekat SB , Ernst A , Zielonka S , Noll G , Prüfer D. 2012 . Interactions among tobacco sieve element occlusion (SEO) proteins . Plant Signaling and Behavior 7 , 1918 - 1920 . Kaplan B , Davydov O , Knight H , Galon Y , Knight MR , Fluhr R , Fromm H. 2006 . Rapid transcriptome changes induced by cytosolic Ca2+ transients reveal ABRE-related sequences as Ca2+ responsive cis elements in Arabidopsis . The Plant Cell 18 , 2733 - 2748 . Kartte S , Seemüller E. 1991 . Histopathology of apple proliferation in Malus taxa and hybrids of different susceptibility . Journal of Phytopathology 131 , 149 - 160 . Kauss H , Jeblick W. 1991 . Induced Ca2+ uptake and callose synthesis in suspension-cultured cells of Cataranthus roseus are decreased by the protein phosphatase inhibitor okadaic acid . Physiologia Plantarum 81 , 309 - 312 . Kehr J , Buhtz A. 2013 . Endogenous RNA constituents of the phloem and their possible roles in long-distance signalling . In: Thompson GA , van Bel AJE, eds. Phloem. Molecular cell biology, systemic communication, biotic interactions . Chichester: Wiley-Blackwell, 186 - 208 . Kempers R , Ammerlaan A , van Bel AJE . 1998 . Symplasmic constriction and ultrastructural features of the sieve element/ companion cell complex in the transport phloem of apoplasmically and symplasmically loading species . Plant Physiology 116 , 271 - 278 . Kinoshita T , Nishimura M , Shimazaki K. 1995 . Cytosolic concentration of Ca2+ regulates the plasma membrane H-ATPase in guard cells of fava bean . The Plant Cell 7 , 1333 - 1342 . Kishimoto U , Takeuchi Y , Ohkawa T , Kami-ike N. 1985 . A kinetic analysis of the electrogenic pump of Chara corallina . III. Pump acivity during action potential . Journal of Membrane Biology 86 , 27 - 36 . Klüsener B , Boheim G , Liß H , Engelberth J , Weiler EW . 1995 . EMBO Journal 14 , 2708 - 2714 . Klüsener B , Weiler EW . 1999 . A calcium-selective channel from root-tip endomembranes of garden cress . Plant Physiology 119 , 1399 - 1405 . Knepper C , Savory EA , Day B. 2011 . Arabidopsis NDR1 is an integrin-like protein with a role in fluid loss and plasma membrane-cell wall adhesion . Plant Physiology 156 , 286 - 300 . Knight H , Trewavas AJ , Knight MR . 1996 . Cold calcium signaling in Arabidopsis involves two cellular pools and a change in calcium signature after acclimation . The Plant Cell 8 , 489 - 503 . Knoblauch M , Noll G , Müller T , Prüfer D , Schneider-Hüther I , Scharner D , van Bel AJE , Peters WS . 2003 . ATP-independent contractile proteins from plants . Nature Materials 2 , 600 - 603 . Knoblauch M , Noll GA , Müller T , Prüfer D , Schneider-Hüther I , Scharner D , van Bel AJE , Peters WS . 2005 . Corrigendum. Nature Materials 4 , 353 . Knoblauch M , Oparka KJ . 2012 . The structure of the phloem-still more questions than answers . The Plant Journal 70 , 147 - 156 . Knoblauch M , Peters WS , Ehlers K , van Bel AJE . 2001 . Reversible calcium-regulated stopcocks in legume sieve tubes . The Plant Cell 13 , 1221 - 1230 . 2012. Forisome performance in artificial sieve tubes . Plant, Cell and Environment 15 , 1419 -1427 Knoblauch M , van Bel AJE . 1998 . Sieve tubes in action . The Plant Cell 10 , 35 - 50 . 2004. Transient knockout of photosynthesis mediated by electrical signals . New Phytologist 161 , 715 - 722 . Krol E , Dziubinska H , Trebacz K. 2003 . Low-temperature-induced transmembrane potential changes in the liverwort Conocephalum conicum . Plant and Cell Physiology 44 , 527 - 533 . Krol E , Dziubinska H , Trebacz K. 2004 . Low-temperature-induced transmembrane potential changes in mesophyll cells of Arabidopsis thaliana, Helianthus annuus and Vicia faba . Physiologia Plantarum 120 , 265 - 270 . Kudla J , Batistic O , Hashimoto K. 2010 . Calcium signals: the lead currency of plant information processing . The Plant Cell 22 , 541 - 563 . Kulikova AL , Puryaseva AP . 2002 . Actin in pumpkin phloem exudate . Russian Journal of Plant Physiology 49 , 54 - 60 . Lacombe B , Pilot G , Michard E , Gaymard F , Sentenac H , Thibaud J-B. 2000 . A shaker-like K+ channel with weak rectification is expressed in both source and sink phloem tissues of Arabidopsis . The Plant Cell 12 , 837 - 851 . Langhans M , Ratajczak R , Lützelschwab M , Michalke W , Wächter R , Fischer-Schliebs E , Ullrich CE. 2001 . Immunolocalization of plasma membrane H+-ATPase and tonoplasttype pyrophosphatase of the sieve element-companion cell complex in the stem of Ricinus communis L. Planta 213 , 11 - 19 . Lautner S , Grams TEE , Matyssek R , Fromm J. 2005 . Characteristics of electrical signals in poplar and responses in photosynthesis . Plant Physiology 138 , 2200 - 2209 . Leckie CP , McAinsh MR , Allen GJ , Sanders D , Hetherington AM . 1998 . Abscisic acid-induced stomatal closure mediated by cyclic ADP-ribose . Proceedings of the National Academy of Sciences , USA 95 , 15837 - 15842 . Lecourieux D , Ranjeva R , Pugin A. 2006 . Calcium in plant defencesignalling pathways . New Phytologist 171 , 249 - 269 . Lee D , Polisensky DH , Braam J. 2005 Genome-wide identification of touch- and darkness regulated Arabidopsis genes: a focus on calmodulin-like and XTH genes . New Phytologist 165 , 429 - 444 . 2003. Inositol hexakiphosphate mobilizes an endomembrane store of calcium in guard cells . Proceedings of the National Academy of Sciences USA 100 , 10091 - 10095 . Mazars C , Thion L , Thuleau P , Graziana A , Knight MR , Moreau M , Ranjeva R . 1997 . Organization of cytoskeleton McAinsh MR , Pittman JK . 2009 . Shaping the calcium signature . New Phytologist 181 , 275 - 294 . McCormack E , Braam J. 2003 . Calmodulins and related potential Ca2+ sensors of Arabidopsis . New Phytologist 159 , 585 - 598 . McCormack E , Tsai YC , Braam J. 2005 Handling calcium signaling: Arabidopsis CaMs and CMLs . Trends in Plant Science 10, 383 - 389 . McLean AM , Hogenhout SA . 2013 . Phytoplasmas and spiroplasmas: the phytopathogenic mollicutes of the phloem . In: Thompson GA , van Bel AJE, eds. Phloem. Molecular cell biology, systemic communication, biotic interactions . Chichester: WileyBlackwell , 293 - 309 . Medina-Ortega KJ , Walker GP . 2013 . Does aphid salivation affect phloem sieve element occlusion in vivo ? Journal of Experimental Botany 64 , 5525 - 5535 . Michard E , Lima PT , Borges F , Silva A , Portes MT , Carvalho JE , Gilliham M , Liu L-H , Obermeyer G , Feijo JA . 2011 . Glutamate receptor-like genes form Ca2+ channels in pollen tubes and are regulated by pistil d -serine . Science 332 , 434 - 437 . Miedema H , Demidchik V , Véry AA , Bothwell JHF , Brownlee C , Davies JM . 2008 . Two voltage-dependent calcium channels co-exist in the apical plasma membrane of Arabidopsis thaliana root hairs . New Phytologist 179 , 378 - 385 . Miles PW . 1999 . Aphid saliva . Biological Reviews of the Cambridge Philosophical Society 74 , 41 - 85 . Minorsky PV , Spanswick RM . 1989 . Electrophysiological evidence for a role for calcium in temperature sensing by roots of cucumber seedlings . Plant, Cell and Environment 12 , 137 - 143 . Monroy AF , Dhindsa RS . 1995 . Low temperature signal transduction: induction of cold acclimation-specific genes of alfalfa by calcium at 25°C . The Plant Cell 7, 321 - 331 . Moyen C , Johannes E. 1996 . Systemin transiently depolarizes the tomato mesophyll cell membrane and antagonizes fusicoccininduced extracellular acidification of mesophyll tissue . Plant, Cell and Environment 19 , 464 - 470 . Mummert H , Gradmann D. 1991 . Action potentials in Acetabularia: measurement and simulation of voltage-gated fluxes . Journal of Membrane Biology 124 , 265 - 273 . 2011 . The Arabidopsis calcium-dependent protein kinase, CPK6, functions as a positive regulator of methyl jasmonate signalling in guard cells . Plant Physiology 155 , 553 - 561 . Munnik T , Vermeer JE . 2010 . Osmotic stress-induced phosphoinositide and inositol phosphate signalling in plants . Plant, Cell and Environment 33 , 665 - 669 . Musetti R , Buxa SV , De Marco F , Loschi A , Polizzotto R , Kogel KH , van Bel AJE . 2013 . Phytoplasma-triggered Ca2+ influx is involved in sieve-tube blockage . Molecular Plant-Microbe Interactions 26 , 379 - 386 . Musetti R , Favali MA . 1999 . Histological and ultrastructural comparative study between Prunus varieties of different susceptibility to plum leptonecrosis . Cytobios 99 , 73 - 82 . Plieth C , Hansen U-P , Knight H , Knight MR . 1999 . Temperature sensing by plants: the primary characteristics of signal perception and calcium response . The Plant Journal 18 , 491 - 497 . Sivaguru M , Fujiwara T , Samaj J , Baluska F , Yang Z , Osawa H , Maeda T , Mori T , Volkmann D , Matsumoto M. 2000 . Aluminiuminduced 1→3-β-d -glucan inhibits cell-to-cell trafficking of molecules van Sambeek JW , Pickard BG , Ulbright CE . 1976 . Mediation of rapid electrical, metabolic, transpirational, and photosynthetic changes by factors released from wounds . II. Mediation of the variation potential by Ricca's factor . Canadian Journal of Botany 54 , 2651 - 2661 . Volk G , Franceschi VR . 2000 . Localization of a calcium-channel-like protein in the sieve element plasma membrane . Australian Journal of Plant Physiology 27 , 779 - 786 . 2004. Proteomics of cucurbit phloem exudates reveals a network of defence proteins . Phytochemistry 65 , 1795 - 1804 . Walker FP , Medina-Ortega KJ . 2012 . Penetration of fava bean sieve element by pea aphid does not trigger forisome dispersal . Entomologia Experimentalis et Applicata 144 , 326 - 335 . Walling LL . 2000 . The myriad plant responses to herbivores . Journal of Plant Growth Regulation 19 , 195 - 216 . Wang YF , Fan L-M , Zhang W-Z , Zhang W , Wu W-H. 2004 . Ca2+- permeable channels in the plasma membrane of Arabidopsis pollen are regulated by actin microfilaments . Plant Physiology 136 , 3892 - 3904 . Wayne R . 1994 . The excitability of plant cells: with special emphasis on characean internodal cells . Botanical Reviews 60 , 265 - 367 . Webb AAR , McAinsh MR , Taylor JE , Hetherington AM . 1996 . Advances in Botanical Research 22 , 45 - 96 . White PJ . 2004 . Calcium signals in root cells: the roles of plasma membrane calcium channels . Biologia 59 , 77 - 83 . White PJ . 2009 . Depolarization-activated calcium channels shape the calcium signatures induced by low-temperature stress . New Phytologist 183 , 6 - 8 . White PJ , Broadley MR . 2003 . Calcium in plants . Annals of Botany 92 , 487 - 511 . White PJ , Ridout MS . 1999 . An energy-barrier model for the permeation of monovalent and divalent cations through the maxi cation channel in the plasma membrane of rye roots . Journal of Membrane Biology 168 , 63 - 75 . Wildon DC , Thain JF , Minchin PEH , Gubb IR , Reilly A , Skipper YD , Doherty HM , O'Donnell PJ , Bowles DJ . 1992 . Electrical signalling and systemic proteinase inhibitor induction in the wounded plant . Nature 360 , 62 - 65 . Will T , Carolan JC , Wilkinson TL . 2013 . Breaching the sieve element-the role of aphid saliva as the molecular interface between aphids and the phloem . In: Thompson GA , van Bel AJE, eds. Phloem. Chichester: Wiley-Blackwell, 310 - 327 . 2009. Aphid saliva counteracts sieve-tube occlusion: a universal phenomenon ? Journal of Experimental Biology 22 , 3305 - 3312 . Will T , Tjallingii WF , Thönnessen A , van Bel AJE . 2007 . Molecular sabotage of plant defense by aphids . Proceedings of the National Academy of Sciences , USA 104 , 10536 - 10541 . Will T , van Bel AJE . 2006 . Physical and chemical interactions between aphids and plants . Journal of Experimental Botany 57 , 729 - 737 . Will T , van Bel AJE . 2008 . Induction as well as suppression; how aphids may exert opposite effects on plant defense . Plant Signaling and Behavior 3 , 427 - 430 . Williams SE , Pickard BG . 1972a. Properties of action potentials in Drosera tentacles . Planta 103 , 222 - 240 . Williams SE , Pickard BG . 1972b. Receptor potentials and action potentials in Drosera tentacles . Planta 103 , 193 - 221 . Williams SE , Pickard BG . 1974 . Connections and barriers between cells of Drosera tentacles in relation to their electrophysiology . Planta 116 , 1 - 6 . Williams SE , Spanswick RM . 1976 . Propagation of the neuroid action potential of the carnivorous plant Drosera . Journal of Comparative Physiology 108 , 211 - 223 . Xie B , Wang X , Zhu M , Zhang Z , Hong Z. 2011 . CalS7 encodes a callose synthase responsible for callose deposition in the phloem . The Plant Journal 65 , 1 - 14 . Yoo BC , Lee JY , Lucas WJ . 2002 . Analysis of the complexity of protein kinases within the phloem sieve tube system . Journal of Biological Chemistry 277 , 15325 - 15332 . Zawadzki T , Davies E , Dziubinska H , Trebacz K. 1991 . Characteristics of action potentials in Helianthus annuus . Physiologia Plantarum 83 , 601 - 604 . Zhang W , Fan L , Wu W. 2007 . Osmo-sensitive and stretch-activated calcium-permeable channels in Vicia faba guard cells are regulated by actin dynamics . Plant Physiology 143 , 1140 - 1151 . Zimmermann MR , Felle HH . 2009 . Dissection of heat-induced systemic signals: superiority of ion fluxes to voltage changes in substomatal cavities . Planta 229 , 539 - 547 . Zimmermann MR , Hafke JB , van Bel AJE , Furch ACU . 2013 . The interaction of xylem and phloem during exudation and wound occlusion in Cucurbita maxima . Plant, Cell and Environment 36 , 337 - 347 . 2009. System potentials, a novel electrical long-distance apoplastic signal in plants, induced by wounding . Plant Physiology 149 , 593 - 600 . Zimmermann MR , Mithöfer A. 2013 . Electrical long-distance signalling in plants . In: Baluska F, ed. Long- distance systemic signalling and communication in plants . Berlin: Springer, 291 - 308 .

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Aart J. E. van Bel, Alexandra C. U. Furch, Torsten Will, Stefanie V. Buxa, Rita Musetti, Jens B. Hafke. Spread the news: systemic dissemination and local impact of Ca2+ signals along the phloem pathway, Journal of Experimental Botany, 2014, 1761-1787, DOI: 10.1093/jxb/ert425