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
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
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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).
channels and other channels involved in
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
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
Location of the Ca2+-permeable channels in sieve
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
Deployment of other channels and pumps involved in
Ultrastructure and plasmodesmal connectivity of sieve
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
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.
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
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
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
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).
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
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?
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