Download 1 Calcium at the Cell Wall

Calcium at the Cell Wall – Cytoplast Interface
Running Title: Calcium at the cell wall cytoplast interface
Peter K. Hepler1 and Lawrence J. Winship2
1. Department of Biology, University of Massachusetts, Amherst, MA 01003
2. School of Natural Science, Hampshire College, Amherst MA 01002
Contact:
Prof. Peter K. Hepler
Department of Biology, University of Massachusetts
611 North Pleasant St.
Amherst, MA 01003
Tel:
413-545-2083
Fax:
413-545-3243
Email: [email protected]
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Abstract:
Attention is given to the role of Ca2+ at the interface between the cell wall and the cytoplast, especially as
seen in pollen tubes. While the cytoplasm directs the synthesis and deposition of the wall, it is less well
appreciated that the wall exerts considerable self control and influences activities of the cytoplasm. Ca2+
participates as a crucial factor in this two way communication. In the cytoplasm, a [Ca2+] above 0.1 μM,
regulates myriad processes, including secretion of cell wall components. In the cell wall Ca2+, at 10 μM10 mM, binds negative charges on pectins and imparts structural rigidity to the wall. The plasma
membrane occupies a pivotal position between these two compartments, where selective channels
regulate influx of Ca2+, and specific carriers pump the ion back into the wall. In addition we draw
attention to different factors, which either respond to the wall or are present in the wall, and usually
generate elevated [Ca2+] in the cytoplasm. These factors include 1) stretch activated channels; 2)
calmodulin; 3) annexins; 4) wall associated kinases; 5) oligogalacturonides; and 6) extracellular ATP.
Together they provide evidence for a rich and multifaceted system of communication between the
cytoplast and cell wall, with Ca2+ as a carrier of information.
Key Word:
Calcium (Ca2+), Cell Wall, Pectin, Pectin Methyl Esterase (PME), Plasma membrane (PM), Pollen Tube
Growth
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A. Introduction:
Historically, the plasma membrane (PM) has been considered to be the interface between the living
protoplasm and the non-living extracellular matrix. While this membrane marks the boundary between
the cell wall to the outside and the cytoplasm to the inside, implying that the cell wall is “non-living” is
misleading. Even though it is deposited outside the PM, the cell wall is formed by the activity of the
cytoplasm, and maintains a close relationship with it as revealed by the presence of structural attachments
to the PM. In addition, the cell wall contains enzymes, and other factors that greatly influence and
modify its structure and composition (Carpita and Gibeaut, 1993). The cell wall thus possesses active
mechanisms for self regulation. Increasingly we are confronted with evidence that the cell wall, due to its
structural properties and composition, influences the behavior of the “living” protoplast (Brownlee, 2002;
Monshausen and Gilroy, 2009). For example, during the process of cell growth itself, the yielding of the
wall dictates the extension of the cell, and for polarized cells, local yielding determines plant cell shape
(Schopfer, 2006). It seems clear therefore that much communication takes place between the cell wall
and the cytoplasm, with the PM occupying a pivotal position in responding to and transmitting
information.
Calcium ions (Ca2+) are an important component of the information system that operates between the
cytoplasm and the cell wall, participating in crucial ways in both compartments. In the pollen tube cell
wall for example, the activity of free Ca2+ is in dynamic steady state, influenced by inward diffusion from
the external medium, where [Ca2+] typically ranges from 10 μM to 10 mM (Picton and Steer, 1983), and
by ion exchange and coordination with wall polysaccharides. Ca2+ participates in cross-linking negative
charges, especially on the carboxylic residues of pectins, imparting significant structural rigidity to the
wall. In the cytoplasm the [Ca2+] is tightly regulated in the vicinity of 0.1 μM, with some spatially and
temporally restricted elevated domains at 1 μM - 10 μM (Rathore et al., 1991; Miller et al., 1992; Pierson
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et al., 1996). As is increasingly well appreciated, Ca2+ in the cytoplasm participates in myriad events
where it acts as a second messenger in a host of signaling pathways.
Given the substantial differences in [Ca2+] between the cell wall and cytoplasm, together with the
realization that the ions are serving rather different functions in these two compartments, it becomes an
interesting question how Ca2+ operates at the interface. Of course it is well appreciated that channels and
pumps on the PM act as gates to control Ca2+ influx and efflux. However, it is often not acknowledged
that the Ca2+ that crosses the PM and enters the cytosol, must first pass through the cell wall. The Ca2+
binding reactions, rather than the concentration of the ion in the medium, will determine the activity of
Ca2+ at the mouth of a selective channel and thus the magnitude of its influx into the cell. Similarly, the
Ca2+ that is pumped out of the cell, and extruded into the cell wall, will immediately influence wall
structure in those local domains near the pumps, being governed by the local [Ca2+] and not that of the
medium. As a consequence there will be local domains with associated fluxes that become crucial to the
growth and development of the plant as a whole.
It will be the purpose of this essay to consider the activity of Ca2+ at the cell wall/cytoplast interface.
While we draw broadly on work that has been gained from studies of different plant cells, we give
emphasis to results from research on the growth of pollen tubes. Although the pollen tube is a specialized
cell, it possesses a cell wall composed of polymers well recognized for their presence in other plant cells,
and obeys the biophysical laws governing cell wall expansion that apply to plant cells in general (Hepler
et al., 2006; Chebli and Geitmann, 2007). The pollen tube PM also contains different pumps and channels
that participate in regulation of Ca2+ import and export (Sze et al., 2006). Although some of the specific
proteins are uniquely expressed in pollen tubes, they are members of larger families with conserved
functions fundamental to all plant cells. But beyond this, the growing pollen tube demonstrates clear and
specific requirements for Ca2+ and in addition demonstrates intracellular profiles, such as the tip-focused
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Ca2+ gradient, which are recognized as essential features (Holdaway-Clarke and Hepler, 2003). The
pollen tube is thus a favorable system in which to explore the role of Ca2+ at the cell wall/cytoplast
interface.
B. Ca2+ and the Cell Wall:
It has long been appreciated that Ca2+ plays a key role in determining the structure and function of the cell
wall (Hepler, 2005). Through cross-linking negatively charged regions in pectins, found primarily in
short stretches of de-methoxylated homogalacturonans, Ca2+ imparts rigidity to pollen tube cell walls.
When the [Ca2+] is too low, the wall is weakened and may even break, whereas at high [Ca2+] the pectin
chains will be cross-linked and aggregated, and the wall maximally rigidified. The experimental control
of [Ca2+] was pivotal in early studies aimed at separating single living cells. Over 50 years ago, the
chelator, EDTA, was used to remove Ca2+ from pectins in the middle lamella, and thus to breakdown
tissues into single cells (Letham, 1958).
In pollen tubes the growing apical wall consists almost entirely of pectin, with cellulose and callose being
located several microns from the apex (Ferguson et al., 1998). As a consequence, when the [Ca2+] is
lowered sufficiently the pollen tube wall completely loses its structural integrity and bursts. The
permissive [Ca2+] for pollen tubes extends between 10 μM to 10 mM (Picton and Steer, 1983). Below 10
μM, the pollen tube bursts because the wall is too weak, and above 10 mM, pollen tube growth frequently
stops likely due to excessive cross-linking. Because a [Ca2+] of 10 µM or less should be sufficient to
satisfy the number of binding sites in the wall it may be more pertinent to speak about the availability of
Ca2+ at the cell wall/cytoplast interface as new wall material is being deposited. Because 20 to 30% of the
newly deposited pectin will be de-esterified (Bosch and Hepler, 2005), there will be always an immediate
need for Ca2+ by the growing pollen tube. Therefore a [Ca2+] of 10 µM is the amount needed in the
medium to ensure that sufficient ions are available at the cell wall/cytoplast interface in order to prevent
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the tube from bursting. But even in other cell types, the Ca2+/pectate bonds play a major role in regulating
the ability of the cell wall to extend. For example, mature cucumber hypocotyls cells, which become
insensitive to the growth promotive effects of expansin, can regain their sensitivity simply through
chelation of the wall Ca2+ with EGTA (Zhao et al., 2008).
Pectins are secreted largely as methoxy-esters (70-80%), and only later de-esterified in the apoplast by the
enzyme pectin methyl esterase (PME), which has also been secreted (Willats et al., 2001; Bosch and
Hepler, 2005). The methoxylation pattern of newly secreted pectins is not known, but it is known that
plant PMEs carry out block-wise de-methoxylation, creating contiguous stretches of galacturonic acid
residues ( Bosch, et al., 2005). The extent and strength of Ca2+ cross-linking will depend on the pattern of
de-methoxylation as well as on the number and availability of the acidic residues. Recent evidence from
in vitro systems under conditions of a high degree of methoxylation, and a low degree of block-wise demethoxylation, or simply degree of blockiness, indicate that pectins associate ionically, with carboxyl
moieties participating in labile binding with free Ca2+ forming plastic gels with low shear strength (Fang,
et al., 2008). As pectins are de-methoxylated block-wise by PME, dimers begin to form in a cooperative
fashion, so that binding strength increases rapidly as the ratio of Ca2+ to available binding sites increases.
The number of consecutive de-methoxylated galacturonic acid residues required to form stable chains in a
modified, or shifted “egg-box” configuration (Braccini, et al, 2001; 2005) has been estimated in various
systems to range from 6 to 20 (Fraeye, et al., 2009).
Ultimately, at high [Ca2+], with a low degree of methoxylation and high degree of blockiness, pectins
reach maximum strength (Fraeye, et al., 2009). Thus at a given, but permissive [Ca2+], pollen tube wall
expansion and growth can be stopped simply by adding exogenous PME (Bosch et al., 2005; Parre and
Geitmann, 2005). Enzymatic de-methoxylation creates an abnormal number of blocky, de-methoxylated
regions over the entire pollen tube tip, concomitantly increasing wall stiffness due to the extent of Ca2+
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cross-linking. From these brief comments it can be appreciated that pollen tube growth is dependent on a
balance between the number of available acidic residues and the [Ca2+]. If there are too few acidic groups
or too little Ca2+ the pollen tube bursts, but if there are too many acidic groups and/or too much Ca2+, the
wall is overly cross-linked and unable to extend.
Another factor that influences Ca2+ binding is the stress that turgor pressure imposes on the wall binding
sites (Boyer, 2009). When Ca2+ cross-linked pectates are placed under physical tension, as imposed by
turgor pressure, e.g. 0.2 MPa, the bonds may lengthen and thus weaken and decrease their affinity for
Ca2+ (Proseus and Boyer, 2007; Boyer, 2009). Dissociation may occur, allowing turgor dependent
expansion. This scenario has been developed from studies on single internode cells of the green alga,
Chara/Nitella, but could also apply to cells of higher plants including pollen tubes. In this scheme it is
also important to recognize the ability of turgor pressure to force polymeric building blocks, notably
pectates, into isolated Characean cell walls, and to achieve normal rates of growth (Proseus and Boyer,
2006). Here the effects are two-fold: firstly, the insertion of new material within the matrix weakens the
existing bonds and facilitates growth by intussusception, and secondly, the new pectates, which would
exhibit a low [Ca2+], locally act as a chelator and remove the ion from load bearing bonds, thus
weakening ionic linkages, and permitting turgor-dependent elongation (Boyer, 2009).
It is now well established that the rate of pollen tube growth oscillates, with changes of up to 6 fold (100
to 600 nm/s) occurring during periods of 20-50 s (Cárdenas et al., 2008). Simply from a consideration of
the above factors, including notably the availability and activity of PME, there will be ongoing changes in
the extensibility of the cell wall. Particularly because the secretion of PME itself oscillates at the apical
cell surface (McKenna et al., 2009), it follows that there will be an associated oscillation in the
appearance of acidic groups, which will lead to changes in the cell wall yielding properties and ultimately
to changes in growth rate. A direct confirmation comes from studies using the extracellular Ca2+ selective
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electrode to monitor fluxes associated with growth. The results reveal an influx of Ca2+ focused at the tip
of the tube (Kühtreiber and Jaffe, 1990), which oscillates (Holdaway-Clarke et al., 1997; Messerli et al.,
1999). Cross-correlation analysis further shows that the increase in influx follows the increase in growth
rate by over a 1/3rd of a growth rate cycle (Holdaway-Clarke et al., 1997; Messerli et al., 1999).
There has been disagreement about what this influx represents, with some investigators thinking that it is
due to movement of Ca2+ directly into the cytoplasm, while other supporting wall binding. Kwack
(1967), applying autoradiography to pollen tubes that had been administered with 45Ca2+, showed
extensive labeling of the tube apex, which he interpreted as Ca2+ binding by the acidic pectic residues. A
similar study by Jaffe et al. (1975) on lily pollen tubes yielded similar results, but the authors argued in
favor of uptake into the cytoplasm. Their reason against wall binding was mainly that it would be
insufficient to account for the magnitude of signal observed. Also the ions were found to be tightly bound
and resistant to washings. However these conclusions do not fit with data on the wall binding of Ca2+. In
studies of mung been hypocotyls the [Ca2+] has been shown to increase during maturation from 80 µmol
per gram dry weight of wall to 122.5 µmol per gram dry weight (Goldberg et al., 1986), which when
extrapolated to the pollen tube accounts for the bulk of influx observed. Thus Holdaway-Clarke et al.
(1997) calculated that one could predict an influx of 35 pmol cm-2 sec-1, which is reasonably close to the
15-20 pmol cm-2 sec-1 that has been measured, especially when you take into account the fact the electrode
is only 50% efficient. By contrast the calculation of the influx needed to sustain the gradient yields
values of 0.5-2 pmol cm-2 sec-1, which is a small fraction of the amount measured. Indeed the small
amount of influx needed to support the intracellular gradient is within the noise of the extracellular flux
measurements, explaining why an influx directly corresponding to the change in the intracellular gradient
has not been observed (Holdaway-Clarke et al., 1997). While there is no question that some Ca2+ does
enter the cytoplasm, it must be recognized that the cell wall, including that of the pollen tube, has
enormous binding capacity and in addition can bind Ca2+ tightly. Recent work on the dimensions and
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dynamics of pollen tube walls (McKenna, et al., 2009) suggest that the Ca2+ flux required may be even
greater than that estimated in Holdaway-Clarke, et al. (1997) since the maximum wall thickness is 0.5
µm, compared to the 0.2 µm used earlier. In addition, extensive work on in vitro pectin systems has
demonstrated three main regions in pectin/Ca2+ state diagrams (Vincent and Williams, 2009). If the most
energetically stable configurations found in vitro also occur in pollen tube walls, it would require 320 to
640 µmol Ca2+ per gram of pectin compared to the 80 to 120 µmol per gram used in earlier calculations.
With improved methods for measuring Ca2+ fluxes we anticipate values on the order of 250 to 500 pmol
cm-2 sec-1.
A second argument against the idea that the bulk of the Ca2+ detected from the influx data directly crosses
the PM and enters the cytoplast stems from the lack of agreement in the phase relationships. Thus the
increase in the intracellular tip-focused gradient increases +10-400 behind the increase in growth rate
(Messerli et al., 2000; Cardenas et al., 2008), whereas the extracellular influx increases +1300 after the
increase in growth (Holdaway-Clarke et al., 1997). If Ca2+ were directly entering the cytoplasm there
should be a close agreement between the intracellular and extracellular signals. To explain the difference
it has been argued that Ca2+ enters the cytoplasm but is immediately sequestered within the ER or other
compartments, and thus not detected by cytosolic Ca2+ indicators (Trewavas and Malhó, 1997; Malhó et
al., 2006). This process, which has been described in animal cells, is referred to as “capacitatively
coupled uptake” or “store operated calcium entry” (Putney et al., 2001; Parekh and Putney, 2005).
However, studies with manganese, an ion that enters cells through Ca2+ channels and quenches indicator
dyes such as indo-1, indicate that the apical signal in pollen tubes is rapidly reduced (Malhó et al., 1995),
providing support for influx from the extracellular medium directly into the cytosol, and not via
capacitative processes.
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We can explain the lag of Ca2+ influx relative to growth if we consider both the spatial and temporal
binding patterns. Ca2+ demand in the wall is a function of the amount of wall material, the degree of
methoxylation and the degree of blockiness. The peak in the amount wall material at the pollen tube tip
(Region I, Figure 1) occurs at -1000 (McKenna, et al., 2009) relative to the peak in growth rate. This
material has a degree of methoxylation of 70 to 80%, with random dispersed carboxyl groups, and a low
degree of blockiness. Ca2+ demand in this region will be relatively low, and binding will be primarily
ionic and labile.
Wall expansion begins to accelerate as the highly methoxylated pectin intercalates and the wall softens.
Continued wall expansion displaces the methoxylated pectin away from the primary site of deposition. At
the same time, PME becomes more active, either as the pro region is cleaved or as inhibitor diffuses away
and begins block-wise de-methoxylation, dramatically increasing both the Ca2+ demand for dimerization
of pectin chains (Region II, Fig 1) and wall stiffness. Shear forces may orient pectin dimers, possibly
leading to anisotropy in wall stiffness. Those Ca2+ that are bound between pairs of pectin chains by
multiple coordination bonds are likely to be less labile than those that are ionically bound to pairs of
carboxyl residues on the exterior of the pectin dimers.
As a result of the elapsed time, the maximum in de-methoxylated pectin occurs well after the peak in
growth, which is thought to be about 1/3rd of a growth cycle based on extracellular Ca2+ influx data
(Holdaway-Clarke et al., 1997). Spatially this would take place at the “shoulder” of the pollen tube tip,
where expansion must eventually slow as the wall becomes less extensible. For this reason, the flux of
Ca2+ into the tube tip is maximal well after the peak in growth. Final wall maturation occurs in Region III
(Figure 1), where dimers further aggregate with small amounts of additional Ca2+ to form the final rigid
wall of the pollen tube shank.
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Although the focus herein is on Ca2+ we must also recognize that cell wall structure and activity will be
influenced by other ions, notably protons. De-methoxylation of the methyl-esters by PME produces
methanol and a carboxyl group. Given the pH of the wall (5-6), and the pK of pectic acid (about 3.7), it is
likely that all the acidic residues are fully dissociated, and available for Ca2+ binding. However there
could be local domains, governed in part by the influx of protons at the tip of the pollen tube and efflux
along the sides of the clear zone, which could influence the local degree of dissociation.
C. Ca2+ and the Cytoplasm: Many studies richly document the extensive role for Ca2+ as a second
messenger in signal transduction. A full review of this topic is beyond the scope of this article, but can be
gained from several recent and insightful reviews (Harper et al., 2004; Bothwell and Ng, 2005; Harper
and Harmon, 2005; Wheeler and Brownlee, 2008; Kim et al., 2009; Mazars et al., 2009; McAinsh and
Pittman, 2009). We will however, focus on a few aspects that seem especially important in the formation
and structural organization of the cell wall. Most notable is the process of secretion itself, which delivers
many of the building blocks for cell wall formation, including notably the non-cellulosic components
(Nebenführ and Staehelin, 2001). Ca2+ would be expected to occupy a major role because it has been
shown to stimulate secretion in both plant and animal cells (Zorec and Tester, 1992; Battey et al., 1999).
In plants a good example can be found in studies of maize coleoptile protoplasts, where measurements by
patch-clamp analysis of whole-cells show that increasing the [Ca2+] causes a marked increase in the total
surface area of the cell, as detected by an increase in membrane capacitance (Sutter et al., 2000). While
noting the agreement between these results and those obtained from animal cells, Sutter et al. (2000)
allow that plant cells appear to be more responsive to small changes in the [Ca2+] than typical animal
cells. Thus half maximal saturation for the maize coleoptile protoplasts is achieved at 0.9 μM and full
saturation at 1.5 μM. These data and several other examples establish a role for Ca2+ in secretion in
plants.
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Secretion in the pollen tube appears to fit into the Ca2+-stimulus paradigm. Firstly, pollen tubes of all
species, which have been investigated, possess a “tip-focused” gradient of free Ca2+ that is tightly
appressed to the apical PM (Holdaway-Clarke and Hepler, 2003). Ca2+ gradients have even been
observed in Arabidopsis grown in vivo (Iwano et al., 2004; 2009). In lily pollen tubes this gradient
reaches from 3–10 μM (Messerli et al., 2000) and declines to a basal level of 0.1-0.2 μM within 20 μm of
the tip (Pierson et al., 1996). Secondly, the region of high [Ca2+] also contains numerous secretory
vesicles (Figure 1), which have been brought forth by the actin cytoskeleton (Lancelle and Hepler, 1992).
Thirdly, experimental perturbations aimed at affecting the Ca2+ gradient indicate a close coupling between
cell elongation and the presence of the tip-focused gradient; growth correlates with a high apical [Ca2+]
(Pierson et al., 1994; 1996). One contrary example involves inhibition of growth with Yariv reagent,
which blocks arabinogalactan protein activity; however, the pollen tubes exhibit a high apical [Ca2+] (Roy
et al., 1999). In this instance, although growth is blocked, secretion continues but is markedly
delocalized, consonant with a similar delocalization in the elevated Ca2+ domains. Fourthly, experiments
in which the position of the gradient has been modified, either by blocking its formation with local
application of inhibitors such as lanthanides, or stimulating its formation with local application of Ca2+
ionophores, reveal that the location of high [Ca2+] defines the region of cell elongation (Malhó and
Trewavas, 1996). A general conclusion has been that the tip-focused gradient, which is sufficiently high
to activate Ca2+ dependent proteins, facilitates the process of secretion in local domains, and thus not only
stimulates growth, but establishes the polarity of the cell.
Additional insight regarding the connection between Ca2+ and secretion comes from studies of oscillatory
pollen tube growth. Because the underlying processes also oscillate with the same period but not usually
the same phase, it has been possible using cross-correlation analysis to establish whether a process differs
in phase from growth, if so by how much and perhaps most importantly, does it precede or follow the
increase in growth rate (Holdaway-Clarke and Hepler, 2003; Hepler et al., 2006). It has been an
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assumption that processes that increase in anticipation of an increase in growth rate, might play a role in
initiating the growth process, whereas processes that increase after an increase in growth rate, either serve
to inhibit the growth event or by some means prepare the cell for the next stimulatory period.
The phase analysis of intracellular Ca2+ vs. growth rate yields a conclusion that comes as a surprise.
Although the intracellular Ca2+ signal clearly oscillates with changes from 0.75 μM to above 3.0 μM, the
cross-correlation analysis unambiguously indicates that its increase follows by +10-400 the increase in
growth rate (Messerli et al., 2000; Cardenas et al., 2008), an observation that also holds for oscillatory
growth of root hairs (Monshausen et al., 2008). Additionally perplexing have been the data showing that
exocytosis itself oscillates, but that it anticipates the increase in growth rate by –1000 in lily pollen tubes
(McKenna et al., 2009). Ca2+ and secretion thus appear to differ with one another by as much as 1400. A
somewhat similar lack of correspondence between the [Ca2+] and secretion comes from the work of
Camacho and Malhó (2003), who showed that application of the non-hydrolyzable GTP analogue,
GTPγS, which activates G-proteins, causes an increase in exocyotsis. However, these treatments led to a
small but detectable decrease in the intracellular [Ca2+].
These results, however, cannot be construed as evidence against a role for Ca2+ in secretion. It must be
emphasized that the growing pollen tube retains a marked tip-focused gradient, which even at its low
point (0.75 μM)(Pierson et al., 1996) is still quite substantial. If the pollen tube has properties similar to
the maize coleoptile protoplast (Sutter et al., 2000), and reaches maximal rates of secretion at 1.5 μM,
then the oscillating gradient even at its low point may be sufficient to saturate the system. We must keep
in mind that the low values of 0.75 μM derive from studies using fluorescent dyes, and are subject to
signal averaging within the microscope system. At the mouth of a channel, where the ion first enters the
cytosol, the [Ca2+] would be expected to be much higher. However, these postulated high values would
go undetected because they will be averaged with much lower numbers within the resolution limit of 0.2
2+
μm. Because secretion will be regulated by the [Ca ] at the surface of the PM, it may be that those
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values always exceed saturation (1.5 μM). Taken together these considerations support the idea that Ca2+
participates in exocytosis, but that the observed oscillations in [Ca2+] do not cause oscillations in
exocytosis because the ion concentration is already above threshold.
If Ca2+ participates in exocytosis in the pollen tube, what proteins are involved in responding to the ion
and facilitating the process? Given the general role of Ca2+ in both plants and animals, it has been
attractive to imagine that proteins similar to those identified in animal cells are also active in plants. One
candidate includes the annexins, which constitute a large family of proteins found in plants and animals
(Bushart and Roux, 2007; Mortimer et al., 2008; Laohavisit et al., 2009). A feature that implicates these
proteins in secretion consists of a 70 amino acid domain that contains a Ca2+ binding region through
which the protein cross-bridges with the negatively charged phospholipids on membranes. Further
pertinent information includes the observation that these proteins are abundant and close to the PM in
cells undergoing active secretion. For example, annexins are abundant at the apex of tip growing cells
such as pollen tubes (Blackbourn et al., 1992), root hairs (Clark et al., 2005), and fern rhizoids (Clark et
al., 1995). Particularly persuasive has been the observation that annexin stimulates Ca2+ dependent
vesicle fusion to the PM of maize root cap protoplasts (Carroll et al., 1998). A second family of proteins
that are likely involved in transmitting the Ca2+ stimulus during exocytosis are the synaptotagmins, which
participate in exocytosis in animal cells (Martens et al., 2007). Earlier studies had identified these
proteins in plants (Kiyosue and Ryan, 1997). Recent work indicates that synaptotagmin 1 in Arabidopsis
plays a major role in controlling membrane stability, where it facilitates the ability of the PM to reseal
following lesions induced by osmotic or freezing stress (Schapire et al., 2008; Yamazaki et al., 2008).
These properties make synaptotagmin 1 a good candidate for a role in exocytosis where membrane
breakage and resealing occurs during vesicle fusion, and release of material into the extracellular matrix.
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Within the framework of pollen tube growth there are likely several processes, in addition to exocytosis,
that are affected by the tip-focused Ca2+ gradient. Indeed some of these processes may respond to the
oscillations in the [Ca2+]. Obvious candidate proteins are CaM and CDPKs, which participate in many
different processes. Although bulk CaM is relatively uniformly distributed throughout the length of the
pollen tube, the activated form, as revealed through the use of fluorescent analogue (TA-CaM) that binds
to the Ca2+ bound form, localizes to the apex of the pollen tube in a manner similar to Ca2+ (Rato et al.,
2004). In addition the results reveal that activated CaM oscillates similarly to Ca2+ itself, and thus is able
to modulate downstream effectors. However, given the breadth and extent of the CaM literature we will
not attempt to comprehensively review this literature, but instead refer the reader to several recent reviews
(Yang and Poovaiah, 2003; Zhang and Lu, 2003; Du and Poovaiah, 2005; Harper and Harmon, 2005;
McCormack et al., 2005; Boursiac and Harper, 2007; Finkler et al., 2007; Ma et al., 2008; Kim et al.,
2009). We will however mention two CaM target proteins that are particularly pertinent to the control of
Ca2+ itself, and thus likely central in the control of pollen tube growth.
These two Ca2+/CaM targets are: 1) the cyclic nucleotide gated channel CNGC-18 (Frietsch et al., 2007),
and 2) the autoinhibited Ca2+ pump, ACA9 (Schiøtt et al., 2004). CNGC-18 localizes to the apical PM,
and is presumably activated by either cyclic AMP or cyclic GMP, and inactivated by Ca2+/CaM (Talke et
al., 2003; Kaplan et al., 2007; Ma and Berkowitz, 2007). Like other plant CNGC channels, it is thought
to be a non-selective cation channel allowing the influx of both divalent and monovalent cations including
Ca2+ and potassium (Talke et al., 2003; Kaplan et al., 2007; Ma and Berkowitz, 2007). The presence of
CNGC-18 fits well with an earlier study showing that transient elevations of cAMP modulate the pollen
tube growth axis (Moutinho et al., 2001). ACA9 also localizes to the PM, but is distributed uniformly
over the length of the pollen tube, and is predicted to be activated Ca2+/CaM. Their different localization
and complementary functions suggest that CNGC-18 and ACA9 closely interact to control the
intracellular [Ca2+]. Thus the opening of the CNGC-18 allows Ca2+ to enter the cell. Because this
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channel is located to the apical PM, Ca2+ influx would be restricted to the tip region, consistent with
numerous observations. However, the elevated [Ca2+] would bind CaM, and close the channel, blocking
further entry of Ca2+ (Hua et al., 2003; Kaplan et al., 2007). In this sense CNGC-18 is likely selfregulatory, and potentially able to restrict a massive build up of Ca2+. Nevertheless, elevated domains are
generated, and these, if allowed to persist and spread, could cause serious physiological problems; for
example, Ca2+ could react with phosphates and block energy (ATP) production. The ACA9 pump thus
becomes an essential factor in reducing the intracellular [Ca2+] to basal levels. While we acknowledge the
presence of Ca2+-pumps on the internal membranes (ER, mitochondria, vacuole), ACA9 serves a key role
in extruding Ca2+ into the cell wall. Although ACA9 is localized over the entire pollen tube, it will only
be active where Ca2+/CaM is high. Normally this would be in the apical domain, where ACA9 would
return Ca2+ specifically to the apical apoplasm (Schiøtt et al., 2004). However, if the pollen tube suffers a
wound anywhere along its length, the inevitable rise in [Ca2+] would locally bind CaM, and activate
ACA9, thus lowering the [Ca2+] and restricting damage.
The Ca2+ dependent protein kinases or the calmodulin domain protein kinases (CDPKs) constitute another
large family of Ca2+ binding proteins uniquely found in plants and some protists (Harper et al., 2004;
Harper and Harmon, 2005). They too are being widely and extensively investigated, and thus will not be
considered in detail here. Briefly we note that several of these kinases are membrane associated through
myristoylation or palmitoylation linkages. In early studies Moutinho et al. (1998) identified a Ca2+dependent protein kinase activity, possibly a CDPK, associated with the pollen tube apex, which appeared
to participate in polarized growth. Recent studies in Arabidopsis have focused on CDPK 17 and 34,
which are similar to one another, and localized to the PM being biased towards the apical domain (Myers
et al., 2009). These appear to have a profound effect on pollen tube growth as evidenced by observations
that double mutants have slower growth rates, and are nearly sterile. Petunia CDPK 1, which is similar to
Arabidopsis CDPK 17/34 and also PM localized, likewise participates in pollen tube growth (Yoon et al.,
16
2006). Although these kinases are crucial target proteins that respond to changes in the [Ca2+], we know
much less about the downstream components. With CDPK 34 alone there may be more than 50 proteins
that are phosphorylated (Myers et al., 2009); determining the specific targets and processes associated
with them constitute important goals in future research.
D. Ca2+ at the PM/Cell Wall Interface:
It has been appreciated for years that extracellular Ca2+ affects the stability and permeability of the plant
cell PM, with concentrations between 0.1 and 1.0 mM being required to retain the integrity and selective
ion transport properties (Hanson, 1984; Hepler, 2005). Diverse processes such as retardation of leaf and
tissues senescence (Hanson, 1984), and closure of stomatal pores are stimulated by extracellular Ca2+ in
the range of 0.1 to 1.0 mM (McAinsh and Pittman, 2009). The likely basis for the effects in guard cells
may derive from the activity of a Ca2+ sensor found on its PM (Han et al., 2003). Plant cells thus have
mechanisms for responding to Ca2+ in the cell wall. These responses may involve changes in the
permeability of the PM itself to Ca2+ and other ions, but they may involve changes in the interaction
between the PM and cell wall that affect many aspects of cell growth and development. In the paragraphs
below we provide specific examples where extracellular Ca2+ and extracellular factors act as
intermediaries between the PM and cell wall, and in most instances induce a rise in intracellular [Ca2+].
1. Mechanical Activation: A Role for Stretch Activated Channels (SACs): It is well known that plant
cells respond to a variety of mechanical stimuli, some of which may be generated endogenously
(Monshausen et al., 2009; Monshausen and Gilroy, 2009). A prime example of the close interplay
between the cell wall and the cytoplasm revolves around the activity of SACs on the PM. Although we
have mentioned channels earlier, i.e., CNGC-18, we discuss SACs here because they respond to
deformations of the PM, which are dependent on preceding changes in the extensibility of the cell wall. In
pollen tubes a candidate SAC has been identified that appears to localize specifically to membrane
17
patches derived from the growing tip of lily pollen tubes, and that selectively conducts Ca2+ (Dutta and
Robinson, 2004). Within the context of oscillatory pollen tube growth this channel emerges as the best
candidate to account for the corresponding oscillations in the tip-focused Ca2+ gradient. It seems
plausible, therefore, that as the apical cell wall relaxes and the pollen tube extends, the PM in this specific
location, would deform due to the force of turgor pressure. The SAC would be activated and Ca2+ entry
would occur, being tightly localized to those regions of the PM experiencing deformation. Not only does
this scenario explain the spatial positioning of the tip-focused Ca2+ gradient, but it accounts for its
temporal properties in relation to changes in growth rate. Thus as the cell extends, and the PM deforms, a
Ca2+ rise occurs, which slightly follows the increase in growth rate, as observed. Similarly as growth
slows, PM deformation lessens, the SAC closes and the tip-focused gradient declines (Dutta and
Robinson, 2004). Although CNGC-18 may contribute to the formation and oscillation of the tip focused
Ca2+ gradient (Frietsch et al., 2007), we think that the SAC, described by Dutta and Robinson (2004) is a
much better candidate. For example, while CNGC-18 localizes to the apical PM, its distribution is rather
broad, extending across the dome and partly down the flank of the pollen tube apex. There is no evidence
that it would give rise to the focused high [Ca2+] that is observed. Also the rise and fall of the tip-focused
Ca2+ gradient, which is tightly coupled to changes in the growth rate, is elegantly explained by an apically
positioned SAC that is activated by wall loosening and turgor-dependent extension of the wall and
associated PM. While the studies of Dutta and Robinson (2004) provide physiological evidence for the
presence of a stretch activated channel in the PM of pollen tubes, more recent work on Arabidopsis
provides molecular evidence for the existence of a SAC. Nakagawa et al. (2007) have isolated and
characterized an Arabidopsis cDNA, MCA1, which complements a lethal phenotype in the yeast
Saccharomyces for a stretch activated Ca2+ channel. In Arabidopsis this channel increases [Ca2+] when
the plasma membrane is deformed.
18
2. CaM: A persuasive case is building for a role for extracellular CaM in the control of plant cell growth
and development. Ma and Sun (1997) showed that an anti-CaM serum, and the CaM antagonist, W-7
strongly inhibited pollen germination and tube growth, and that the inhibitory effects could be reversed by
addition, up to 1.0 μM, of purified cauliflower CaM. In further studies Ma et al. (1999), show that
heterotrimeric G-proteins may be involved in the signal transduction scheme for CaM. Using
microinjection they show that addition of GDP-β-S or an anti-Gα antibody, both of which block
heterotrimeric G-proteins, also slow pollen tube growth. In addition they report that anti-CaM inhibition
can be reversed by cholera toxin, a G-protein agonist, or that CaM stimulation can be inhibited by
pertussis toxin, a G-protein antagonist. Taken together they suggest that heterotrimeric G-proteins
participate in the signal transduction pathway that involves extracellular CaM as a regulator of pollen tube
growth. A possible downstream target has been identified in suspension-culture cells of Angelica; this is
a 21 kDa CaM binding protein that is localized in the cell wall (Mao et al., 2005).
More recent studies indicate that CaM, although present in the apoplasm, specifically binds to the outer
surface of the PM, and that it stimulates the influx of Ca2+. Binding at the PM has been shown most
convincingly with a quantum dot-CaM probe, using both fluorescence and transmission electron
microscopy. With ion imaging, Shang et al. (2005) first showed that extracellular addition of purified
cauliflower CaM stimulates Ca2+ entry into pollen grains. In an extension of that study, Wang et al.
(2009) report that the quantum dot-CaM probe also stimulates Ca2+ influx. In parallel with results noted
above, Shang et al. (2005) further show that application of a CaM anti-serum, or the CaM antagonist W-7,
inhibits the increase in intracellular Ca2+. Patch clamp studies further indicate that Ca2+ influx occurs
through a voltage gated channel, which based on its characteristics appears to belong to the category of
hyperpolarization-activated Ca2+-permeable channels that participate in growth regulations at potentials
more negative than -100 to -150 mV (Shang et al., 2005).
19
Because CaM is synthesized in the cytosol it is a puzzle how it is delivered to the apoplasmic space, or
how its concentration might change to cause the reported changes in intracellular Ca2+. It is conceivable
that during secretion CaM leaks out as a result of microlesions in the PM. Because exocytosis oscillates
so too would the transport of CaM to the wall. For the pollen tube this would translate into an oscillatory
profile for extracellular CaM in the apical apoplasm. In turn this could generate a positive feedback loop
wherein the process of exocytosis delivers more CaM to the wall, which then causes further Ca2+ influx
into the apical cytosol.
3. Annexins: The annexins are soluble, multifunctional proteins usually found in the cytosol, but also
associated with membranes and sometimes integral components of the membrane (Bushart and Roux,
2007; Mortimer et al., 2008; Laohavisit et al., 2009). They bind Ca2+, hydrolyze nucleotides, interact with
membrane phospholipids, exhibit peroxidase activity, bind to the actin cytoskeleton, and in general seem
capable of myriad functions. We previously made reference to their role in facilitating secretion; here we
draw attention to the observations showing that these proteins are themselves secreted and that they can
affect the activity of the PM from the apoplasm (Laohavisit et al., 2009). Somewhat similarly to CaM,
annexins can facilitate the influx of Ca2+ into the cytosol. However in this instance they appear to do so
by inserting in the membrane and constructing conductance channels through which Ca2+ can pass
(Laohavisit et al., 2009). The annexins, together with CaM, appear able to establish a positive feedback
loop of reinforcing signals. Indeed the annexins may facilitate their own secretion and through their
subsequent activity as a stimulator of Ca2+ influx, they may be able to promote further secretion or at least
bias secretion to the regions where the annexins are active.
4. WAKs: The cell wall associated kinases or WAKs are potentially key elements in the continuum
between the cell wall and the cytoplasm (Kohorn, 2000; Kohorn et al., 2006a; 2006b). These proteins,
which occur widely in plants including pollen tubes, consist of a single transmembrane domain with a
20
serine/threonine kinase on the cytoplasmic side and an extracellular fragment that associates with the wall
polymers in the apoplasm. In the wall the WAKs bind to pectins, in a manner that is dependent on the
Ca2+ cross-linked or “egg-box” configuration of de-esterified galacturonic acid residues (Decreux and
Messiaen, 2005). Reducing the [Ca2+] in the wall, or more precisely the Ca2+/monovalent ion ratio that
retards egg-box formation, specifically inhibits the binding of WAK1. The association between pectin
and WAKs may occur very early in development as studies show that these components appear together
in the Golgi apparatus and are secreted as a complex (Kohorn et al., 2006a). Although we lack a great
deal of information about how these kinases operate some recent evidence suggests that they may affect
sugar metabolism (Kohorn et al., 2006b). Specifically it has been reported that mutants in WAK2, which
exhibit a dwarf phenotype, are defective in vacuolar invertase activity. Because cells experience a slight
decrease in turgor pressure as a loosened wall extends, Kohorn et al. (2006b) suggest that the reduced
invertase activity may prevent mutant cells from restoring normal turgor pressure in association with cell
elongation process. While much remains to be discovered the WAKs emerge as extremely interesting
candidates for transducing a signal from the cell wall to the cytoplasm that is influenced by the Ca2+ poise
within the wall.
5. Oligogalacturonides (OGs): An important development has been the realization that certain cell wall
fragments, notably the α 1, 4 OGs, derived from pectins, serve as signaling agents in response to plant
pathogens and also for normal development (Lecourieux et al., 2006). Although we are not aware of a
specific information that applies directly to pollen tubes, given the pectin rich cell wall and the likely
presence of the appropriate OGs, their role in the pollen tube growth seems entirely plausible. Very
briefly these OGs, with a degree of polymerization of 10 to 15 and exhibiting the Ca2+ cross-linked egg
box configuration, are able to stimulate an intracellular increase in the [Ca2+]. For example, in studies in
aequorin transformed Arabidopsis cells, the application of OGs induce a sharp rise in Ca2+. This rise can
be blocked by lanthanum, which is a Ca2+ channel blocker or by tetrabromobenzotriazole, which is a
21
protein kinase inhibitor, indicating that you can prevent the cellular response either by preventing Ca2+
from entering the cell at the channel or by blocking a kinase activity inside the cell (Moscatiello et al.,
2006). Moscatiello et al. (2006) further performed a transcript analysis following OG application and
their results draw attention to changes in cell wall modifying factors as well as to factors involved in the
biosynthesis of jasmonate. To some degree there appears to be a an intersection between the WAK
requirement, mentioned in the previous section, and OGs discussed here; both involve a role for Ca2+
cross-linked pectin fragments exhibiting the egg-box configuration and a degree of polymerization of 10
to 20. Additional work on these pectic wall fragments could provide important insight about the ability of
cell wall to transmit signals to the cytoplasm.
6. Extracellular ATP: Finally we draw attention to extracellular ATP (eATP), which is yet another factor
that is able to modulate intracellular Ca2+ (Demidchik et al., 2003; Tang et al., 2003; Jeter et al., 2004;
Kim et al., 2006; Song et al., 2006; Foresi et al., 2007; Roux and Steinebrunner, 2007; Demidchik et al.,
2009). As with CaM we normally think of ATP as residing entirely in the cytoplasm in the mM level and
where it serves as a universal energy currency for the cell. However it is well known in animal cells and
now emerging in plants including pollen tubes (Steinebrunner et al., 2003) that ATP occurs in the
extracellular or apoplasmic domain, where its concentration (0.3-40 μM) is much less than in the
cytoplasm (Roux and Steinebrunner, 2007). Consequently eATP probably does not energize reactions,
rather it appears to act as a signaling agent. Specifically eATP has been shown to increase the
intracellular [Ca2+].
Because ATP is synthesized in the cytoplasm a question arises on how it moves to the apoplasmic space.
Different possibilities emerge including leakage through wound sites, or osmotic and mechanical stress
lesions, and through secretion itself, where the ATP will be trapped in vesicles along with other material
that is being exocytosed. Different studies show that the highest [eATP] occurs in association with
22
amplified exocytosis, as for example at the tip of root hairs (Kim et al., 2006; Wu et al., 2008). Once in
the apoplasm eATP is partly inactivated by the enzyme apyrase. It might seem that these conditions
would completely destroy the activity of eATP; however that is not true because studies on pollen
germination reveal that grains with much reduced apyrase activity are prevented from growing
(Steinebrunner et al., 2003). From these observations Roux and Steinbrenner (2007) suggest that the
[eATP] must be at an intermediary level between low and high extremes to support normal growth. The
activity of eATP also depends upon the availability of Ca2+ in the extracellular matrix. If Ca2+ is chelated,
eATP cannot stimulate an increase in the intracellular [Ca2+] (Demidchik et al., 2003; Jeter et al., 2004).
With all of the agents that stimulate an increase in [Ca2+] much remains to be learned about the identity
and function of the downstream targets. A recent study makes a compelling link between the rise in
intracellular [Ca2+] induced by eATP, and the stimulation of an NADPH oxidase-dependent increase in
reactive oxygen species. Here the train of events appears to flow from eATP to the activation of a PM
NADPH oxidase, with an increase in reactive oxygen species. The later agent then stimulates the opening
of a Ca2+ channel, which causes the intracellular elevation of this ion (Demidchik et al., 2009). We
acknowledge that research on reactive oxygen species attracts a great deal of interest these days owing to
the central role that these toxic factors pay as signaling agents in controlling response to pathogens, as
well as normal growth and development (Mori and Schroeder, 2004; Carol and Dolan, 2006). Although
we start to gain information on the pathway for eATP activity, we nevertheless lack knowledge of the
presumed receptor. Continued research on this exciting topic is clearly warranted.
E. Conclusion:
A considerable amount of information increasingly shows the existence of an extensive interplay between
the cytoplasm and the cell wall, which contributes to the control of plant growth and development. We
have long known that the cytoplasm gives rise to the cell wall through the secretion of the building
23
blocks. More recently we realize that the cell wall can have a profound effect on the cytoplasm with Ca2+
participating in different regulatory pathways. Ca2+ has recently held our attention because of its central
role as a second messenger to control many different processes. Now it becomes crucial to consider the
interaction between the cytoplasmic Ca2+ and cell wall Ca2+ with the intervening PM occupying a pivotal
position in the process of information transfer. The conversation, however, is definitely two ways; the
cell wall through its ability to bind and control the [Ca2+] at the surface of the PM, influences the entry of
the ion into the cytoplasm. But, through the activity of pumps the cytoplasm can extrude Ca2+ into the
wall and affect local structure and activity of wall components. With the increasing number of factors
that exert further control over this process, we come to recognize a complexity that could not imagined a
few years ago.
In the last section of this brief essay we have drawn attention to a few factors, namely SACs, CaM,
annexins, WAKs, OGs, and ATP that either occur in the apoplasm, or interact with the cell wall to
produce a signal that requires Ca2+ or influences its translocation in ways that affect growth and
development. With the exception of the WAKs, a common theme appears to emerge that ties these
factors together, namely the ability to stimulate an increase in the intracellular [Ca 2+]. Although we have
ideas about the specific signatures of the Ca2+ responses for SACs, we know much less about the other
factors that modulate the intracellular [Ca2+]. For example, are the Ca2+ elevations restricted to specific
locations in the cell, do they oscillate and if so their amplitude and period, and are they temporally brief
or prolonged? This information could provide clues about the downstream targets, and begin to make it
possible to sort out different pathways. We also imagine that additional factors will emerge. When
considered as a whole it becomes evident that there is an extensive system involving the interaction
between the cell wall and the cytoplast, with Ca2+ as an intermediary. Resolving the details will provide
insight about the control of plant growth and development.
24
Acknowledgments:
We thank our colleagues at the University of Massachusetts at Amherst for stimulating discussion on the
ideas presented in this article. This work was supported by a grant from the USA National Science
Foundation (MCB-0847867).
25
Figure 1: Ca2+ flux at the pollen tube tip. Ca2+ diffuses into the pollen tube wall (large green arrows) at
different rates across the expanding tip, driven by gradients created by different Ca2+ binding mechanisms
in the wall. In zone I, a moderate flux of Ca2+ (green circles) binds ionically to carboxyl groups on newly
secreted, loosely associated, highly methoxylated pectin chains (red lines). More pectin arrives
continuously in this zone by exocytosis of secretory vesicles (red circles). Pectin methylesterase (PME) is
also secreted in inactive form (yellow circles). As the pollen tube wall expands, stretch-activated channels
(blue boxes) open creating a small Ca2+ influx (narrow green arrows) that sustains a dynamic Ca2+
gradient in the cytoplasm, created by Ca2+ uptake by the endoplasmic reticulum. In zone II PME becomes
active (yellow stars) and de-methoxylates pectins in block-wise fashion, leading to tight association of
pectin dimers through coordination with Ca2+. Ca2+ demand for this process is very high. In zone III
pectin dimers form aggregates, also bound with Ca2+, creating a mature, inextensible pollen tube wall.
26
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