- freely available
Plants 2013, 2(4), 615-634; doi:10.3390/plants2040615
Abstract: Calcium (Ca2+) plays important roles in plant growth, development, and signal transduction. It is a vital nutrient for plant physical design, such as cell wall and membrane, and also serves as a counter-cation for biochemical, inorganic, and organic anions, and more particularly, its concentration change in cytosol is a ubiquitous second messenger in plant physiological signaling in responses to developmental and environmental stimuli. Actin cytoskeleton is well known for its importance in cellular architecture maintenance and its significance in cytoplasmic streaming and cell division. In plant cell system, the actin dynamics is a process of polymerization and de-polymerization of globular actin and filamentous actin and that acts as an active regulator for calcium signaling by controlling calcium evoked physiological responses. The elucidation of the interaction between calcium and actin dynamics will be helpful for further investigation of plant cell signaling networks at molecular level. This review mainly focuses on the recent advances in understanding the interaction between the two aforementioned signaling components in two well-established model systems of plant, guard cell, and pollen.
Higher plants are equipped with signaling systems that play essential roles to fulfill their life cycles. Plant signaling transduction can be divided into several steps: the perception of developmental and environmental stimuli, the transduction of cellular signal by complex networks, and the physiological and morphological responses. Numerous ions, molecules, genes, proteins, and physiological processes and biochemical reactions are involved in a typical signaling cascade in a cell (e.g., guard cell ABA signaling) . Biochemical, molecular, and genetic experiments have supported a comprehensive network of plant signaling even for limited plant physiological process carried by specific cells, e.g., guard cell invoked stomatal movements for transpiration, or pollen germination, and tube growth during pollination [2,3]. This review focuses on the significance of the two important signaling components, Ca2+ and actin cytoskeleton, especially on the interaction between them, in guard cells and pollens.
Ca2+ is an indisputably important element for both cell structure and signaling during plant growth and development, and also the accumulating data demonstrate its indubitable significance in plant nutrition. Furthermore, recent researches support the concept of cytosolic free Ca2+ ([Ca2+]cyt) as an intracellular second messenger that mediates developmental and environmental stimuli into plant physiological and biochemical responses [4,5]. The investigations on the latter topic as how Ca2+ as one of the most important regulator in plant signaling have been carried out from whole-plant to single protein levels . [Ca2+]cyt is maintained lower than 100 nM in resting plant cells while Ca2+ in the extracellular and organellar levels is at millimolar level . The elevation or oscillation of free Ca2+ concentration in the cytosol acts as the [Ca2+]cyt signal, which is the result of stimulus triggered Ca2+ influx from extracellular and release from endo-organelles (e.g., vacuole and endoplasmic reticulum) through Ca2+-permeable channels on plasmamembrane and endomembrane, respectively [6,7,8,9]. Many research groups have been attempting to discover upstream components and downstream effectors of Ca2+ signaling network.
Actin dynamics play important roles in many cellular signaling cascades in plants. In almost all eukaryotic cells, actin exists in two forms: filamentous actin (F-actin) and monomeric actin or globular actin (G-actin). F-actin filaments (with 5 to 7 nm diameter) play important roles in cell shaping, division, growth, vesicle and organelle movements, cell death, as well as signaling in responses to biotic and abiotic stresses [10,11,12,13]. G-actin can polymerize into F-actin with a characteristic of architecture and polarity in cytosol and could form actin filament bundles or cables as actin cytoskeleton. The actin dynamics as polymerization and de-polymerization between the G-actin and F-actin is an essential process for actin function, and could be regulated by developmental and environmental signals . The actin dynamics needs the existence of enough G-actin accompanies with visible F-actin bundles. Indeed, in Arabidopsis pollens, the G-actin is estimated at very high concentration to about 50 µM , and only very small amount of a plant cell’s G-actin polymerizes into F-actin . The superabundance of unpolymerized G-actin in plant cells suggests they are readily available for quick polymerization in response of stimuli , and also indicates the highly dynamic characteristic of actin network.
2. Why Guard Cells and Pollens
Not only cells are the structural and functional unit of all organisms but also vital responder to internal and external stimuli. Hence, the study of signaling network at cellular level is very important. Plants contain many specialized cells, for example, guard cells and pollens. The characteristic of a specialized cell is controlled by expression of a particular set of genes, while keeping the others repressed, and epigenetic regulation is also involved for their differentiation processes during development. The difference between guard cells and pollens has been identified by large-scale transcriptomic and proteomic analyses and the hypothesis has been proved [18,19,20,21,22,23]. These findings indicate that the signaling pathways maybe different among the different plant cell types or even different developmental stages of same cell type. Consequently, the comparison the signaling network of different specialized plant cells and the cells at different stages of similar cell type will be very helpful for understanding the common signaling mechanisms in response to developmental and environmental stimuli.
Leaves are the main photosynthetic organs of plant and are important for plant vegetative growth. In addition, leaves are also important for transpiration, and play important roles in response to biotic and abiotic stresses to cope with the environment and plant survivability [24,25,26]. The absorption of atmospheric carbon dioxide (CO2) (a substrate for carbohydrate biosynthesis by photosynthetic reaction), and the loss of water vapor by transpiration, are regulated by stomata (the microscopic pores present on the leaf surfaces). Stomata are consisted of guard cells surrounded in pairs, and are separated functional complex with no plasmodesmata connected with the contacted epidermal cells. Stomata are also major entry points for pathogen invasion. The opening and closing of stoma are important for gas exchange as well as to cope with the environment and plant survivability in response to phytohormone and environmental stresses [27,28,29]. Many internal and external (biotic and abiotic) stimuli are well known regulators of stomatal aperture, including hormones (e.g., ABA, auxin, and ethylene), blue and red light, water status, CO2 concentration, circadian clock, temperature, and pathogen [2,30,31,32]. The stomatal movements are controlled by the volume change of guard cells, and the volume change is the result of turgor alteration. During stomatal opening, guard cells accumulate solutes such as potassium, anions, and malate, and the increased osmolality enables water uptake, swelling of guard cells that in turn causes opening of stomata. In the process of stomatal closing, guard cells lose osmotic solute by efflux mechanism, and malate metabolizes into osmotically inactive starch that in turn facilitates shrinking of guard cells and closing of stomata [2,31,32,33]. Because of their physiological importance, guard cells have been well studied at whole-plant, whole-leaf, cellular, subcellular, and molecular levels by using different investigation tools, and they have been developed as the first model system for plant cell signaling. Both, Ca2+ and actin dynamics, have been suggested to be involved in guard cell singling networks.
After the transition from vegetative growth into reproductive growth stage, the fertilization is the most important process for sexual reproduction in flowering plants, and that begins with landing of the mature pollen on the top of a suitable pistil, followed by pollen hydration, germination, and pollen tube polar growth, and fertilization of the female gametophyte . During the processes of pollen germination and tube elongation, many factors, such as Ca2+, K+, H+, Cl−, NO, ROS, and cytoskeletal components, are involved in the regulatory network [34,35,36,37,38,39]. Among these regulatory factors, Ca2+ has been indicated as a crucial player in pollen signaling [4,40,41], and actin organization patterns and actin dynamics have been implicated in cellular signaling as well as in cellular architecture [11,42,43]. As pollens and pollen tubes are representatives for the investigation of haploid and polarized growth of plant cells, together with their significance in double fertilization, pollens and pollen tubes have been regarded as very important systems for reproduction and cellular signaling studies.
Both, guard cells and pollens, are well established model systems for investigation of plant signaling networks, so the similarities and differences between them will give us common signaling characteristics of plant cells, and the accordance of specific cell functioning and signaling. About 20 years ago, the elevation of [Ca2+]cyt was shown to trigger the fragmentation of actin bundles in pollen tubes and actin reorganization in guard cells [44,45]. Research findings indicate that the disruption of actin also affects the physiological processes during pollen germination, pollen tube growth, and stomatal movements [34,43,46,47,48]. The actin dynamics also regulate Ca2+-permeable channel activity and [Ca2+]cyt signal in pollens and guard cells [41,49,50]. These data suggest the existence of a complicated regulatory feedback loop between the two important signaling elements, [Ca2+]cyt and actin.
3. Means to Study Ca2+ and Actin Signaling
Without the aid of developed microscopy, biochemical, and genetic techniques for plant cell investigation, it would be impossible to detect the changes in the levels of [Ca2+]cyt, the structure or/and dynamics of actin, and to determine the roles of Ca2+ and actin in diverse physiological and developmental processes in guard cell and pollen signaling.
Calcium imaging is the very powerful and visible technique to study the change of [Ca2+]cyt in living plant cells. [Ca2+]cyt concentration change shows complex patterns, such as spiking or oscillation in response to upstream regulators and, thus, to trigger the downstream cellular responses [5,7]. In response to developmental or environmental cues, [Ca2+]cyt concentration can be varied from 100 nM (for maintenance of resting cells) to above μM level (during activated state of the cells). As high concentrations of [Ca2+]cyt is toxic to living cells, the elevation of [Ca2+]cyt is brief and/or subcellularly restricted within the physiological range. A set of small-molecules, Ca2+ sensing fluorescent dyes (e.g., Fluo-3, Fura-2) have been widely used to detect [Ca2+]cyt changes in plant cells [41,50,51,52,53], and the advantage of these dyes is that they can easily be applied to all kinds of plant cells [52,54,55]. FRET-based Ca2+ sensors, such as yellow cameleon (YC) 2.1 and higher versions of yellow cameleon (e.g., YC 3.6), are considered as a more sensitive and efficient method than to fluorescent dyes [9,56]. To detect the relative or quantitative change of [Ca2+]cyt, a fluorescent microscope is needed for Ca2+ sensing fluorescent dye application, and FRET-based sensor methods require a confocal microscope.
It is well known that, Ca2+ channels located on the plasmamembrane and endomembrane facilitate the Ca2+ influx and [Ca2+]cyt rise that, in turn, cause changes of [Ca2+]cyt. Based on above facts, these channels could be regarded as direct upstream elements for generation of [Ca2+]cyt in plant cells. Patch clamp technique enables the recording of Ca2+ channels in plant cells [6,7,8,57]. Due to the poor selectivity of plant cell Ca2+ channels for the cations, a Ca2+ channel is referred as Ca2+-permeable channel [58,59,60]. By using electrophysiological means (the activation and gating mechanisms), several kinds of Ca2+-permeable channels have been recorded in the plasma membrane, tonoplast, endoplasmic reticulum, chloroplast, and nuclear membranes of plant cells, including depolarization-activated, hyperpolarization-activated, voltage-independent, stretch-activated, and cyclic nucleotide gated (CNG) Ca2+ channels, as well as their pharmacological characteristics [7,50,60,61,62]. In addition, the application of stretch-activated Ca2+ channel blocker, Gd3+, stops the osmotic stress induced [Ca2+]cyt elevation, indicating the opening of plasma membrane Ca2+ channels is essential to initiate the [Ca2+]cyt signaling . Several gene families have been suggested as Ca2+ channel candidates, in Arabidopsis, that include cyclic nucleotide gated channel family (CNGC, 20 members), glutamate receptor homolog family (GLR, 20 members), and a two-pore channel (TPC1) [8,61,63,64,65].
To accomplish biochemical and physiological responses of the [Ca2+]cyt signals triggered by internal or external stimuli, downstream Ca2+ sensors play vital roles in the perception and decoding of Ca2+ signals. There are four gene families with EF-hand Ca2+ binding domain that have been suggested as Ca2+ sensors in plants, and these families include calmodulin family, calmodulin-like protein (CML) family, Ca2+-dependent protein kinase (CDPK or CPK) family, and the calcineurin B-like protein (CBL) family [66,67]. The large number of calcium sensor proteins share different Ca2+-binding characteristics, and with diverse subcellular localizations and distinct downstream signaling interactions that enable these proteins to decode the information related to Ca2+ oscillations and spikes and subsequently to process of the information into appropriate cellular function.
To study the function and organization of actin during plant development and signaling, pharmacological agent(s) (drug) that causes disruption of actin filaments (e.g., latrunculin B), causes stabilization of actin filaments (e.g., phalloidin), and blocks actin polymerization (cytochalasin), were used previously [68,69,70,71]. Besides pharmacological agents, fluorescent methods were also usually used to visualize actin and were applied for actin imaging, prior to the discovery of fluorescent proteins . Fluorescent proteins have the higher efficiency than fluorescent dyes to visualize actin organization and actin dynamics in living plant cells. GFP fused with actin binding domain or actin regulatory proteins have been demonstrated to bind F-actin in vivo [59,73,74,75]. Furthermore, a green to red photo-convertible probe, mEosFp::FABD-mTn, has been shown to work with higher precision for labeling F-actin and can be used for visualization of actin dynamics and interactions with microtubules and other organelles . Lifeact-mEGFP was also suggested as a new versatile probe of F-actin without changing actin dynamics nor the physiological activities of living plant cells [77,78]. These actin labeling and imaging techniques supplied powerful tools for study of physiological roles of plant actin skeleton and actin dynamics in living cells.
4. Crosstalk of Actin and Ca2+ in Plant Cells
During developmental processes and in response to various stresses, most often, plant cells show both change of [Ca2+]cyt and alteration of actin structure. For instance, during stomatal movements, Ca2+ signal activation is needed for both opening and closure, and rapid actin rearrangement is also essential for normal stomatal function [2,28,44,79,80]. The same phenomena have also been reported in other plant cells, such as root hairs during their growth [81,82], xylem cells during their differentiation [83,84,85], and in pollens during their germination and tube elongation [34,45]. Below, we elucidate the signaling crosstalk between Ca2+ and actin in guard cell and pollen systems.
4.1. Ca2+ and Actin in Guard Cells
In guard cell signaling networks, multiple stimuli that result in change of stomatal aperture mostly utilize [Ca2+]cyt as a second messenger. In the well-established stomatal movement system, for ABA induced stomatal closing and ABA inhibited stomatal opening processes, [Ca2+]cyt elevation or oscillation could usually be seen, even in Ca2+-independent signaling pathway (e.g., pH pathway) [2,31,86,87,88,89,90,91]. A recent report also indicates that [Ca2+]cyt plays a positive regulatory role in ABA activation of pHcyt . [Ca2+]cyt elevation has been shown to inactivate inward K+ channels and activate slow anion channels  and, thus, leading to stomatal closure . In auxin, blue light and Cyclic GMP induced stomatal opening, [Ca2+]cyt elevations have been observed [94,95,96].
In addition to ABA, other abiotic stimuli such as high CO2, cold shock, ROS, and biotic-stress by pathogens also cause [Ca2+]cyt elevations and stomatal closure [57,97,98,99]. The removal of extracellular Ca2+ using EGTA abolishes [Ca2+]cyt elevations in response to H2O2, cyclic GMP and hypo-osmolality, indicating plasma membrane Ca2+ influx is essential to initiate the [Ca2+]cyt-mediated signaling [50,57,100]. These reports suggest that the importance of Ca2+ influx through the Ca2+-permeable channels on the plasma membrane for generation of [Ca2+]cyt, and its role in stomatal movements, are response to both abiotic and biotic stresses.
Both CNGC1 and CNGC2, two members of CNGC family, are permeable to Ca2+, but also permeate K+ and are regulated by cyclic nucleotides, calmodulin, and pathogens. The activation of these CNGC channels in response to pathogen generates Ca2+ (Ba2+), current in Arabidopsis guard cell protoplasts, suggesting a possible role of CNG channels in Ca2+-based signaling in this cell type [101,102,103,104,105]. AtTPC1 channel also has been indicated to be involved in Ca2+ signaling cascades . Recently, AtTPC1 has been shown to encode a vacuolar two pore channel 1 and its function has been implicated in Ca2+ signaling and stomatal closure [63,107,108].
In guard cell signaling several Ca2+ sensors have been identified that sense and amplify the Ca2+ signal. In Arabidopsis thaliana, CPK23 exhibits a stoma phenotype under water stress. T-DNA knockout mutant plants of CPK23 gene show enhanced drought resistance and decrease stomatal aperture, while its overexpression lines show hypersensitivity to drought and wider stomatal aperture ; CPK3 and 6 are involved in ABA regulation of guard cell slow-type currents , CPK6 also is a positive regulator in methyl jasmonate signaling in guard cells ; CPK4 and CPK11 phosphorylate ABF4 and ABF1 (two members of the ABA-RESPONSIVE ELEMENT BINDING FACTORS (ABFs)) in vitro, and act as positive regulators of stomatal movements in calcium-mediated ABA signaling pathways . SLAC1, a slow anion channel on the plasma membrane, is important for stomatal closing, could be activated by coexpression with CPK21 and CPK23 heterogeneously [113,114,115]. In addition to CPKs, other calcium sensors have been identified. CBL1 and CBL9, two members of CBLs, have been shown to play important roles on guard cell ABA signaling and interact with CIPK23. The cbl1 cbl9 double mutant plants show hypersensitive to ABA regulation of stomatal movements and more resistance to drought treatment . Extracellular application of calmodulin induces stomatal closure via a G-protein, nitric oxide, and H2O2 invoked pathway [117,118], suggesting the possible importance of calmodulin in regulation of guard cell signaling downstream of Ca2+ signal. CML9 and CML4 are involved in stress responses including ABA-inhibited seed germination and young seedling growth [119,120], however, the function of this family in guard cell Ca2+ signaling still needs further investigation.
Actin dynamics plays important role during stomatal movements. During light inducted stomatal opening and the dark or ABA induced stomatal closing, the actin reorganizes between a radial pattern (opening status) and a randomly oriented and short-fragmented pattern (closure status) [44,46], and the stomatal opening induced by fusicoccin is delayed with the pretreatment with phalloidin (a pharmacological agent that prevents de-polymerization of actin filaments) . The ABA-insensitive mutant (abi1-1), which is impaired in ABA induced stomatal closure, fails to depolymerize actin and stomatal closure [121,122,123]. In wild type Arabidopsis plant, a small guanosine triphosphatase (GTPase) protein AtRac1 is inactivated by ABA, and the inactivation of AtRac1 GTPases induces disruption of the guard cell actin cytoskeleton and stomatal closure, while in abi1-1, neither AtRAC1 inactivation nor actin cytoskeleton disruption nor stomatal closure happens , indicating ABI1 functions upstream of AtRAC1 and actin in guard cell ABA signaling. ABA induced actin disruption could be dismissed by removal of external Ca2+ with application of EGTA, and treatment of Ca2+ could mimic the ABA induced actin rearrangement is guard cells, suggesting the Ca2+ regulation of actin dynamic pathway is important in ABA-mediated stomatal movements . Osmotic stress is also involved in actin dynamics and stomatal movements . In Arabidopsis, ARP2 (Actin Related Protein) encodes ARPC2 subunit of the ARP2/3 complex . The arp2 mutant plants are less sensitive to ABA and CaCl2-induced stomatal closure and do not show actin reorganization in response to ABA. Upon addition of cytochalasin D (actin modifying agent that induces de-polymerization of actin filaments) arp2 mutant guard cells show similar ABA-mediated stomatal closure as wild type, indicating that ABA acts through ARP2/3 complex in ABA disruption or de-polymerization of actin . Recently, a plant specific actin binding protein, SCAB1, has been identified and that interacts with F-actin and plays a regulatory role in ABA-mediated stomatal response . These findings indicate the importance of actin dynamics in stomatal movements, but it appears that actin skeleton is rearranged passively.
Accumulating evidences show that actin is also an active regulator of stomatal movements. Cytochalasin D, which could depolymerize the F-actin, activates K+in channels and enhances light-induced stomatal opening. Similarly, application of actin filament stabilizer, phalloidin, inhibits of K+in currents and light-induced stomatal opening . Cytochalasin D also restores the hyperosmolarity inhibition of K+in currents , suggesting the depolymerized actin is a positive activator of K+in channels during stomatal movements. More importantly, actin depolymeirzation induced by Cytochalasin D or hypo-tonic external solution, also activates guard cell plasma membrane localized Ca2+-permeable channels (both voltage-dependent and stretch-activated Ca2+-permeable channels) and currents at both single channel and whole-cell recording levels, suggesting the actin dynamics is downstream of osmotic stress but an upstream regulator of [Ca2+]cyt signal [49,50]. Actin dynamics has also been shown to regulate both ion release from vacuole and Ca2+ entry mediated by plasma membrane localized Ca2+-permeable channels in guard cells during ABA induced stomatal closing . Altogether, these findings indicate that both [Ca2+]cyt and actin dynamics are regulated by a complex inter-regulatory loop in guard cells during stomatal movements.
4.2. Calcium and Actin in Pollens and Pollen Tubes
Ca2+ also plays important roles in pollen germination and in pollen tube growth, and the crucial effect of this ion during these processes has been described in the 1960s . With the development of advanced biological techniques, now we know much more about the role of Ca2+ signaling during pollen germination and tube growth, which includes influx of Ca2+ and other ions in pollens, and the tip-focused Ca2+ gradient (in pollen tubes, the [Ca2+]cyt ranges from 2–10 μM in the apical region, 20 μm at the tip, and 20–200 nm in the shank) and Ca2+ oscillations, apical Ca2+ entry and other osmotic ion influx through ion channels. As well, the contribution of Ca2+ in polar growth of the tube has been firmly established, and inhibition of external Ca2+ entry in pollen or pollen tube or disruption of Ca2+ gradient in the growing tube has been shown to prohibit the normal germination and elongation [8,41,130,131,132,133,134,135,136,137].
Entry of Ca2+ from outside for generation of [Ca2+]cyt is important for pollen germination, as well as for pollen tube growth and orientation [40,129,131,137,138]. By using Ca2+-selective vibrating probes, pharmacological treatments, together with electrical fields and ionophoretic microinjection method and the comparison of [Ca2+]cyt dynamic between non-growing and growing pollen tubes, it has been shown that Ca2+ channels (e.g., voltage-gated Ca2+ channels) in the plasma membrane of the pollen tubes are important for regulation of [Ca2+]cyt and tube growth [132,139,140,141]. By using patch clamp technique, stress-activated Ca2+ channels have been identified in both lily pollens and pollen tubes , and voltage-dependent Ca2+-permeable channels of pollens also have been identified in different plant species [142,143,144,145]. These reports indicate that the activation of these channels is important for [Ca2+]cyt generation . Furthermore, genes encoding Ca2+-permeable channels have been identified in pollens. The CNGC18, a plasma membrane localized cyclic nucleotide-gated channel of Arabidopsis, is expressed at the tip of the pollen tube and is essential for normal tip growth. In addition, expression of CNGC18 has been shown to accumulate Ca2+ in E. coli, indicating that CNDC18 may be a Ca2+-permeable channel . D-Serine activation of pollen Ca2+ currents, [Ca2+]cyt generation and tube growth, together with the data from Ca2+-specific vibrating probe and physiological assays by using glutamate receptor like genes, indicating that GLRs are functional Ca2+-channels important for pollen tube elongation .
Downstream of [Ca2+]cyt in pollen and pollen tube signaling, multiple Ca2+ sensors have been identified that transduce the [Ca2+]cyt signals into physiological and biochemical responses. Growing pollen tubes exhibit higher activity of the CDPK kinase in the apical region, whereas nongrowing tubes show uniform kinase activity. Moreover, the modification of the kinase activity changes the orientation of the tubes, and the increased [Ca2+]cyt enhances the kinase activity that, in turn, leads to reorientation of pollen tube . Microinjection of fluorescence labelled calmodulin into pollen tubes indicated evenly distribution of calmodulin, but the higher calmodulin activity was observed in the apex, and that was correlative with [Ca2+]cyt-concentration distribution .These data suggest that CDPKs may play important roles in pollen germination and tube growth. In Petunia, overexpression of PM localized PiCDPK1 seriously affected the polarity of pollen-tube growth and [Ca2+]cyt . CDPK24 and 32, CBL2, and CBL3 were identified by a large-scale screening of Ca2+ sensors in pollen tubes and were found to be involved in regulating pollen-tube function . CPK17 and CPK34 play important roles in Ca2+ signaling to increase the growth rate of pollen-tube tip . These works open the door to access the complex signaling network downstream of [Ca2+]cyt. However, up to now, as researchers have limiting knowledge on overall pollen expressing Ca2+ sensors, we need to go a long way to clarify and to identify the downstream components and their interacting partners of Ca2+ signaling network in pollens and pollen tubes.
Actin cytoskeleton in pollens and pollen tubes has been emphasized since long time, and has been implicated in protoplasmic streaming and vesicle trafficking, which are essential for pollen-tube tip growth [150,151]. Pollen-tube actin filaments can be categorized into three distinct patterns: longitudinal actin cables found in the pollen tube shank; dense actin structures called actin fringes or collars at the subapex; and highly dynamic, but less abundant actin filaments found at the extreme distal pollen-tube tip [152,153]. Up to now, it has been found that the alteration of actin dynamics or change of actin pattern in living cells by cytochalasin B, phalloidin, profillin (monomeric actin-binding protein), actin-binding proteins (e.g., villins and myosins), and small G proteins, results in disruption of normal pollen germination and pollen-tube growth [15,37,39,59,69,71,72,150,151,153,154,155,156,157,158,159,160,161,162,163]. In addition, multiple reports indicate association between actin pattern and Ca2+ gradient [45,161,164]. F-actin co-localizes with CDPK, and CDPK does not interact with actin directly, and actin binding proteins such as 135-kilodalton actin-bundling protein and myosin XI could be regulated by [Ca2+]cyt in regulation of actin bundling, suggesting that actin might be indirectly regulated by Ca2+ , these findings indicate that Ca2+ is an essential element in regulation of actin dynamics. On the contrary, actin also has been shown as a positive regulator of [Ca2+]cyt in pollens. In the study by Wang et al., 2004, treatment of F-actin de-polymerization reagents, cytochalasin B and D, activate Ca2+-permeable channels in the plasma membrane of Arabidopsis pollen protoplasts, and also triggers [Ca2+]cyt elevation . In addition, the authors have shown that the activation of Ca2+-channels and [Ca2+]cyt could be repressed by pretreatment of phalloidin, indicating that the regulation of Ca2+ influx through Ca2+-permeable channels is dependent on the actin dynamics and the de-polymerization status of actin is a positive regulator for generation of [Ca2+]cyt .
5. Conclusions and Perspectives
During the development and physiological response to external stimuli processes, both [Ca2+]cyt and actin network dynamics are important messengers of signaling in guard cells, pollens, and other plant cells. In this review, we briefly described individual roles of each mentioned messengers and compared the relationship of these two messengers in the two growing plant cell types: the reversible growing model cell, guard cell, and the tip polar growing model cell, pollen. Accumulating data show the existence of the inter-regulation of [Ca2+]cyt and actin in these plant cells, and the importance of interaction between them for the normal function of these cells for the plant life. But the fundamental mechanism for the interaction between these two signaling components is still largely unknown, the [Ca2+]cyt regulation of actin dynamics looks easier to understand, which is primarily dependent on the downstream Ca2+ sensors. In contrast, the actin regulation of [Ca2+]cyt via activation/deactivation of Ca2+-permeable channels is not very clear at molecular level. Is it the reason of abundance of G-actin, or the ratio of G-actin/F-actin in cytosol? In human, gelsolin is important for actin regulation of Ca2+ channels [166,167]. In plants, gelsolin, myosin XI, and the villin/gelsolin/fragmin superfamily proteins also exist, and they are important for actin dynamics [39,74,154,158,161,163,168,169], but their function in regulation of Ca2+ channel activation and [Ca2+]cyt generation still needs further investigation.
This work was supported by the Natural Science Foundation of China (31170236 and 31271506 to W.Z.) and open fund of State Key Laboratory of Plant Physiology and Biochemistry (SKLPPBKF11003 to W.Z.).
Conflicts of Interest
The authors declare no conflict of interest.
- Li, S.; Assmann, S.M.; Albert, R. Predicting essential components of signal transduction networks: A dynamic model of guard cell abscisic acid signaling. PLoS Biol. 2006, 4, e312. [Google Scholar] [CrossRef]
- Kim, T.H.; Bohmer, M.; Hu, H.; Nishimura, N.; Schroeder, J.I. Guard cell signal transduction network: Advances in understanding abscisic acid, CO2, and Ca2+ signaling. Annu. Rev. Plant Biol. 2010, 61, 561–591. [Google Scholar]
- Guan, Y.; Guo, J.; Li, H.; Yang, Z. Signaling in pollen tube growth: Crosstalk, feedback, and missing links. Mol. Plant 2013, 6, 1053–1064. [Google Scholar] [CrossRef]
- Hepler, P.K. Calcium: A central regulator of plant growth and development. Plant Cell 2005, 17, 2142–2155. [Google Scholar] [CrossRef]
- Dodd, A.N.; Kudla, J.; Sanders, D. The language of calcium signaling. Annu. Rev. Plant Biol. 2010, 61, 593–620. [Google Scholar] [CrossRef]
- Schroeder, J.I.; Thuleau, P. Ca2+ channels in higher plant cells. Plant Cell 1991, 3, 555–559. [Google Scholar]
- White, P.J. Calcium channels in higher plants. Biochim. Biophys. Acta 2000, 1465, 171–189. [Google Scholar] [CrossRef]
- Hedrich, R. Ion channels in plants. Physiol. Rev. 2012, 92, 1777–1811. [Google Scholar] [CrossRef]
- Allen, G.J.; Chu, S.P.; Harrington, C.L.; Schumacher, K.; Hoffmann, T.; Tang, Y.Y.; Grill, E.; Schroeder, J.I. A defined range of guard cell calcium oscillation parameters encodes stomatal movements. Nature 2001, 411, 1053–1057. [Google Scholar] [CrossRef]
- Day, B.; Henty, J.L.; Porter, K.J.; Staiger, C.J. The pathogen-actin connection: A platform for defense signaling in plants. Annu. Rev. Phytopathol. 2011, 49, 483–506. [Google Scholar] [CrossRef]
- Volkmann, D.; Baluška, F. Actin cytoskeleton in plants: From transport networks to signaling networks. Microsc. Res. Tech. 1999, 47, 135–154. [Google Scholar] [CrossRef]
- Henty-Ridilla, J.L.; Shimono, M.; Li, J.; Chang, J.H.; Day, B.; Staiger, C.J. The plant actin cytoskeleton responds to signals from microbe-associated molecular patterns. PLoS Pathog. 2013, 9, e1003290. [Google Scholar] [CrossRef]
- Hussey, P.J.; Ketelaar, T.; Deeks, M.J. Control of the actin cytoskeleton in plant cell growth. Annu. Rev. Plant Biol. 2006, 57, 109–125. [Google Scholar] [CrossRef]
- Blanchoin, L.; Boujemaa-Paterski, R.; Henty, J.L.; Khurana, P.; Staiger, C.J. Actin dynamics in plant cells: A team effort from multiple proteins orchestrates this very fast-paced game. Curr. Opin. Plant Biol. 2010, 13, 714–723. [Google Scholar] [CrossRef]
- Staiger, C.J.; Poulter, N.S.; Henty, J.L.; Franklin-Tong, V.E.; Blanchoin, L. Regulation of actin dynamics by actin-binding proteins in pollen. J. Exp. Bot. 2010, 61, 1969–1986. [Google Scholar] [CrossRef]
- Snowman, B.N.; Kovar, D.R.; Shevchenko, G.; Franklin-Tong, V.E.; Staiger, C.J. Signal-mediated depolymerization of actin in pollen during the self-incompatibility response. Plant Cell 2002, 14, 2613–2626. [Google Scholar] [CrossRef]
- Staiger, C.J.; Blanchoin, L. Actin dynamics: Old friends with new stories. Curr. Opin. Plant Biol. 2006, 9, 554–562. [Google Scholar] [CrossRef]
- Ahmad, A.; Zhang, Y.; Cao, X.-F. Decoding the epigenetic language of plant development. Mol. Plant 2010, 3, 719–728. [Google Scholar] [CrossRef]
- Stange, L. Plant cell differentiation. Annu. Rev. Plant Physiol. 1965, 16, 119–140. [Google Scholar] [CrossRef]
- Zhou, L.; Fu, Y.; Yang, Z. A genome-wide functional characterization of Arabidopsis regulatory calcium sensors in pollen tubes. J. Integr. Plant Biol. 2009, 51, 751–761. [Google Scholar] [CrossRef]
- Wang, P.; Song, C.P. Guard-cell signalling for hydrogen peroxide and abscisic acid. New Phytol. 2008, 178, 703–718. [Google Scholar] [CrossRef]
- Zhao, Z.; Zhang, W.; Stanley, B.A.; Assmann, S.M. Functional proteomics of Arabidopsis thaliana guard cells uncovers new stomatal signaling pathways. Plant Cell 2008, 20, 3210–3226. [Google Scholar] [CrossRef]
- Leonhardt, N.; Kwak, J.M.; Robert, N.; Waner, D.; Leonhardt, G.; Schroeder, J.I. Microarray expression analyses of arabidopsis guard cells and isolation of a recessive abscisic acid hypersensitive protein phosphatase 2C mutant. Plant Cell 2004, 16, 596–615. [Google Scholar] [CrossRef]
- Terashima, I.; Hanba, Y.T.; Tholen, D.; Niinemets, Ü. Leaf functional anatomy in relation to photosynthesis. Plant Physiol. 2011, 155, 108–116. [Google Scholar] [CrossRef]
- Gates, D.M. Transpiration and leaf temperature. Annu. Rev. Plant Physiol. 1968, 19, 211–238. [Google Scholar] [CrossRef]
- Beattie, G.A.; Lindow, S.E. The secret life of foliar bacterial pathogens on leaves. Annu. Rev. Phytopathol. 1995, 33, 145–172. [Google Scholar] [CrossRef]
- Sirichandra, C.; Wasilewska, A.; Vlad, F.; Valon, C.; Leung, J. The guard cell as a single-cell model towards understanding drought tolerance and abscisic acid action. J. Exp. Bot. 2009, 60, 1439–1463. [Google Scholar] [CrossRef]
- Schroeder, J.I.; Allen, G.J.; Hugouvieux, V.; Kwak, J.M.; Waner, D. Guard cell signal transduction. Annu. Rev. Plant Physiol. Plant Mol. Biol. 2001, 52, 627–658. [Google Scholar] [CrossRef]
- Melotto, M.; Underwood, W.; Koczan, J.; Nomura, K.; He, S.Y. Plant stomata function in innate immunity against bacterial invasion. Cell 2006, 126, 969–980. [Google Scholar] [CrossRef]
- Acharya, B.; Assmann, S. Hormone interactions in stomatal function. Plant Mol. Biol. 2009, 69, 451–462. [Google Scholar] [CrossRef]
- Pandey, S.; Zhang, W.; Assmann, S.M. Roles of ion channels and transporters in guard cell signal transduction. FEBS Lett. 2007, 581, 2325–2336. [Google Scholar] [CrossRef]
- Zhang, W.; He, S.Y.; Assmann, S.M. The plant innate immunity response in stomatal guard cells invokes g-protein-dependent ion channel regulation. Plant J. 2008, 56, 984–996. [Google Scholar] [CrossRef]
- Zhang, W. Roles of heterotrimeric g proteins in guard cell ion channel regulation. Plant Signal. Behav. 2011, 6, 986–990. [Google Scholar] [CrossRef]
- Taylor, L.P.; Hepler, P.K. Pollen germination and tube growth. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1997, 48, 461–491. [Google Scholar] [CrossRef]
- Fu, Y.; Wu, G.; Yang, Z. Rop gtpase-dependent dynamics of tip-localized f-actin controls tip growth in pollen tubes. J. Cell Biol. 2001, 152, 1019–1032. [Google Scholar] [CrossRef]
- Hill, A.E.; Shachar-Hill, B.; Skepper, J.N.; Powell, J.; Shachar-Hill, Y. An osmotic model of the growing pollen tube. PLoS One 2012, 7, e36585. [Google Scholar]
- Gu, Y.; Fu, Y.; Dowd, P.; Li, S.; Vernoud, V.; Gilroy, S.; Yang, Z. A Rho family GTPase controls actin dynamics and tip growth via two counteracting downstream pathways in pollen tubes. J. Cell Biol. 2005, 169, 127–138. [Google Scholar] [CrossRef]
- Michard, E.; Lima, P.T.; Borges, F.; Silva, A.C.; Portes, M.T.; Carvalho, J.E.; Gilliham, M.; Liu, L.H.; Obermeyer, G.; Feijo, J.A. Glutamate receptor-like genes form Ca2+ channels in pollen tubes and are regulated by pistil D-serine. Science 2011, 332, 434–437. [Google Scholar] [CrossRef]
- Zhang, H.; Qu, X.; Bao, C.; Khurana, P.; Wang, Q.; Xie, Y.; Zheng, Y.; Chen, N.; Blanchoin, L.; Staiger, C.J.; et al. Arabidopsis VILLIN5, an actin filament bundling and severing protein, is necessary for normal pollen tube growth. Plant Cell 2010, 22, 2749–2767. [Google Scholar] [CrossRef]
- Fan, L.M.; Wang, Y.F.; Wang, H.; Wu, W.H. In vitro Arabidopsis pollen germination and characterization of the inward potassium currents in Arabidopsis pollen grain protoplasts. J. Exp. Bot. 2001, 52, 1603–1614. [Google Scholar] [CrossRef]
- Wang, Y.F.; Fan, L.M.; Zhang, W.Z.; Zhang, W.; Wu, W.H. Ca2+-permeable channels in the plasma membrane of Arabidopsis pollen are regulated by actin microfilaments. Plant Physiol. 2004, 136, 3892–3904. [Google Scholar] [CrossRef]
- Klyachko, N.L.; Kulikova, A.L.; Erokhina, M.A. Plant polysome binding to the actin cytoskeleton as a target for physiological regulation. Cell Biol. Int. 2003, 27, 217–218. [Google Scholar] [CrossRef]
- Fu, Y. The actin cytoskeleton and signaling network during pollen tube tip growth. J. Integr. Plant Biol. 2010, 52, 131–137. [Google Scholar] [CrossRef]
- Kim, M.; Hepler, P.K.; Eun, S.O.; Ha, K.S.; Lee, Y. Actin filaments in mature guard cells are radially distributed and involved in stomatal movement. Plant Physiol. 1995, 109, 1077–1084. [Google Scholar]
- Kohno, T.; Shimmen, T. Accelerated sliding of pollen tube organelles along characeae actin bundles regulated by Ca2+. J. Cell Biol. 1988, 106, 1539–1543. [Google Scholar] [CrossRef]
- Gao, X.Q.; Chen, J.; Wei, P.C.; Ren, F.; Wang, X.C. Array and distribution of actin filaments in guard cells contribute to the determination of stomatal aperture. Plant Cell Rep. 2008, 27, 1655–1665. [Google Scholar] [CrossRef]
- Feijó, J.A.; Sainhas, J.; Holdaway-Clarke, T.; Cordeiro, M.S.; Kunkel, J.G.; Hepler, P.K. Cellular oscillations and the regulation of growth: The pollen tube paradigm. BioEssays 2001, 23, 86–94. [Google Scholar]
- Li, L.J.; Ren, F.; Gao, X.Q.; Wei, P.C.; Wang, X.C. The reorganization of actin filaments is required for vacuolar fusion of guard cells during stomatal opening in arabidopsis. Plant Cell Environ. 2013, 36, 484–497. [Google Scholar] [CrossRef]
- Zhang, W.; Fan, L.M. Actin dynamics regulates voltage-dependent calcium-permeable channels of the vicia faba guard cell plasma membrane. J. Integr. Plant Biol. 2009, 51, 912–921. [Google Scholar] [CrossRef]
- Zhang, W.; Fan, L.M.; Wu, W.H. Osmo-sensitive and stretch-activated calcium-permeable channels in Vicia faba guard cells are regulated by actin dynamics. Plant Physiol. 2007, 143, 1140–1151. [Google Scholar] [CrossRef]
- Elliott, D.C.; Petkoff, H.S. Measurement of cytoplasmic free calcium in plant protoplasts. Plant Sci. 1990, 67, 125–131. [Google Scholar] [CrossRef]
- Sebastiani, L.; Lindberg, S.; Vitagliano, C. Cytoplasmic free Ca2+ dynamics in single tomato (lycopersicon esculentum) protoplasts subjected to chilling temperatures. Physiol. Plant 1999, 105, 239–244. [Google Scholar] [CrossRef]
- Kader, M.A.; Lindberg, S.; Seidel, T.; Golldack, D.; Yemelyanov, V. Sodium sensing induces different changes in free cytosolic calcium concentration and pH in salt-tolerant and -sensitive rice (oryza sativa) cultivars. Physiol. Plant 2007, 130, 99–111. [Google Scholar] [CrossRef]
- Bothwell, J.H.F.; Brownlee, C.; Hetherington, A.M.; Ng, C.K.Y.; Wheeler, G.L.; McAinsh, M.R. Biolistic delivery of Ca2+ dyes into plant and algal cells. Plant J. 2006, 46, 327–335. [Google Scholar] [CrossRef]
- Swanson, S.J.; Choi, W.-G.; Chanoca, A.; Gilroy, S. In vivo imaging of Ca2+, ph, and reactive oxygen species using fluorescent probes in plants. Annu. Rev. Plant Biol. 2011, 62, 273–297. [Google Scholar] [CrossRef]
- Miyawaki, A.; Griesbeck, O.; Heim, R.; Tsien, R.Y. Dynamic and quantitative Ca2+ measurements using improved cameleons. Proc. Natl. Acad. Sci. USA 1999, 96, 2135–2140. [Google Scholar] [CrossRef]
- Pei, Z.-M.; Murata, Y.; Benning, G.; Thomine, S.; Klusener, B.; Allen, G.J.; Grill, E.; Schroeder, J.I. Calcium channels activated by hydrogen peroxide mediate abscisic acid signalling in guard cells. Nature 2000, 406, 731–734. [Google Scholar] [CrossRef]
- Fairley-Grenot, K.A.; Assmann, S.M. Permeation of Ca2+ through K+ channels in the plasma membrane of Vicia faba guard cells. J. Membr. Biol. 1992, 128, 103–113. [Google Scholar]
- Chen, C.Y.; Wong, E.I.; Vidali, L.; Estavillo, A.; Hepler, P.K.; Wu, H.-M.; Cheung, A.Y. The regulation of actin organization by actin-depolymerizing factor in elongating pollen tubes. Plant Cell 2002, 14, 2175–2190. [Google Scholar] [CrossRef]
- Piñeros, M.; Tester, M. Calcium channels in higher plant cells: Selectivity, regulation and pharmacology. J. Exp. Bot. 1997, 48, 551–577. [Google Scholar] [CrossRef]
- Jammes, F.; Hu, H.-C.; Villiers, F.; Bouten, R.; Kwak, J.M. Calcium-permeable channels in plant cells. FEBS J. 2011, 278, 4262–4276. [Google Scholar] [CrossRef]
- Swarbreck, S.; Colaco, R.; Davies, J. Plant calcium-permeable channels. Plant Physiol. 2013. [Google Scholar] [CrossRef]
- Peiter, E.; Maathuis, F.J.; Mills, L.N.; Knight, H.; Pelloux, J.; Hetherington, A.M.; Sanders, D. The vacuolar Ca2+-activated channel TPC1 regulates germination and stomatal movement. Nature 2005, 434, 404–408. [Google Scholar] [CrossRef]
- Finn, J.T.; Grunwald, M.E.; Yau, K.-W. Cyclic nucleotide-gated ion channels: An extended family with diverse functions. Ann. Rev. Physiol. 1996, 58, 395–426. [Google Scholar] [CrossRef]
- Lacombe, B.; Becker, D.; Hedrich, R.; DeSalle, R.; Hollmann, M.; Kwak, J.M.; Schroeder, J.I.; Le Novere, N.; Nam, H.G.; Spalding, E.P.; et al. The identity of plant glutamate receptors. Science 2001, 292, 1486–1487. [Google Scholar]
- Hashimoto, K.; Kudla, J. Calcium decoding mechanisms in plants. Biochimie 2011, 93, 2054–2059. [Google Scholar] [CrossRef]
- McCormack, E.; Braam, J. Calmodulins and related potential calcium sensors of Arabidopsis. New Phytol. 2003, 159, 585–598. [Google Scholar] [CrossRef]
- Cooper, J.A. Effects of cytochalasin and phalloidin on actin. J. Cell Biol. 1987, 105, 1473–1478. [Google Scholar] [CrossRef]
- Gibbon, B.C.; Kovar, D.R.; Staiger, C.J. Latrunculin b has different effects on pollen germination and tube growth. Plant Cell 1999, 11, 2349–2363. [Google Scholar]
- Eun, S.O.; Lee, Y. Stomatal opening by fusicoccin is accompanied by depolymerization of actin filaments in guard cells. Planta 2000, 210, 1014–1017. [Google Scholar] [CrossRef]
- Staiger, C.J.; Yuan, M.; Valenta, R.; Shaw, P.J.; Warn, R.M.; Lloyd, C.W. Microinjected profilin affects cytoplasmic streaming in plant cells by rapidly depolymerizing actin microfilaments. Curr. Biol. 1994, 4, 215–219. [Google Scholar] [CrossRef]
- Schmit, A.C.; Lambert, A.M. Microinjected fluorescent phalloidin in vivo reveals the F-actin dynamics and assembly in higher plant mitotic cells. Plant Cell 1990, 2, 129–138. [Google Scholar]
- Kost, B.; Spielhofer, P.; Chua, N.-H. A GFP-mouse talin fusion protein labels plant actin filamentsin vivoand visualizes the actin cytoskeleton in growing pollen tubes. Plant J. 1998, 16, 393–401. [Google Scholar] [CrossRef]
- Klahre, U.; Friederich, E.; Kost, B.; Louvard, D.; Chua, N.-H. Villin-like actin-binding proteins are expressed ubiquitously in Arabidopsis. Plant Physiol. 2000, 122, 35–48. [Google Scholar] [CrossRef]
- Wang, Y.-S.; Motes, C.M.; Mohamalawari, D.R.; Blancaflor, E.B. Green fluorescent protein fusions to Arabidopsis fimbrin 1 for spatio-temporal imaging of F-actin dynamics in roots. Cell Motil. Cytoskeleton 2004, 59, 79–93. [Google Scholar] [CrossRef]
- Schenkel, M.; Sinclair, A.; Johnstone, D.; Bewley, J.D.; Mathur, J. Visualizing the actin cytoskeleton in living plant cells using a photo-convertible mEos::FABD-mTn fluorescent fusion protein. Plant Methods 2008, 4, 21. [Google Scholar] [CrossRef]
- Vidali, L.; Rounds, C.M.; Hepler, P.K.; Bezanilla, M. Lifeact-megfp reveals a dynamic apical F-actin network in tip growing plant cells. PLoS One 2009, 4, e5744. [Google Scholar]
- Era, A.; Tominaga, M.; Ebine, K.; Awai, C.; Saito, C.; Ishizaki, K.; Yamato, K.T.; Kohchi, T.; Nakano, A.; Ueda, T. Application of lifeact reveals F-actin dynamics in Arabidopsis thaliana and the liverwort, marchantia polymorpha. Plant Cell Physiol. 2009, 50, 1041–1048. [Google Scholar] [CrossRef]
- Eun, S.-O.; Lee, Y. Actin filaments of guard cells are reorganized in response to light and abscisic acid. Plant Physiol. 1997, 115, 1491–1498. [Google Scholar]
- Hwang, J.U.; Suh, S.; Yi, H.; Kim, J.; Lee, Y. Actin filaments modulate both stomatal opening and inward K+-channel activities in guard cells of Vicia faba L. Plant Physiol. 1997, 115, 335–342. [Google Scholar]
- Cárdenas, L.; Vidali, L.; Domı́nguez, J.; Pérez, H.; Sánchez, F.; Hepler, P.K.; Quinto, C. Rearrangement of actin microfilaments in plant root hairs responding to Rhizobium etli nodulation signals. Plant Physiol. 1998, 116, 871–877. [Google Scholar] [CrossRef]
- Herrmann, A.; Felle, H.H. Tip growth in root hair cells of Sinapis alba L.: Significance of internal and external Ca2+ and pH. New Phytol. 1995, 129, 523–533. [Google Scholar] [CrossRef]
- Wightman, R.; Turner, S.R. The roles of the cytoskeleton during cellulose deposition at the secondary cell wall. Plant J. 2008, 54, 794–805. [Google Scholar] [CrossRef]
- Fukuda, H.; Kobayashi, H. Dynamic organization of the cytoskeleton during tracheary-element differentiation. Dev. Growth Differ. 1989, 31, 9–16. [Google Scholar]
- Herbette, S.; Cochard, H. Calcium is a major determinant of xylem vulnerability to cavitation. Plant Physiol. 2010, 153, 1932–1939. [Google Scholar] [CrossRef]
- De Silva, D.L.R.; Hetherington, A.M.; Mansfield, T.A. Synergism between calcium ions and abscisic acid in preventing stomatal opening. New Phytol. 1985, 100, 473–482. [Google Scholar] [CrossRef]
- Schroeder, J.I.; Hagiwara, S. Cytosolic calcium regulates ion channels in the plasma membrane of Vicia faba guard cells. Nature 1989, 338, 427–430. [Google Scholar] [CrossRef]
- Wang, X.Q.; Ullah, H.; Jones, A.M.; Assmann, S.M. G protein regulation of ion channels and abscisic acid signaling in Arabidopsis guard cells. Science 2001, 292, 2070–2072. [Google Scholar] [CrossRef]
- Fan, L.M.; Zhang, W.; Chen, J.G.; Taylor, J.P.; Jones, A.M.; Assmann, S.M. Abscisic acid regulation of guard-cell K+ and anion channels in Gβ- and RGS-deficient Arabidopsis lines. Proc. Natl. Acad. Sci. USA 2008, 105, 8476–8481. [Google Scholar]
- Allan, A.C.; Fricker, M.D.; Ward, J.L.; Beale, M.H.; Trewavas, A.J. Two transduction pathways mediate rapid effects of abscisic acid in commelina guard cells. Plant Cell 1994, 6, 1319–1328. [Google Scholar]
- Netting, A.G. pH, abscisic acid and the integration of metabolism in plants under stressed and non-stressed conditions. II. Modifications in modes of metabolism induced by variation in the tension on the water column and by stress. J. Exp. Bot. 2002, 53, 151–173. [Google Scholar] [CrossRef]
- Gonugunta, V.K.; Srivastava, N.; Puli, M.R.; Raghavendra, A.S. Nitric oxide production occurs after cytosolic alkalinization during stomatal closure induced by abscisic acid. Plant Cell Environ. 2008, 31, 1717–1724. [Google Scholar]
- Blatt, M.R. Ca2+ signalling and control of guard-cell volume in stomatal movements. Curr. Opin. Plant Biol. 2000, 3, 196–204. [Google Scholar]
- Irving, H.R.; Gehring, C.A.; Parish, R.W. Changes in cytosolic ph and calcium of guard cells precede stomatal movements. Proc. Natl. Acad. Sci. USA 1992, 89, 1790–1794. [Google Scholar] [CrossRef]
- Shimazaki, K.-I.; Kinoshita, T.; Nishimura, M. Involvement of calmodulin and calmodulin-dependent myosin light chain kinase in blue light-dependent H+ pumping by guard cell protoplasts from Vicia faba L. Plant Physiol. 1992, 99, 1416–1421. [Google Scholar] [CrossRef]
- Curvetto, N.; Darjania, L.; Delmastro, S. Effect of two camp analogs on stomatal opening in Vicia faba: Possible relationship with cytosolic calcium concentration. Plant Physiol. Biochem. 1994, 32, 365–372. [Google Scholar]
- McAinsh, M.R.; Clayton, H.; Mansfield, T.A.; Hetherington, A.M. Changes in stomatal behavior and guard cell cytosolic free calcium in response to oxidative stress. Plant Physiol. 1996, 111, 1031–1042. [Google Scholar]
- Speth, E.B.; Melotto, M.; Zhang, W.; Assmann, S.M.; He, S.Y. Crosstalk in Pathogen and Hormonal Regulation of Guard Cell Signaling. In Signal Crosstalk in Plant Stress Responses; Wiley-Blackwell: Oxford, UK, 2009; pp. 96–112. [Google Scholar]
- Ma, W.; Yoshioka, K.; Berkowitz, G.A. Cyclic nucleotide gated channels and ca-mediated signal transduction during plant innate immune response to pathogens. Plant Signal. Behav. 2007, 2, 548–550. [Google Scholar] [CrossRef]
- Cousson, A.; Vavasseur, A. Putative involvement of cytosolic Ca2+ and GTP-binding proteins in cyclic-GMP-mediated induction of stomatal opening by auxin in Commelina communis L. Planta 1998, 206, 308–314. [Google Scholar] [CrossRef]
- Ali, R.; Ma, W.; Lemtiri-Chlieh, F.; Tsaltas, D.; Leng, Q.; von Bodman, S.; Berkowitz, G.A. Death don’t have no mercy and neither does calcium: Arabidopsis cyclic nucleotide gated channel2 and innate immunity. Plant Cell 2007, 19, 1081–1095. [Google Scholar] [CrossRef]
- Qi, Z.; Verma, R.; Gehring, C.; Yamaguchi, Y.; Zhao, Y.; Ryan, C.A.; Berkowitz, G.A. Ca2+ signaling by plant Arabidopsis thaliana PEP peptides depends on AtPEPR1, a receptor with guanylyl cyclase activity, and cGMP-activated Ca2+ channels. Proc. Natl. Acad. Sci. USA 2010, 107, 21193–21198. [Google Scholar]
- Yoshioka, K.; Moeder, W.; Kang, H.G.; Kachroo, P.; Masmoudi, K.; Berkowitz, G.; Klessig, D.F. The chimeric Arabidopsis cyclic nucleotide-gated ion channel11/12 activates multiple pathogen resistance responses. Plant Cell 2006, 18, 747–763. [Google Scholar] [CrossRef]
- Leng, Q.; Mercier, R.W.; Yao, W.; Berkowitz, G.A. Cloning and first functional characterization of a plant cyclic nucleotide-gated cation channel. Plant Physiol. 1999, 121, 753–761. [Google Scholar] [CrossRef]
- Walker, R.K.; Berkowitz, G.A. Detection of reactive oxygen species downstream of cyclic nucleotide signals in plants. Methods Mol. Biol. 2013, 1016, 245–252. [Google Scholar] [CrossRef]
- Furuichi, T.; Cunningham, K.W.; Muto, S. A putative two pore channel attpc1 mediates Ca2+ flux in Arabidopsis leaf cells. Plant Cell Physiol. 2001, 42, 900–905. [Google Scholar] [CrossRef]
- Islam, M.M.; Munemasa, S.; Hossain, M.A.; Nakamura, Y.; Mori, I.C.; Murata, Y. Roles of attpc1, vacuolar two pore channel 1, in Arabidopsis stomatal closure. Plant Cell Physiol. 2010, 51, 302–311. [Google Scholar] [CrossRef]
- Rienmuller, F.; Beyhl, D.; Lautner, S.; Fromm, J.; Al-Rasheid, K.A.; Ache, P.; Farmer, E.E.; Marten, I.; Hedrich, R. Guard cell-specific calcium sensitivity of high density and activity SV/TPC1 channels. Plant Cell Physiol. 2010, 51, 1548–1554. [Google Scholar] [CrossRef]
- Ma, S.-Y.; Wu, W.-H. AtCPK23 functions in Arabidopsis responses to drought and salt stresses. Plant Mol. Biol. 2007, 65, 511–518. [Google Scholar] [CrossRef]
- Mori, I.C.; Murata, Y.; Yang, Y.; Munemasa, S.; Wang, Y.-F.; Andreoli, S.; Tiriac, H.; Alonso, J.M.; Harper, J.F.; Ecker, J.R.; et al. CDPKs CPK6 and CPK3 function in ABA regulation of guard cell S-type anion- and Ca2+-permeable channels and stomatal closure. PLoS Biol. 2006, 4, e327. [Google Scholar] [CrossRef]
- Munemasa, S.; Hossain, M.A.; Nakamura, Y.; Mori, I.C.; Murata, Y. The Arabidopsis calcium-dependent protein kinase, CPK6, functions as a positive regulator of methyl jasmonate signaling in guard cells. Plant Physiol. 2011, 155, 553–561. [Google Scholar] [CrossRef]
- Zhu, S.-Y.; Yu, X.-C.; Wang, X.-J.; Zhao, R.; Li, Y.; Fan, R.-C.; Shang, Y.; Du, S.-Y.; Wang, X.-F.; Wu, F.-Q.; et al. Two calcium-dependent protein kinases, cpk4 and cpk11, regulate abscisic acid signal transduction in arabidopsis. Plant Cell Online 2007, 19, 3019–3036. [Google Scholar] [CrossRef]
- Geiger, D.; Scherzer, S.; Mumm, P.; Marten, I.; Ache, P.; Matschi, S.; Liese, A.; Wellmann, C.; Al-Rasheid, K.A.; Grill, E.; et al. Guard cell anion channel SLAC1 is regulated by CDPK protein kinases with distinct Ca2+ affinities. Proc. Natl. Acad. Sci. USA 2010, 107, 8023–8028. [Google Scholar] [CrossRef]
- Geiger, D.; Scherzer, S.; Mumm, P.; Stange, A.; Marten, I.; Bauer, H.; Ache, P.; Matschi, S.; Liese, A.; Al-Rasheid, K.A.; et al. Activity of guard cell anion channel SLAC1 is controlled by drought-stress signaling kinase-phosphatase pair. Proc. Natl. Acad. Sci. USA 2009, 106, 21425–21430. [Google Scholar] [CrossRef]
- Franz, S.; Ehlert, B.; Liese, A.; Kurth, J.; Cazalé, A.-C.; Romeis, T. Calcium-dependent protein kinase CPK21 functions in abiotic stress response in Arabidopsis thaliana. Mol. Plant 2011, 4, 83–96. [Google Scholar] [CrossRef]
- Cheong, Y.H.; Pandey, G.K.; Grant, J.J.; Batistic, O.; Li, L.; Kim, B.-G.; Lee, S.-C.; Kudla, J.; Luan, S. Two calcineurin B-like calcium sensors, interacting with protein kinase CIPK23, regulate leaf transpiration and root potassium uptake in Arabidopsis. Plant J. 2007, 52, 223–239. [Google Scholar] [CrossRef]
- Li, J.-H.; Liu, Y.-Q.; Lü, P.; Lin, H.-F.; Bai, Y.; Wang, X.-C.; Chen, Y.-L. A signaling pathway linking nitric oxide production to heterotrimeric G protein and hydrogen peroxide regulates extracellular calmodulin induction of stomatal closure in Arabidopsis. Plant Physiol. 2009, 150, 114–124. [Google Scholar] [CrossRef]
- Chen, Y.-L.; Huang, R.; Xiao, Y.-M.; Lü, P.; Chen, J.; Wang, X.-C. Extracellular calmodulin-induced stomatal closure is mediated by heterotrimeric G protein and H2O2. Plant Physiol. 2004, 136, 4096–4103. [Google Scholar] [CrossRef]
- Delk, N.A.; Johnson, K.A.; Chowdhury, N.I.; Braam, J. CML24, regulated in expression by diverse stimuli, encodes a potential Ca2+ sensor that functions in responses to abscisic acid, daylength, and ion stress. Plant Physiol. 2005, 139, 240–253. [Google Scholar] [CrossRef]
- Magnan, F.; Ranty, B.; Charpenteau, M.; Sotta, B.; Galaud, J.-P.; Aldon, D. Mutations in AtCML9, a calmodulin-like protein from Arabidopsis thaliana, alter plant responses to abiotic stress and abscisic acid. Plant J. 2008, 56, 575–589. [Google Scholar] [CrossRef]
- Eun, S.O.; Bae, S.H.; Lee, Y. Cortical actin filaments in guard cells respond differently to abscisic acid in wild-type and abi1-1 mutant Arabidopsis. Planta 2001, 212, 466–469. [Google Scholar] [CrossRef]
- Leung, J.; Bouvier-Durand, M.; Morris, P.; Guerrier, D.; Chefdor, F.; Giraudat, J. Arabidopsis ABA response gene ABI1: Features of a calcium-modulated protein phosphatase. Science 1994, 264, 1448–1452. [Google Scholar]
- Murata, Y.; Pei, Z.-M.; Mori, I.C.; Schroeder, J. Abscisic acid activation of plasma membrane Ca2+ channels in guard cells requires cytosolic NAD(p)H and is differentially disrupted upstream and downstream of reactive oxygen species production in abi1-1 and abi2-1 protein phosphatase 2C mutants. Plant Cell 2001, 13, 2513–2523. [Google Scholar]
- Lemichez, E.; Wu, Y.; Sanchez, J.-P.; Mettouchi, A.; Mathur, J.; Chua, N.-H. Inactivation of AtRac1 by abscisic acid is essential for stomatal closure. Genes Dev. 2001, 15, 1808–1816. [Google Scholar] [CrossRef]
- Liu, K.; Luan, S. Voltage-dependent K+ channels as targets of osmosensing in guard cells. Plant Cell 1998, 10, 1957–1970. [Google Scholar]
- Jiang, K.; Sorefan, K.; Deeks, M.J.; Bevan, M.W.; Hussey, P.J.; Hetherington, A.M. The ARP2/3 complex mediates guard cell actin reorganization and stomatal movement in Arabidopsis. Plant Cell 2012, 24, 2031–2040. [Google Scholar] [CrossRef]
- Zhao, Y.; Zhao, S.; Mao, T.; Qu, X.; Cao, W.; Zhang, L.; Zhang, W.; He, L.; Li, S.; Ren, S.; et al. The plant-specific actin binding protein SCAB1 stabilizes actin filaments and regulates stomatal movement in Arabidopsis. Plant Cell 2011, 23, 2314–2330. [Google Scholar] [CrossRef]
- MacRobbie, E.A.C.; Kurup, S. Signalling mechanisms in the regulation of vacuolar ion release in guard cells. New Phytol. 2007, 175, 630–640. [Google Scholar] [CrossRef]
- Brewbaker, J.L.; Kwack, B.H. The essential role of calcium ion in pollen germination and pollen tube growth. Am. J. Bot. 1963, 50, 859–865. [Google Scholar]
- Pierson, E.S.; Miller, D.D.; Callaham, D.A.; van Aken, J.; Hackett, G.; Hepler, P.K. Tip-localized calcium entry fluctuates during pollen tube growth. Dev. Biol 1996, 174, 160–173. [Google Scholar] [CrossRef]
- Feijó, J.A.; Malhó, R.; Obermeyer, G. Ion dynamics and its possible role during in vitro pollen germination and tube growth. Protoplasma 1995, 187, 155–167. [Google Scholar] [CrossRef]
- Pierson, E.S.; Miller, D.D.; Callaham, D.A.; Shipley, A.M.; Rivers, B.A.; Cresti, M.; Hepler, P.K. Pollen tube growth is coupled to the extracellular calcium ion flux and the intracellular calcium gradient: Effect of BAPTA-type buffers and hypertonic media. Plant Cell 1994, 6, 1815–1828. [Google Scholar]
- Malho, R.; Trewavas, A.J. Localized apical increases of cytosolic free calcium control pollen tube orientation. Plant Cell 1996, 8, 1935–1949. [Google Scholar]
- Fan, L.M.; Wu, W.H.; Yang, H.Y. Identification and characterization of the inward K+ channel in the plasma membrane of Brassica pollen protoplasts. Plant Cell Physiol. 1999, 40, 859–865. [Google Scholar] [CrossRef]
- Miller, D.D.; Callaham, D.A.; Gross, D.J.; Hepler, P.K. Free Ca2+ gradient in growing pollen tubes of Lillium. J. Cell Sci. 1992, 101, 7–12. [Google Scholar]
- Schiøtt, M.; Romanowsky, S.M.; Bækgaard, L.; Jakobsen, M.K.; Palmgren, M.G.; Harper, J.F. A plant plasma membrane Ca2+ pump is required for normal pollen tube growth and fertilization. Proc. Natl. Acad. Sci. USA 2004, 101, 9502–9507. [Google Scholar]
- Jaffe, L.A.; Weisenseel, M.H.; Jaffe, L.F. Calcium accumulations within the growing tips of pollen tubes. J. Cell Biol. 1975, 67, 488–492. [Google Scholar] [CrossRef]
- Malhó, R.; Feijó, J.A.; Pais, M.S.S. Effect of electrical fields and external ionic currents on pollen-tube orientation. Sex. Plant Reprod. 1992, 5, 57–63. [Google Scholar] [CrossRef]
- Malho, R.; Read, N.D.; Trewavas, A.J.; Pais, M.S. Calcium channel activity during pollen tube growth and reorientation. Plant Cell 1995, 7, 1173–1184. [Google Scholar]
- Kühtreiber, W.M.; Jaffe, L.F. Detection of extracellular calcium gradients with a calcium-specific vibrating electrode. J. Cell Biol. 1990, 110, 1565–1573. [Google Scholar] [CrossRef]
- Wu, X.; Chen, T.; Zheng, M.; Chen, Y.; Teng, N.; Samaj, J.; Baluska, F.; Lin, J. Integrative proteomic and cytological analysis of the effects of extracellular Ca2+ influx on Pinus bungeana pollen tube development. J. Proteome Res. 2008, 7, 4299–4312. [Google Scholar] [CrossRef]
- Dutta, R.; Robinson, K.R. Identification and characterization of stretch-activated ion channels in pollen protoplasts. Plant Physiol. 2004, 135, 1398–1406. [Google Scholar] [CrossRef]
- Shang, Z.-L.; Ma, L.-G.; Zhang, H.-L.; He, R.-R.; Wang, X.-C.; Cui, S.-J.; Sun, D.-Y. Ca2+ influx into lily pollen grains through a hyperpolarization-activated Ca2+-permeable channel which can be regulated by extracellular cam. Plant Cell Physiol. 2005, 46, 598–608. [Google Scholar] [CrossRef]
- Qu, H.Y.; Shang, Z.L.; Zhang, S.L.; Liu, L.M.; Wu, J.Y. Identification of hyperpolarization-activated calcium channels in apical pollen tubes of Pyrus pyrifolia. New Phytol. 2007, 174, 524–536. [Google Scholar] [CrossRef]
- Wu, J.; Shang, Z.; Jiang, X.; Moschou, P.N.; Sun, W.; Roubelakis-Angelakis, K.A.; Zhang, S. Spermidine oxidase-derived H2O2 regulates pollen plasma membrane hyperpolarization-activated Ca2+-permeable channels and pollen tube growth. Plant J. 2010, 63, 1042–1053. [Google Scholar] [CrossRef]
- Frietsch, S.; Wang, Y.-F.; Sladek, C.; Poulsen, L.R.; Romanowsky, S.M.; Schroeder, J.I.; Harper, J.F. A cyclic nucleotide-gated channel is essential for polarized tip growth of pollen. Proc. Natl. Acad. Sci. USA 2007, 104, 14531–14536. [Google Scholar] [CrossRef]
- Moutinho, A.; Love, J.; Trewavas, A.J.; Malhó, R. Distribution of calmodulin protein and mRNA in growing pollen tubes. Sex. Plant Reprod. 1998, 11, 131–139. [Google Scholar] [CrossRef]
- Yoon, G.M.; Dowd, P.E.; Gilroy, S.; McCubbin, A.G. Calcium-dependent protein kinase isoforms in Petunia have distinct functions in pollen tube growth, including regulating polarity. Plant Cell 2006, 18, 867–878. [Google Scholar] [CrossRef]
- Myers, C.; Romanowsky, S.M.; Barron, Y.D.; Garg, S.; Azuse, C.L.; Curran, A.; Davis, R.M.; Hatton, J.; Harmon, A.C.; Harper, J.F. Calcium-dependent protein kinases regulate polarized tip growth in pollen tubes. Plant J. 2009, 59, 528–539. [Google Scholar] [CrossRef]
- Franke, W.; Herth, W.; VanDerWoude, W.; Morré, D.J. Tubular and filamentous structures in pollen tubes: Possible involvement as guide elements in protoplasmic streaming and vectorial migration of secretory vesicles. Planta 1972, 105, 317–341. [Google Scholar] [CrossRef]
- Mascarenhas, J.P.; Lafountain, J. Protoplasmic streaming, cytochalasin B, and growth of the pollen tube. Tissue Cell 1972, 4, 11–14. [Google Scholar] [CrossRef]
- Lovy-Wheeler, A.; Wilsen, K.; Baskin, T.; Hepler, P. Enhanced fixation reveals the apical cortical fringe of actin filaments as a consistent feature of the pollen tube. Planta 2005, 221, 95–104. [Google Scholar] [CrossRef]
- Gebert, M.; Dresselhaus, T.; Sprunck, S. F-actin organization and pollen tube tip growth in Arabidopsis are dependent on the gametophyte-specific armadillo repeat protein ARO1. Plant Cell 2008, 20, 2798–2814. [Google Scholar] [CrossRef]
- Zhang, Y.; Xiao, Y.; Du, F.; Cao, L.; Dong, H.; Ren, H. Arabidopsis VILLIN4 is involved in root hair growth through regulating actin organization in a Ca2+-dependent manner. New Phytol. 2011, 190, 667–682. [Google Scholar] [CrossRef]
- Zhu, L.; Zhang, Y.; Kang, E.; Xu, Q.; Wang, M.; Rui, Y.; Liu, B.; Yuan, M.; Fu, Y. MAP18 regulates the direction of pollen tube growth in Arabidopsis by modulating f-actin organization. Plant Cell 2013, 25, 851–867. [Google Scholar] [CrossRef]
- Nakayasu, T.; Yokota, E.; Shimmen, T. Purification of an actin-binding protein composed of 115-kDa polypeptide from pollen tubes of lily. Biochem. Biophys. Res. Commun. 1998, 249, 61–65. [Google Scholar] [CrossRef]
- Yokota, E.; Muto, S.; Shimmen, T. Inhibitory regulation of higher-plant myosin by Ca2+ ions. Plant Physiol. 1999, 119, 231–240. [Google Scholar] [CrossRef]
- Yokota, E.; Muto, S.; Shimmen, T. Calcium-calmodulin suppresses the filamentous actin-binding activity of a 135-kilodalton actin-bundling protein isolated from lily pollen tubes. Plant Physiol. 2000, 123, 645–654. [Google Scholar] [CrossRef]
- Yokota, E.; Takahara, K.-I.; Shimmen, T. Actin-bundling protein isolated from pollen tube of lily. Biochemical and immunocytochemical characterization. Plant Physiol. 1998, 116, 1421–1429. [Google Scholar] [CrossRef]
- Yokota, E.; Shimmen, T. The 135-kDa actin-bundling protein from lily pollen tubes arranges F-actin into bundles with uniform polarity. Planta 1999, 209, 264–266. [Google Scholar] [CrossRef]
- Yokota, E.; Tominaga, M.; Mabuchi, I.; Tsuji, Y.; Staiger, C.J.; Oiwa, K.; Shimmen, T. Plant villin, lily P-135-ABP, possesses G-actin binding activity and accelerates the polymerization and depolymerization of actin in a Ca2+-sensitive manner. Plant Cell Physiol. 2005, 46, 1690–1703. [Google Scholar] [CrossRef]
- Vidali, L.; Burkart, G.M.; Augustine, R.C.; Kerdavid, E.; Tüzel, E.; Bezanilla, M. Myosin XI is essential for tip growth in Physcomitrella patens. Plant Cell 2010, 22, 1868–1882. [Google Scholar] [CrossRef]
- Qu, X.; Zhang, H.; Xie, Y.; Wang, J.; Chen, N.; Huang, S. Arabidopsis villins promote actin turnover at pollen tube tips and facilitate the construction of actin collars. Plant Cell 2013, 25, 1803–1817. [Google Scholar] [CrossRef]
- Wang, Y.; Chen, T.; Zhang, C.; Hao, H.; Liu, P.; Zheng, M.; Baluska, F.; Samaj, J.; Lin, J. Nitric oxide modulates the influx of extracellular Ca2+ and actin filament organization during cell wall construction in Pinus bungeana pollen tubes. New Phytol. 2009, 182, 851–862. [Google Scholar] [CrossRef]
- Putnam-Evans, C.; Harmon, A.C.; Palevitz, B.A.; Fechheimer, M.; Cormier, M.J. Calcium-dependent protein kinase is localized with F-actin in plant cells. Cell Motil. Cytoskeleton 1989, 12, 12–22. [Google Scholar] [CrossRef]
- Montalbetti, N.; Li, Q.; Timpanaro, G.A.; González-Perrett, S.; Dai, X.-Q.; Chen, X.-Z.; Cantiello, H.F. Cytoskeletal regulation of calcium-permeable cation channels in the human syncytiotrophoblast: Role of gelsolin. J. Physiol. 2005, 566, 309–325. [Google Scholar] [CrossRef]
- Lader, A.S.; Kwiatkowski, D.J.; Cantiello, H.F. Role of gelsolin in the actin filament regulation of cardiac L-type calcium channels. Am. J. Physiol. Cell Physiol. 1999, 277, C1277–C1283. [Google Scholar]
- Su, H.; Wang, T.; Dong, H.; Ren, H. The villin/gelsolin/fragmin superfamily proteins in plants. J. Integr. Plant Biol. 2007, 49, 1183–1191. [Google Scholar] [CrossRef]
- Tao, Z.; Ren, H. Regulation of gelsolin to plant actin filaments and its distribution in pollen. Sci. China C Life Sci. 2003, 46, 379–388. [Google Scholar] [CrossRef]
© 2013 by the authors; licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution license (http://creativecommons.org/licenses/by/3.0/).