Abstract Changes in cytosolic Ca2+ concentration ([Ca2+]cyt) serve to transmit information in eukaryotic cells. The involvement of this second messenger in plant cell growth as well as osmotic and water relations is well established. After almost 40 years of intense research on the coding and decoding of plant Ca2+ signals, numerous proteins involved in Ca2+ action have been identified. However, we are still far from understanding the complexity of Ca2+ networks. New in vivo Ca2+ imaging techniques combined with molecular genetics allow visualization of spatio-temporal aspects of Ca2+ signalling. In parallel, cell biology together with protein biochemistry and electrophysiology are able to dissect information processing by this second messenger in space and time. Here, we focus on the time-resolved changes in cellular events upon Ca2+ signals, concentrating on the two best-studied cell types, pollen tubes and guard cells. We put their signalling networks side by side, compare them with those of other cell types, and discuss rapid signalling in the context of Ca2+ transients and oscillations to regulate ion homeostasis. Anion channel, Ca2+ gradient, Ca2+ signalling, CBL/CIPKs, CPKs, guard cell, live-cell imaging, OST1, pollen tube growth, SLAH3 Introduction Calcium ions (Ca2+) play a universal role as a second messenger in cells across kingdoms of life. Although Ca2+ signalling in plants and animals is very similar in general, plants evolved unique Ca2+ signalling components. Information from biotic as well as abiotic stimuli can be encoded and transmitted via alterations in the free cytosolic Ca2+ concentration [Ca2+]cyt, on either a single cell or a tissue level. The resting [Ca2+]cyt level from various cell types has been determined to be ~100–300 nM in most animal and plant cells. Changes in plant [Ca2+]cyt levels upon various stimuli are very diverse in space and time, with differences in amplitude and duration that can result in steady transitions as well as rapid transients lasting only seconds, or rhythmic alterations (Trewavas, 1999; Evans et al., 2001). Specific physiological outputs are postulated to be encoded by these different Ca2+ dynamics, also known as ‘Ca2+ signatures’ (Webb et al., 1996; McAinsh and Hetherington, 1998; Ng and McAinsh, 2003). In plants, it is well known that different types of environmental stresses provoke distinct stimulus- and tissue-specific Ca2+ signatures (Kilian et al., 2007 ; Zhu et al., 2013; Keinath et al., 2015; Liu et al., 2017; Yuan et al., 2017). Evidence for the Ca2+ signature hypothesis in plants was shown experimentally by imposing defined Ca2+ alterations via electric or abiotic stimuli which evoked distinct gene expression profiles (Whalley et al., 2011; Whalley and Knight, 2013; Choi et al., 2014). The generation/coding of this Ca2+ signature is thought to be facilitated by Ca2+-permeable channels allowing a [Ca2+]cyt increase, while Ca2+-ATPases and Ca2+/H+ transporters revert [Ca2+]cyt to the resting state. For a detailed description on Ca2+ translocation as well as mechanisms for Ca2+ sequestration, see recent reviews (McAinsh and Pittman, 2009; Bose et al., 2011; Spalding and Harper, 2011; Choi et al., 2017). While the plasma membrane has been primarily studied for the latter function, the contribution of various cellular compartments has recently received new attention. Ca2+ release from intracellular stores is associated with osmotically induced expansion of the guard cell (GC) cytosol, for instance (Voss et al., 2016). A comprehensive review on the contribution of organelles to plant intracellular Ca2+ signalling by Costa et al. (2018) can be found in this Special Issue. Downstream of a [Ca2+]cyt increase, processing is achieved by Ca2+-binding proteins that sense and relay the information. In Arabidopsis, >250 proteins with Ca2+-binding motifs have been identified (Day et al., 2002) and proposed to decrypt [Ca2+]cyt alterations. Typical Ca2+-decoding proteins harbour domains with ‘so-called’ EF-hand motifs that undergo conformational changes upon binding of Ca2+, thereby exposing a hydrophobic core which allows an association with other proteins in order to modulate their activity (Kawasaki et al., 1998; Hoeflich and Ikura, 2002). The major classes of Ca2+-decoding proteins are Ca2+-dependent protein kinases (CDPKs or CPKs), calmodulins (CaMs), CaM-like proteins (CaMLs or CMLs), and the CBL/CIPK system (Sheen, 1996; Luan et al., 2002; Hrabak et al., 2003; McCormack et al., 2005; Luan, 2009; DeFalco et al., 2010). The latter system consists of the Ca2+ sensors, called CBLs (calcineurin B-like proteins), and their interacting kinases called CIPKs (CBL-interacting protein kinases). The model plant Arabidopsis encodes 10 CBL Ca2+ sensors and 26 CIPKs, which are able to interact with each other to give rise to multiple CBL–CIPK combinations depending on their expression profiles and subcellular localizations (Sanyal et al., 2015; Mao et al., 2016). The CPKs comprise 34 members in Arabidopsis and are unique in their function as sensor–responder proteins exhibiting both a Ca2+-binding moiety and kinase activity. CaMs are sensor proteins unable to modify target proteins post-transcriptionally, but alter target protein function upon interaction. The common belief is that Ca2+-binding proteins convert the information encoded in the Ca2+ signature into a stimulus-specific response by specifically interacting with downstream targets. How specificity is achieved during Ca2+-mediated signal transduction is an important biological question that has been subject to intense research ever since Ca2+ was recognized as a second messenger. Different Ca2+ affinities, distinct subcellular localizations, and specific targets of the Ca2+ decoders are currently hypothesized to convey specificity within Ca2+ signalling networks (Dammann et al., 2003; Batistic et al., 2010; Curran et al., 2011; Boudsocq et al., 2012). On the cellular level, two cell types, namely pollen tubes (PTs) and GCs, have been extensively studied to understand the cellular events of information processing upon changes in [Ca2+]cyt. Easy accessibility, the fact that they are electrically isolated cells, and easy monitoring of osmotic-driven PT growth as well as quick GC movement have rendered these two cell types very popular model cell systems to study Ca2+ signalling. In this review we will mainly concentrate on the spatio-temporal aspects of Ca2+ signalling in these two cell types and compare them with those of other model cell systems such as root hairs or mesophyll cells. It is conceivable that the tubular PTs make spatial analysis of Ca2+ signalling easier than in GCs. While the evident morphological feature of tubular PTs allows for spatially defined domains, it is worth noting that Ca2+ signals in the comparably small GCs can occur locally too, and need to be considered in stomatal research in the future. Spatial differences in [Ca2+]cyt might enable Ca2+ decoders with distinct affinities to be activated locally, even in GCs. The temporal aspects of this review focus on [Ca2+]cyt changes in the time frame of seconds and minutes rather than long-term changes associated with circadian rhythms or nutrient alterations that have been discussed by experts elsewhere (Webb, 2003; Dodd et al., 2005; Malhó et al., 2006; Liu et al., 2017). The role of reactive oxygen species (ROS; i.e. H2O2) associated with Ca2+ signalling is largely omitted here but has been reviewed recently (Mazars et al., 2010; Steinhorst and Kudla, 2013; Gilroy et al., 2014; Wudick and Feijó, 2014; Murata et al., 2015; Choi et al., 2017). We mainly focus on the integration of Ca2+ signals in abscisic acid (ABA)-dependent stomatal closure and PT growth due to intense research during the last decade as knowledge on Ca2+-decoding proteins strongly increased. For detailed information about GC calcium signalling in response to other stimuli, we refer readers to Roelfsema and Hedrich (2010), (Murata et al. (2015), and Yuan et al. (2017). The impact of Ca2+ on GC metabolic changes has been recently reviewed (Santelia and Lawson, 2016) and will also not be dealt with here. We will draw special attention to the following questions: (i) when do cell type-specific Ca2+ signals occur in space and time; (ii) which proteins facilitate Ca2+ decoding; and (iii) how is this Ca2+ information channelled into a specific response? Finally, we discuss the parallels of Ca2+-dependent and independent signalling networks in GCs and PTs. Based on this comparison, new signalling components and new molecular interaction partners are proposed. Pollen tubes: a model for plant tip growth As early as the mid-20th century evidence accumulated that Ca2+ is essential for proper PT growth and germination (Brewbaker and Kwack, 1963; Brink, 1924). A Ca2+ concentration gradient within growing PTs was visualized by autoradiograms, demonstrating a >100-fold 45Ca2+ accumulation at the tip versus the PT shank (Jaffe et al., 1975). This pioneering work by Lionel Jaffe, Manfred Weissenseel, and co-workers postulated an important role for this Ca2+ gradient in PT growth. Later, [Ca2+]cyt gradients were imaged ratiometrically by using organic dyes and were manipulated by means of Ca2+-permeable channel blockers which emphasized the importance of a channel-mediated Ca2+ influx at the tip (Obermeyer and Weisenseel, 1991; Rathore et al., 1991; Holdaway-Clarke et al., 1997). The increase of [Ca2+]cyt on one side of the PT tip by locally applying the Ca2+ ionophore A23187, or Ca2+ uncaging, resulted in re-orientation of the PT growth axis towards this side, representing strong evidence for Ca2+ to steer the PT growth direction (Malho and Trewavas, 1996; Moutinho et al., 1998). Numerous reports highlight that tip [Ca2+]cyt correlates with growth velocity (Rathore et al., 1991; Pierson et al., 1996; Michard et al., 2008). Genetically encoded Ca2+ indicators are nowadays commonly used to visualize the high tip-focused [Ca2+]cyt gradient which controls growth speed and tube orientation (Iwano et al., 2004; Michard et al., 2008; Gutermuth et al., 2013). Using genetically encoded probes is an elegant way to measure the [Ca2+]cyt regime; however, this is only applicable to species that are amenable to genetic transformation. Synthetic [Ca2+]cyt reporter dyes such as Indo 1 (BAPTA-conjugates of fluorescein) or Fura-2 are also valuable tools for [Ca2+]cyt imaging after pressure injection into Lilium longiforum PTs, for instance (Obermeyer and Weisenseel, 1991; Holdaway-Clarke et al., 1997; Roy et al., 1999). This approach recently resulted in data that diminish the governing role of the Ca2+ gradient and award the tip-focused pH gradient a key role in PT growth control (Winship et al., 2017). In line with Winship et al. (2017), the cytosolic Ca2+ increase that accompanies oscillatory growth of root hairs, another model cell system for polar growth, was actually demonstrated to limit turgor-driven expansion after each burst of elongation (Monshausen et al., 2008). Ca2+-permeable channels in pollen tubes Genetic evidence indicates that members of the glutamate receptor-like channels (GLRs) (Michard et al., 2011) and cyclic nucleotide-gated channels (CNGCs) form Ca2+-permeable channels in PTs (Gao et al., 2016). According to Gao et al. (2016), who compared many GLR and CNGC knock-out mutants, the most important Ca2+-permeable channel in PTs is CNGC18. This Ca2+-permeable channel was reported to facilitate Ca2+ currents in animal cell lines activated by cyclic nucleotides (Gao et al., 2014). The current characteristics and biophysical channel properties of CNGC18 (Gao et al., 2016) resemble those of hyperpolarization-activated Ca2+-permeable channels identified in pollen grain and PT tip protoplasts via the patch-clamp technique (Shang et al., 2005; Qu et al., 2007; Wu et al., 2007). Our own voltage-clamp experiments with double-barrelled microelectrodes inserted into growing PTs demonstrated that a hyperpolarization pulse activated a [Ca2+]cyt increase at the PT tip only, indicating that Ca2+-permeable channels (either ligand gated or hyperpolarization activated) reside exclusively there or are rendered active only there (Gutermuth et al., 2018). Surprisingly, CNGC18 was shown to localize predominantly to the shank of the PT rather than the tip (Boisson-Dernier et al., 2009); however, green fluorescent protein (GFP)/yellow fluorescent protein (YFP) labelling included a tip localization (Frietsch et al., 2007; Zhou et al., 2014). The interaction with and activation of CNGC18 by pollen-expressed CPK32 was recently reported (Zhou et al., 2014). In line with CNGC18 function as a Ca2+-permeable channel (Frietsch et al., 2007; Gao et al., 2014, 2016), Zhou et al. (2014) measured small Ca2+ currents by co-expression of CPK32 with CNGC18 in Xenopus laevis oocytes. It was also shown by the same authors in yeast two-hybrid assays that CNGC18 specifically interacted with CPK32 but not with CPK34. However, CPK34 was previously shown to phosphorylate several members of the CNGC family, including CNGC18 in vitro (Curran et al., 2011). This discrepancy, and the fact that our group was not able to reproduce the activation of CNGC18 by CPK32 in oocytes, awaits pollen CPK32 signalling to be confirmed and discussed. Besides GLRs and CNGCs, the putative hyperosmolality-gated calcium-permeable channels (OSCAs) (Yuan et al., 2014), namely OSCA1.4, OSCA1.7, OSCA2.1, and OSCA2.2, are highly expressed in Arabidopsis thaliana PTs. The involvement of mechanosensitive Ca2+-permeable channels in the tip-focused [Ca2+]cyt gradient was proposed in pollen physiology early on (Feijó et al., 2001). Whether the aforementioned proteins or the plant homologue of the recently identified animal mechanosensitive Ca2+-channel PIEZO (Li et al., 2014), which is also highly expressed in pollen, contribute to Ca2+ fluxes in the male gametophyte remains to be explored. For a comprehensive review on plant Ca2+-permeable channels please refer to the review by José A. Feijó and colleagues in this Special Issue (Wudick et al., 2018). Temporal aspects of pollen tube Ca2+ signalling Rhythmic growth and Ca2+ behaviour is a perfect experimental system to study the spatio-temporal aspects of Ca2+ signalling in oscillatory tip-growing cells, such as PTs and root hairs. In both oscillatory growing cells, cross-correlation analysis revealed [Ca2+]cyt peaks lagging growth spurts, a sequence of events that might argue against a Ca2+-dependent exocytosis-driven growth mechanism (Roy et al., 1999; Sutter et al., 2012). A lag phase as high as 10–15 s for [Ca2+]cyt following the PT growth regime was attributed to technical limitations of the extracellular vibrating ion selective electrodes used. This technique was later discussed to be affected by the [Ca2+]cyt buffer capacity of the cell wall especially caused by Ca2+ binding to pectins of newly secreted cell wall material (Holdaway-Clarke et al., 1997; Messerli et al., 1999; Holdaway-Clarke and Hepler, 2003). Genetically encoded or microinjected probes to visualize [Ca2+]cyt should largely rule out this cell wall buffering artefact. However, the use of intracellular biosensors still revealed a phase shift where the rise in [Ca2+]cyt lagged behind growth by 1–4 s in oscillatory growing cells (Messerli et al., 2000; Lassig et al., 2014). In contrast, movement of exocytosis vesicles to the PT tip was shown to precede growth by 5–10 s (Parton et al., 2001; Coelho and Malhó, 2006; McKenna et al., 2009). The existence and role of plant Ca2+ microgradients is completely unexplored but could account for the discrepancy in cross-correlation analysis between [Ca2+]cyt and growth in tip growing cells. Physically targeting Ca2+ reporters to the plasma membrane could enable investigation of Ca2+ signalling in close proximity to the membrane in the future. Another effective approach already applied is using high-affinity Ca2+ probes (Nanos), which enable visualization of cellular Ca2+ dynamics which could not be resolved before, by using Yellow Cameleon YC3.6 technology (Choi et al., 2014; Waadt et al., 2017). Generally, it is assumed that Ca2+ has power over exocytosis as in animal neurotransmitter secretion (Neher and Sakaba, 2008). [Ca2+]cyt elevations within microdomains in animals is well known to control membrane-delimited processes such as growth (Clapham, 2007; Wei et al., 2012). The genetic regulation of plant cell exocytosis by the exocyst complex and Rop GTPases or their interacting proteins, to facilitate growth in PTs, has been partially unravelled (Woollard and Moore, 2008; Yang and Lavagi, 2012; Žárský et al., 2013). The exocyst subunit SEC3a interacts with specific membrane lipids at the PT tip, and its subcellular localization predicts the site of vesicle fusion (Bloch et al., 2016). ROP1 activity at the tip plasma membrane oscillates ahead of growth as well as the tip [Ca2+]cyt peaks, and might control influx of Ca2+ across the tip plasma membrane (Gu et al., 2005; Hwang et al., 2005). One of the key players in chemotactic PT guidance are the LURE proteins, cysteine-rich attractant peptides secreted from synergid cells (Okuda et al., 2009). The LURE receptors PRK6 and MDIS1 together with their co-receptors MIK1 and MIK2 serve to guide the exocytosis-co-ordinated mechanisms (ROP1 Rho GTPase signalling) for tip growth at the PT apex (Takeuchi and Higashiyama, 2016; Wang et al., 2016; Luo et al., 2017; X. Zhang et al., 2017). However, it still remains largely elusive how Ca2+ signalling regulates exocytosis-driven cell elongation (Himschoot et al., 2015) and how LURE receptors govern this process. Spatial aspects of Ca2+ decoders in pollen physiology The tip-focused Ca2+ gradient in steady or oscillatory growing PTs strongly suggests that Ca2+ signalling predominantly takes place at the apex, where up to 3 µM [Ca2+]cyt can occur (Pierson et al., 1994, 1996). We assume [Ca2+]cyt to regulate spatially a multitude of activities through Ca2+-decoding proteins at the PT tip (Konrad et al., 2011). It is thus of great importance to know the subcellular localization/distribution of Ca2+-binding and -decoding proteins to understand their physiological role in the context of a proposed function (Simeunovic et al., 2016). In the following paragraphs, we will focus on the spatial aspects and physiological role(s) of CPKs, CaMs, CMLs, and the CBL/CIPK network in pollen physiology. CPKs CPKs are disproportionally highly represented in PTs, with 12 out of 34 CPK family members being (highly) expressed, some being exclusively found in this cell type. To pin down their site of action spatially, and to generate a complete map of their subcellular localization, we expressed pollen CPKs in their native environment, the Arabidopsis PTs, by generating stable Arabidopsis Col-0 lines expressing CPKs with a C-terminal YFP fusion (Fig. 1). Generally, CPKs are proteins that lack transmembrane helices and reside in the cytoplasm unless they are post-transcriptionally modified via N-myristoylation and S-acylation (also known as palmitoylation) for membrane tethering. CPK26, 4, and 11 (from subgroup I) were localized to the cytoplasm (Fig. 1), with the latter two CPKs accumulating towards the apex of PTs. This is suggestive of a prevalent physiological role for CPK4 and CPK11 at the apex of Arabidopsis PTs. CPKs of subgroup III, namely CPK14, CPK24, and CPK32, are predicted to possess N-myristoylation and S-acylation sites and show a dual subcellular localization (Fig. 1). YFP fluorescence of CPK14, CPK32, and CPK24 was observed at the plasma membrane, especially at the shank 10–30 µm behind the tip. Additionally, however, it was also observed in the sperm cell membrane or vegetative nucleus in the case of CPK14/CPK32 or CPK24, respectively (Fig. 1). CPK16, a member of subgroup IV, is characterized by a very similar plasma membrane localization pattern to CPK14, CPK32, and CPK24, but shows only faint additional sperm cell membrane localization (Fig. 1). CPK6, CPK2, and CPK20 from subgroup I are also N-myristoylated and S-acylated. While CPK6 reveals relatively strong localization in endomembrane systems of unknown origin and in the sperm cell membrane, it additionally shows clear but dispersed plasma membrane localization towards the apex (Fig. 1). Plasma membrane localization of CPK2 and CPK20 as well as CPK17 and CPK34 at the tip (as reported by Myers et al., 2009) could be confirmed here in Arabidopsis PTs (Fig. 1). Generally, our subcellular localization studies presented here confirmed the results of our previous transient transformation approach in tobacco pollen (Gutermuth et al., 2013) and additionally revealed unprecedented differences of potential importance for Arabidopsis pollen research. N-Myristoylation and S-acylation of CPKs (and CBL discussed later) seems very important and requires a small excursion into this post-transcriptional modification at this point. In general, tethering of proteins to membranes by N-terminal S-acylation is a common (~2% of the proteome) post-transcriptional mechanism to recruit signalling components to membranes (Hemsley et al., 2013; Hemsley, 2015). These post-transcriptional modifications, in which long-chain fatty acids are attached to the N-terminus, are crucial, since N-myristoylation was demonstrated even to over-ride other targeting signals (Stael et al., 2011). While N-myristoylation is thought to be irreversible, S-acylation seems predominantly reversible (Hemsley, 2015). When glycine to alanine substitutions (G2A) and cysteine to serine substitutions within the N-terminal consensus sequence for N-myristoylation and S-acylation sites are performed, they result in a cytosolic CPK (and CBL) localization (Mehlmer et al., 2010; Stael et al., 2011; Gutermuth et al., 2013; Lu and Hrabak, 2013). Different combinations of N- and S-acyl anchors might cause the proteins to associate with distinct membrane lipid compositions, potentially bringing putative interacting partners together in distinct membrane regions. Interestingly, a detailed map of membrane lipids visualized with lipid marker proteins transiently expressed in tobacco PTs exhibited a manifestation of such distinct lipid zones (Ischebeck et al., 2011; Potocký et al., 2014). Experimental evidence for specific protein targeting (as can be seen in Fig. 1) to distinct lipid micro- or macrodomains via combinations of lipid modifications remains to be shown. PTs would be an ideal model cell to investigate this hypothesis. Fig. 1. View largeDownload slide Subcellular localization of CPKs in Arabidopsis pollen tubes. Subcellular localization of pollen-expressed CPKs in stably transformed Col-0 PTs. Expression of CPKs with a C-terminal YFP fusion in PTs was driven by the LeLat52 or AtTub4 promoter. (A) Representative epifluorescence images are presented. The corresponding CPKs are indicated in close proximity in the figure. (B) A magnification of the PT expressing CPK6:YFP is shown. Scale bar=20 µm. Fig. 1. View largeDownload slide Subcellular localization of CPKs in Arabidopsis pollen tubes. Subcellular localization of pollen-expressed CPKs in stably transformed Col-0 PTs. Expression of CPKs with a C-terminal YFP fusion in PTs was driven by the LeLat52 or AtTub4 promoter. (A) Representative epifluorescence images are presented. The corresponding CPKs are indicated in close proximity in the figure. (B) A magnification of the PT expressing CPK6:YFP is shown. Scale bar=20 µm. Among the Ca2+-binding proteins, CPKs and their function in Ca2+ decoding within PTs have been studied best (Konrad et al., 2011). One of the earliest and most severe pollen CPK phenotypes reported was the mutant with double loss of function of CPK17 and CPK34 (Myers et al., 2009). Single CPK17 and CPK34 mutants exhibit normal fertility rates, while the double mutant was reported to have a near sterile phenotype assigned to the male gametophyte. The cpk17 cpk34 PT growth phenotype is striking, but the molecular identity of CPK target proteins in vivo remains to be identified (Myers et al., 2009). To address this question, a protein interaction screen with selected CPKs (CPK1, 10, 16, and 34) was performed. By this means, a huge set of proteins, including ion channels, was identified as being targeted by CPK34 and less by CPK16, in a Ca2+-dependent manner and with different substrate affinities (Curran et al., 2011). Curran et al. (2011) elegantly showed distinct substrate preference, phosphorylation sites, and turnover rates of the four CPKs tested, pointing to the long-held assumption that the vast amount of Ca2+ decoders target discrete pollen proteins at distinct subcellular localization sites. Our group could recently identify anion transport to be governed by [Ca2+]cyt in PTs. The very closely related CPK2 and CPK20 are able to target the S-type anion channel SLAH3 (slow anion channel homologue 3) in vitro and in vivo. These two CPKs control SLAH3-mediated anion efflux exclusively at the PT tip to promote polar growth (Gutermuth et al., 2013). SLAH3 activation by CPK2 and CPK20 was demonstrated by anion current measurements in Xenopus oocytes and A. thaliana PTs, interaction studies in oocytes and growing PT tips (BiFC), as well as through FRET-FLIM (fluorescence resonance energy transfer–fluorescence lifetime imaging) in Arabidopsis protoplasts (Gutermuth et al., 2013). In our follow-up study, we have demonstrated R-type anion channels from the ALMT (Aluminum-activated Malate Transporter) family to be controlled in the same fashion. We identified CPK2, CPK20, and CPK6 to be the major Ca2+ decoders for ALMT12/13/14 and SLAH3 activation (Gutermuth et al., 2018). In a triple CPK2/20/6 mutant, we observed abolished Ca2+-dependent activation of SLAH3 and ALMT channels and in turn diminished PT growth in vitro and in vivo (Gutermuth et al., 2018). Anion efflux via SLAH3 and ALMT12/13/14 at the very tip is consistent with CPK activation by the tip-focused Ca2+ gradient with ubiquitous plasma membrane anion channel localization (Gutermuth et al., 2013, 2018). An important interplay between Ca2+ and K+ homeostasis for pollen germination and tube growth has been firmly established (Fan et al., 2001; Wang et al., 2008; Wang and Wu, 2013, 2017; Wang et al., 2015). Trans-phosphorylation of CPK24 by CPK11 was shown to be a prerequisite for the deactivation of the Shaker pollen inward rectifying K+ channel (SPIK) by CPK24 (Zhao et al., 2013). CPK24 function at the plasma membrane for SPIK regulation is compatible with our results of a dual subcellular localization of this kinase in A. thaliana PTs (Fig. 1). However, the main subcellular localization of CPK24 in the vegetative nucleus implies functions that have not been addressed to date (Fig. 1; Gutermuth et al., 2013). A cytosolic localization of CPK11 preferentially in the apex of Arabidopsis PTs (Fig. 1) along with its moderate Ca2+ sensitivity (Boudsocq et al., 2012) suggest a predominant regulation of SPIK in the apex. The subcellular localization of the two major K+ channels in pollen, SPIK and stelar K+ outward rectifier SKOR, remains to be shown and could shed light on the site and direction of K+ fluxes at the PT tip, which are still controversially discussed (Messerli et al., 1999; Robinson and Messerli, 2002; Michard et al., 2017). Differences in Ca2+ signalling networks to control SPIK seem to occur in PTs and grains, indicating specific regulatory networks at distinct developmental stages of the male gametophyte. In pollen grains or PTs, activation or inactivation of SPIK by elevated [Ca2+]cyt has been shown by patch-clamp (Obermeyer and Kolb, 1993; Zhao et al., 2013). CMLs and CaMs From the >50 CMLs and six loci encoding typical CaMs in the Arabidopsis genome (Luan et al., 2002), database analysis (Genevestigator) designates at least 20 CML (2, 3, 4, 6, 7, 8, 13, 15, 16, 17, 21, 25, 26, 28, 29, 31, 33, 34, and 39) and three CaM (2, 4, and 7) genes expressed in pollen. The physiological role of most of them is unknown. Both CMLs and CaMs, lack transmembrane domains or membrane targeting signals, which is consistent with their cytoplasmic distribution in tobacco PTs (Zhou et al., 2009). Endomembrane localization was reported for CML4/5 proteins with a potential function in vesicle transport within endomembrane systems (Ruge et al., 2016). A combination of biochemical and protoplast imaging experiments revealed AtCML30 and AtCML3 to be targeted to mitochondria and peroxisomes, respectively (Chigri et al., 2012). Recently, CML36 was shown to interact with the plasma membrane Ca2+ pump ACA8 to stimulate its activity (Astegno et al., 2017) in agreement with its CaM-binding site (Bonza et al., 2000). This is consistent with regulatory domains for autoinhibition of ACAs. Binding of calcium-bound CaM to this domain releases this inhibition, thereby activating the pump (Tidow et al., 2012). Generally, the autoinhibited Ca2+-ATPases (ACAs) are thought to provide the basis for regulation of Ca2+ homeostasis. Pharmacological experiments indicate a role for endoplasmatic reticulum- (ER) localized CPA-sensitive Ca2+-ATPases during PT growth (Iwano et al., 2009). However, a direct role for the ER to serve as a capacitor/buffer of [Ca2+]cyt was not shown as the Ca2+ concentration in the ER lumen did not reveal a difference along the PT axis. Ubiquitous plasma membrane localization of ACA9 in PTs is thought to contribute to the formation of the apical Ca2+ gradient by extruding Ca2+ behind the tip (Schiott et al., 2004). Multiple mechanisms may exist to control Ca2+ sequestration by ACAs in addition to CML interaction. Fine-tuning of the Ca2+ signature by ACA8 via CBL9/CIPK14 was demonstrated in transiently transformed Nicotiana benthamiana leaves during mechanical wounding as well as extracellular ATP application (Costa et al., 2017). Interestingly, it was found that Flg22-induced Ca2+-permeable channel activity was diminished in whole seedlings of aca8 aca10 double mutants (Frei dit Frey et al., 2012) pointing to a well-balanced mechanism of Ca2+ homeostasis feeding back to sustain Ca2+-permeable channel activity. This regulatory system for ACA regulation via CML still remains to be addressed in PTs. It would also be interesting to investigate whether pollen CaMs are able to feed back on the Ca2+ gradient, as it was reported that CaM binding to the C-terminal IQ domain of CNGCs represents a common feature for their regulation (Hua et al., 2003; DeFalco et al., 2016; Fischer et al., 2017). Reports of CaM function in PTs are still rare; however, by using fluorescein-labelled CaM microinjected in PTs, a higher CaM activity was visualized in the apical region although the protein itself had a uniform distribution (Moutinho et al., 1998; Rato et al., 2004). CML24 and CML25 localize in the cytosol, and loss-of-function mutants exhibited diminished PT growth in vitro and in vivo. Loss of CML25 function was reported to alter Ca2+-dependent K+ channel regulation and actin cable formation (Yang et al., 2014; Wang et al., 2015). CML39 and CML15 are exclusively expressed in pollen, and CML15 was demonstrated by biochemical means to act as a Ca2+ sensor there. However, their physiological roles still remain to be explored (Vanderbeld and Snedden, 2007; Ogunrinde et al., 2017). The CIPK/CBL network The role of the CIPK/CBL network in pollen Ca2+ signalling has not been investigated in detail, but it is expected to contribute to PT growth regulation, as database (Genevestigator) analysis (Hruz et al., 2008) indicates 3–5 CBLs (CBL2, 3, 5, 8, and 9) and at least 10–13 CIPKs (CIPK1, 9–15, 18–20, 23, and 24) to be expressed in pollen. Several CIPKs have been shown to be localized in the cytosol of transiently transformed tobacco PTs when expressed alone (Zhou et al., 2015). A genetic study of CBL–CIPK function in PTs points to a role for the CBL2/3–CIPK12 module in vacuolar morphology and tip growth (Steinhorst et al., 2015). The effect of CBL2/3–CIPK12 interaction at the tonoplast of PTs was shown to depend on the CIPK12 kinase activity. The impact of the CBL2/3–CIPK12 Ca2+ dependency on the observed vacuolar phenotype has not however been shown. It would be very interesting to analyse the Ca2+ dependency of the CBL/CIPK module described by Steinhorst et al. (2015) because the very fragmented vacuoles in the PTs are literally absent from the apex and localize to distal regions with resting [Ca2+]cyt. The physiological role of additional plasma membrane localization of CBL2/3 at the PT tip, as demonstrated by CBL2/3–mCherry fusion proteins (Steinhorst et al., 2015), still remains to be investigated. Two other plasma membrane tip-localized CBL proteins, CBL1 and CBL9, have been characterized to play a role in K+ homeostasis and, in turn, normal PT growth under K+-limiting conditions (Mähs et al., 2013). Surprisingly, the pollen CBL1 overexpression phenotype depended on CBL1 plasma membrane localization but was independent of its Ca2+ binding (Mähs et al., 2013). The length and morphology of cipk19-1 PTs was impaired in vitro and in vivo, and CIPK19, CIPK14, CIPK10, and CIPK12, but not CIPK11, overexpression in tobacco PTs resulted in wider tubes with very high tip [Ca2+]cyt (Zhou et al., 2015). As the activity of GC-expressed SLAH3 is under the control of the CIPK23–CBL1 complex (Maierhofer et al., 2014), this kinase could also be a putative regulator of SLAH3 in PTs, assuming a co-localization of the three components. Ca2+-(in)dependent mechanisms of guard cell motion Focused research on GC osmo-mechanics goes back to the mid-19th century, and extensive work between the 1960s and 1980s has drawn much attention to the role of ion fluxes in the osmotic-driven movement of stomata (MacRobbie, 1970; Schnabl and Raschke, 1980; Zeiger, 1983). The transported ions for osmo-control of stomatal aperture were mainly characterized to be K+, Cl–, and malate (Humble and Raschke, 1971; Van Kirk and Raschke, 1978a, b). The phytohormone ABA has been demonstrated to reduce leaf transpiration through control of stomatal pore size (Little and Eidt, 1968; Mittelheuser and Vansteve, 1969; Wright and Hiron, 1969). One of the first indications that Ca2+ plays a role in stomatal movement came from experiments in which the application of extracellular Ca2+ concentrations [Ca2+]ext affected stomatal aperture (Schwartz, 1985). Low levels of [Ca2+]ext favoured stomatal opening, while 1–10 mM [Ca2+]ext or treatment with the Ca2+ ionophore A32187 evoked stomatal closure (Willmer and Mansfield, 1969; De Silva et al., 1985; MacRobbie, 1986; Inoue and Katoh, 1987; Gilroy et al., 1990; McAinsh et al., 1990). The hypothesis of a [Ca2+]cyt rise associated with stomatal closure was substantiated when McAinsh et al. (1990) detected a gradual increase in GC [Ca2+]cyt immediately after ABA application. This increase preceded stomatal closure by ~6 min. Two types of plasma membrane anion channels termed R-type (rapid-type) and S-type (slow-type) anion channels were shown to be activated in a Ca2+-dependent manner by patch-clamp studies in the same year (Hedrich et al., 1990), strengthening the Ca2+-dependent stomatal closure hypothesis. Later, however, live-cell Ca2+ imaging demonstrated that ABA-induced stomatal closure is only associated with a rise in GC [Ca2+]cyt in <50% of the cells in Arabidopsis (Hubbard et al., 2012), and ~70% in Commelina communis or Nicotiana tabacum (McAinsh et al., 1990; Marten et al., 2007). Interestingly, the degree of these Ca2+ responses was reported to be temperature dependent in C. communis (Allan et al., 1994) and dependent on high humidity growth conditions in Arabidopsis (Hubbard et al., 2012). Even in those tobacco GCs exhibiting [Ca2+]cyt upward deflections upon ABA exposure, a delay in Ca2+ responses with respect to anion channel activation was recorded in ~50% of the cells (Marten et al., 2007). This mismatch between anion channel activity and increase in [Ca2+]cyt, together with the Vicia faba results, emphasized the existence and importance of a Ca2+-independent mechanism for stomatal closure. Activation of anion channels is the key step to initiate stomatal closure because anion efflux causes a GC depolarization, which in turn activates voltage-dependent K+ outward channels for net efflux of K+ salts. Nowadays anion channel activation has turned out to be an important readout in GC research within reverse genetic studies and signal transduction analysis (Negi et al., 2008; Vahisalu et al., 2008; Geiger et al., 2009, 2010, 2011; Meyer et al., 2010; Brandt et al., 2012, 2015; Hedrich, 2012; Roelfsema et al., 2012). Today, more than ever, research on mechanisms for ABA-dependent stomatal closure becomes increasingly important with respect to global climate changes (Trenberth et al., 2013) which negatively affect food production. ABA is referred to as a stress hormone which, among other physiological roles, prevents excessive plant water loss by triggering stomatal closure. Time-resolved analysis of the fast stomatal closure response has received little attention lately, although genetically encoded [Ca2+]cyt reporters such as R-GECO1 are available to monitor GC [Ca2+]cyt with a good dynamic range (Fig. 2). In Arabidopsis GC research, stomatal aperture phenotypes are routinely quantified 0.5–2 h after application of stimuli. The current literature is thus very limited in time-resolved analysis on the cellular and molecular events early after a stomatal closing stimulus. This is especially the case in reverse genetic studies of most stomata phenotype mutants. We think this scientific gap has to be addressed much more rigorously in the future by combining reverse genetics, electrophysiology, and Ca2+ imaging. It is of great importance because the fast stomatal closure response is usually accomplished within a few minutes (5–15 min) by various types of biotic or abiotic stresses including fungal (Koers et al., 2011), bacterial (Güzel Deger et al., 2015), or phytohormone (ABA) treatment (Levchenko et al., 2005) as tracked in individual cells. Fig. 2. View largeDownload slide Ca2+ imaging in individual guard cells with R-GECO1. p35S::NES:R-GECO1 expression in stably transformed N. tabacum epidermal peels. Ca2+ imaging displays spatio-temporal fluctuations in cytosolic Ca2+ concentrations ([Ca2+]cyt) under control conditions (1 mM CaCl2, 50 mM MES, pH 5.8). (A) Time-lapse Ca2+ imaging series (interval time=5 s) with an R-GECO1 signal intensity in false colour code. (B) Representative brightfield and fluorescence image at t=0. (C) Simultaneous kymograph (false colour) and intensity over time analysis (white line) from the guard cell displayed in (A) and (B), as indicated by the dashed square. The signal change in the upper and lower half of the kymograph represents spontaneous [Ca2+]cyt changes at different sites within the cell. The white trace represents the mean [Ca2+]cyt changes of the whole cell. Note that the mean [Ca2+]cyt changes differ from the local [Ca2+]cyt changes. Fig. 2. View largeDownload slide Ca2+ imaging in individual guard cells with R-GECO1. p35S::NES:R-GECO1 expression in stably transformed N. tabacum epidermal peels. Ca2+ imaging displays spatio-temporal fluctuations in cytosolic Ca2+ concentrations ([Ca2+]cyt) under control conditions (1 mM CaCl2, 50 mM MES, pH 5.8). (A) Time-lapse Ca2+ imaging series (interval time=5 s) with an R-GECO1 signal intensity in false colour code. (B) Representative brightfield and fluorescence image at t=0. (C) Simultaneous kymograph (false colour) and intensity over time analysis (white line) from the guard cell displayed in (A) and (B), as indicated by the dashed square. The signal change in the upper and lower half of the kymograph represents spontaneous [Ca2+]cyt changes at different sites within the cell. The white trace represents the mean [Ca2+]cyt changes of the whole cell. Note that the mean [Ca2+]cyt changes differ from the local [Ca2+]cyt changes. Electrophysiological studies on V. faba GCs demonstrated no requirement for Ca2+ elevation in ABA-induced stomatal closure responses (Levchenko et al., 2005; Hubbard et al., 2012). Nevertheless, V. faba stomata are able to close in response to [Ca2+]ext. However, we still do not know any physiological stimulus that would cause an abrupt increase in [Ca2+]ext. Interestingly, V. faba GCs seem to be less sensitive to [Ca2+]ext compared with those of Arabidopsis (Allen et al., 2000; Iwai et al., 2003). Anion current recordings in intact V. faba GCs, or protoplasts thereof, revealed a maximum anion channel activation within 1.5–2.5 min after cytosolic ABA application (Levchenko et al., 2005), which is consistent with the activation and autophosphorylation of a Ca2+-independent kinase in that time frame (Li and Assmann, 1996; Li et al., 2000; Takahashi et al., 2007). The regulatory network of ABA-dependent guard cell signalling By and large, protein (de)-phosphorylation is possibly the most widespread post-translational modification and has been proven by pharmacological (Schmidt et al., 1995), genetic, and biochemical (Geiger et al., 2009) approaches to be crucial for the ABA signal transduction pathway. Phenotypes of Arabidopsis knock-out mutants lacking ABA receptors (Ma et al., 2009; Park et al., 2009), protein kinases (Mustilli et al., 2002; Mori et al., 2006; Hubbard et al., 2012), or protein phosphatase 2C (PP2C)-type phosphatases (Leung et al., 1994, 1997; Meyer et al., 1994) uncovered the receptors and kinases as positive regulators and the PP2Cs as negative regulators in the stomatal closure response. In the absence of ABA, the PP2Cs inhibit the activity of the Ca2+-independent SnRK2 protein kinase open stomata1 (OST1/SnRK2.6) (Mustilli et al., 2002; Yoshida et al., 2006; Geiger et al., 2009; Lee et al., 2009; Umezawa et al., 2009; Vlad et al., 2009). Interestingly, OST1 lack-of-function mutants do exhibit one of the most striking wilting phenotypes reported to date, which is consistent with recent data showing that OST1 is one of the major ABA signalling components not only in the Ca2+-independent but also in the Ca2+- and CO2-dependent stomatal closure pathway (Xue et al., 2011; Acharya et al., 2013; Brandt et al., 2015). OST1 (auto)phosphorylation has been biochemically reported to occur within 1–3 min after ABA exposure. Within the same time frame, the Ca2+-independent V. faba orthologue AAPK is activated in vivo (Li and Assmann, 1996; Takahashi et al., 2007) and activation of S- and R-type anion channels in V. faba and S-type anion channels in N. tabacum can be recorded (Roelfsema et al., 2004; Levchenko et al., 2005; Marten et al., 2007). This leads to the assumption that ABA receptors, kinases, and PP2Cs are the building blocks of an ABA signalling cascade (Fujii and Zhu, 2009; Nishimura et al., 2010) to fine-tune anion channel activity for stomatal aperture control. In 2008, an Arabidopsis mutant called slac1 (S-type anion channel related 1) was described to show strongly reduced S-type anion channel currents in GCs (Negi et al., 2008; Vahisalu et al., 2008). Despite this genetic evidence, it was only the co-expression of SLAC1 with OST1 in Xenopus oocytes that confirmed SLAC1 as a functional S-type anion channel (Geiger et al., 2009). The SLAC1 N-terminus is phosphorylated by OST1 at Ser120 which results in macroscopic anion currents reminiscent of S-type anion currents in GC protoplasts (Geiger et al., 2009; Lee et al., 2009). A large quantity of reverse genetic studies within the last two decades on the model plant Arabidopsis accumulated a compelling amount of evidence for the importance of a Ca2+-dependent signalling pathway in ABA-induced stomatal closure, reviewed multiple times over the past few years (Roelfsema and Hedrich, 2010; Steinhorst and Kudla, 2013; Murata et al., 2015; Edel and Kudla, 2016). Interestingly, the Ca2+-dependent and -independent branches of the ABA signalling pathway seem to be interconnected, as the triple snrk2.2/snrk2.3/ost1 mutant was impaired in Ca2+ activation of GC S-type anion channels and, in turn, stomatal closure (Brandt et al., 2015). Several Ca2+-dependent kinases could be identified to play a role in stomatal closure upon different stimuli. CPK3, 4, 5, 6, and 11, initially found in a functional genetic screen for innate immune responses (Boudsocq et al., 2010), were later identified to regulate stomatal aperture. Single and even more pronounced double mutants of CPK4 and CPK11 as well as CPK3 and CPK6 were characterized by diminished stomatal closure phenotypes (Mori et al., 2006; Zhu et al., 2007). Impaired Ca2+-induced stomatal closure was also found in cpk10 and cpk7/8/32 mutants (Hubbard et al., 2012). Interestingly, the ABA-dependent stomatal response and flg22-induced depolarization response in mesophyll cells of a cpk3/5/6/11 quadruple mutant was wild type like (Güzel Deger et al., 2015). Nevertheless, CPK3, 5, 6 and 11 are essential for the flg22-induced ROS burst (Güzel Deger et al., 2015). In contrast, the cpk5/6/11/23 quadruple mutant exhibited complete ABA and Ca2+ insensitivity regarding stomatal closure and S-type anion channel activation (Brandt et al., 2015). This points to an important role for CPK23 in the Ca2+-dependent branch of the ABA signalling pathway in GCs. From the CIPK/CBL network, only CBL1, CIPK15, and CIPK23 have so far been described to play a role in stomatal response to ABA (Guo et al., 2002; Albrecht et al., 2003; Cheong et al., 2003, 2007). However, whereas cbl1 knock-out plants were less tolerant to salt and drought stress, cipk15 and cipk23 mutants exhibited an increased ABA sensitivity. Detailed expression studies revealed that the lack of single kinases such as CIPK23 or CPK23 deregulates the equilibrium between activating and deactivating components (kinases and phosphatases) of the ABA signalling network (Geiger et al., 2010; Maierhofer et al., 2014), basically impeding the interpretation of cipk or cpk mutant phenotype data. The ABA-dependent activation of S-type anion channels is followed by a potassium efflux from GCs. Although GC K+ channels are thought to be mainly regulated by the membrane potential, Ca2+ seems to be involved in modulation of channel activity. Thereby inward and outward rectifying potassium channels are regulated oppositely. The activity of the main GC K+ efflux channel GORK (Ache et al., 2000) was shown to be increased by CPK33 (Corratgé-Faillie et al., 2017) or CPK21 (van Kleeff et al., 2018), thus promoting stomatal closure. In contrast, the K+ uptake channel KAT1, known to be involved in stomatal opening, was shown to be negatively regulated by OST1 (Acharya et al., 2013). In response to yeast elicitor (YEL), CPK6 also seems to be important for KAT1 deactivation enabling stomatal closure (Ye et al., 2013). GC proton pumps are an additional transport system counteracting stomatal closure as the H+-ATPase AHA1 was found to be crucial for stomatal opening upon blue light (Yamauchi et al., 2016) and the Arabidopsis mutant ost2 in the AHA1 gene (resulting in constitutive activation of AHA1) exhibited an ABA-insensitive phenotype (Merlot et al., 2007). As a consequence, a tight regulation of H+-ATPase activity is needed for proper stomatal function. This was shown to take place at the large cytosolic C-terminus that acts as an autoinhibitory domain (Jahn et al., 1997). Regarding AHA2, phosphorylation of specific sites within this regulatory region either promotes or prevents activation of the H+-ATPase by binding of a 14-3-3 protein (Fuglsang et al., 1999, 2007; Kinoshita and Shimazaki, 1999; Svennelid et al., 1999). However, knowledge about the kinases involved in this regulation is very limited, but calcium seems to be a crucial factor at least for the deactivation of AHA2. The calcium-stimulated kinase PKS5 (also known as CIPK11) mediates the specific phosphorylation of Ser392, thereby preventing the 14-3-3 protein from binding and thus inhibiting H+-ATPase action (Fuglsang et al., 2007). Spatial aspects of Ca2+ decoders in guard cells Targeting of CPKs to the plasma membrane or membranes of peroxisomes, ER, or mitochondria is achieved by N-terminal N-myristoylation and S-acylation (Dammann et al., 2003; Harper and Harmon, 2005). We should note that N-terminal N-myristoylation and S-acylation of CPKs in pollen results in distinct subcellular membrane localizations (Fig. 1). Targeting of CBL/CIPK pairs seems also to depend on N-myristoylation and S-acylation of CBLs (Batistic et al., 2010). Differential N-terminal lipid anchors of CBL1, CBL4, CBL5, and CBL9 lead to a localization at the plasma membrane, while CBL4 and CBL5 can also be found in the cytoplasm and at the nucleus. On the other hand, CBL2, CBL3, CBL6, and CBL10 localize to the tonoplast, and CBL7 and CBL8 are nuclear and cytoplasmic calcium sensors (Batistic et al., 2010). Tonoplast localization seems to depend on different states of CBL S-acylation (Zhang et al., 2017). Regarding CIPK14, it was shown by bimolecular fluorescence complementation (BiFC) experiments via N. benthamiana infiltration that its localization shifts from the tonoplast to the plasma membrane depending on its interaction with CBL2 and CBL3 or CBL8, respectively (Batistic et al., 2010). Thus, it seems likely that CBLs define the subcellular localization of the bound CIPK (Batistic et al., 2010); however, BiFC experiments are prone to unspecific interactions in plants by tethering the split YFP halves irreversibly together and should be generally interpreted with caution. In addition to N-terminal acylation, it should be considered that phosphorylation of substrates might also cause a change in subcellular localization. A mutation in the phosphorylation site of two cotton Cys2/His2-type zinc-finger proteins which are targeted by AtCPK11 results in a re-localization of these two proteins from the nucleus to the cytosol (Qin et al., 2016). However, a re-distribution of proteins upon [Ca2+]cyt signals cannot be excluded for signalling components. For instance, ABA-dependent phosphorylation of the transcription factor ABF4 by CPK32 has been demonstrated (Choi et al., 2005), which is possibly achieved by ABA-dependent re-localization of CPK32 from the membrane to the nucleus (Karva, 2009). The possibility that spatial re-localization of N-myristoylated and S-acylated proteins may contribute to regulation of multiple targets at different subcellular locations has been discussed (Hurst and Hemsley, 2015). These re-distributions of proteins emphasize the importance of time-resolved imaging in Ca2+ signalling research to track components of the signalling network in space and time. Adaptability of the ABA ‘signalosome’ How Ca2+-dependent and independent kinases are integrated into the ABA signalling pathway has been intensively studied during the last few years by means of electrophysiological and biochemical techniques. In the heterologous Xenopus oocyte expression system, anion channel activation by kinases was found via co-expression of SLAC1 with OST1, CPK3, 5, 6, 21, 23, or CIPK23/CBL1 (Geiger et al., 2009, 2010; Brandt et al., 2012, 2015; Scherzer et al., 2012; Maierhofer et al., 2014). Interaction between core components of the ABA signalling network regulating anion channel activity was found independent of the presence of ABA (Fujii and Zhu, 2009; Nishimura et al., 2010). This points to a stimulus-independent pre-assembly of a multiprotein complex, previously entitled the ‘ABA signalosome’ (Nishimura et al., 2010), that would differ from the formation of a simple sequential signalling cascade in spatial as well as temporal aspects. It can be speculated that the ABA signalosome allows for fast initiation of channel activation when general repressors of ABA signalling such as phosphatases are inactivated or removed from the signalling complex upon ABA perception without the need for targeting processes. Thereby not all signalling components have to be part of the same signalosome but different compositions could co-exist depending on the tissue or stimulus. Where this signalosome/multiprotein complex is situated is still debatable since it contains plasma membrane and cytosolic components. Thus, OST1 seems to be an important downstream signal component that is tightly regulated by upstream components of the fast ABA signalling pathway (Acharya et al., 2013). ABA binding by members of the cytosolic RCAR1/PYR/PYL ABA receptor family (Ma et al., 2009; Park et al., 2009) induces a conformational change of the receptors, rendering them in an active state (Hao et al., 2011; Miyakawa et al., 2013). PYR1 and PYL1–PYL3 were found to be homodimers that monomerize after ABA binding, whereas PYL4–PYL10 are thought to be monomers in the presence or absence of ABA (Miyakawa et al., 2013). After binding of ABA to the monomeric ABA receptor, a complex is formed with group-A PP2Cs, including ABI1, ABI2, HAB1, or HAB2 (Hao et al., 2011; Soon et al., 2012). Interestingly, OST1 and the other GC-expressed SnRK2 kinases SnRK2.2 and 2.3 co-immunoprecipitate with 9 out of 14 ABA receptors and ABI1 in protein extracts of Arabidopsis independently of the presence of exogenous ABA (Nishimura et al., 2010). The crystal structure of the complex between PYL2, PP2C (HAB1), and OST1 (Soon et al., 2012) revealed that the kinase activity of OST1 is inhibited by HAB1 in the absence of ABA, through dephosphorylation of a serine in the activation loop, which blocks the catalytic cleft. Binding of ABA to PYL2 blocks PP2C and in turn releases OST1 from the complex. The following autophosphorylation of the serine in the activation loop of OST1 initiates kinase activation and phosphorylation of downstream targets (Soon et al., 2012), including transcription factors (Sirichandra et al., 2010), NADPH oxidases (Sirichandra et al., 2009), and anion channels such as ALMT12 (Imes et al., 2013) and SLAC1 (Geiger et al., 2009; Lee et al., 2009). OST1 function might be regulated additionally because it can form homo- and heteromers with SnRK2.2, SnRK2.3, OST1, and SnRK2.8 in an ABA-dependent manner. The ABA signalling cascade becomes even more complex when taking in account the fact that 113 of the 126 possible ABA–receptor–PP2C combinations are functional (Tischer et al., 2017). In addition to the PP2Cs, several PP2A-type protein phosphatase regulatory subunits were found to interact with OST1 (Waadt et al., 2015), and a reciprocal regulation between the TOR kinase, ABA receptors, and SnRK2 kinases was demonstrated (Wang et al., 2018). Thus, a tight regulation of the ABA signalosome was found, with PP2Cs having a crucial dual function. On the one hand, ABI1 was described to interact with and deactivate the kinase OST1 (Yoshida et al., 2006; Geiger et al., 2009; Lee et al., 2009; Vlad et al., 2009; Soon et al., 2012). On the other hand, PP2Cs are also able to dephosphorylate and thereby deactivate the channel directly (Maierhofer et al., 2014; Brandt et al., 2015). Among the SLAC1-activating kinases, one has to distinguish between the different kinase families regarding the influence of PP2Cs on channel activity. Similar to OST1, the CIPK23/CBL1 complex seems to be under the control of the ABA signalling pathway, as ABI2 can directly interact and dephosphorylate CIPK23 (Léran et al., 2015), thereby inhibiting its kinase activity. Interestingly, this ABA-dependent regulation of kinase activity could not be observed for CPK6 (Brandt et al., 2015), indicating that its activity depends solely on the Ca2+ concentration, which is remarkable as CPK6 possesses a high basal kinase activity even in the absence of Ca2+ (Scherzer et al., 2012). However, the activation of SLAC1 by all different kinase families can be inhibited in an ABA-dependent manner by PP2C-mediated dephosphorylation of SLAC1 (Maierhofer et al., 2014; Brandt et al., 2015). Experimental evidence indicates that CPK3, CPK6, CPK21, and CPK23 as well as CIPK23/CBL1 activate SLAC1 with diverging Ca2+ dependencies (Geiger et al., 2010; Brandt et al., 2012; Scherzer et al., 2012; Maierhofer et al., 2014). CPK3 and CPK21 are inactive in the absence of Ca2+ and reach their half-maximal activity at Ca2+ concentrations of 0.25 µM and 0.28 µM, respectively (Geiger et al., 2010; Scherzer et al., 2012). CIPK23 has a low core activity of 20% in the presence of its Ca2+-binding cofactor CBL1 or CBL9, and reaches a half-maximal activity at 1.56 µM Ca2+ (Maierhofer et al., 2014). Some of the CPK family members, such as CPK6 and CPK23, harbour only a weak Ca2+ dependency and a high core activity of 50% and 60%, respectively (Geiger et al., 2010; Scherzer et al., 2012). That is why the Arabidopsis CPK family can be divided into Ca2+-dependent and -independent subgroups (Boudsocq et al., 2012). Regarding CPK6, other studies revealed a strong Ca2+ dependency with a half maximal activity at 0.51 µM calcium (Laanemets et al., 2013). However, when comparing these different results, one has to consider that the substrate may have a strong influence on the phosphorylation properties of the kinase. This might explain the different calcium dependencies of CPK6 when phosphorylating the artificial substrate Syntide-2 (Laanemets et al., 2013) or the physiologically relevant target SLAC1 (Scherzer et al., 2012). The molecular mechanism of S-type anion channel regulation via phosphorylation is still unknown (Maierhofer et al., 2014). Putative phosphorylation sites were found in the N- and C-terminal part of SLAC1. It could be demonstrated that different kinases target different phosphorylation sites. The Ca2+-independent kinase OST1 seems to phosphorylate specifically Ser120 in the SLAC1 N-terminus (Geiger et al., 2009; Maierhofer et al., 2014; Brandt et al., 2015). In contrast to OST1, the tested Ca2+-dependent kinases CPK6, CPK23, and CIPK23/CBL1 could still activate SLAC1 when Ser120 was mutated to alanine (Maierhofer et al., 2014; Brandt et al., 2015). For the latter three kinases, mutation of the N-terminal Ser59 resulted in an inhibition or reduction of SLAC1-mediated currents, indicating that this site seems to be important for channel activation by all three kinase families (Brandt et al., 2012; Maierhofer et al., 2014). However, phosphorylation mimetic mutants of SLAC1 Ser59 and/or Ser120 failed to mediate macroscopic S-type anion currents (Maierhofer et al., 2014). In vitro kinase assays revealed that not only the N- but also the C-terminal domain can be phosphorylated (Geiger et al., 2009; Lee et al., 2009) and phosphorylation mimetic mutants of putative phosphorylation sites in the SLAC1 C-terminus uncovered SLAC1 T513D as a constitutively active channel mutant. Additional N-terminal phosphorylation mimetic mutations of Ser59 or Ser120 had no further influence on current amplitude, emphasizing the important role of Thr513 for channel activation (Maierhofer et al., 2014). In addition to SLAC1, its homologue SLAH3 is the second S-type anion channel expressed in GCs of Arabidopsis (Geiger et al., 2011; Zheng et al., 2014). In contrast to SLAC1, SLAH3 cannot be activated by kinases from the SnRK2 family, but both channels were shown to be activated by a similar set of kinases including CPKs as well as CIPK23/CBL1 in an ABA-dependent manner (Geiger et al., 2009, 2010, 2011; Scherzer et al., 2012,Maierhofer et al., 2014). Additionally, both GC-expressed S-type anion channels play a role in stomatal closure in response to pathogen attack, as Arabidopsis mutants lacking SLAC1 and SLAH3 are not able to close their stomata upon flg22 treatment (Güzel Deger et al., 2015). In contrast, the single mutants were only partially impaired (slac1) or completely unaffected (slah3). Recent studies show that SLAC1 and SLAH3 can interact with and inhibit the inward-rectifying potassium channel KAT1, known to be involved in stomatal opening (Zhang et al., 2016). Both the C-terminus of KAT1 and the N-terminus of SLAC1 are involved in this inhibitory effect, preventing the re-opening of stomata under stress conditions. This suppression of stomatal opening during stomatal closure is an important step that is supported by the finding that the activity of the H+-ATPase AHA2 is regulated by the Ca2+-dependent kinase CIPK11 (Fuglsang et al., 2007). It is thought that the ABA-triggered membrane depolarization via anion channel activity activates the outward rectifying voltage-gated K+ channel GORK, maintaining the efflux of ions from GCs to close stomata. However, recent studies revealed that GORK activity is not solely dependent on the membrane potential but is also down-regulated by components of the ABA signalosome, such as ABI2 and PP2CA (Lefoulon et al., 2016). The question of which kinase is responsible for the ABA-dependent GORK activation still has to be addressed in future studies. As OST1 failed to activate GORK (Lefoulon et al., 2016), Ca2+-dependent kinases seem to be good candidates. Just recently, S-type anion channel-activating kinase CPK21 was shown to phosphorylate GORK in vitro (van Kleeff et al., 2018). Additionally, CPK33 was found to enhance GORK currents when co-expressed in oocytes, and cpk33 GCs are impaired in Ca2+-induced stomatal closure (Corratgé-Faillie et al., 2017). Whether or not this regulation is under the control of the ABA signalling network is still unclear. In general, these findings indicate that the crosstalk between different pathways can be integrated into one signalosome governing anion and cation channels in the plasma membrane of GCs. Parallels between the Ca2+ signalosome of guard cells and pollen tubes to control anion channels: master regulators of movement and growth An apparent phytohormone-based nastic physiology of GCs seems very different from the chemotactic-driven polar growth process of PTs. However, tight control over ion channel regulation is crucial and has strong consequences for osmotic-driven motion/movement in both cell types. The parallels in ion fluxes during stomatal closure and PT growth are really striking. Large anion efflux together with reduced H+-ATPase pumping result in regulation of membrane transport in GCs and the PT apex by shifting the membrane voltage to depolarized potentials. Both anion channel and K+ channel activity are highly Ca2+ dependent, and Ca2+ influx is probably mediated by the same classes of Ca2+-permeable channels. Evidence accumulates that the mechanism for Ca2+-dependent activation of anion channels and regulation of K+ channels in GCs and PTs exhibits very similar regulatory modules. We highlight here the functional similarities within both signalling networks with respect to anion and K+ channel regulation. The scientific focus of research on the signalling networks of both cell systems in the past has been slightly different. While many interacting partners of the GC signalosome have been biochemically identified and were studied in great detail, this area in pollen research is still lagging behind. In contrast, the timely aspect of Ca2+ signalling in PTs has recently been in focus, while GC research is lagging behind in this respect. This opens up the possibility for a comparison and speculations about missing links for Ca2+ signalling and ion channel regulation between the two cell types. The molecular identity of Ca2+-permeable channels upstream of the Ca2+ signalling cascades are only currently being reported for GCs and PTs. Studies based on reverse genetics indicate CNGCs and GLRs or CNGCs and OSCAs to represent Ca2+-permeable channels in PTs and GCs, respectively (Michard et al., 2011; Wang et al., 2013; Gao et al., 2014, 2016; Yuan et al., 2014). Electrophysiological characteristics of plasma membrane Ca2+-permeable channels in PTs and GCs are very similar. In both cell systems, membrane hyperpolarization increases Ca2+ inward currents (Shang et al., 2005; Qu et al., 2007; Wu et al., 2007). These hyperpolarization-activated Ca2+-permeable channels were reported to be activated by ROS (Pei et al., 2000; Breygina et al., 2016). However, CNGC5, CNGC6, as well as OSCA1, the Ca2+-permeable channels described in GCs so far, seem not to play a role in ABA-dependent stomatal closure (Wang et al., 2013; Yuan et al., 2014) where ROS are thought to be crucial signalling molecules (Sierla et al., 2016). Therefore, mechanisms for Ca2+-permeable channel activation via the cytosolic ABA receptors in GCs or via the plasma membrane leucine repeat-rich (LRR) receptors responsible for PT guidance is still completely unknown (Fig. 3). Proteins downstream of LURE perception to trigger PT re-orientation still await identification. Mechanistic parallels to the well-described BIK1-mediated immune signalling pathway upon flagellin perception (Couto et al., 2016) or similarities to the chitin receptor complex (CERK1/LYK5) and its downstream signalling (Cao et al., 2014; Yamaguchi et al., 2017) in GCs and mesophyll cells might help to understand the molecular events upon LURE sensing in PTs. Activation of the LURE receptors PRK6 and MDIS1/MIK1 and MIK2 could result in direct signal transmission via their kinase domain, a signalling mechanism that was discussed to be responsible for chitin signalling by CERK1 (Suzuki et al., 2016). Another possibility could be that LURE receptors interact with other LRR co-receptors reported to occur in flagellin signalling. A re-localization of the LURE receptor PRK6 to the site of LURE exposure was shown (Takeuchi and Higashiyama, 2016) and it is likely that the [Ca2+]cyt gradient will change accordingly; however, this still remains to be shown. Time-resolved Ca2+ imaging experiments addressing this question would not only shed light on the Ca2+ signalling network for chemotactic guidance but would also resolve the spatio-temporal aspects of Ca2+ signalling for growth of PTs in general. Fig. 3. View largeDownload slide Comparison of ABA-induced stomatal closure and pollen tube growth signalosomes. In guard cells, ABA perception by ABA receptors (PYR/PYL/RCAR) leads to the inhibition of phosphatases from the PP2C and PP2A family that in turn alleviates the deactivation of Ca2+-independent SnRK2 kinases (SnRK2.2, SnRK2.3, and OST1), Ca2+-dependent CIPK23/CBL1, and anion channels (SLAC1, SLAH3, and ALMT12). About 50% of ABA responses are accompanied by a rise in cytosolic Ca2+ that renders CIPK23/CBL1 or CPKs (3, 6, 21, 23) active and in turn results in activation of S-type anion channels SLAH3 and SLAC1. SnRK2 kinases activate SLAC1 and the R-type anion channel ALMT12. The efflux of anions depolarizes the plasma membrane, thereby activating voltage-gated outward rectifying potassium channels (Kout) for potassium efflux. The loss of osmolytes finally initiates turgor-driven stomatal closure. At the same time, stomatal opening is prevented by the deactivation of inward rectifying potassium channels (Kin) by direct interaction with SLAC1 or SLAH3. Pollen tube growth relies on the activity of an overlapping set of anion channels (SLAH3, ALMT12, ALMT13, and ALMT14) controlled by similar Ca2+-dependent kinases (CPKs). Binding of LURE1 to its receptors (MDIS1/MIK1/MIK2 or PRK6/PRK3) probably changes anion channel activity. Whether LURE1 perception changes kinase and anion channel activity via regulation of Ca2+ channels and PP2C/PP2As remains to be investigated. The grey arrows and question marks indicate that the regulation of Ca2+ channels (CNGC/GLR in pollen tubes; CNGC/OSCA in guard cells) involved in Ca2+ increases during stomatal closure and pollen tube growth is still unresolved. Please note that the influence of reactive oxygen species (ROS) on Ca2+ signalling in both cell types is excluded from the model. Fig. 3. View largeDownload slide Comparison of ABA-induced stomatal closure and pollen tube growth signalosomes. In guard cells, ABA perception by ABA receptors (PYR/PYL/RCAR) leads to the inhibition of phosphatases from the PP2C and PP2A family that in turn alleviates the deactivation of Ca2+-independent SnRK2 kinases (SnRK2.2, SnRK2.3, and OST1), Ca2+-dependent CIPK23/CBL1, and anion channels (SLAC1, SLAH3, and ALMT12). About 50% of ABA responses are accompanied by a rise in cytosolic Ca2+ that renders CIPK23/CBL1 or CPKs (3, 6, 21, 23) active and in turn results in activation of S-type anion channels SLAH3 and SLAC1. SnRK2 kinases activate SLAC1 and the R-type anion channel ALMT12. The efflux of anions depolarizes the plasma membrane, thereby activating voltage-gated outward rectifying potassium channels (Kout) for potassium efflux. The loss of osmolytes finally initiates turgor-driven stomatal closure. At the same time, stomatal opening is prevented by the deactivation of inward rectifying potassium channels (Kin) by direct interaction with SLAC1 or SLAH3. Pollen tube growth relies on the activity of an overlapping set of anion channels (SLAH3, ALMT12, ALMT13, and ALMT14) controlled by similar Ca2+-dependent kinases (CPKs). Binding of LURE1 to its receptors (MDIS1/MIK1/MIK2 or PRK6/PRK3) probably changes anion channel activity. Whether LURE1 perception changes kinase and anion channel activity via regulation of Ca2+ channels and PP2C/PP2As remains to be investigated. The grey arrows and question marks indicate that the regulation of Ca2+ channels (CNGC/GLR in pollen tubes; CNGC/OSCA in guard cells) involved in Ca2+ increases during stomatal closure and pollen tube growth is still unresolved. Please note that the influence of reactive oxygen species (ROS) on Ca2+ signalling in both cell types is excluded from the model. The core components of the signalosome for anion channel activation in GCs are the ABA receptors (RCAR/PYR1/PYL), PP2Cs, and Ca2+-dependent and independent kinases (Fig. 3). A very tight inter-relationship between the cytosolic ABA receptors and the PP2Cs has been established, allowing the phosphatases to be defined as co-receptors (Tischer et al., 2017). Such a coupling of receptors and phosphatases in PTs has not yet been reported; however, database analysis (Genevestigator) points to the expression of some of these GC core signalosome components such as HAB1, PP2CA, ABI1, and the kinase-associated protein phosphatase KAPP, and PYL2, PYL3, PYL9, and PYR1 in male gametophytes. While CPKs and CIPKs activate the anion channels in GCs, the PP2C ABI1 was reported to decrease SLAC1 activity directly (Brandt et al., 2015) and repress the activity of anion channel-activating kinases (Geiger et al., 2009, 2010, 2011; Brandt et al., 2012, 2015; Maierhofer et al., 2014) (Fig. 3). A similar role for pollen-expressed phosphatases has not been demonstrated yet, but the aforementioned phosphatases and members of the PP2C clade D might represent good candidates (Fig. 3). Although experimental evidence validating this hypothesis is missing, it is obvious that known anion channel phosphorylation by CPKs and SnRKs has to be counteracted by phosphatases. GCs and PTs mediate anion efflux through R- and S-type anion channels, which represent positive regulators for stomatal closure and PT growth. It is striking that SLAC/SLAHs and ALMTs are similarly activated by SnRK and/or CPK kinases in both cell types (Fig. 3). Activation of anion channels in growing PTs is strictly polarized and delimited to the very tip (Gutermuth et al., 2013, 2018). This is achieved by both a polarized CPK localization and a standing tip-focused Ca2+ gradient, which has been characterized as oscillating frequently (Gutermuth et al., 2013). Spatio-temporal differences in Ca2+ signalling within GCs might lead to local ion channel regulation as Ca2+ imaging of GCs with the genetically encoded R-GECO1 visualizes spontaneous Ca2+ variations, regular oscillations, or wave-like [Ca2+]cyt dynamics at varying positions within the cells (Fig. 2; Supplementary Movie S1 at JXB online). The role of local [Ca2+]cyt dynamics for GC physiology has not yet been investigated. However, these local [Ca2+]cyt elevations should be taken into account when the physiological role of Ca2+-decoding proteins with a certain Ca2+ affinity are discussed. Furthermore, the plasma membrane itself is thought to possess spatial structural differences called nanodomains with a specific lipid composition (Zappel and Panstruga, 2008). These so-called detergent-resistant membranes (DRMs) display a clear enrichment in signalling proteins (Peskan et al., 2000; Mongrand et al., 2004; Shahollari et al., 2004) including members of the ABA signalling pathway (Demir et al., 2013). Biochemical studies using N. benthamiana infiltration revealed a localization of the highly Ca2+-dependent kinase CPK21 in DRMs and its downstream target SLAH3 in the detergent-soluble fraction (DSF) of the plasma membrane. Interestingly, a transition of SLAH3 to DRMs was found after co-expression of the channel with CPK21. Moreover, in the presence of the phosphatase ABI1, both CPK21 and SLAH3 are shifted from the DRM into the DSF, thereby loosening kinase–channel complex formation. When the ABA receptor RCAR1, ABI1, CPK21, and SLAH3 are co-expressed together, the interaction between CPK21 and SLAH3 occurs in an ABA-dependent manner (Demir et al., 2013). Thus, it is assumed that under drought or salt stress conditions, SLAH3 is transferred to specialized signalling platforms in the plasma membrane of GCs where it can be activated by CPK21. A similar scenario might occur in tip-growing cells as lipid nanodomains/DRMs were characterized and visualized histochemically at the PT (Liu et al., 2009; Moscatelli et al., 2015) and root hair apex (Ovecka et al., 2010; Zhao et al., 2015). For PTs, an interaction upon co-expression of SLAH3 with CPK2 (BiFC) in the tip of these cells could be observed, while expression of SLAH3 alone revealed ubiquitous plasma membrane localization (Gutermuth et al., 2013). Similarly to CPK21, the tip-localized CPK2 is N-myristoylated and S-acylated to determine membrane association. Whether local signal propagation within DRMs is a common mechanism for membrane-delimited Ca2+ signalling is completely unexplored in plant cells. Future studies need to address the question of whether the ABA-dependent shift of the SLAH3/CPK21-containing signalosome of GCs in lipid nanodomains coincides with the formation of Ca2+ microdomains. Ca2+ microdomains in animal cells are local Ca2+ gradients in close proximity to the membrane caused by Ca2+-permeable channel activity (Clapham, 2007; Wei et al., 2012). The underlying biophysical mechanisms for the generation of Ca2+ microdomains by Ca2+-permeable channels is the same in animal and plant cells (Naraghi and Neher, 1997; Bauer, 2001). We can thus assume that they exist in plant cells as well and that they could play a role in spatially defined local Ca2+ gradients. A prerequisite to trigger Ca2+ signalling within these Ca2+ microdomains would be the localization of Ca2+-binding proteins in close proximity to these Ca2+ hotspots. New live-cell imaging techniques with the opportunity for high spatio-temporal resolution (Godin et al., 2014; Cox, 2015) might enable research related to this topic in the future. In addition to Ca2+-dependent anion channel regulation by overlapping CPKs (Geiger et al., 2009, 2010, 2011; Scherzer et al., 2012; Gutermuth et al., 2013, 2018), K+ channels are similarly regulated in GCs and PTs. The activity of hyperpolarization-activated Kin channels in GCs (KAT1) and PTs (SPIK) depends on kinase interaction (Fig. 3). KAT1 activity can be suppressed by ABA-dependent SnRKs in GCs (Sato et al., 2009) and SPIK is deactivated by CPK11/CPK24 in a co-operative manner (Zhao et al., 2013). Furthermore, KAT1 activity can be inhibited by direct interaction with SLAH3 and SLAC1 (Zhang et al., 2016). One very important role for the signalosome to control anion channels in GCs is the regulation of the membrane potential. ABA receptor-mediated Ca2+-dependent and/or independent signalling results in anion channel activation which consequently leads to a membrane depolarization (Fig. 3). The importance for membrane depolarization in GCs lies in regulation of the depolarization-activated K+-channel GORK, which is also under the control of kinases and phosphatases of the GC signalosome (Lefoulon et al., 2016). Additionally, activity of the outward rectifying potassium channels GORK and SKOR expressed in GCs or PTs, respectively, was shown to be directly enhanced by ROS accompanying Ca2+ signalling (Garcia-Mata et al., 2010; Tran et al., 2013; Lassig et al., 2014; Breygina et al., 2016). Whether a membrane potential depolarization in PTs is mediated by SLAH3 and/or ALMTs has not yet been reported; however, it seems likely because the amplitude and direction of anion currents and fluxes would be sufficient to do so (Zonia et al., 2002; Gutermuth et al., 2013). Concluding remarks Several stomatal closure stimuli, including ABA, bacterial and fungal elicitors, result in anion channel and K+ channel regulation via phosphorylation and dephosphorylation events in GCs. This is achieved by a signalosome consisting of a temporary multiprotein complex of ABA receptors, PP2Cs, and Ca2+-dependent (CPKs and CIPKs) and independent (SnRKs) kinases to steer anion and K+ channels, described in detail within the GC and signalosome sections. In pollen physiology, only kinases have so far been identified to regulate ion channels. The mechanistic link between the recently identified LURE receptors and ion channel regulation is completely unknown to date. Due to the similarities between GC and PT Ca2+ networks for ion channel regulation, new Ca2+ signalling network hypotheses can be tested. The signalosomes of both cell types display many parallels, and we are just at the tip of the iceberg to uncover the mechanisms for ion transport regulation to control electric and osmotic adjustments for cell function. The delicate co-operation between a Ca2+-dependent and -independent signalling network is just starting to be established. Supplementary data Supplementary data are available at JXB online. Movie S1. Time-lapse imaging of [Ca2+]cyt dynamics in N. tabacum stomata with R-GECO1. Acknowledgements We would like to acknowledge the Deutsche Forschungsgemeinschaft for financial support to KR. K. (DFG KO3657/2-3). We thank Tilman Güthoff for GC Ca2+ imaging. For comments on the manuscript and English proofreading, we would like to thank Professor Dirk Becker and Dr Frances Sussmilch, respectively. References Acharya BR , Jeon BW , Zhang W , Assmann SM . 2013 . Open Stomata 1 (OST1) is limiting in abscisic acid responses of Arabidopsis guard cells . 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Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: firstname.lastname@example.org This article is published and distributed under the terms of the Oxford University Press, Standard Journals Publication Model (https://academic.oup.com/journals/pages/open_access/funder_policies/chorus/standard_publication_model)
Journal of Experimental Botany – Oxford University Press
Published: Aug 3, 2018
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