Journal of Experimental Botany

Feijó, José A; Wudick, Michael M

doi: 10.1093/jxb/ery279pmid: N/A

Ca2+-binding protein, Ca2+ channel, Ca2+ signaling, calcium, calmodulin, guard cells, mitochondria, nodulation, pollen tubes, stress tolerance Ca2+ signaling is critically important for cell and developmental biology. Despite long-standing issues still holding back the field, an increasingly large repertoire of genes and mechanisms has now been described. The unanticipated complexity revealed by genomics is giving way to a renaissance in our understanding, and the characterization of novel molecular mechanisms. The reviews in this special issue bring together research focused on specific structures, including mitochondria, pollen tubes and guard cells, as well as on the processes of ion homeostasis and salt stress tolerance, and nodulation. Back in 1995, a British scientist was driving past a research institute in the south of France when his attention was caught by an unusual road sign: ‘Le Calcium C’est La Vie’. A bizarre sight for any driver, a picture of it swiftly moved into the talks given by Anthony Trewavas, a leading Ca2+ signaling researcher, to signify the importance and relevance of calcium (Ca2+) as the most versatile signaling second messenger, involved in practically all aspects of cell and developmental biology from egg activation to cell apoptosis. Eventually the French pronouncement made it into the title of an essay about the nature and mechanisms behind Ca2+ waves in plants (Trewavas, 1999). The present special issue takes on the symbolic urgency of this road sign to highlight the centrality of Ca2+ signaling in practically every scenario that can be classed as ‘experimental botany’. Key considerations for the reviews are novelty and the simultaneous need to address long-standing issues still holding back the field. For example, we lack fundamental knowledge on the apparent absence of any ligand-operated Ca2+ storage system, we are only just beginning to reveal the molecular identity of Ca2+ channels, and strong disagreement still reigns about Ca2+ channel gating and regulation. Despite this, an increasingly large repertoire of genes and mechanisms has been described in recent years, and there is a growing body of researchers contributing breakthroughs on many fronts, from the identification of bona fide Ca2+ channels in plants to the definition of putative Ca2+-signaling networks. Gene discovery, unanticipated complexity Back in ‘Le Calcium C’est La Vie’ days, the feeling of excitement was similar. A number of Ca2+-binding proteins, putative transducers of the basic Ca2+ signals, were discovered by a combination of biochemistry and the first genetic screens, which were designed for the most essential aspects of plant biology. These revealed gene/protein families specific to plants, like the Calcium Dependent Protein Kinases (CPKs or CDPKs) and the CBL–CIPK (Calcineurin B-like protein and CBL-Interacting Protein Kinase) pairs. Some ‘usual suspects’, such as animal homologs of calmodulin and the CMLs (Calmodulin-Like proteins) were also confirmed as playing important roles (reviews in Harper et al., 2004; Hepler, 2005). The field went ahead quickly on the basis of what looked like the roadmap for a true Ca2+ signature and signaling paradigm in plants, as was occurring in the animal field. The advent of genomics, and the consequent reverse genetics approaches, brought tremendous speed to the gene discovery process, but rather than confirming a paradigm this revealed a great deal of unanticipated complexity, with members of most Ca2+-signaling protein families running into the dozens. Genomics also brought about a need to revise many pharmacological approaches due to the absence of homologs to the mammalian genes in light of which those assays were designed and interpreted. And there was the conundrum of what were the Ca2+ channels in plants, as no obvious family emerged from the Arabidopsis genome and multiple forward genetics screens over a decade or so failed to bring consensus about their genetic identity. Plants do it differently There is now something of a renaissance in our understanding of many of these issues: there is some agreement about at least five families of Ca2+-permeable channels (Swarbreck et al., 2013) and the involvement of differently coded Ca2+ signaling in various aspects of plant physiology seems beyond doubt (Dodd et al., 2010; Edel and Kudla, 2015). A recent analysis of the evolutionary trends of Ca2+ signaling in plants (Edel et al., 2017) focused on the fact that, when compared to animals, the available repertoire of genes coding for Ca2+-influx mechanisms in plants is reduced, and therefore the available machinery must shoulder a greater burden in terms of fulfilling the same signaling functions. The authors elaborate that this limitation on channel diversity is compensated by larger and more-diverse families of Ca2+-binding signaling proteins capable of contributing to the amplification and integration of the primary Ca2+ signals. These are provocative conclusions that may be falsified if new families of channels are found, but suggest, as perhaps the most reasonable explanation for present findings, that plants ‘do it differently’. So although the animal paradigms served us well in searching for conservation of function, the time is ripe to assume that (i) even when the same molecular mechanisms are present, they may result from convergent evolution with adaptation to the very different contexts of plant physiology, and thus (ii) the same function may be achieved through different associations and regulatory mechanisms. Box 1 brings together the key elements of the novel molecular mechanisms described in the reviews in this special issue. Channels and stores Of all the gene families documented as coding for Ca2+-permeable channels, the ones for which there are more data available are the Glutamate Receptor-Like (GLRs) and the Cyclic Nucleotide Gated channels (CNGCs). GLRs made it to center stage directly from their genomic identification during the assembly of the Arabidopsis genome (Lam et al., 1998). This is not surprising as there was little expectation of their existence in organisms without an organized nervous system. In Arabidopsis, the family has twenty genes divided into three clades and high functional redundancy, making this family more numerous than its homolog in our own human nervous system. A decade of primary screens allowed some advances in defining their physiological roles (reviews in Davenport, 2002; Konrad et al., 2011; Forde and Roberts, 2014). Multiple functions have been attributed to GLRs, but the field was shaken by the demonstration that they may be involved in the conductance of long-range electrical signaling in response both to herbivore (Mousavi et al., 2013) and aphid (Vincent et al., 2017) feeding. Wudick et al. (2018) take a different perspective, and rather focus on the point that given the current uncertainties on regulation by oligomerization, ligand gating, ion specificity and association with other proteins, data from this kind of screening will always be difficult to interpret in terms of channel function. Further structural and evolutionary arguments are raised to make the case that elucidation of the molecular properties of these channels is needed for full understanding of their biological function, as GLRs stand as a good example of the limitations inherent to strictly translating mammalian knowledge of function and regulation. Equally with 20 gene copies, but contrary to GLRs, some single mutant CNGCs seem highly unique in their phenotypes. CNGC18 was one of the first to be characterized (Frietsch et al., 2007), with its single mutation resulting in an extremely strong pollen tube/reproductive phenotype. Other members show similarly strong phenotypes from single mutations, which is remarkable given the multitude of members, for which one would expect a high degree of redundancy. Another puzzling fact is our lack of knowledge on the pathways for synthesis and degradation of any type of cyclic nucleotides in plants. Yet, of relevance, CNGC15 was found to be essential for the generation of Ca2+ signatures in the nuclei of Medicago root cells during Rhizobium infection (Charpentier et al., 2016). This was the last and most elusive member of the cascade of proteins involved in the propagation of Ca2+ signals triggered by Nod factors along the root hair, where the nodulation transcriptional program is triggered in the nucleus upon a specific number of Ca2+ elevations. Nuclear Ca2+ oscillations have been known for a long time (e.g. Pauly et al., 2000), and in this issue Charpentier (2018) contextualizes the nodulation signal based on all the reported nuclear Ca2+ signaling phenomena described in plants. The nodulation case study is then used as a template to discuss the origin of nuclear signals in diverse contexts and the mechanisms of downstream transcriptional regulation, clearly suggesting a role for the nuclear envelope as an important Ca2+ store capable of generating specific transcriptional triggering signatures, namely through CNGCs. The whole issue of Ca2+ stores is taken to a new level in the review by Costa et al. (2018). These authors bring together what we know about the main intracellular Ca2+ stores: the vacuole, endoplasmic reticulum, Golgi, peroxisomes, apoplast, and the double membrane organelles, the mitochondria and plastids. Special attention is given to the latter two, as the authors have been at the forefront of the molecular characterization of the channels involved in Ca2+ transport from mitochondria and plastids. Some GLRs (3.4 and 3.5) have distinct peptide signals that target these organelles, and the authors were pioneers in showing that to be the case and so implicating them in Ca2+ homeostasis (Teardo et al., 2015). More profoundly the team has been at the forefront in characterizing the mitochondrial channel uniporter (MCU) in plants (Teardo et al., 2017). These transporters were long sought, their existence implied by a number of mitochondrial Ca2+ pathologies, and first demonstrated by Rizzuto’s team (De Stefani et al., 2011). Given their importance for cytosolic Ca2+ homeostasis in mammalian cells, their discovery in plants bears promise of equally relevant functions. Besides thorough coverage of the molecular mechanisms operating in all these organelles and how they make functional Ca2+ stores, Costa et al. (2018) also offer arguably the most extensive and comprehensive published account of Ca2+-imaging sensors (and methods for each), with critical comparisons from the leading group in the world in this area. Codes, networks and stress The hallmark of Ca2+ signaling is the formation of unique spatial and temporal patterns of cytosolic concentration changes that carry specific information. These are collectively known as Ca2+ signatures, and include oscillations, elevations, standing waves and, more rarely, standing gradients. The holy grail of the field is to know exactly how these patterns encode information, and how specific proteins that bind Ca2+ with different affinities and kinetics are able to decode them, resulting in specific modifications (e.g. phosphorylation/de-phosphorylation) of other downstream proteins. Konrad et al. (2018) focus on two systems with Ca2+ oscillation either on a standing gradient (the pollen tube) or spatially distributed (guard cells/stomata) to infer common patterns and different properties that could help explain the network of interactions, feedback loops and pattern-generation mechanisms. Both systems have been extensively used for Ca2+-signaling research, but the meaning of their Ca2+ signatures remains elusive. Pollen tubes possess arguably the most robust and conspicuous standing Ca2+ gradients of any cell at their growing tip, and when germinated in vitro display oscillations in many species. However, this is not always the case, and there are no sound data showing that they exist in vivo (Damineli et al., 2017). Guard cells, on the other hand, stand together with nodulation as one of the two examples where a certain number of elevations have been shown and suggested to have a physiological function, in this case the closure of the stomata (Allen et al., 2000). Konrad et al. (2018) cover all the known families of Ca2+-binding proteins, but with a bias for the CPKs, the area in which the authors have contributed most significantly. Some original data are presented on Ca2+ dynamics during fast stomata closure. A comparison between the ionic regulation of these two systems has been published before (Michard et al., 2017), the originality here being the greater molecular detail and definition of a set of behaviors collectively designated ‘signalosomes’. Comparison of the signalosomes is used to establish correlations between genetics, spatial and temporal patterns, and biochemistry; these are then built into a comparative model that suggests that pollen tubes and stomata seem to operate through the same sort of functional units to generate the two macroscopic outputs of these cells, growth and closure, respectively. These kind of parallels are useful as a narrative and to inspire experiments to test the underlying hypotheses in terms of temporal delays, which can be measured with ever-increasing efficiency as new probes become available (see Costa et al., 2018, for probe choice) and as the group has recently shown (Guttermuth et al., 2018). Concluding the issue, Manishankar et al. (2018) review the very competitive field of Ca2+ signaling during salt stress. Salt stress is simultaneously one of the most profound abiotic stress problems and one of the most successful stories in which non-biased genetic screens have led to the discovery of completely unsuspected and original molecular mechanisms in plants. The first such mutants were of the class SOS (salt overly sensitive; Liu and Zhu, 1997) and gave rise to one of the most dynamic fronts of research on Ca2+ decoding, involving the CBL–CIPK sensor (Kudla et al., 1999). This sensor arguably constituted the first identified pathway for ion homeostasis in plants and is triggered by Ca2+ binding giving rise to numerous and eloquent reviews on the subject (e.g. Edel and Kudla, 2015). The huge number of possible combinations between the members of the two families (10 CBLs×26 CIPKs in Arabidopsis) constitutes a formidable challenge such that all combinations are tested under specific screens. Nevertheless, the prospect that some of these combinations might bear the right kinetics and affinities to make them ‘the’ specific sensor for a certain Ca2+ signature is tantalizing. Manishankar et al. (2018) cover the abundant literature that relates to specific CBL–CIPKs as being associated with specific kinds of salt stress responses, namely for potassium, nitrogen molecules, magnesium, metals and anions, and argue that CBL–CIPKs have a ‘…coordinated role for Ca2+ signaling in plant nutrition’. As with the review by Konrad et al. (2018), the core of the system consists of the phosphorylation of specific ion channels that in return affect Ca2+ concentration, providing the feedback loop for Ca2+ binding to the kinase or kinase complex, respectively. Conclusion The representation of novel molecular mechanisms provided in Box 1 highlights how much progress the Ca2+-signaling field is experiencing. In addition, it shows the fragmentation that has occurred into each specialist area, which calls for a more systems-oriented perspective to integrate these different parts. The reviews in this issue provide challenging perspectives on ways to reach this goal, but achieving it would lay the ground for the next steps where the formation of waves and the decoding of specific signatures still lack defined molecular mechanisms. Box 1. Ca2+ signaling in the plant cell Unified representation of Ca2+ signaling in the plant cell, with different types of organization color coded by quadrant of the ‘textbook’ diagram. Moving clockwise: (i) structures—mitochondria (Costa et al., 2018), pollen tubes (Wudick et al., 2018), and guard cells (Konrad et al., 2018); (ii) processes —ion homeostasis and salt stress tolerance (Manishankar et al., 2018), and nodulation (Charpentier, 2018). Acknowledgments The reviews in this issue followed from interactions at a symposium on Ca2+ channels and signaling organized during the International Botanical Congress (IBC) in Shenzhen, China, in 2017, with support from Journal of Experimental Botany and the New Phytologist Trust. Work in the author’s lab is supported by the US National Science Foundation (MCB 1616437/2016 and MCB 1714993/2017) and the University of Maryland. References Allen GJ , Chu SP, Schumacher K, et al. 2000 . Alteration of stimulus-specific guard cell calcium oscillations and stomatal closing in Arabidopsis det3 mutant . Science 289 , 2338 – 2342 . Google Scholar Crossref Search ADS PubMed WorldCat Charpentier M . 2018 . 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This is an Open Access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted reuse, distribution, and reproduction in any medium, provided the original work is properly cited. © The Author(s) 2018. Published by Oxford University Press on behalf of the Society for Experimental Biology.

doi: 10.1093/jxb/ery153pmid: 29684179

Abstract Animal ionotropic glutamate receptors (iGluRs) are ligand-gated channels whose evolution is intimately linked to that of the nervous system, where the agonist glutamate and co-agonists glycine/d-serine act as neurotransmitters or neuromodulators. While iGluRs are specialized in neuronal communication, plant glutamate receptor-like (GLR) homologs have evolved many plant-specific physiological functions, such as sperm signaling in moss, pollen tube growth, root meristem proliferation, innate immune, and wound responses. GLRs have been associated with Ca2+ signaling by directly channeling its extracellular influx into the cytosol. Nevertheless, very limited information on functional properties of GLRs is available, and we mostly rely on structure/function data obtained for animal iGluRs to interpret experimental results obtained for plant GLRs. Yet, a deeper characterization and better understanding of plant GLRs is progressively unveiling original and different functions when compared with their mammalian counterparts. Here, we review the function of plant GLRs comparing their predicted structure and physiological roles with those of the well-documented roles of iGluRs. We conclude that interpreting GLR function based on comparison with their animal counterparts calls for caution, especially when presuming physiological roles and the mode of action for plant GLRs, and when comparing iGluRs in neuronal tissues with those in peripheral, non-neuronal tissues. Ca2+ signaling, cation channel, electric signaling, GLR, glutamate receptor-like channel, iGluR, ionotropic glutamate receptor, structure–function Introduction Ionotropic glutamate receptors (iGluRs) have been identified in all three domains of life, and their presence in the animal branch ranges from ctenophores to vertebrates. Homologs of iGluRs, known as glutamate receptor-like (GLR) channels, were also found in the genomes of Chlamydomonas, chlorophytes, mosses, ferns, gymnosperms, and flowering plants (De Bortoli et al., 2016) (Fig. 1). It is noteworthy that GLRs are absent from yeast, eubacteria, archaebacteria, and fungi (Chiu et al., 1999), suggesting kingdom-specific roles for iGluRs and GLRs in animals and plants, respectively. Indeed, in mammals, iGluRs play an important role in integrative cognitive processes such as memory and learning, and they have been linked to the pathology of depression and psychosis as well as neurodegenerative diseases, including Alzheimer’s disease (Reinders et al., 2016). The evolution of iGluRs is intimately linked to central nervous system complexification (Jorgensen, 2014). Remarkably, iGluRs are believed to be fundamental for neuronal signaling, development, and plasticity in phyla as ancient as cnidarians, such as hydra (Pierobon, 2012), and are conserved in the more evolved phyla of nematodes and vertebrates (Fig. 1). Fig. 1. View largeDownload slide Phylogram of selected glutamate receptors from bacteria, plants, and animals. The tree shows the phylogenetic relationship of chosen glutamate receptors from Arabidopsis thaliana (At), the nematode Caenorhabditis elegans (Ce, °), the unicellular green alga Chlamydomonas rheinhardtii (Cr, *), the fruit fly Drosophila melanogaster (Dm, ¶), the gymnosperm Ginkgo biloba (Gb), Homo sapiens (Hs), the filamentous green alga Klebsormidium flaccidum (Kf,¨), the liverwort Marchantia polymorpha (Mp, ∞), the sea anemone Nematostella vectensis (Nv, ˇ), the moss Physcomitrella patens (Pp, ‡), the ctenophore Pleurobrachia bachei (Pb, ^), and the cyanobacterium Synechocystis sp. (GluR0, ˜). The tree was generated with the Clustal Omega Neighbor–Joining method software (Sievers et al., 2011). The scale bar indicates substitutions per site. Sequences used can be found in Supplementary Fig. S1 at JXB online. Fig. 1. View largeDownload slide Phylogram of selected glutamate receptors from bacteria, plants, and animals. The tree shows the phylogenetic relationship of chosen glutamate receptors from Arabidopsis thaliana (At), the nematode Caenorhabditis elegans (Ce, °), the unicellular green alga Chlamydomonas rheinhardtii (Cr, *), the fruit fly Drosophila melanogaster (Dm, ¶), the gymnosperm Ginkgo biloba (Gb), Homo sapiens (Hs), the filamentous green alga Klebsormidium flaccidum (Kf,¨), the liverwort Marchantia polymorpha (Mp, ∞), the sea anemone Nematostella vectensis (Nv, ˇ), the moss Physcomitrella patens (Pp, ‡), the ctenophore Pleurobrachia bachei (Pb, ^), and the cyanobacterium Synechocystis sp. (GluR0, ˜). The tree was generated with the Clustal Omega Neighbor–Joining method software (Sievers et al., 2011). The scale bar indicates substitutions per site. Sequences used can be found in Supplementary Fig. S1 at JXB online. Plants are characterized by the absence of a nervous system or any structure allowing for neuron-like electric signaling, and thus the discovery of plant GLRs was an unexpected revelation (Lam et al., 1998). Since then, and contrary to their overly specialized animal counterparts, plant GLRs have been shown to be implicated in a vast variety of cellular processes and different aspects of plant and cell physiology. For instance, GLRs have been linked to carbon and nitrogen metabolism (Kang and Turano, 2003; Kang et al., 2004), abscisic acid (ABA) biosynthesis and signaling (Kang and Turano, 2003; Kang et al., 2004; Kong et al., 2015), water loss (Kang et al., 2004; Lu et al., 2014), root gravitropism (Miller et al., 2010), lateral root initiation (Vincill et al., 2013), root development (Singh et al., 2016), innate immune responses (Kang et al., 2004; Kwaaitaal et al., 2011; Li et al., 2013; Manzoor et al., 2013; Forde and Roberts, 2014), stomatal closure (Cho et al., 2009), pollen tube growth (Michard et al., 2011; Wudick et al., 2018), self-incompatibility (Iwano et al., 2015), wound-induced leaf to leaf electric signaling (Mousavi et al., 2013), seed germination (Kong et al., 2015), response to aphid feeding (Vincent et al., 2017), and moss sperm signaling (Ortiz-Ramírez et al., 2017). Despite the broad physiological relevance of GLRs, there is no clear understanding about many of their fundamental regulatory aspects, namely concerning their endogenous ligand(s), gating, their ion selectivity, or the subcellular localization of most members. Exceptions for some of these aspects include AtGLR1.4 and 3.4 and the moss PpGLR1, which were shown to have a poor ionic selectivity when heterologously expressed in mammalian cells or Xenopus oocytes (Vincill et al., 2012; Tapken et al., 2013; Ortiz-Ramírez et al., 2017). Although initially expected to be plasma membrane channels, some plant GLRs were subsequently detected in chloroplasts and mitochondria (Teardo et al., 2011, 2015), as well as in the sperm cell (endo)membranes and the vacuolar membrane (Table 1) (Wudick et al., 2018), suggesting a diversification of their localization in plants. The lack of any precise functional characterization of GLRs makes it impossible to understand the cellular and molecular mechanisms behind the growing number of phenotypes described in the literature. In the absence of plant-specific data regarding the structure, function, and regulation of GLRs, results are often—and understandably—interpreted in the context of and compared with animal iGluRs. Excellent reviews with a focus on GLR function and phylogeny have recently been published (e.g. Price et al., 2012; Forde and Roberts, 2014; De Bortoli et al., 2016; Weiland et al., 2016), along with original data demonstrating some unique features of early land plant GLRs (Ortiz-Ramírez et al., 2017). In this review, we place an emphasis on the emerging differences between iGluRs and GLRs. Focusing on the scarce functional data available for plant GLRs and comparing GLR sequences, their predicted structures and functional domains with the respective parts from iGluRs, we want to raise awareness of obvious differences between both. These comparisons reflect the profoundly different evolutionary path taken by plants and animals, and are informative of the major specialization and functionalization steps that took place during the development of different traits and adaptations. Ultimately, we aim at challenging the interpretation of plant GLR function based on findings from iGluRs when taking first principles into consideration to explain the available data. Table 1. Prediction of N-terminal transmembrane domain (N-TMD) and documented localization AtGLR Predicted N-TMD? Plant localization AtGLR1.1 7/18 ++ AtGLR1.2 13/18 +++ AtGLR1.3 9/18 ++ AtGLR1.4 15/18 +++ Pa AtGLR2.1 14/18 +++ Vb AtGLR2.2 6/18 + AtGLR2.3 8/18 ++ AtGLR2.4 13/18 +++ AtGLR2.5 4/18 + AtGLR2.6 13/18 +++ AtGLR2.7 12/18 ++ AtGLR2.8 7/18 ++ AtGLR2.9 9/18 ++ AtGLR3.1 17/18 +++ AtGLR3.2 9/18 ++ Pc AtGLR3.3 5/18 + Pc, Sb AtGLR3.4 9/18 ++ C, Pc–f AtGLR3.5.1 3/18 + Mg AtGLR3.5.2 1/18 + Cg AtGLR3.5.3 6/12 + AtGLR3.6 7/18 ++ AtGLR3.7 11/18 ++ AtGLR Predicted N-TMD? Plant localization AtGLR1.1 7/18 ++ AtGLR1.2 13/18 +++ AtGLR1.3 9/18 ++ AtGLR1.4 15/18 +++ Pa AtGLR2.1 14/18 +++ Vb AtGLR2.2 6/18 + AtGLR2.3 8/18 ++ AtGLR2.4 13/18 +++ AtGLR2.5 4/18 + AtGLR2.6 13/18 +++ AtGLR2.7 12/18 ++ AtGLR2.8 7/18 ++ AtGLR2.9 9/18 ++ AtGLR3.1 17/18 +++ AtGLR3.2 9/18 ++ Pc AtGLR3.3 5/18 + Pc, Sb AtGLR3.4 9/18 ++ C, Pc–f AtGLR3.5.1 3/18 + Mg AtGLR3.5.2 1/18 + Cg AtGLR3.5.3 6/12 + AtGLR3.6 7/18 ++ AtGLR3.7 11/18 ++ Predicted likelihood of bearing an N-TMD: high (+++), medium (++), low (+), based on results from 18 different prediction programs (aramemnon.uni-koeln.de). Multiple AtGLR3.5 splicing variants were included due to their differential localization. C, chloroplast; M, mitochondrion, P, plasma membrane; S, sperm cell; V, vacuole; aTapken et al. (2013); bWudick et al. (2018); cVincill et al. (2013); dMeyerhoff et al. (2005); eTeardo et al. (2011); fVincill et al. (2012); gTeardo et al. (2015). View Large Table 1. Prediction of N-terminal transmembrane domain (N-TMD) and documented localization AtGLR Predicted N-TMD? Plant localization AtGLR1.1 7/18 ++ AtGLR1.2 13/18 +++ AtGLR1.3 9/18 ++ AtGLR1.4 15/18 +++ Pa AtGLR2.1 14/18 +++ Vb AtGLR2.2 6/18 + AtGLR2.3 8/18 ++ AtGLR2.4 13/18 +++ AtGLR2.5 4/18 + AtGLR2.6 13/18 +++ AtGLR2.7 12/18 ++ AtGLR2.8 7/18 ++ AtGLR2.9 9/18 ++ AtGLR3.1 17/18 +++ AtGLR3.2 9/18 ++ Pc AtGLR3.3 5/18 + Pc, Sb AtGLR3.4 9/18 ++ C, Pc–f AtGLR3.5.1 3/18 + Mg AtGLR3.5.2 1/18 + Cg AtGLR3.5.3 6/12 + AtGLR3.6 7/18 ++ AtGLR3.7 11/18 ++ AtGLR Predicted N-TMD? Plant localization AtGLR1.1 7/18 ++ AtGLR1.2 13/18 +++ AtGLR1.3 9/18 ++ AtGLR1.4 15/18 +++ Pa AtGLR2.1 14/18 +++ Vb AtGLR2.2 6/18 + AtGLR2.3 8/18 ++ AtGLR2.4 13/18 +++ AtGLR2.5 4/18 + AtGLR2.6 13/18 +++ AtGLR2.7 12/18 ++ AtGLR2.8 7/18 ++ AtGLR2.9 9/18 ++ AtGLR3.1 17/18 +++ AtGLR3.2 9/18 ++ Pc AtGLR3.3 5/18 + Pc, Sb AtGLR3.4 9/18 ++ C, Pc–f AtGLR3.5.1 3/18 + Mg AtGLR3.5.2 1/18 + Cg AtGLR3.5.3 6/12 + AtGLR3.6 7/18 ++ AtGLR3.7 11/18 ++ Predicted likelihood of bearing an N-TMD: high (+++), medium (++), low (+), based on results from 18 different prediction programs (aramemnon.uni-koeln.de). Multiple AtGLR3.5 splicing variants were included due to their differential localization. C, chloroplast; M, mitochondrion, P, plasma membrane; S, sperm cell; V, vacuole; aTapken et al. (2013); bWudick et al. (2018); cVincill et al. (2013); dMeyerhoff et al. (2005); eTeardo et al. (2011); fVincill et al. (2012); gTeardo et al. (2015). View Large The ontology of the animal iGluR and plant GLR family Glutamate receptors are part of the superfamily of gated channels, which include the eubacterial proton-gated K+ channel KscA, and derivative eukaryotic voltage- and ligand-gated channels (Wo and Oswald, 1995; Price et al., 2012). In mammals, four iGluR clades have been differentiated based on their divergent sequences and ligand specificities (Traynelis et al., 2010) (Fig. 1). The most ancestral clade, present as early as in cnidarian species, consists of N-methyl-d-aspartate (NMDA) receptors that differentiated into α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) and kainate receptors (Chiu et al., 1999; Price et al., 2012; De Bortoli et al., 2016; Greer et al., 2017). A fourth group is comprised of two delta receptors (δ-receptors), which share sequence and structural homology with iGluRs, but cannot be activated by glutamate. In addition to different ligand affinities, the members from each iGluR clade also differ by their (de)activation kinetics, as well as their ionic selectivities (Traynelis et al., 2010), giving to each group of iGluRs a specific role in the highly complex and evolved nervous system. Based on their sequence homology and the analyzed sequence region, plant GLRs were shown to be similar to both NMDA and non-NMDA receptor-like iGluRs, suggesting a divergence of iGluRs/GLRs prior to their clade differentiation (Weiland et al., 2016). Sequence alignments and analyses of higher plant GLRs revealed that they group in three clades (Chiu et al., 2002), with some authors proposing a fourth clade in monocotyledons (Chen et al., 2016; De Bortoli et al., 2016). Early land plants, such as mosses, ferns, and gymnosperms (e.g. ginkgo and pine tree with 9 and 40 GLRs, respectively), code for channels that exclusively cluster in clade 3, while clades 1 and 2 are only found in angiosperms (Price et al., 2012; De Bortoli et al., 2016) (Fig. 1). Functional data for plant GLRs is very limited, making it difficult to assess their overall or clade-specific role. Early studies focused on pharmacology and point mutations to address structure–function relationships in iGluRs by patch-clamp (Traynelis et al., 2010). More recent advances resolving iGluR structures by crystallography (Sobolevsky et al., 2009; Chen et al., 2014; Karakas and Furukawa, 2014; Yelshanskaya et al., 2016) and cryo-electron microscopy (Zhao et al., 2016; Twomey et al., 2016; Lü et al., 2017) positioned these proteins among the best-characterized ion channels at the molecular level (reviewed in Dawe et al., 2015; Karakas et al., 2015; Sobolevsky, 2015; Mayer, 2016; Zhu and Gouaux, 2017). The level of homology between plant GLRs and iGluRs has been accepted as a sufficient basis (Chiu et al., 2002) to warrant inferences about their general structure derived from available data on iGluRs (Fig. 2). In the next paragraphs we will discuss the different GLR domains, including the N-terminal domain, the ligand-binding and pore domain, and the C-terminal domain (Fig. 2). Fig. 2. View largeDownload slide Putative tetrameric membrane structure of AtGLRs based on the structure of iGluRs. Schematic membrane structure of four AtGLR subunits (I–IV). As exemplified in (I), each subunit consists of an amino-terminal domain (ATD) that forms a clamshell-shaped binding site for modulators with homology to bacterial leucine/isoleucine/valine-binding protein (LIVBP) domains. It is followed by the ligand-binding domain (LBD), which has a similar structure but the two lobes are composed of two segments (S1 and S2) from the soluble terminus and the external loop, respectively, and bear a lysine/arginine/ornithine-binding protein (LAOBP)-like domain with homology to the periplasmic binding protein-like II superfamily. The three membrane-spanning helices (M1–M3) of the transmembrane domains (TMDs) and a membrane re-entrant loop, forming the pore region (P) of the channel, are followed by the cytosolic carboxy-terminal domain (CTD). After Shepherd and Huganir (2007). Not drawn to scale. Fig. 2. View largeDownload slide Putative tetrameric membrane structure of AtGLRs based on the structure of iGluRs. Schematic membrane structure of four AtGLR subunits (I–IV). As exemplified in (I), each subunit consists of an amino-terminal domain (ATD) that forms a clamshell-shaped binding site for modulators with homology to bacterial leucine/isoleucine/valine-binding protein (LIVBP) domains. It is followed by the ligand-binding domain (LBD), which has a similar structure but the two lobes are composed of two segments (S1 and S2) from the soluble terminus and the external loop, respectively, and bear a lysine/arginine/ornithine-binding protein (LAOBP)-like domain with homology to the periplasmic binding protein-like II superfamily. The three membrane-spanning helices (M1–M3) of the transmembrane domains (TMDs) and a membrane re-entrant loop, forming the pore region (P) of the channel, are followed by the cytosolic carboxy-terminal domain (CTD). After Shepherd and Huganir (2007). Not drawn to scale. Structural and functional domains: iGluRs versus GLRs The N-terminal signal peptide: to cleave or not to cleave? Members of the iGluR family of ligand-gated ion channels generally possess an N-terminal transmembrane anchor sequence that acts as a signal peptide allowing the nascent protein to enter the secretory pathway. Following the insertion of the protein into the endoplasmic reticulum (ER) membrane, the signal peptide is typically cleaved off and thus not part of the mature iGluR (He et al., 2016). Based on in silico analyses of 18 different topology prediction programs (aramemnon.uni-koeln.de), 16 out of 20 AtGLRs have a medium to high probability (predicted by ≥6 of 18 programs) of bearing an N-terminal transmembrane domain, which could function as a signal peptide (Table 1). Interestingly, among the AtGLRs with a low probability (predicted by <6 programs), there are two AtGLRs with predicted (AtGLR2.5) or documented (AtGLR3.5) localization outside the secretory pathway, namely in mitochondria and/or chloroplasts (Teardo et al., 2015). Signal sequences mediating the targeting towards those organelles are typically soluble and eventually cleaved (Kim and Hwang, 2013), and therefore would not integrate into a membrane. Whether these putative signal peptide sequences are cleaved is as yet not well studied, as very few conclusive experimental data are available to address this issue in plants. Experiments addressing this question either failed (Li et al., 2006) or it was found that the first transmembrane domain alone was not necessarily sufficient for ER targeting (Davenport, 2002). Localization of AtGLRs along the secretory pathway has only been shown for a few members, including AtGLR1.4 (Tapken et al., 2013), AtGLR3.2, and AtGLR3.3 (Vincill et al., 2013, Wudick et al., 2018), as well as AtGLR3.4 (Meyerhoff et al., 2005; Teardo et al., 2011; Vincill et al., 2012, 2013). For AtGLR3.4 and an alternatively spliced AtGLR3.5, localization in membranes of either chloroplast or mitochondria, respectively, was documented in addition to their plasma membrane localization (Teardo et al., 2011, 2015), while AtGLR2.1 is the first AtGLR to be localized to the vacuolar membrane (Table 1) (Wudick et al., 2018). The amino-terminal domain (ATD) The ATD is located in the soluble N-terminus of glutamate receptors, and structures of several iGluR ATDs have been determined by crystallography, including members of the AMPA, kainate, and NMDA receptor family (reviewed in Dawe et al., 2015; Karakas et al., 2015; Sobolevsky, 2015; Mayer, 2016; Zhu and Gouaux, 2017). They were shown to form a clamshell-like binding pocket with homology to bacterial leucine/isoleucine/valine-binding protein (LIVBP) domains (Acher and Bertrand, 2005), which is absent in the bacterial iGluR0 (reviewed in Furukawa, 2012) (Fig. 2). The ATD is best studied in NMDA receptors where it is thought to allow regulation by allosteric modulators such as polyamines, Zn2+ ions, or specific agents such as ifenprodil (Paoletti et al., 1997; Masuko et al., 1999). So far no such regulation has been described for non-NMDA receptors, which might suggest that this layer of regulation is not present in AMPA and kainate receptors. Subunit assembly and multimerization of iGluRs in the ER as well as synaptic transmission were also shown to depend on the N-termini for members of the AMPA (Díaz-Alonso et al., 2017), kainate (Sheng et al., 2017), and NMDA receptor families (Traynelis et al., 2010). Moreover, an alternatively spliced version of the ATD from the NMDA receptor GRIN1 was shown to affect the allosteric modification of the channel (Traynelis et al., 1995). The ATD region additionally plays a role in iGluR gating (see below), as structure resolution by crystallography and cryo-electron microscopy revealed large movements of the region during pore opening. The opening is triggered by ligand binding, after which the channel spontaneously closes in a ligand-independent manner and transiently stays ligand insensitive. This mechanism, which is also referred to as desensitization, relies on specific molecular conformational changes involving the ATD domain (reviewed in Dawe et al., 2015; Karakas et al., 2015; Sobolevsky, 2015; Mayer, 2016; Zhu and Gouaux, 2017). Nevertheless, desensitization has not so far been unequivocally described for plant GLRs at the electrophysiological level, and there are no data indicating that the conformational changes in iGluRs are identical to those of plant GLRs. Sequence comparisons revealed from the beginning an overall similarity between Arabidopsis GLRs and iGluRs (Lam et al., 1998), and subsequent phylogenetic analyses indicated that both families diverged prior to the divergence of the iGluR classes (Chiu et al., 1999). However, a later study refined the analyses by separately evaluating the relatedness of the first third (or N-terminal region) and the last two-thirds (or C-terminal regions) of AtGLR proteins to iGluRs (Turano et al., 2001). While supporting the conclusion that the C-terminal regions were most closely related to iGluRs, the authors described that the N-terminal regions of AtGLRs from clade 3 showed more homology to members of the subfamily C of G-protein-coupled receptors (GPCRs), which contain metabotropic glutamate receptors (mGluRs) and γ-aminobutyric acidB (GABA-B) receptors (Turano et al., 2001). Accordingly, Gene Ontology annotation revealed a possible implication in GPCR-mediated ligand signaling for some AtGLR3s (Roy and Mukherjee, 2016). Strikingly, this distinct ATD is absent in iGluRs but appears to be conserved in plant GLRs closely related to clade 3 AtGLRs, including the moss Physcomitrella patens, the liverwort Marchantia polymorpha, and the gymnosperm Ginkgo biloba (Fig. 3), while being absent in clade 1 and 2 AtGLRs. Interestingly, AtGLR3.5 is the only member containing all amino acids of the domain’s consensus motif. It has been reported that the conserved residues in this domain constitute a binding pocket for the glycine moiety (H2N-RCH-COOH) of amino acids (Acher and Bertrand, 2005), and the binding of glutamate was shown for rat mGluR1 (Morikawa et al., 2000). Based on these findings, it is tempting to speculate that members of the AtGLR3 clade and other GLRs grouping with this clade are modulated or activated differently, not only when compared with iGluRs, but also when compared with members of clade 1 and 2 of AtGLRs. Though lacking further characterization, several alternatively spliced versions of different AtGLRs affecting the ATD are annotated (apps.araport.org/thalemine) and—if translated into functional proteins—are expected to change the modulation of the channels. An interaction of the N-terminus with auxiliary proteins similar to that described in iGluRs has not yet been reported for AtGLRs. Box 1 > AtGLRs can have different target sequences and (additional) subcellular localizations > They have a conserved iGluR/GLR ATD structure (clamshell domain), but are functionally different > In NMDA receptors, the ATD is important for binding of allosteric modulators > The ATD of AtGLR3s bears a motif that is also found in metabotropic glutamate receptors (mGluRs) where it functions as a binding pocket for glycine moieties ( i.e. amino acids) Fig. 3. View largeDownload slide Predicted amino-terminal domain (ATD) in the N-terminus of clade 3-like GLRs and metabotropic glutamate receptors (mGluRs). Sequence alignment of the N-terminal domains of clade 3-like GLRs from Arabidopsis thaliana (At), Physcomitrella patens (Pp), Marchantia polymorpha (Mp), and Ginkgo biloba (Gb), as well as the human metabotropic glutamate receptor 1 (HsmGluR1). The residues of the consensus sequence (on the top and bold letters with green highlight) are predicted to form a binding pocket for glycine moieties. Conservation of plant ATDs: *fully conserved residue; : conservation between groups of strongly similar properties; . conservation between groups of weakly similar properties. Fig. 3. View largeDownload slide Predicted amino-terminal domain (ATD) in the N-terminus of clade 3-like GLRs and metabotropic glutamate receptors (mGluRs). Sequence alignment of the N-terminal domains of clade 3-like GLRs from Arabidopsis thaliana (At), Physcomitrella patens (Pp), Marchantia polymorpha (Mp), and Ginkgo biloba (Gb), as well as the human metabotropic glutamate receptor 1 (HsmGluR1). The residues of the consensus sequence (on the top and bold letters with green highlight) are predicted to form a binding pocket for glycine moieties. Conservation of plant ATDs: *fully conserved residue; : conservation between groups of strongly similar properties; . conservation between groups of weakly similar properties. The receptor or ligand-binding domain (LBD) iGluRs are ligand-gated channels, which upon binding of the agonist open their pore. Decades of point mutation analyses revealed two protein segments, S1 and S2, that are composed of two highly conserved 10 amino acid motifs, which are directly involved in ligand binding and flank the membrane pore-forming domain of the channel (Traynelis et al., 2010) (Fig. 2). The 3D structure of the S1 and S2 stretches is buried inside a receptor domain, which, like the ATD, folds into a clamshell-like structure (reviewed in Dawe et al., 2015; Karakas et al., 2015; Sobolevsky, 2015; Mayer, 2016; Zhu and Gouaux, 2017). This structure is homologous to lysine/arginine/ornithine-binding protein (LAOBP)-like domains from members of the periplasmic-binding protein-like II superfamily in bacteria (Acher and Bertrand, 2005). When the ligand binds, the clamshell closes, which induces a larger trans-conformational rearrangement of the tetrameric channel, and leads to the opening of the pore (Mayer, 2016; Zhu et al., 2016; Twomey and Sobolevsky, 2018). Animal iGluRs show strong ligand specificity. Depending on their subunit composition, NMDA and delta receptors bind either to glutamate/aspartate or to glycine/d-serine, while AMPA and kainate receptors exclusively bind to glutamate/aspartate (Traynelis et al., 2010). The binding displays low affinity, which occurs typically at ~0.1–3 µM for endogenous ligands (Traynelis et al., 2010), and its specificity and affinity are determined by the structure formed by S1 and S2 inside the clamshell. The S1 and S2 domains are highly conserved among iGluRs, showing the highest conservation within each of the iGluR clades. Strikingly, a similar sequence conservation is missing in S1- and S2-like domains of AtGLRs (De Bortoli et al., 2016). The gating of AtGLRs is still not well characterized (Forde and Roberts, 2014) and was initially assessed by employing the pharmacology from iGluRs. The first indication of possible GLR activity in plants was detected as an effect on hypocotyl growth in relation to light upon application of the iGluR antagonist 6,7-dinitroquinoxaline-2,3-dione (DNQX; Lam et al., 1998). However, while the AMPA receptor-specific antagonist DNQX is typically used at concentrations of 10 µM when studying iGluRs (affinity <1 µM; Traynelis et al., 2010), much higher concentrations of ~0.1–0.4 mM were initially used in plants. Accordingly, later studies in plants showed a lack of specificity for antagonists of the animal NMDA or AMPA receptor class, with or without associated agonists. Examples include studying the effects of cyanquixaline (6-cyano-7-nitroquinoxaline-2,3-dione) (CNQX), DNQX, and 2-amino-5-phosphonovalerate (AP5) on tobacco pollen tubes and protoplasts (Michard et al., 2011), of DNQX on whole plants (Dubos et al., 2003), of 500 µM CNQX, DNQX, or 2,3-dihydroxy-6-nitro-7-sulfamoyl-benzo[f]quinoxaline-2,3-dione (NBQX) on leaves (Meyerhoff et al., 2005), of 100 µM DNQX on vesicles from the inner chloroplast membrane of spinach containing AtGLR clade 3-like receptors (Teardo et al., 2010), or of AP5 and DNQX on leaves from glr3.3 mutant plants treated with the potential ligand glutathione (Li et al., 2013). Moreover, the palette of putative endogenous AtGLR agonists is not yet well defined. First proof of a plant GLR activity was shown by using the cycad-derived agonist β -N-methylamino-l-alanine (BMAA) to block light-induced hypocotyl shortening and cotyledon expansion in Arabidopsis (Brenner et al., 2000). Subsequent studies revealed that plant GLRs seem to respond to a wide range of amino acids, expanding beyond the name-giving glutamate and including non-conventional amino acids, such as d-serine, and the tripeptide glutathione (Michard et al. 2011; Weiland et al., 2016). Pharmacological analyses of AtGLRs are consistent with the idea of them being different from non-NMDA receptors in general and divergent within plant GLR clades in particular. In animals it was shown that in addition to glutamate, many other amino acids and naturally occurring molecules are able to activate AMPA and kainate receptors (Traynelis et al., 2010), making it difficult to pinpoint specific mechanisms related to each agonist. So far only AtGLR1.4 and 3.4 have been characterized as ligand-gated channels in heterologous systems [for a thorough review on AtGLR (ant)agonists, see Weiland et al., 2016]. AtGLR1.4 can be activated by methionine, tryptophan, phenylalanine, leucine, tyrosine, asparagine, and threonine, but, when studied in seedlings, only methionine was found to have a depolarizing effect on the membrane potential, which was absent in a glr1.4 mutant (Tapken et al., 2013). It is antagonized by eight natural amino acids, arginine being the most efficient, as well as by the synthetic antagonists DNQX, Philanthoxin, and MK-801, although at high concentrations (≥100 µM; Tapken et al., 2013). By introducing point mutations, the authors proved the involvement of the LBD in ligand binding to AtGLR1.4 (Tapken et al., 2013). Upon heterologous expression in HEK cells, AtGLR3.4 could be activated by asparagine, l-serine, and glycine (Stephens et al., 2008). Of note, the ligand concentrations used in these experiments ranged widely between micromolar (AtGLR1.4) and millimolar (AtGLR3.4) concentrations. Intriguingly, and in contrast to their animal homologs, some GLRs also seem to be active without the presence of a ligand (Michard et al., 2011; Ortiz-Ramírez et al., 2017, Wudick et al., 2018). In tobacco pollen tube protoplasts, the antagonist CNQX inhibited currents, which were recorded in the absence of any ligand in the bath solution (Michard et al., 2011). Even more strikingly, AtGLR3.2 and 3.3, as well as PpGLR1 from the moss P. patens, induced ionic currents without the presence of any ligand when expressed in COS-7 cells (Ortiz-Ramírez et al., 2017, Wudick et al., 2018). In the case of PpGLR1, the currents were inhibited by the iGluR antagonists AP5 and CNQX, suggesting that the channel was open in a basal state configuration, without any apparent ligand (Ortiz-Ramírez et al., 2017). Altogether, the data suggests that gating of plant GLRs is different from that observed in animal iGluRs. Further functional characterization and structural analyses are necessary to understand fully the gating, or apparent lack thereof, in plant GLRs. The pore: driving ionic selectivity The ionic pore is formed at the interface of the four subunits of the functional channel (Fig. 2). On the protein sequence level, the pore region is composed of ~20 residues located between the first and second transmembrane domains, forming a membrane re-entrant loop (Traynelis et al., 2010). While the prokaryotic iGluR0 is a highly potassium-selective channel and shows the typical ‘GYGD’ selectivity filter motif (Chen et al., 1999), which is also found in rotifer AvGluR1 (Janovjak et al., 2011), related animal iGluRs lost that motif and they are non-selective cation channels (Traynelis et al., 2010). Some iGluRs, especially from the NMDA family, were shown to be Ca2+ permeable (Traynelis et al., 2010). The pore from an AMPA receptor in an open state has recently been resolved by cryo-electron microscopy imaging (Twomey and Sobolevsky, 2018). Of special note, RNA editing mechanisms affecting residues from the pore (and the extracellular loop) region of iGluRs have been described to modulate this Ca2+ permeability, and channel properties in general (Barbon and Barlati, 2011). The Q/R editing site in the pore region of AMPA and kainate receptors represents some of the best-characterized residues with implication in several diseases (Barbon and Barlati, 2011). A similar kind of editing has not so far been documented for AtGLRs. Unfortunately, the poor conservation of the pore region between animal and plant glutamate receptors makes it impossible to derive information on GLR selectivity (De Bortoli et al., 2016). In addition, the pore sequence, and particularly the residues dictating the Ca2+ permeability in NMDA channels, are poorly conserved in AtGLRs, suggesting an overall different selectivity for GLRs, and possibly large selectivity differences between plant GLRs (De Bortoli et al., 2016). With a few exceptions, the selectivity of AtGLRs is unknown. Ca2+ imaging and studies on the membrane electric polarization of leaf cells in response to amino acids in wild-type and glr3.3/3.4 knock-down plants suggested that those channels are Ca2+ permeable and could be blocked by the broad cation channel inhibitor gadolinium (Gd3+, Kim et al., 2001). In pollen tubes, d-serine increased Ca2+ influxes as measured by using external ion-specific electrodes, as well as cytosolic Ca2+ concentrations monitored with the yellow cameleon Ca2+ probe YC3.6 (Michard et al., 2011). The first molecular data were obtained by swapping the pore of 17 AtGLRs with those of AMPA and kainate receptors, followed by the characterization of the HEK cell-expressed chimera by the patch-clamp technique (Tapken and Hollmann, 2008). Two chimeras containing the pores of AtGLR1.1 and 1.4 were functional, both displaying a non-selective cationic conductance. Furthermore, expression in HEK cells of AtGLR3.4 was found to generate a cationic current, partly carried by Ca2+ (Vincill et al., 2012). A similar result was described for AtGLR1.4 upon expression in Xenopus oocytes (Tapken et al., 2013). Recent characterization of the moss channel PpGLR1 and of AtGLR3.2 and 3.3 showed that these channels are permeable to cations, including Ca2+, which could be partially inhibited by Gd3+ (Ortiz-Ramírez et al., 2017; Wudick et al., 2018). In summary, essential questions such as AtGLR permeability to Ca2+, relative selectivity for different ions, anion permeability, and the existence of a mechanism similar to the magnesium block of NMDA receptors (Mayer, 2017) to avoid a toxic Ca2+ influx into the cytosol remain unanswered and need to be addressed in the future. The ‘gate’ region of the channel In a simplistic way, structural changes induced by agonist binding to the LBD are transmitted to the ‘gate’ region of the protein, which is occluding the pore in a non-ligand-bound state, leading to the opening of the pore of the channel (Twomey and Sobolevsky, 2018). Wider conformational changes occur in response to receptor–gate interactions, including large rearrangements between the subunits of the channel tetramer, and the twist of the NTD domain (reviewed in Dawe et al., 2015; Karakas et al., 2015; Sobolevsky, 2015; Mayer, 2016; Zhu and Gouaux, 2017). In iGluRs, the ‘gate’ is highly conserved and represented by a ‘SYTANLAA’ amino acid motif (Traynelis et al., 2010) (Fig. 4). Targeted mutations in the ‘gate’ motif of GRIA2 revealed that the second alanine is responsible for ‘gate’ opening (Moore et al., 2013). Fig. 4. View largeDownload slide Graphical representation of the sequence conservation for the GLR ‘gate’ motif in different species. Representation of aligned sequences (numbers given) from the ‘gate’ region of all human iGluRs (HsGluRs, 18), Arabidopsis thaliana (AtGLRs, 20), and Pleurobrachia bachei (PbGLRs, 11) (left column), and a clade-specific representation of AtGLRs (right column). The ‘gate’ motif of all nine ginkgo GbGLRs, the two moss PpGLRs, and the algal MpGLR represented in Fig. 1 are identical to the AtGLR3 consensus motif, except for a perfectly conserved serine (S) at position 5. AtGLR3.7 is the only AtGLR3 with an asparagine (N) in this position. Note that two PbGLRs did not display any ‘gate’ motif and were not considered here. Logos were created with WebLogo3.5 (Crooks et al., 2004). Fig. 4. View largeDownload slide Graphical representation of the sequence conservation for the GLR ‘gate’ motif in different species. Representation of aligned sequences (numbers given) from the ‘gate’ region of all human iGluRs (HsGluRs, 18), Arabidopsis thaliana (AtGLRs, 20), and Pleurobrachia bachei (PbGLRs, 11) (left column), and a clade-specific representation of AtGLRs (right column). The ‘gate’ motif of all nine ginkgo GbGLRs, the two moss PpGLRs, and the algal MpGLR represented in Fig. 1 are identical to the AtGLR3 consensus motif, except for a perfectly conserved serine (S) at position 5. AtGLR3.7 is the only AtGLR3 with an asparagine (N) in this position. Note that two PbGLRs did not display any ‘gate’ motif and were not considered here. Logos were created with WebLogo3.5 (Crooks et al., 2004). Sequence comparisons between animal iGluRs and plant GLRs reveal that the ‘gate’ motif is not completely conserved (Fig. 4). In AtGLRs, only the tyrosine (T), alanine (A), and leucine (L) in position 3, 4, and 6, respectively, are fully conserved, and most variability is observed in position 5 (Fig. 4). Interestingly, within Arabidopsis, AtGLR3s show the most conserved ‘gate’ motif, that—except for AtGLR3.7—follows the ‘SYTASLTS’ sequence, which is also conserved in the two moss and all nine ginkgo GLRs (Fig. 4). While AtGLR2s show additional variation in position 7, AtGLR1s show by far the most divergent ‘gate’ motif, with further variations in positions 1 and 3 (Fig. 4). Interestingly, in iGluRs, an A to T switch in position 8 of the motif, the so-called Lurcher mutation, results in a constitutively open channel (Zuo et al., 1997; Klein and Howe, 2004). Since several plant GLRs display a serine in position 8, which like threonine is a polar, hydrophilic amino acid, it is tempting to speculate that at least some of them could also display channel activity in the absence of any agonists. Indeed, the moss PpGLR1 as well as AtGLR3.2 and 3.3 were recently shown to be active without any ligand (Ortiz-Ramírez et al., 2017; Wudick et al., 2018), indicating that ligand-dependent channel activation might not be necessary in plant GLRs or may have been lost in other species. An extreme example of the latter can be found for Pleurobrachia bachei, which displays very poor conservation of the ‘gate’ motif (Fig. 4). Box 2 > Unlike their homologous domains in iGluRs, S1/S2 segments are poorly conserved in GLRs > There is an apparent broad ligand spectrum for AtGLRs > The pore region is poorly conserved in plant GLRs > The divergent ‘gate’ motif might reflect that some GLRs (i.e. PpGLR1, AtGLR3.2, and AtGLR3.3) might function without ligand-induced activation From the ‘gate’ to the cytosolic C-terminus: a role in gating and desensitization A region including the S2 lobe of the LBD and preceding the last transmembrane domain of AMPA receptors was shown to undergo alternative splicing events, yielding either ‘FLIP’ or ‘FLOP’ variants of the channel. Though the physiological role of this event is still not totally understood, the two splicing variants display different kinetic properties and show differential interaction with auxiliary proteins binding in that region (Greger et al., 2017). An impact on the early trafficking of the channel was also reported (Coleman et al., 2006). With the exception of the N-terminal region of AtGLR3.5 (see above), alternative splicing events have not been studied for plant GLRs, but the occurrence of several splicing variants covering the respective regions in AtGLRs (apps.araport.org/thalemine) might be indicative of a similar way to modify the channel activity/specificity. Interestingly, in the case of AtGLR2.4, the most abundant (and hence the often only annotated) splicing variant is completely lacking the ‘gate’ motif. While present in both iGluRs and GLRs, the last transmembrane domain is absent in the prokaryotic iGluR0 (Chen et al., 1999). In addition to serving as an ‘anchor’ for the C-terminal domain that is essential for iGluR post-translational regulation (see below), it is also involved in conformational changes of the channel during gating, as revealed by point mutation analyses in iGluRs. More specifically, tryptophan-scanning mutagenesis of the last transmembrane domain revealed its impact on gating (for both NMDA and AMPA receptors) and desensitization (for AMPA receptors) (Amin et al., 2017). Importantly, this desensitization is essential for iGluR function in general, and their clade-specific function in particular (Traynelis et al., 2010; Popescu, 2012). In leaf cells, an amino acid-specific desensitization-like phenomenon has been reported, which was affected in glr3.3 and glr3.4 knock-down plants, suggesting that a desensitization-like mechanism might also exist in plants (Stephens et al., 2008). Nevertheless, neither AtGLR1.4 nor 3.4 showed desensitization kinetics when heterologously expressed (Vincill et al., 2012; Tapken et al., 2013). The cytosolic C-terminal domain The intracellular C-terminus of iGluRs is the most diverse part of the receptors, in terms of both amino acid sequence and length (Traynelis et al., 2010). Despite the apparent lack of functional domains, distinct motifs such as ER retention sites can be found (Traynelis et al., 2010). Additionally, some AMPA receptors also harbor a C-terminal consensus motif, which allows interaction with proteins bearing a Psd-95/DlgA/ZO1 (PDZ)-domain, such as GRIP, ABP/GRIP2, PICK1, and others, which are involved in trafficking and recycling of the receptor (Collingridge et al., 2004; Shepherd and Huganir, 2007; Traynelis et al., 2010). Interestingly, iGluR interaction/modification through PDZ domains is apparently not well conserved or only occurred in higher organisms, since none of the Drosophila iGluRs from the neuromuscular junction have such a domain (Kim et al., 2012). Accordingly, so far neither homologs nor proteins with PDZ domains were identified in Arabidopsis. The presence of 14-3-3 protein-binding sites in the C-terminus of several iGluRs has also been documented. While interaction with 14-3-3 proteins and the kainate receptor GRIK2 slowed down the receptor’s decay kinetics (Sun et al., 2013), interaction with members of the NMDA receptor family impacted the trafficking and isoform-specific interaction (Chung et al., 2015) or led to increased surface expression of the channels (Chen and Roche, 2009). Accordingly, deletion of the C-terminus did not abolish the overall function but rather affected the regulation of different iGluRs (Traynelis et al., 2010). Similar to their mammalian counterparts, AtGLRs show highly variable C-terminal sequences and lengths (Fig. 5). While AtGLR1s display the shortest C-termini (18 amino acids), the lengths of the C-termini are most divergent in AtGLR2s (between 41 and 113 amino acids; Fig. 5). Multiple putative ER retention/retrieval motifs with documented function in plants (i.e. KKxx, KxKxx, RR, RxR, RxxR, and Φxx[K/R/D/E]Φ) (Boulaflous et al., 2009; Gao et al., 2014) are present in all AtGLR C-termini (Fig. 5). Moreover, 14-3-3 protein mode I (Rxx[S/T]xP) or mode II (Rxxx[S/T]xP) binding sites can be found in the C-termini of six AtGLRs from all three clades (Table 2). For three members (AtGLR1.4, 2.9, and 3.7), an interaction with 14-3-3 proteins was shown experimentally (Chang et al., 2009; Shin et al., 2011) (Table 2). Since binding of 14-3-3 proteins relies on phosphorylation of the serine/threonine residue in the center of the binding motif, phosphorylation events are also likely to occur in the GLR C-terminus. An interaction of the C-termini with other auxiliary proteins is possible but not yet documented. Box 3 > AtGLR C-termini are most variable in terms of sequence and length > They contain putative ER retention motifs and 14-3-3 protein-binding motifs > They lack regulatory PDZ domains that can be found in iGluRs Fig. 5. View largeDownload slide Graphical representation of the length and sequence of AtGLR C-termini. The graph depicts the lengths and sequences of all 20 AtGLR C-termini. While clade 1 termini (magenta) are overall the shortest, the length of clade 2 C-termini (orange) is most divergent. The amino acid number is given for each peptide. Lengths of clade 3 C-termini are contoured in blue. Putative ER retention motifs of the KKxx, KxKxx, RR, RxR, RxxR, or Φxx[K/R/D/E]Φ type are underlined. The lengths were determined with the help of multiple topology prediction programs at aramemnon.uni-koeln.de. Fig. 5. View largeDownload slide Graphical representation of the length and sequence of AtGLR C-termini. The graph depicts the lengths and sequences of all 20 AtGLR C-termini. While clade 1 termini (magenta) are overall the shortest, the length of clade 2 C-termini (orange) is most divergent. The amino acid number is given for each peptide. Lengths of clade 3 C-termini are contoured in blue. Putative ER retention motifs of the KKxx, KxKxx, RR, RxR, RxxR, or Φxx[K/R/D/E]Φ type are underlined. The lengths were determined with the help of multiple topology prediction programs at aramemnon.uni-koeln.de. Table 2. AtGLRs with a putative C-terminal 14-3-3 protein-binding motif Gene Position Peptide Mode Interacting 14-3-3 protein AtGLR1.3 840 FVRSIH[T]SPLD II – AtGLR1.4 850 EIRPSP[T]TPNR II At4g05685, AtGRF1a AtGLR2.6 832 DNMRAP[T]SPPI I – AtGLR2.9 921 SEERFT[T]QPII I At1g78300, AtGRF2b AtGLR3.6 861 GSIRRR[S]SPSA I or II – AtGLR3.7 860 RMERTS[S]MPRA I At1g78300, AtGRF2b Gene Position Peptide Mode Interacting 14-3-3 protein AtGLR1.3 840 FVRSIH[T]SPLD II – AtGLR1.4 850 EIRPSP[T]TPNR II At4g05685, AtGRF1a AtGLR2.6 832 DNMRAP[T]SPPI I – AtGLR2.9 921 SEERFT[T]QPII I At1g78300, AtGRF2b AtGLR3.6 861 GSIRRR[S]SPSA I or II – AtGLR3.7 860 RMERTS[S]MPRA I At1g78300, AtGRF2b The table lists AtGLRs with predicted C-terminal 14-3-3 protein-binding motifs and their position. Conserved residues are in bold, and putatively phosphorylated residues in brackets. The motif modes (I Rxx[S/T]xP; or II, Rxxx[S/T]xP) and experimental evidence for interaction are given. aShin et al. (2011); bChang et al. (2009). View Large Table 2. AtGLRs with a putative C-terminal 14-3-3 protein-binding motif Gene Position Peptide Mode Interacting 14-3-3 protein AtGLR1.3 840 FVRSIH[T]SPLD II – AtGLR1.4 850 EIRPSP[T]TPNR II At4g05685, AtGRF1a AtGLR2.6 832 DNMRAP[T]SPPI I – AtGLR2.9 921 SEERFT[T]QPII I At1g78300, AtGRF2b AtGLR3.6 861 GSIRRR[S]SPSA I or II – AtGLR3.7 860 RMERTS[S]MPRA I At1g78300, AtGRF2b Gene Position Peptide Mode Interacting 14-3-3 protein AtGLR1.3 840 FVRSIH[T]SPLD II – AtGLR1.4 850 EIRPSP[T]TPNR II At4g05685, AtGRF1a AtGLR2.6 832 DNMRAP[T]SPPI I – AtGLR2.9 921 SEERFT[T]QPII I At1g78300, AtGRF2b AtGLR3.6 861 GSIRRR[S]SPSA I or II – AtGLR3.7 860 RMERTS[S]MPRA I At1g78300, AtGRF2b The table lists AtGLRs with predicted C-terminal 14-3-3 protein-binding motifs and their position. Conserved residues are in bold, and putatively phosphorylated residues in brackets. The motif modes (I Rxx[S/T]xP; or II, Rxxx[S/T]xP) and experimental evidence for interaction are given. aShin et al. (2011); bChang et al. (2009). View Large From structure to function: implications of plant GLRs GLRs play a role in Ca2+ signaling Although the pivotal role of Ca2+ signaling during plant development and (a)biotic stress responses has been acknowledged and documented for years, the discovery and characterization of the molecular nature of possible Ca2+ transport systems spanned decades and is still ongoing. GLRs were one of the first molecular candidates that were shown to transport Ca2+ (Michard et al., 2011; Vincill et al., 2012), and since have been linked to Ca2+ influx into leaf cells and during plant immune responses (reviewed by Forde and Roberts, 2014), for example. With documented plasma membrane localizations, and considering the fact that mutations in GLR genes affect chemotaxis and immune response pathways, it is plausible that GLRs are involved in plasma membrane-elicited pathways that are triggered by ligands from environmental cues. Following the pathogen–plant interaction studies involving GLR mutants (Kang et al., 2006; Li et al., 2013; Manzoor et al., 2013), subsequent studies tried to understand whether GLRs could recognize microbe-associated molecular patterns (MAMPs), which often trigger the innate immune response pathways. Despite being addressed by an indirect approach when using pharmacology to block GLRs, it was shown that AtGLRs partly participate in the MAMP recognition and consequent apoplastic Ca2+ influxes that are necessary for the activation of downstream signaling events [mitogen-activated protein kinase (MAPK)] related to plant defense (Kwaaitaal et al., 2011). It was further shown that GLRs regulate pollen tube growth by controlling Ca2+ fluxes at the pollen tube tip (Michard et al., 2011; Wudick et al., 2018) and participate in sperm cell guidance (Ortiz-Ramírez et al., 2017). More specifically, it was shown that the amino acid d-serine acts as an agonist for pollen GLRs, which are believed to be involved in pollen tube growth control (Michard et al., 2011). Moreover, AtGLRs are involved in generating the Ca2+ wave in response to aphid feeding in leaves (Vincent et al., 2017). The main physiological role of iGluRs is voltage control. What about GLRs? All documented physiological implications of AtGLRs so far are linked to their capability to permeate Ca2+. Strikingly, in animals, this role is mostly attributed to NMDA receptors, while non-NMDA receptors show a much more diversified ion selectivity (Traynelis et al., 2010). As exemplified before, animal iGluRs have been extensively studied in neurons, highlighting their role in regulating membrane potential and fast excitatory synaptic transmission (Traynelis et al., 2010). In synapses, upon agonist binding, iGluR transmembrane domains change their conformation, opening a pore on the membrane that allows the influx of K+, Na+, and/or Ca2+, depending on the selectivity of the channel (Willard and Koochekpour, 2013). The influx of cations, consequently, depolarizes the membrane, elevating the membrane potential to values closer to the depolarized voltage threshold, which in turn activates voltage-gated channels in the vicinity, eventually generating an action potential (Traynelis et al., 2010). Studies in mouse giant synapses revealed that knocking down the only iGluR expressed in those cells almost totally abolished excitatory post-synaptic currents and delayed the onset of action potential generation (Seol and Kuner, 2015). This work exemplarily showed that iGluRs are the first responders to neurotransmitter-conveyed signals by bringing the membrane potential close to the threshold, thus priming the cell to fire an action potential. Despite being crucial for cell physiology, membrane potential fluctuations have been underappreciated for a very long time (Blackiston et al., 2009; Gallé et al., 2015). Interestingly, observations in plants also point to the importance of membrane potential and changes thereof for plant physiology. For instance, mesophyll cells were shown to change their membrane potential after dark and light transitions (Shabala and Newman, 1999). It was further shown in guard cells that a stimulus-induced membrane depolarization triggered an activation of anion channels, which further depolarized the membrane, resulting in an increase of the osmotic potential and subsequent closure of the stomata (Hedrich, 2012). It is interesting to note that some of the stimuli triggered a Ca2+-mediated response and that changes in the membrane voltage played an essential role in this process, during both the signaling (through voltage-gated channels) and the response (increase of water potential) (Hedrich, 2012). Even though plants do not have such specialized cells as neurons, they do have excitable cells capable of generating and propagating electrical signals (Gallé et al., 2015; Hedrich et al., 2016). In fact, bio-electric phenomena in plants were widely characterized more than a century ago (i.e. Bose, 1913). The closest structure to axons in plants is the phloem, a symplastic tissue, displaying a low-resistance path for electric signaling. During their development, the phloem cells undergo partial programmed cell death resulting in the degeneration of the central vacuole, the nucleus, and the common plastids, thereby creating a low resistance cytoplasm dominated by an electrolyte of ~100 mM K+ and thus more favorable to conduct electrical signals (Gallé et al., 2015). It was recently shown that AtGLRs are crucial to evoke and propagate an electrical signal along the phloem in response to herbivory and that mutations in AtGLR3.1, 3.2, 3.3, and 3.6 caused reduced duration of surface potential changes in Arabidopsis leaves after caterpillar-induced wounding (Mousavi et al., 2013). Strikingly, the propagation of the electrical signal to neighboring leaves was dependent on AtGLR3.3 and 3.6, showing the implication of those genes in conveying the electrical signal over a long distance (Mousavi et al., 2013). Moreover, AtGLR3.5 was shown to block the spatial distribution of those long-distance electrical signals, preventing the signal from being fully systemic. Conversely, in glr3.5 mutants, wound-induced action potentials transversed the entire plant, propagating the electrical signal to non-neighboring leaves (Salvador-Recatalà, 2016). Overall, those results showed that AtGLRs can modulate the shape of the electrical signal and its intensity, and further control its propagation to defined organs, thus providing evidence for a tight regulation of AtGLR-mediated electrical signals that may be as complex as in neurons. Beyond the neurons: conclusions and outlook for a complex family In addition to playing a pivotal role in the central nervous system, iGluRs are also expressed in many peripheral, non-neuronal cells. For instance, in bones, iGluRs were shown to influence the dynamic remodeling of this tissue (Hinoi et al., 2004; Xie et al., 2016). In the pancreas, iGluRs are important for the ion signaling in the islet β-cells that control insulin release (Hinoi et al., 2004), while in kidneys iGluRs stimulate vasodilation in the glomerulus, which may have an impact in water/salt balance, potentially regulating blood pressure (Hogan-Cann and Anderson, 2016). Interestingly, it was shown that peripheral iGluRs act in heteromers different from those in the central nervous system, and that their kinetics, especially of desensitization, appear to be slower (Traynelis et al., 2010). Consequentially, understanding the physiological role of iGluRs outside the nervous system becomes more and more important. It is noteworthy that the characterization of these peripheral iGluRs faces limitations and problems similar to the study of plant GLRs. (i) Peripheral iGluRs seem to be involved in Ca2+ signaling, but the molecular mechanism are not understood, though it has been proposed that they could modulate the basal Ca2+ level in the cell. (ii) The amino acid concentration in the periphery of cells that express those iGluRs is much higher than their half-activation concentration observed in the brain, particularly in red blood cells or lymphocytes that evolve in an environment of constantly high amino acid concentrations (Zhou et al., 2013). So how does the gating of those channels work? (iii) Complex subunit arrangements and multimer formation might occur and alter channel properties. In this review, we discussed that GLRs and iGluRs, although sharing a common structure, display major sequence divergences, especially in key domains such as the receptor, the pore, and the ‘gate’. Recent electrophysiological characterization of PpGLR1, AtGLR3.2, and AtGLR3.3 (Ortiz-Ramírez et al., 2017; Wudick et al., 2018) showed their ligand-independent activity, providing evidence for functional differences between iGluRs and at least these specific GLRs. In addition to the GLR diversity (20 members in Arabidopsis, four clades in higher plants), AtGLRs can apparently undergo various alternative splicing processes at the mRNA level, and have the ability to form clade-overarching functional heteromers (Price et al., 2013; Vincill et al., 2013). The aforementioned differences between iGluRs and plant GLRs make it obvious that more functional data are required to understand this complex channel family profoundly on the cellular level. Generally, functional comparisons between GLRs and iGluRs should be done very carefully, since both families seem to have at least as many things in common as things that separate them. Supplementary data Supplementary data are available at JXB online. Fig. S1. Sequences used to generate the phylogram, gate motif logos, and ATD regions. Acknowledgements JAF’s lab is supported by the US National Science Foundation (MCB 1616437/2016 and MCB1714993/2017) and the University of Maryland. 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doi: 10.1093/jxb/ery160pmid: 29718301

Abstract The universality of calcium as an intracellular messenger depends on the dynamics of its spatial and temporal release from calcium stores. Accumulating evidence over the past two decades supports an essential role for nuclear calcium signalling in the transduction of specific stimuli into cellular responses. This review focuses on mechanisms underpinning changes in nuclear calcium concentrations and discusses what is known so far about the origin of the nuclear calcium signals identified, primarily in the context of microbial symbioses and abiotic stresses. Abiotic stress, biotic stress, calcium, ion channel, nucleus, symbioses Introduction Intracellular calcium regulates a wide variety of cellular processes in eukaryotes (Berridge et al., 2000; White and Broadley, 2003; Dodd et al., 2010). Understanding how such a versatile secondary messenger encodes a specific cellular response requires precise knowledge of the dynamics of its release with both spatial and temporal resolution. Within the same cell, diverse stimuli can induce specific calcium signals varying in amplitude, duration, and frequency, in the cytoplasm or organelles (Hetherington and Brownlee, 2004; McAinsh and Pittman, 2009; Stael et al., 2012; Nomura and Shiina, 2014). Since its first detailed description by the botanist Robert Brown in 1831, the nucleus is now recognized as one of the most elaborate organelles, keeper of the genetic information but also functionally highly dynamic in co-ordinating cellular activities (Meier and Somers, 2011; Meier et al., 2017; Zeng et al., 2017). Over the past two decades, reports of nuclear calcium-dependent processes and nuclear-localized calcium-binding proteins highlight the importance of the spatial location of calcium signals within the nucleus (van Der Luit et al., 1999; Perruc et al., 2004; Ishida et al., 2008; Defalco et al., 2010; Lachaud et al., 2010; Miller et al., 2013; Huang et al., 2017). Despite the demonstration that plant nuclei possess calcium autonomy (Pauly et al., 2000; Mazars et al., 2009), the precise origin of most nuclear calcium signals observed in plant organs or isolated cells is often subject to debate (Table 1), mainly due to the nuclear architecture itself. The nuclear lumen is bounded by the nuclear envelope, which consists of an outer nuclear membrane (ONM) and an inner nuclear membrane (INM). The ONM is contiguous with the membrane of the endoplasmic reticulum (ER) (Staehelin, 1997), the lumen of which serves as a calcium store (Bush et al., 1989; Iwano et al., 2009; Capoen et al., 2011; Bonza et al., 2013). Table 1. Nuclear calcium signals observed in planta Physiological process Elicitor Biological system Species Ca2+ probe Ca2+ signal Ca2+ store References Biotic: root endosymbioses Nod and Myc factors Roots Medicago truncatula, Lotus japonicus RGecoNES- GGecoNLS, NLS-YC2.1, NLS-YC3.6 Cytoplasmic and nuclear Ca2+ oscillation Nuclear envelope lumen Kelner (2018); Sieberer et al. (2009); Krebs et al. (2012) Biotic: plant defence Cryptogein, hairpin, flg22, oligogalac-turonides Leaf cell suspension N. tabacum cv. xanthi and N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic, or nuclear Ca2+ signals ND Lecourieux et al. (2005) Abiotic stress Acidic pH-dependent mechanical stimulation Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Neutral and alkaline pH dependent temperature Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Cold shock Seedlings N. plumbaginifolia Aequorin or nuclear- targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Cold shock Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Wind Seedlings N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Hyperosmotic stress Roots A. thaliana YC3.6NLS or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Huang et al. (2017) Hyperosmotic stress BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Pauly et al. (2001) Hyperosmotic stress Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Abiotic or biotic stress Wounding Leaf N. benthamiana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2017) Sphingolipid BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Xiong et al. (2008); Lachaud et al. (2010) Hormone signalling Jasmonates BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Walter et al. (2007) Ja-Ile BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient nuclear Ca2+ signals ND Walter et al. (2007) l-Glutamate signalling l-Glu Roots A. thaliana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2013) ATP signalling ATP Roots A. thaliana YC3.6NLS, PM-YC3.6-LTI6b, or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Krebs et al. (2012); Costa et al. (2013) Physiological process Elicitor Biological system Species Ca2+ probe Ca2+ signal Ca2+ store References Biotic: root endosymbioses Nod and Myc factors Roots Medicago truncatula, Lotus japonicus RGecoNES- GGecoNLS, NLS-YC2.1, NLS-YC3.6 Cytoplasmic and nuclear Ca2+ oscillation Nuclear envelope lumen Kelner (2018); Sieberer et al. (2009); Krebs et al. (2012) Biotic: plant defence Cryptogein, hairpin, flg22, oligogalac-turonides Leaf cell suspension N. tabacum cv. xanthi and N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic, or nuclear Ca2+ signals ND Lecourieux et al. (2005) Abiotic stress Acidic pH-dependent mechanical stimulation Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Neutral and alkaline pH dependent temperature Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Cold shock Seedlings N. plumbaginifolia Aequorin or nuclear- targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Cold shock Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Wind Seedlings N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Hyperosmotic stress Roots A. thaliana YC3.6NLS or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Huang et al. (2017) Hyperosmotic stress BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Pauly et al. (2001) Hyperosmotic stress Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Abiotic or biotic stress Wounding Leaf N. benthamiana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2017) Sphingolipid BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Xiong et al. (2008); Lachaud et al. (2010) Hormone signalling Jasmonates BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Walter et al. (2007) Ja-Ile BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient nuclear Ca2+ signals ND Walter et al. (2007) l-Glutamate signalling l-Glu Roots A. thaliana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2013) ATP signalling ATP Roots A. thaliana YC3.6NLS, PM-YC3.6-LTI6b, or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Krebs et al. (2012); Costa et al. (2013) ND not determined. View Large Table 1. Nuclear calcium signals observed in planta Physiological process Elicitor Biological system Species Ca2+ probe Ca2+ signal Ca2+ store References Biotic: root endosymbioses Nod and Myc factors Roots Medicago truncatula, Lotus japonicus RGecoNES- GGecoNLS, NLS-YC2.1, NLS-YC3.6 Cytoplasmic and nuclear Ca2+ oscillation Nuclear envelope lumen Kelner (2018); Sieberer et al. (2009); Krebs et al. (2012) Biotic: plant defence Cryptogein, hairpin, flg22, oligogalac-turonides Leaf cell suspension N. tabacum cv. xanthi and N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic, or nuclear Ca2+ signals ND Lecourieux et al. (2005) Abiotic stress Acidic pH-dependent mechanical stimulation Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Neutral and alkaline pH dependent temperature Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Cold shock Seedlings N. plumbaginifolia Aequorin or nuclear- targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Cold shock Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Wind Seedlings N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Hyperosmotic stress Roots A. thaliana YC3.6NLS or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Huang et al. (2017) Hyperosmotic stress BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Pauly et al. (2001) Hyperosmotic stress Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Abiotic or biotic stress Wounding Leaf N. benthamiana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2017) Sphingolipid BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Xiong et al. (2008); Lachaud et al. (2010) Hormone signalling Jasmonates BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Walter et al. (2007) Ja-Ile BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient nuclear Ca2+ signals ND Walter et al. (2007) l-Glutamate signalling l-Glu Roots A. thaliana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2013) ATP signalling ATP Roots A. thaliana YC3.6NLS, PM-YC3.6-LTI6b, or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Krebs et al. (2012); Costa et al. (2013) Physiological process Elicitor Biological system Species Ca2+ probe Ca2+ signal Ca2+ store References Biotic: root endosymbioses Nod and Myc factors Roots Medicago truncatula, Lotus japonicus RGecoNES- GGecoNLS, NLS-YC2.1, NLS-YC3.6 Cytoplasmic and nuclear Ca2+ oscillation Nuclear envelope lumen Kelner (2018); Sieberer et al. (2009); Krebs et al. (2012) Biotic: plant defence Cryptogein, hairpin, flg22, oligogalac-turonides Leaf cell suspension N. tabacum cv. xanthi and N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic, or nuclear Ca2+ signals ND Lecourieux et al. (2005) Abiotic stress Acidic pH-dependent mechanical stimulation Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Neutral and alkaline pH dependent temperature Isolated nuclei from BY2 cells N. tabacum L. cultivar Aequorin Transient nuclear Ca2+ signal Nuclear envelope lumen Xiong et al. (2004) Cold shock Seedlings N. plumbaginifolia Aequorin or nuclear- targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Cold shock Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Wind Seedlings N. plumbaginifolia Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Van der Luit et al. (1999) Hyperosmotic stress Roots A. thaliana YC3.6NLS or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Huang et al. (2017) Hyperosmotic stress BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Pauly et al. (2001) Hyperosmotic stress Root elongation zone A. thaliana GGecoNES- RGecoNLS Transient cytoplasmic and nuclear Ca2+ signals ND Kelner et al. (2018) Abiotic or biotic stress Wounding Leaf N. benthamiana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2017) Sphingolipid BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Xiong et al. (2008); Lachaud et al. (2010) Hormone signalling Jasmonates BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient cytoplasmic or nuclear Ca2+ signals ND Walter et al. (2007) Ja-Ile BY2 cells N. tabacum L. cultivar Aequorin or nuclear-targeted aequorin Transient nuclear Ca2+ signals ND Walter et al. (2007) l-Glutamate signalling l-Glu Roots A. thaliana YC3.6NLS Transient nuclear Ca2+ signals ND Costa et al. (2013) ATP signalling ATP Roots A. thaliana YC3.6NLS, PM-YC3.6-LTI6b, or YC3.6NES Transient cytoplasmic or nuclear Ca2+ signals ND Krebs et al. (2012); Costa et al. (2013) ND not determined. View Large The connection between the nucleoplasm and the cytoplasm is mediated by nucleopore complexes (NPCs) which form on the nuclear envelope at fusion sites between the INM and ONM (Meier et al., 2017). NPCs are large aqueous channels composed of units of ~30 proteins called nucleoporins (NUPs), which are organized with octagonal symmetry (Meier et al., 2017). Ions and molecules of masses lower than 40–60 kDa can freely diffuse through the NPCs, while larger molecules can be actively transported through NPCs (Grünwald and Singer, 2012). This nuclear architecture suggests that there could be multiple routes to generate nuclear calcium signals. Considering the permeability of NPCs, passive diffusion of calcium from the cytoplasm to the nucleoplasm via NPCs appears to be a simple way to generate calcium signals in the nucleoplasm. Alternatively, calcium could be released directly from the nuclear envelope lumen, either into the nucleoplasm via the activation of calcium channels in the INM, or via the activation of calcium channels in the ONM and the subsequent diffusion of calcium via NPCs to the nucleoplasm. Finally, in a ‘controlled-access highway’ scenario, a first cytoplasmic calcium signal could be required to activate nuclear-localized calcium channels directly or indirectly via the action of a secondary component. In this review, I present the most recent progress in nuclear calcium-dependent signalling, especially in the context of microbial symbioses, and I highlight the diversity of nuclear calcium signals observed in various biological contexts and the known function of nuclear calcium signals. Generation of nuclear calcium signals in root symbioses The perception of diffusible nodulation (Nod) and mycorrhizal (Myc) factors released by rhizobial and arbuscular mycorrhizal symbionts, respectively, induces calcium oscillations in and around the nucleus of root hair, epidermal, and cortical cells from host plants (Ehrhardt et al., 1996; Miwa et al., 2006a; Capoen et al., 2011; Sieberer et al., 2012; Genre et al., 2013; Granqvist et al., 2015; Sun et al., 2015). Legume mutants impaired in the generation of these calcium oscillations are defective for both root-nodule and arbuscular-mycorrhizal (AM) symbioses, demonstrating that nuclear calcium oscillations activate endosymbiotic programmes in response to microbial signals (Parniske, 2008; Charpentier and Oldroyd, 2013). Over the past years, forward and reverse genetic screens led to the identification of the nuclear-localized ion channels and the calcium pump required to generate the calcium oscillations. In Medicago truncatula, those are encoded by the calcium pump, a SERCA-type calcium ATPase (MCA8) (Capoen et al., 2011); the potassium-permeable channel, DOES NOT MAKE INFECTIONS 1 (DMI1) (Ané et al., 2004); and the calcium channel, CYCLIC NUCLEOTIDE GATED CHANNEL 15 (CNGC15) (Charpentier et al., 2016). The two channels (DMI1 and CNGC15) interact, implying co-ordinated ion movements (Charpentier et al., 2016). Mathematical modelling predicts that these three components (MCA8, DMI1, and CNGC15) are sufficient to generate nuclear calcium oscillations. In this model, a secondary messenger, hypothetically a cyclic nucleotide, binds to CNGC15, triggering its activation and releasing calcium; DMI1 allows movement of potassium ions to balance the transmembrane charge balance induced by calcium movement, and calcium would be returned to the store by the action of MCA8 (Charpentier et al., 2016). DMI1, MCA8, and CNGC15 localize to both the ONM and INM (Capoen et al., 2011; Charpentier et al., 2016), raising the possibility of calcium oscillations being produced on either side of the nuclear envelope or simultaneously on both sides. The nuclear calcium oscillations induced by symbiotic factors were analysed using microinjected calcium-responsive dyes (Ehrhardt et al., 1996; Wais et al., 2000) or transgenic roots expressing cytoplasmic- or nuclear-localized ‘cameleon’ constructs, which allow Förster resonance energy transfer (FRET)-based measurements (Miwa et al., 2006b; Sieberer et al., 2009). Although useful to assay calcium oscillations in symbiotically defective mutants (Miwa et al., 2006b; Charpentier et al., 2016), or to gain insights into the cellular activation of the nuclear calcium oscillations in vivo (Sieberer et al., 2012), these approaches could not resolve the spatial dynamics of calcium release at the nuclear envelope. Since the development of the first genetically encoded calcium indicators (Shimomura et al., 1962; Knight et al., 1991; Miyawaki et al., 1997; Baird et al., 1999), improved single fluorescent protein-based calcium sensors have been generated (Zhao et al., 2011). The ‘genetically encoded calcium indicators for optical measurement’ (GECOs) can monitor calcium changes with emission at 446 nm (Blue-GECO1), 513 nm (Green-GECO1), or 600 nm (Red-GECO1) (Zhao et al., 2011). By expressing both the Red-GECO1 fused to a nuclear exclusion signal and the Green-GECO1 fused to a nuclear localization signal, the cytoplasmic and nuclear calcium dynamics could be measured in transgenic M. truncatula roots responding to symbiotic factors (Kelner et al., 2018). This showed that symbiotic factor-induced calcium oscillations preferentially start inside the nucleus, demonstrating that nuclear calcium signals can be generated directly in the nucleoplasm by calcium released from the lumen of the nuclear envelope. It further suggests that if the activation of the complex CNCG15–DMI1 requires a secondary messenger, this secondary messenger is either produced inside the nucleus or diffuses into the nucleus via the NPCs. To date, the nature of this secondary messenger is still unknown. Although CNGCs can be gated by cyclic nucleotide monophosphates (cNMPs) (Balagué et al., 2003; Gao et al., 2012, 2016), and adenylyl cyclase activity has been detected in M. sativa root extracts (Carricarte et al., 1988), a requirement for cNMPs in the induction of nuclear calcium oscillation by symbiosis factors remains to be determined. Recent studies may provide clues about the unidentified secondary messenger(s) signalling pathway leading to the activation of nuclear calcium oscillations during initiation of root symbioses. In legume–rhizobial symbioses, Nod factors are perceived at the plasma membrane by a heterocomplex of lysine motif receptor-like kinases including Nod Factor Receptor 1 (NFR1) and NFR5 (called NFP in M. truncatula) (Madsen et al., 2003, 2011; Radutoiu et al., 2003, 2007; Broghammer et al., 2012) (Fig. 1). NFR1 and NFR5 interact with the receptor-like kinase SYMRK (called DMI2 in M. truncatula) (Stracke et al., 2002; Ried et al., 2014) that is essential for induction of calcium oscillations by both Nod and Myc factors (Wais et al., 2000; Genre et al., 2013; Sun et al., 2015). Recently, the receptor like-kinases have been connected to two pathways which could potentially be involved in activation of the the nuclear calcium oscillations. Fig. 1. View largeDownload slide Nodulation factor signal transduction. Perception of rhizobial Nod factors (NFs) at the plasma membrane is mediated by the lysine motif receptor-like kinases, the Lotus japonicus Nod factor receptor 1 (NFR1) and NFR5, and Medicago truncatula LYK3 and NFP. NFR1 and NFR5 associate with the putative co-receptor-like kinase M. truncatula DMI2 and L. japonicus SYMRK. DMI2 interacts with the 3-hydroxy-3-methyl-glutaryl-coenzyme A reductase (HGMR1), which produces mevalonate, an activator of nuclear Ca2+ oscillations. Soybean NFR1 associates with and phosphorylates RGS proteins, which positively regulate nodulation. Mevalonate and the G-protein signalling pathway might be required to stimulate the production of an as yet unidentified secondary messenger. The secondary messenger then activates the CNGC15–DMI1 complex, leading to Ca2+ release in the nucleus. MCA8 pumps Ca2+ back into the lumen of the nuclear envelope. The calcium and calmodulin-dependent kinase (CCaMK) is activated by the nuclear calcium oscillation and transduces the information to downstream components such as the transcription factor CYCLOPS. In addition to the NF-induced nuclear calcium oscillation, the mastoparan analogue, Mas7, potentially activates the production of the symbiotically induced secondary messenger to activate the nuclear-localized calcium channel, as well as having side activation on an as yet identified calcium channel. Fig. 1. View largeDownload slide Nodulation factor signal transduction. Perception of rhizobial Nod factors (NFs) at the plasma membrane is mediated by the lysine motif receptor-like kinases, the Lotus japonicus Nod factor receptor 1 (NFR1) and NFR5, and Medicago truncatula LYK3 and NFP. NFR1 and NFR5 associate with the putative co-receptor-like kinase M. truncatula DMI2 and L. japonicus SYMRK. DMI2 interacts with the 3-hydroxy-3-methyl-glutaryl-coenzyme A reductase (HGMR1), which produces mevalonate, an activator of nuclear Ca2+ oscillations. Soybean NFR1 associates with and phosphorylates RGS proteins, which positively regulate nodulation. Mevalonate and the G-protein signalling pathway might be required to stimulate the production of an as yet unidentified secondary messenger. The secondary messenger then activates the CNGC15–DMI1 complex, leading to Ca2+ release in the nucleus. MCA8 pumps Ca2+ back into the lumen of the nuclear envelope. The calcium and calmodulin-dependent kinase (CCaMK) is activated by the nuclear calcium oscillation and transduces the information to downstream components such as the transcription factor CYCLOPS. In addition to the NF-induced nuclear calcium oscillation, the mastoparan analogue, Mas7, potentially activates the production of the symbiotically induced secondary messenger to activate the nuclear-localized calcium channel, as well as having side activation on an as yet identified calcium channel. One signalling pathway could involve heterotrimeric G-protein complexes made up of Gα, Gβ, and Gγ subunits, that interact with RGS (regulator of G-protein signalling). In soybean, RGS, Gβ, and Gγ positively regulate both nodule formation and induction of expression of early nodulation genes (Choudhury and Pandey, 2013, 2015). RGS interacts with and is phosphorylated by NFR1; the phosphorylated RGS positively regulates nodulation (Fig. 1). In line with this, the G-protein agonist, mastoparan 7 (Mas7), induces expression of early nodulation genes (Pingret et al., 1998; Charron et al., 2004). Mas7 also induces calcium oscillations that were independent of NFP and DMI2 (Sun et al., 2007). The calcium oscillations were blocked by an inhibitor of phospholipase D (PLD). These observations suggest that Mas7 acts downstream of both NFP and the DMI2 receptor-like kinase to activate PLD signalling directly or indirectly. One model, therefore, could be that G-protein signalling downstream of the receptor-like kinase could activate the production of a secondary messenger which triggers nuclear-localized calcium oscillation. However, Mas7-induced nuclear calcium spiking was independent of the potassium ion channel DMI1. Furthermore, the calcium oscillations were observed in the nuclear region but, unlike induction of Nod factor, the oscillations were also observed in the cytoplasm distant from the nucleus (Sun et al., 2007). These differences from Nod factor-induced signalling indicate that the specific symbiotic secondary messenger could have a different or more restricted targeting than Mas7 or that it is less potent than Mas7-induced signalling. A possible alternative signalling pathway for production of the secondary messenger came from work in M. truncatula. The M. truncatula 3-hydroxy 3-methylglutaryl-CoA reductase 1 (HMGR1) was found to interact with the DMI2 receptor-like kinase (Kevei et al., 2007) (Fig. 1). HMGR1 is required both for the M. truncatula–Sinorhizobium meliloti symbiosis, and for the nuclear calcium oscillations induced in response to Nod factor and AM fungal exudates (Kevei et al., 2007; Venkateshwaran et al., 2015). The direct product of HMGR1, mevalonate (MVA), is sufficient to activate nuclear calcium oscillations, and these require the DMI1 potassium channel, placing MVA upstream of activation of the nuclear calcium channel (Venkateshwaran et al., 2015). Surprisingly, MVA-induced nuclear calcium oscillations required NFP and partially depended on DMI2, suggesting that MVA could act in synergy with additional components to generate a secondary messenger that activates the calcium channel. In this scenario, the production of the secondary messenger would require the Nod factor receptors. Considering that NFR1 does phosphorylate RGS to regulate root nodule symbiosis positively, the production of a secondary messenger could require the synergistic activation of both the MVA and G-protein signalling pathway via the receptor complex (Fig. 1). Further studies are required to assess this hypothesis and unravel the exact nature of the signalling component required to activate the nuclear calcium oscillation. Origin of nuclear calcium signals in diverse biological contexts Nuclear calcium signals have been detected using nuclear-localized calcium probes or with isolated nuclei, in response to several stimuli, such as plant defence elicitors, mechanical and abiotic stresses, hormones, or ATP (Table 1). These calcium signals differ in frequency, duration, and amplitude compared with the symbiotic factor-induced calcium oscillations. They mainly show a single transient calcium peak varying in amplitude and duration, depending on the stimuli and experimental system used. This suggests that the mechanism of generating these nuclear calcium signals differs from the symbiotic factor-induced nuclear calcium oscillations. Differences could be due to the activation of distinct ion channels/pumps associated with either the same or a different calcium store. Alternatively, if the same ion channels/pumps and store are to be used, different mechanisms of regulation of calcium flow would be required to explain the production of various nuclear calcium signals. In cell suspension cultures and roots, all the nuclear calcium signals observed are associated with transient cytoplasmic calcium signals, with the exception of jasmonate-isoleucine-induced nuclear calcium signalling. However, in all these analyses, the nuclear and cytoplasmic calcium changes were recorded in independent cell lines expressing either a nuclear-localized or a cytoplasmic calcium probe (van Der Luit et al., 1999; Pauly et al., 2001; Lecourieux et al., 2005; Walter et al., 2007; Xiong et al., 2008; Lachaud et al., 2010; Krebs et al., 2012; Costa et al., 2013, 2017; Huang et al., 2017). The lack of simultaneous analyses of cytoplasmic and nuclear calcium changes make it impossible to ascertain the origin of the nuclear calcium signals observed. Even simple diffusion to the nucleoplasm of cytoplasmic calcium released from diverse calcium stores cannot be excluded. With isolated nuclei, mechanical stretch and increased temperature can generate a specific nuclear calcium release (Xiong et al., 2004). This suggests that, as with symbiotic factor-induced nuclear calcium oscillations, the calcium is likely to be mobilized from a store in the lumen of the nuclear envelope; to date, there are no genetic clues to the nature of ion channels/pumps involved in this calcium release in response to stretch or temperature. However, the activation by both stimuli is controlled differentially upon acidic and alkaline pH, suggesting that multiple calcium mobilizing systems might co-exist on the plant nuclear envelope. In animals, temperature-activated ion channels are typically encoded by transient receptor potential (TRP) ion channels (Hilton et al., 2015), but genome sequence analyses revealed that TRP channels are absent from land plants (Wheeler and Brownlee, 2008); thus other ion channels have to fulfil this role. In contrast, numerous mechanosensitive (MCS) ion channels have been characterized in plants (Hamilton et al., 2015), and one of these, MSL10, localizes to both the plasma membrane and the endoplasmic reticulum (ER) (Veley et al., 2014). Additionally, voltage-dependent activation of calcium channels has been recorded using the membrane from the ER of higher plants (Klüsener et al., 1995; Grygorczyk and Grygorczyk, 1998). Based on a pharmacology study (Klüsener et al., 1995), it was proposed that the voltage-dependent calcium release channel shares properties with an MCS channel. These data demonstrate that an MCS channel and an MCS like-calcium release channel are present in the ER. It has not yet been shown that MCS calcium channels are present in the nuclear envelope, but they are likely candidates for stretch-induced calcium channels on the nuclear membrane. An alternative way to explore whether plant nuclear calcium signals are generated independently from cytoplasmic signals is to deplete cytoplasmic calcium using animal calcium-binding proteins such as parvalbumin or calbindin. Buffering the cytoplasmic calcium with parvalbumin signals did not interfere with the salt stress-induced nuclear calcium signals in Arabidopsis thaliana root cells, demonstrating that the salt stress-induced nuclear calcium release does not originate from cytoplasmic passive diffusion and so is likely to be due to calcium release from ER/nuclear calcium stores (Huang et al., 2017). This result was confirmed by the simultaneous analysis of the salt stress-induced cytoplasmic and nuclear calcium signals in the root elongation zone of A. thaliana, using transformed plants expressing both cytoplasmic-localized Green GECO1.2 and nuclear-localized Red-GECO1.2 (Kelner et al., 2018). Although the nucleoplasmic calcium release occurred 2–3 s after the cytoplasmic signal, the magnitude of the amplitude of the calcium signal suggests that passive calcium diffusion from the cytoplasm is unlikely to be the origin of the salt stress-induced nuclear calcium. Nuclear calcium signals: regulation of gene expression In animals, nuclear calcium regulates activation of kinases (Deisseroth et al., 1998), gene expression (Hardingham et al., 1997; Zhang et al., 2009), neuroprotection (Zhang et al., 2007), memory formation (Kang et al., 2001), and cell proliferation (Rodrigues et al., 2007). By specifically buffering either nuclear or cytoplasmic calcium, or inducing specific nuclear calcium responses, it was demonstrated that nucleoplasmic calcium signals specifically regulate these functions (Hardingham et al., 1997; Rodrigues et al., 2007). A similar approach was used to demonstrate the contribution of nuclear calcium in the expression of plant genes induced by salt and osmotic stresses (Huang et al., 2017). Expressing parvalbumin targeted to nuclei or to the cytoplasm repressed or increased, respectively, the calcium-dependent expression of genes induced by NaCl and sorbitol (Huang et al., 2017). In line with a major role for nuclear calcium signals in regulating gene expression (Lenzoni et al., 2018), several calcium-responsive cis-elements have been identified (Kaplan et al., 2006; Whalley et al., 2011). To modulate gene transcription, calcium can activate calcium-binding and calmodulin-binding transcription factors, which can act directly on gene expression (Reddy et al., 2000; Kim and Kim, 2006; Popescu et al., 2007). Calcium can act indirectly via calcium and/or calcium- and calmodulin-regulated kinases (Schulz et al., 2013; Bender et al., 2018) and calcium-binding ATP/GDP protein (Guan et al., 2013). Several nuclear-localized calcium-binding proteins are required for abiotic or biotic stress responses; these include calmodulins (Choi et al., 2005; Bender et al., 2014), calcium-dependent protein kinases (Dammann et al., 2003; Choi et al., 2005; Ishida et al., 2008; Boudsocq et al., 2010), calcium- and calmodulin-dependent protein kinase (CCaMKs) (Singh and Parniske, 2012), and calcium-binding ATP/GDP protein (Guan et al., 2013). This highlights nuclear calcium as an important component of signal transduction, that can directly modulate or activate nuclear-localized calcium-binding proteins, and thus enable the cell to react to environmental changes as well as symbiotic and pathogenic microbes. Decoding symbiotic calcium oscillations Among the calcium-regulated kinases, CCaMK has the unique feature to bind calcium both directly through its three EF-hand domains and indirectly via binding of calcium/calmodulin (Patil et al., 1995; Swainsbury et al., 2012). CCaMK is the prime decoder of symbiotic factor-induced nuclear calcium oscillations. Autoactive forms of CCaMK are sufficient to induce spontaneous nodulation and the formation of the arbuscular mycorrhizal pre-penetration apparatus (Gleason et al., 2006; Tirichine et al., 2006; Hayashi et al., 2010; Takeda et al., 2012; Miller et al., 2013). Using a combination of modelling and expression of mutant versions, the mechanism of activation of CCaMK by calcium has been identified (Miller et al., 2013). At basal cellular calcium concentrations, calcium bound to EF-hand domains suppresses the kinase activity of CCaMK by stabilizing the inactive state of CCaMK, caused by autophosphorylation promoting hydrogen bonding between the kinase and the calmodulin-binding domains. During calcium oscillations, the calcium concentration rises and promotes calcium/calmodulin binding to CCaMK, blocking the autophosphorylation and fostering phosphorylation of target proteins, the most important being the transcription factor CYCLOPS (Yano et al., 2008). The phosphomimic version of CYCLOPS is sufficient to induce spontaneous nodulation in the absence of the symbionts (Singh et al., 2014). Therefore, one of the prime roles of the symbiotic factor-induced nuclear calcium oscillation is to activate the CCaMK/CYCLOPS complex and thus switch on the endosymbiotic programme by regulated gene expression in root nodules and AM-infected roots (Singh et al., 2014; Pimprikar et al., 2016; Cerri et al., 2017). Concluding remarks and future perspectives The characterization of the nuclear envelope as a calcium store, the identification of specific nuclear calcium signals, the presence of ion channels in the INM, and the identification of nuclear-localized calcium-binding proteins all attest to the fact that calcium release within the nucleus does play an essential role in transducing specific information. To date, only a few nuclear calcium signals have been clearly defined and only one plant nuclear-localized calcium channel has been genetically identified. The nuclear calcium signals observed differ in amplitude, duration, and frequency, raising the question of whether other as yet unidentified calcium channels might co-exist in the nuclear envelope. Notably, mechanical and temperature stimuli induce calcium release from the nuclear envelope lumen, and this suggests the presence of temperature-activated and stretch-activated calcium channels. Interestingly among the MCS family (Basu and Haswell, 2017), two calcium-permeable MCS channels, MCA1 and MCA2, mediate cold-induced cytosolic calcium increase (Mori et al., 2018). This suggests that not only mechanical stimuli but also temperature could target as yet unidentified MCS channel(s) at the nuclear envelope. Future work will be required to determine whether known MCS channels are localized to the nuclear envelope or a new class of MCS channel is involved in the stretch- and temperature-induced nuclear calcium signal observed. Although it has been established that the nuclear envelope is the store for calcium released into the nucleoplasm in response to mechanical stimuli, temperature shock, and symbiotic factors, the calcium store being used for many of the observed nuclear calcium signals (Table 1) still remains to be determined. Additionally, several of these studies on nuclear calcium changes have been done using isolated cultured cells, isolated nuclei, or global analysis at the organ level. To understand the role and the origin of nuclear calcium signals in situ, spatio-temporal resolution will be required to identify which cell type within the tissue is responding to those stimuli. Importantly a combination of high-resolution microscopy and multicolour imaging of multiple subcellular compartments will allow a precise understanding of the origin and dynamics of nuclear calcium signalling in the cell. Increasing the resolution and understanding of which store is being used to produce the nuclear calcium signal will facilitate the identification of novel calcium channels via forward or reverse genetic screens. Finally, it remains to be shown whether diverse calcium signals do encode specific information and how those calcium signatures are decoded to transduce specific processes within the same cell. 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doi: 10.1093/jxb/ery185pmid: 29767757

Abstract Calcium (Ca2+) is among the most important intracellular messengers in living organisms. Understanding the players and dynamics of Ca2+ signalling pathways in plants may help to unravel the molecular basis of their exceptional flexibility to respond and adapt to different stimuli. In the present review, we focus on new tools that have recently revolutionized our view of organellar Ca2+ signalling as well as on the current knowledge regarding the pathways mediating Ca2+ fluxes across intracellular membranes. The contribution of organelles and cellular subcompartments to the orchestrated response via Ca2+ signalling within a cell is also discussed, underlining the fact that one of the greatest challenges in the field is the elucidation of how influx and efflux Ca2+ transporters/channels are regulated in a concerted manner to translate specific information into a Ca2+ signature. Aequorin, calcium-permeable channels, calcium-permeable transporters, Cameleon, endomembranes, genetically encoded calcium indicators, organelle Introduction Changes in Ca2+ levels within plant cells can be considered as hallmarks of a plethora of processes such as growth, differentiation, regulation of stomatal opening, induction of pathogen defence responses, establishment of plant–microbe symbioses, and stress adaptation. Indeed, each of these processes is associated with specific ‘Ca2+ signatures’, arising from variations of Ca2+ concentration characterized by a unique amplitude, frequency, and duration within the cytosol and, in some cases, in a given intracellular compartment (see, for example, Trewavas et al., 1996; Evans et al., 2001; Xiong et al., 2006; Monshausen, 2012; Whalley and Knight, 2013). Thus, the concentration of free Ca2+ in the cytosol ([Ca2+]cyt) is crucial for Ca2+-based signalling. Tight regulation of the [Ca2+]cyt is mandatory because sustained increases above approximately 10–4 M can lead to protein and nucleic acid aggregation and to precipitation of phosphates, thus causing damage to membranes and organelles, ultimately leading to a generalized cytotoxicity. On the other hand, rapid and transient increases of the cytosolic Ca2+ concentration via Ca2+ channels, mediating either Ca2+ influx of the ion from the extracellular milieu or the temporary release of the ion from intracellular stores, have enabled Ca2+ to function as a versatile second messenger in basically all physiological systems (Dodd et al., 2010). Upon stimulation, [Ca2+]cyt increases from ~10–7 M to 10–6 M, relaying an external stimulus to the intracellular milieu and allowing triggering of specific biological responses. A set of proteins that undergo Ca2+ binding-induced conformational changes help the cells to decode the signal by responding to the stimulus-induced increases in [Ca2+]cyt (McAinsh and Pittman, 2009). Intracellular organelles may contribute to the regulation of free Ca2+ homeostasis in the cytosol, since a fast response of Ca2+ levels to environmental cues is ensured by compartmentalization of this cation within the plant cell (Stael et al., 2012; Nomura and Shiina, 2014). In fact, in parallel with the actions of Ca2+ influx and Ca2+ efflux systems across the plasma membrane (PM), Ca2+ sequestration into and release from the intracellular compartments are equally important to maintain the transient nature of Ca2+ signals (Trewavas et al., 1996; Kudla et al., 2010). Ca2+ can be mobilized from storage compartments such as the cell wall/apoplast, the vacuole, and the endoplasmic reticulum (ER), whereas the nucleus, as well as chloroplasts and mitochondria, can also generate intraorganellar Ca2+ signals (Stael et al., 2012). Changes in free [Ca2+] in a given organelle in turn may influence its function. Ca2+-based signalling systems have long been described as oversimplified linear pathways (with a stimulus generating a transient [Ca2+]cyt elevation that in turn leads to a specific response). Since plants can be challenged by several stimuli at the same time—most of which involve changes in [Ca2+]cyt—the final response often implies a complex network of intersecting signal transduction pathways, each specific for a given stimulus. Thus, it is becoming increasingly evident that Ca2+ signalling systems are intrinsically complex networks comprising many interconnected nodes and hubs (Dodd et al., 2010). In the following sections, an overview of the recently developed toolkit to measure time-resolved organellar Ca2+ signalling in intact plants as well as plant cell suspension cultures is provided, along with discussion of the possible Ca2+-permeable channels in the various organelles. We will give special emphasis to the bioenergetic organelles mitochondria and chloroplasts, as well as to peroxisomes and the ER, whereas we advise readers to consult the review on nuclear Ca2+ signalling that is published in the present special issue for information on the participation of the nucleus in the Ca2+ signalling network (Charpentier, 2018). General molecular players of plant Ca2+ signalling In plants, shaping of the Ca2+ signature with defined spatial and temporal characteristics and specificity in Ca2+-based signalling is achieved through the interplay of Ca2+ signatures together with Ca2+-binding proteins that act to decode or interpret increases in the Ca2+ level (e.g. Tang and Luan, 2017). Ca2+ signatures are decoded by Ca2+-binding sensor proteins that act either as primary responders or as signal relays (DeFalco et al., 2009; Zhu, 2016; Tang and Luan, 2017). Ca2+-dependent protein kinases (CDPKs) are primary responders, while calmodulins (CaMs), calmodulin-like proteins (CMLs), and calcineurin B-like proteins (CBLs) are part of the latter group. These Ca2+ sensors trigger a downstream signalling cascade that culminates in changes in gene and protein expression, metabolic activity, and development (see, for example, Lenzoni et al., 2018). Excellent, recent reviews underline the crucial role of the above protein families in global Ca2+ signalling (Ranty et al., 2016; Simeunovic et al., 2016; Tang and Luan, 2017; Kudla et al., 2018), therefore the present review only briefly mentions their contribution to Ca2+signalling. The other crucial proteins for Ca2+ signalling are those involved in the transport of this ion across biological membranes, namely transporters (active or passive) and channels, that mediate flux of ions against or down the electrochemical gradient, respectively. These proteins include Ca2+-ATPases, cation/proton exchangers (CAXs), and cation/Ca2+ exchangers (CCXs) that are emerging players in an increasing range of cellular and physiological functions (Bose et al., 2011; Frei dit Frey et al., 2012; Pittman and Hirschi, 2016; Costa et al., 2017; Corso et al., 2018). Ca2+-permeable channels include members of the glutamate-like receptor family (Swarbreck et al., 2013; Forde and Roberts, 2014; Steinhorst and Kudla, 2014), of cyclic nucleotide-gated channels (CNGCs) (Dietrich et al., 2010; DeFalco et al., 2016), and of mechanosensitive channels (MSCs) (Hamilton et al., 2015). Unconventional Ca2+ transporting annexin1 is also a possible player (Davies, 2014). In addition, organelle-specific channels such as the vacuolar two-pore channel TPC1 (Peiter et al., 2005; Choi et al., 2014, 2017; Kiep et al., 2015; Vincent et al., 2017a; Hedrich et al., 2018) and the mitochondrial calcium uniporter (Wagner et al., 2015a, 2016; Teardo et al., 2017) contribute to shaping Ca2+ signalling. We will refer to the above-mentioned Ca2+-transporting molecules in the context of specific organellar Ca2+ signalling in the following sections. Overview of toolkits to measure plant organellar Ca2+ concentrations in vivo Analysis of Ca2+ dynamics in living plants was initially addressed by using Ca2+-sensitive dyes (e.g. Fura-2, Fura-2 dextran, and Ca2+ Green Dextran) loaded in guard cells, pollen tubes, and root hairs (McAinsh et al., 1995; Ehrhardt et al., 1996; Holdaway-Clarke et al., 1997). The use of these dyes allowed fundamental discoveries to be made, but they present some limitations due to their requirement to be loaded or manually injected and also because their use suffers from low throughput and variability, and is prone to artefacts. Hence, we feel comfortable to say that analysis of Ca2+ dynamics in living plants was revolutionized by the introduction of the genetically encoded Ca2+ indicators (GECIs) (Pérez Koldenkova and Nagai, 2013) that permitted non-invasive monitoring of free Ca2+ levels, enabling real-time, spatially and temporally resolved imaging of Ca2+ levels in different cell types and organisms, and even in specific subcellular compartments by specific targeting of GECIs to organelles (Stael et al., 2012; Costa and Kudla, 2015). Furthermore, the possibility to calibrate GECIs may allow information on absolute concentrations for different ions to be obtained (see, for example, Lanquar et al., 2014 for Zn2+ and Waadt et al., 2017 for Ca2+). The first subcompartmental (cytosolic) GECI, exploitable for in vivo measurements, was obtained for aequorin in plants (Knight et al., 1991). A year later, aequorin was specifically expressed in animal mitochondria via fusion with the signal sequence-encoding part of a mitochondria-located protein. This study revealed for the first time that mammalian mitochondria can accumulate high concentrations of Ca2+ upon stimulation of the cells with histamine, an agonist of the inositol triphosphate receptor located in the ER (Rizzuto et al., 1992). Following these studies, this methodology became widely accepted as a general tool to measure organellar Ca2+ changes in the animal field (Brini et al., 1999; Rudolf et al., 2003; Ottolini et al., 2014; Bagur and Hajnoczky, 2017) and to establish the presence of high Ca2+ concentration microdomains that are generated at the ER–mitochondria contact site level (Rizzuto et al., 1993, 1998). In plants, thus far the two main Ca2+ indicators used are aequorin and Cameleon (Knight and Knight, 1995; Mithöfer and Mazars, 2002; Costa and Kudla, 2015) and, importantly, both display a binding affinity for Ca2+ that renders them useful to detect changes in Ca2+ concentrations in the ranges that occur physiologically (Palmer and Tsien, 2006). As mentioned above, the first GECIs to be developed were the aequorin-based probes, which allowed monitoring of Ca2+ dynamics by photon emission measurements in transformed plants after reconstitution of the aequorin holoenzyme with the exogenously applied prosthetic group coelenterazine (Knight et al., 1991, 1992; Sai and Johnson, 2002; Logan and Knight, 2003). It has been an extraordinary tool to determine the Ca2+ dynamics triggered by different stimuli at the level of cell populations or entire plants, forming the basis of our understanding of the in vivo dynamics of free Ca2+ in plants. Since aequorin is largely insensitive to variations of pH and Mg2+ (Brini, 2008), it can be used as a reliable sensor to monitor [Ca2+] changes even in organelles or subcompartments with acidic pH. Furthermore, its bioluminescent properties, high signal-to-noise ratio, and lack of damaging excitation light make it an excellent tool to measure Ca2+ levels in chlorophyll-containing tissues even for long time intervals (Martí et al., 2013). Aequorin-based sensors are available for different plant organelles, such as the vacuole (Knight et al., 1996), the nucleus (van Der Luit et al., 1999), the Golgi apparatus (Ordenes et al., 2012), mitochondria (Logan and Knight, 2003), and plastids/chloroplasts (Johnson et al., 1995; Mehlmer et al., 2012; Sello et al., 2016). Concerning these latter organelles, aequorin chimeras have been targeted to the different chloroplast subcompartments, namely the stroma (Johnson et al., 1995; Sai and Johnson, 2002), the outer and inner membranes of the envelope (Mehlmer et al., 2012), and the thylakoid lumen and membrane (Sello et al., 2018) (Table 1). Aequorin was also targeted to the apoplastic space (Gao et al., 2004). Moreover, the development of novel bioluminescence resonance energy transfer (BRET)-based green fluorescent protein (GFP)–aequorin reporters, initially designed for Ca2+ imaging in animal cells (Baubet et al., 2000; Rogers et al., 2005), has overcome one of the major limitations of aequorin (i.e. its low amount of emitted light), thus allowing visualization of Ca2+ signals propagating over long distances in intact plants (Xiong et al., 2014). Table 1. Summary of available genetically encoded Ca2+ indicators used in plants Name Version Type Peaks of excitation/ emission (mn) In vitro KDfor Ca2+ Subcellular localization References Cameleon YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol and nucleus Nagai et al. (2004); Mori et al. (2006) NES-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol Krebs et al. (2012) NLS-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Krebs et al. (2012) NUP-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Costa et al. (2017) 4mt-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Mitochondria Loro et al. (2012) PM-YC3.6-LTI6b Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Plasma membrane Krebs et al. (2012); Iwano et al. (2015) 2Bam4-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Chloroplasts and plastids Loro et al. (2016) Nano65 Ratiometric CFP/cpVenus Ex 440/Em 480/530 65 nM Cytosol and nucleus Horikawa et al. (2010); Choi et al. (2014) SP-YC4.6-ER Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Endoplasmic reticulum Nagai et al. (2004); Iwano et al. (2009); Tian et al. (2014) 2Bam4-YC4.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Chloroplasts and plastids Loro et al. (2016) 4mt-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Mitochondria Loro et al. (2013) D3cpv-KVK-SKL Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Peroxisomes Palmer et al. (2006); Costa et al. (2010) TP-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Tonoplast Krebs et al. (2012 CRT-D4ER Ratiometric CFP/Citrine Ex 440/Em 480/530 195 μM Endoplasmic reticulum Palmer et al. (2006); Bonza et al. (2013) Twitch Twitch 3 Ratiometric CFP/cpCit174 Ex 440/Em 480/530 250 nM Cytosol and nucleus Thestrup et al. (2014); Waadt et al. (2017) Geco R-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Cytosol and nucleus Zhao et al. (2011); Ngo et al. (2014); Keinath et al. (2015) NR-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Nucleus Zhao et al. (2011); Kelner et al. (2018) NR-Geco1.2 Intensiometric mApple Ex 561/Em 600 1.2 μM Nucleus Wu et al. (2013); Kelner et al. (2018) CG-Geco1 Intensiometric cpGFP Ex 488/Em 515 749 nM Cytosol Zhao et al. (2011); Kelner et al. (2018) CG-Geco1.2 Intensiometric cpGFP Ex 488/Em 515 1.15 μM Cytosol Zhao et al. (2011); Kelner et al. (2018) R-Geco1-mTurquoise Ratiometric mApple/ mTurqouise Ex 405/561/Em 480/600 NA Cytosol and nucleus Waadt et al. (2017) GCaMP GCaMP3 Intensiometric cpGFP Ex 488/Em 515 542 nM Cytosol and nucleus Vincent et al. (2017a, b) GCaMP6f Intensiometric cpGFP Ex 488/Em 515 375 nM Cytosol and nucleus Waadt et al. (2017) GCaMP6s Intensiometric cpGFP Ex 488/Em 515 144 nM Cytosol and nucleus Liu et al. (2017) MatryoshCaMP6s Ratiometric cpGFP/ LSSmOrange Ex 440/Em 515/600 197 nM Cytosol and nucleus Ast et al. (2017) Case Case12 Intensiometric cpGFP Ex 488/Em 515 1 μM Cytosol and nucleus Souslova et al. (2007); Zhu et al. (2013) Aequorin Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol and nucleus Knight et al. (1991) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus van Der Luit et al. (1999) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast stroma Johnson et al. (1995) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Logan and Knight (2003) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Golgi Ordenes et al. (2012) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Vacuole/tonoplast Knight et al. (1996) YFP–aequorin CYA Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol Mehlmer et al. (2012) NYA Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus Mehlmer et al. (2012) YA Bioluminescence No Ex/Em 465 7.2–13 μM Plasma membrane Mehlmer et al. (2012) CHYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast/plastid stroma Mehlmer et al. (2012); Sello et al. (2016) MYA Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Mehlmer et al. (2012) OEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast outer envelope Mehlmer et al. (2012); Sello et al. (2016) IEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast inner envelope Mehlmer et al. (2012) TL-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid lumen Sello et al. (2018) TM-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid membrane Sello et al. (2018) GFP5–aequorin pchitGFP5:AQ Bioluminescence No Ex/Em 465 NA Apoplast Gao et al. (2004) GFP–aequorin G5A Bioluminescence resonance energy transfer No Ex/Em 515 NA Cytosol and nucleus Baubet et al. (2000); Xiong et al. (2014) Name Version Type Peaks of excitation/ emission (mn) In vitro KDfor Ca2+ Subcellular localization References Cameleon YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol and nucleus Nagai et al. (2004); Mori et al. (2006) NES-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol Krebs et al. (2012) NLS-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Krebs et al. (2012) NUP-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Costa et al. (2017) 4mt-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Mitochondria Loro et al. (2012) PM-YC3.6-LTI6b Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Plasma membrane Krebs et al. (2012); Iwano et al. (2015) 2Bam4-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Chloroplasts and plastids Loro et al. (2016) Nano65 Ratiometric CFP/cpVenus Ex 440/Em 480/530 65 nM Cytosol and nucleus Horikawa et al. (2010); Choi et al. (2014) SP-YC4.6-ER Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Endoplasmic reticulum Nagai et al. (2004); Iwano et al. (2009); Tian et al. (2014) 2Bam4-YC4.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Chloroplasts and plastids Loro et al. (2016) 4mt-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Mitochondria Loro et al. (2013) D3cpv-KVK-SKL Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Peroxisomes Palmer et al. (2006); Costa et al. (2010) TP-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Tonoplast Krebs et al. (2012 CRT-D4ER Ratiometric CFP/Citrine Ex 440/Em 480/530 195 μM Endoplasmic reticulum Palmer et al. (2006); Bonza et al. (2013) Twitch Twitch 3 Ratiometric CFP/cpCit174 Ex 440/Em 480/530 250 nM Cytosol and nucleus Thestrup et al. (2014); Waadt et al. (2017) Geco R-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Cytosol and nucleus Zhao et al. (2011); Ngo et al. (2014); Keinath et al. (2015) NR-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Nucleus Zhao et al. (2011); Kelner et al. (2018) NR-Geco1.2 Intensiometric mApple Ex 561/Em 600 1.2 μM Nucleus Wu et al. (2013); Kelner et al. (2018) CG-Geco1 Intensiometric cpGFP Ex 488/Em 515 749 nM Cytosol Zhao et al. (2011); Kelner et al. (2018) CG-Geco1.2 Intensiometric cpGFP Ex 488/Em 515 1.15 μM Cytosol Zhao et al. (2011); Kelner et al. (2018) R-Geco1-mTurquoise Ratiometric mApple/ mTurqouise Ex 405/561/Em 480/600 NA Cytosol and nucleus Waadt et al. (2017) GCaMP GCaMP3 Intensiometric cpGFP Ex 488/Em 515 542 nM Cytosol and nucleus Vincent et al. (2017a, b) GCaMP6f Intensiometric cpGFP Ex 488/Em 515 375 nM Cytosol and nucleus Waadt et al. (2017) GCaMP6s Intensiometric cpGFP Ex 488/Em 515 144 nM Cytosol and nucleus Liu et al. (2017) MatryoshCaMP6s Ratiometric cpGFP/ LSSmOrange Ex 440/Em 515/600 197 nM Cytosol and nucleus Ast et al. (2017) Case Case12 Intensiometric cpGFP Ex 488/Em 515 1 μM Cytosol and nucleus Souslova et al. (2007); Zhu et al. (2013) Aequorin Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol and nucleus Knight et al. (1991) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus van Der Luit et al. (1999) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast stroma Johnson et al. (1995) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Logan and Knight (2003) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Golgi Ordenes et al. (2012) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Vacuole/tonoplast Knight et al. (1996) YFP–aequorin CYA Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol Mehlmer et al. (2012) NYA Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus Mehlmer et al. (2012) YA Bioluminescence No Ex/Em 465 7.2–13 μM Plasma membrane Mehlmer et al. (2012) CHYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast/plastid stroma Mehlmer et al. (2012); Sello et al. (2016) MYA Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Mehlmer et al. (2012) OEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast outer envelope Mehlmer et al. (2012); Sello et al. (2016) IEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast inner envelope Mehlmer et al. (2012) TL-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid lumen Sello et al. (2018) TM-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid membrane Sello et al. (2018) GFP5–aequorin pchitGFP5:AQ Bioluminescence No Ex/Em 465 NA Apoplast Gao et al. (2004) GFP–aequorin G5A Bioluminescence resonance energy transfer No Ex/Em 515 NA Cytosol and nucleus Baubet et al. (2000); Xiong et al. (2014) The in vitro KD values for Ca2+ of the different sensors are those reported in the original works. For the bioluminescent aequorin sensors, the reported in vitro KD values are 13 μM (Kendall et al., 1992) and 7.2 μM (Brini et al., 1995). Other available recently generated Arabidopsis lines expressing GECO variants of Ca2+ sensors are reported in Waadt et al. (2017). View Large Table 1. Summary of available genetically encoded Ca2+ indicators used in plants Name Version Type Peaks of excitation/ emission (mn) In vitro KDfor Ca2+ Subcellular localization References Cameleon YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol and nucleus Nagai et al. (2004); Mori et al. (2006) NES-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol Krebs et al. (2012) NLS-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Krebs et al. (2012) NUP-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Costa et al. (2017) 4mt-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Mitochondria Loro et al. (2012) PM-YC3.6-LTI6b Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Plasma membrane Krebs et al. (2012); Iwano et al. (2015) 2Bam4-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Chloroplasts and plastids Loro et al. (2016) Nano65 Ratiometric CFP/cpVenus Ex 440/Em 480/530 65 nM Cytosol and nucleus Horikawa et al. (2010); Choi et al. (2014) SP-YC4.6-ER Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Endoplasmic reticulum Nagai et al. (2004); Iwano et al. (2009); Tian et al. (2014) 2Bam4-YC4.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Chloroplasts and plastids Loro et al. (2016) 4mt-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Mitochondria Loro et al. (2013) D3cpv-KVK-SKL Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Peroxisomes Palmer et al. (2006); Costa et al. (2010) TP-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Tonoplast Krebs et al. (2012 CRT-D4ER Ratiometric CFP/Citrine Ex 440/Em 480/530 195 μM Endoplasmic reticulum Palmer et al. (2006); Bonza et al. (2013) Twitch Twitch 3 Ratiometric CFP/cpCit174 Ex 440/Em 480/530 250 nM Cytosol and nucleus Thestrup et al. (2014); Waadt et al. (2017) Geco R-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Cytosol and nucleus Zhao et al. (2011); Ngo et al. (2014); Keinath et al. (2015) NR-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Nucleus Zhao et al. (2011); Kelner et al. (2018) NR-Geco1.2 Intensiometric mApple Ex 561/Em 600 1.2 μM Nucleus Wu et al. (2013); Kelner et al. (2018) CG-Geco1 Intensiometric cpGFP Ex 488/Em 515 749 nM Cytosol Zhao et al. (2011); Kelner et al. (2018) CG-Geco1.2 Intensiometric cpGFP Ex 488/Em 515 1.15 μM Cytosol Zhao et al. (2011); Kelner et al. (2018) R-Geco1-mTurquoise Ratiometric mApple/ mTurqouise Ex 405/561/Em 480/600 NA Cytosol and nucleus Waadt et al. (2017) GCaMP GCaMP3 Intensiometric cpGFP Ex 488/Em 515 542 nM Cytosol and nucleus Vincent et al. (2017a, b) GCaMP6f Intensiometric cpGFP Ex 488/Em 515 375 nM Cytosol and nucleus Waadt et al. (2017) GCaMP6s Intensiometric cpGFP Ex 488/Em 515 144 nM Cytosol and nucleus Liu et al. (2017) MatryoshCaMP6s Ratiometric cpGFP/ LSSmOrange Ex 440/Em 515/600 197 nM Cytosol and nucleus Ast et al. (2017) Case Case12 Intensiometric cpGFP Ex 488/Em 515 1 μM Cytosol and nucleus Souslova et al. (2007); Zhu et al. (2013) Aequorin Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol and nucleus Knight et al. (1991) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus van Der Luit et al. (1999) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast stroma Johnson et al. (1995) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Logan and Knight (2003) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Golgi Ordenes et al. (2012) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Vacuole/tonoplast Knight et al. (1996) YFP–aequorin CYA Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol Mehlmer et al. (2012) NYA Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus Mehlmer et al. (2012) YA Bioluminescence No Ex/Em 465 7.2–13 μM Plasma membrane Mehlmer et al. (2012) CHYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast/plastid stroma Mehlmer et al. (2012); Sello et al. (2016) MYA Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Mehlmer et al. (2012) OEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast outer envelope Mehlmer et al. (2012); Sello et al. (2016) IEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast inner envelope Mehlmer et al. (2012) TL-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid lumen Sello et al. (2018) TM-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid membrane Sello et al. (2018) GFP5–aequorin pchitGFP5:AQ Bioluminescence No Ex/Em 465 NA Apoplast Gao et al. (2004) GFP–aequorin G5A Bioluminescence resonance energy transfer No Ex/Em 515 NA Cytosol and nucleus Baubet et al. (2000); Xiong et al. (2014) Name Version Type Peaks of excitation/ emission (mn) In vitro KDfor Ca2+ Subcellular localization References Cameleon YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol and nucleus Nagai et al. (2004); Mori et al. (2006) NES-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Cytosol Krebs et al. (2012) NLS-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Krebs et al. (2012) NUP-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Nucleus Costa et al. (2017) 4mt-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Mitochondria Loro et al. (2012) PM-YC3.6-LTI6b Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Plasma membrane Krebs et al. (2012); Iwano et al. (2015) 2Bam4-YC3.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 250 nM Chloroplasts and plastids Loro et al. (2016) Nano65 Ratiometric CFP/cpVenus Ex 440/Em 480/530 65 nM Cytosol and nucleus Horikawa et al. (2010); Choi et al. (2014) SP-YC4.6-ER Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Endoplasmic reticulum Nagai et al. (2004); Iwano et al. (2009); Tian et al. (2014) 2Bam4-YC4.6 Ratiometric CFP/cpVenus Ex 440/Em 480/530 58 nM/14.4 μM Chloroplasts and plastids Loro et al. (2016) 4mt-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Mitochondria Loro et al. (2013) D3cpv-KVK-SKL Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Peroxisomes Palmer et al. (2006); Costa et al. (2010) TP-D3cpv Ratiometric CFP/cpVenus Ex 440/Em 480/530 600 nM Tonoplast Krebs et al. (2012 CRT-D4ER Ratiometric CFP/Citrine Ex 440/Em 480/530 195 μM Endoplasmic reticulum Palmer et al. (2006); Bonza et al. (2013) Twitch Twitch 3 Ratiometric CFP/cpCit174 Ex 440/Em 480/530 250 nM Cytosol and nucleus Thestrup et al. (2014); Waadt et al. (2017) Geco R-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Cytosol and nucleus Zhao et al. (2011); Ngo et al. (2014); Keinath et al. (2015) NR-Geco1 Intensiometric mApple Ex 561/Em 600 482 nM Nucleus Zhao et al. (2011); Kelner et al. (2018) NR-Geco1.2 Intensiometric mApple Ex 561/Em 600 1.2 μM Nucleus Wu et al. (2013); Kelner et al. (2018) CG-Geco1 Intensiometric cpGFP Ex 488/Em 515 749 nM Cytosol Zhao et al. (2011); Kelner et al. (2018) CG-Geco1.2 Intensiometric cpGFP Ex 488/Em 515 1.15 μM Cytosol Zhao et al. (2011); Kelner et al. (2018) R-Geco1-mTurquoise Ratiometric mApple/ mTurqouise Ex 405/561/Em 480/600 NA Cytosol and nucleus Waadt et al. (2017) GCaMP GCaMP3 Intensiometric cpGFP Ex 488/Em 515 542 nM Cytosol and nucleus Vincent et al. (2017a, b) GCaMP6f Intensiometric cpGFP Ex 488/Em 515 375 nM Cytosol and nucleus Waadt et al. (2017) GCaMP6s Intensiometric cpGFP Ex 488/Em 515 144 nM Cytosol and nucleus Liu et al. (2017) MatryoshCaMP6s Ratiometric cpGFP/ LSSmOrange Ex 440/Em 515/600 197 nM Cytosol and nucleus Ast et al. (2017) Case Case12 Intensiometric cpGFP Ex 488/Em 515 1 μM Cytosol and nucleus Souslova et al. (2007); Zhu et al. (2013) Aequorin Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol and nucleus Knight et al. (1991) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus van Der Luit et al. (1999) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast stroma Johnson et al. (1995) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Logan and Knight (2003) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Golgi Ordenes et al. (2012) Aequorin Bioluminescence No Ex/Em 465 7.2–13 μM Vacuole/tonoplast Knight et al. (1996) YFP–aequorin CYA Bioluminescence No Ex/Em 465 7.2–13 μM Cytosol Mehlmer et al. (2012) NYA Bioluminescence No Ex/Em 465 7.2–13 μM Nucleus Mehlmer et al. (2012) YA Bioluminescence No Ex/Em 465 7.2–13 μM Plasma membrane Mehlmer et al. (2012) CHYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast/plastid stroma Mehlmer et al. (2012); Sello et al. (2016) MYA Bioluminescence No Ex/Em 465 7.2–13 μM Mitochondria Mehlmer et al. (2012) OEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast outer envelope Mehlmer et al. (2012); Sello et al. (2016) IEYA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast inner envelope Mehlmer et al. (2012) TL-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid lumen Sello et al. (2018) TM-YA Bioluminescence No Ex/Em 465 7.2–13 μM Chloroplast thylakoid membrane Sello et al. (2018) GFP5–aequorin pchitGFP5:AQ Bioluminescence No Ex/Em 465 NA Apoplast Gao et al. (2004) GFP–aequorin G5A Bioluminescence resonance energy transfer No Ex/Em 515 NA Cytosol and nucleus Baubet et al. (2000); Xiong et al. (2014) The in vitro KD values for Ca2+ of the different sensors are those reported in the original works. For the bioluminescent aequorin sensors, the reported in vitro KD values are 13 μM (Kendall et al., 1992) and 7.2 μM (Brini et al., 1995). Other available recently generated Arabidopsis lines expressing GECO variants of Ca2+ sensors are reported in Waadt et al. (2017). View Large The application of ratiometric Ca2+ reporter proteins that are based on combinations of GFP-related proteins (Cameleons) has greatly advanced the spatio-temporal resolution and sensitivity of Ca2+ signalling studies. Cameleons are Förster resonance energy transfer (FRET)-based indicator proteins, which harbour cyan and yellow fluorescent proteins (CFP and YFP, or spectral variants thereof) linked together by the Ca2+-binding protein CaM and the CaM-binding peptide M13 (Costa and Kudla, 2015). Binding of Ca2+ to each of the four helix–loop–helix structures of the EF-hand motifs present in CaM (one Ca2+ ion/EF-hand motif) leads to a conformational change resulting in a reduced distance between CFP and YFP and an increase in FRET. FRET, and thus the [Ca2+] increases, can be conveniently measured by the increase in the ratio between the emission intensity of  YFP and CFP upon CFP excitation (Miyawaki et al., 1997). Since the Ca2+ recordings with such ratiometric proteins completely rely on ratio shifts, these measurements are not influenced by the actual cellular expression level of the indicators and can also correct for focus changes. Cameleon sensors are available for different intracellular compartments and even for simultaneous measurement of Ca2+ dynamics in different subcellular compartments (Krebs et al., 2012; Costa and Kudla, 2015). Other GFP-based Ca2+ biosensors, such as, for example, Case12, GCaMP3, GCaMP6 (Zhu et al., 2014; Liu et al., 2017; Vincent et al., 2017a, b), as well as the green and red variant of GECO1 (G-Geco1 and R-Geco1), have also been successfully applied to measure real-time in vivo changes of Ca2+ in the cytosol and nucleus (Ngo et al., 2014; Keinath et al., 2015; Waadt et al., 2017; Kelner et al., 2018). Furthermore, a strategy of a novel, dual-FP (fluorescent protein) biosensor with large dynamic range based on employment of a single FP-cassette that nests a stable reference FP (large Stokes shift LSSmOrange) within a reporter FP (circularly permuted GFP) has been recently set up (Ast et al., 2017). This strategy has been applied to obtain a novel probe from GCaMP6 (Ast et al., 2017). The R-GECO1 and GCaMP biosensors were found to exhibit a significantly higher signal change compared with Cameleon YC3.6 in response to several stimuli (Keinath et al., 2015; Kleist et al., 2017), and the high fluorescent yield of GCaMPs renders these single-FP Ca2+ sensors particularly suited to whole-tissue imaging, which is often required in studies of plant biotic interactions (Vincent et al., 2017b). In addition, the use of red-shifted sensors opens the way to apply distinct Ca2+ probes localized to different compartments simultaneously, for example together with Cameleon or other GFP-based GECIs, as recently reported by Kelner and colleagues who monitored the cytosolic and nuclear Ca2+ dynamics by simultaneously expressing the CG-Geco1 and NR-Geco1 sensors (Kelner et al., 2018). On the other hand, it has to be noted that single-FP sensors cannot quantify absolute [Ca2+] as simply as FRET-based sensors. However, for example in the case of GCaMP6, it has been experimentally determined that exciting GCaMP6 at 410 nm leads to fluorescence emission, which is not Ca2+ dependent. As a consequence, the ratio between 474 nm and 410 nm excitation wavelengths is proportional to [Ca2+] (Patron et al., 2014). In summary, in spite of the wide variety of currently available GECIs, for the moment aequorin-based probes still remain the method of choice when accurate quantifications of Ca2+ levels are needed (Ottolini et al., 2014). As mentioned above, organelle-targeted, bioluminescent or fluorescent GECIs, summarized in Fig. 1, have greatly advanced the field of organellar Ca2+ signalling in both animals and plants. In the following sections, results obtained exploiting these various probes in different organelles will be reported and compared. Table 1 summarizes the measured affinities for Ca2+ (expressed as KD values) of different, organelle-targeted GECIs. Fig. 1. View largeDownload slide Overview of organelle-targeted genetically encoded Ca2+ indicators. Available probes based on aequorin (Knight et al., 1991; Johnson et al., 1995; Knight et al., 1996; van Der Luit et al., 1999; Logan and Knight, 2003; Mehlmer et al., 2012; Ordenes et al., 2012; Sello et al., 2018), on Cameleon (Mori et al., 2006; Costa et al., 2010; Krebs et al., 2012; Loro et al., 2012, 2013, 2016; Bonza et al., 2013), on R-Geco1 (Ngo et al., 2014; Keinath et al., 2015; Waadt et al., 2017), and on GCaMP3 or 6 (Ast et al., 2017; Liu et al., 2017; Vincent et al., 2017a, b; Waadt et al., 2017) used in different cell compartments are summarized. See text and Table 2 for details regarding the Ca2+ concentration values within the organelles, and Table 1 for the affinities of the above probes for Ca2+. Fig. 1. View largeDownload slide Overview of organelle-targeted genetically encoded Ca2+ indicators. Available probes based on aequorin (Knight et al., 1991; Johnson et al., 1995; Knight et al., 1996; van Der Luit et al., 1999; Logan and Knight, 2003; Mehlmer et al., 2012; Ordenes et al., 2012; Sello et al., 2018), on Cameleon (Mori et al., 2006; Costa et al., 2010; Krebs et al., 2012; Loro et al., 2012, 2013, 2016; Bonza et al., 2013), on R-Geco1 (Ngo et al., 2014; Keinath et al., 2015; Waadt et al., 2017), and on GCaMP3 or 6 (Ast et al., 2017; Liu et al., 2017; Vincent et al., 2017a, b; Waadt et al., 2017) used in different cell compartments are summarized. See text and Table 2 for details regarding the Ca2+ concentration values within the organelles, and Table 1 for the affinities of the above probes for Ca2+. The main Ca2+ storage compartment: the vacuole Plant vacuoles are large organelles with a diameter of 20–40 µm, occupying 80–90% of the cell volume in mature plant cells. Rather than being just the plant counterparts of animal lysosomes, they actually fulfil many different roles, such as the temporary storage of primary metabolites, or the permanent accumulation of secondary metabolites, including potentially toxic compounds (Kruger and Schumacher, 2017; Shimada et al., 2018). Similarly to animal lysosomes, they store high concentrations of Ca2+ and Na+ (Peiter, 2011). In the central vacuole, the concentration of Ca2+ can reach values as high as 50 mM. Nevertheless, most of it is present in bound form (the free vacuolar Ca2+ concentration ranges from 0.2 mM to 1–5 mM; Table 2) (Felle, 1989) and therefore is not readily available for Ca2+ signalling (Conn and Gilliham, 2010). However, vacuolar Ca2+ might indirectly affect the signalling by influencing the activity of ion transporters localized on the vacuolar membrane (Peiter, 2011). In an early pioneering study, the targeting of an aequorin probe to the cytosolic face of the vacuolar membrane provided evidence for the participation of the vacuole in Ca2+ signalling activated by cold (Knight et al., 1996). More recently, a Cameleon-based tonoplast-targeted sensor has also been generated, but, besides labelling the tonoplast, it was also present in the cytosol, restricting the actual usefulness of such a sensor (Krebs et al., 2012). A Ca2+ sensor localized to the vacuolar lumen would be extremely useful for the understanding of those Ca2+ signalling events in which a contribution from the internal stores has been hypothesized (see also below). Unfortunately, the low pH and the high Ca2+ concentration of the vacuolar lumen make difficult to use the currently available GECIs to monitor Ca2+ dynamics efficiently inside this organelle. Nevertheless, in mammalian cells, a more acidic pH-resistant Ca2+ sensor, GEM-GECO1 (Horikawa, 2015), has been successfully targeted to the lysosomal lumen (Albrecht et al., 2015), making it possible to study in vivo lysosomal Ca2+ dynamics triggered by histamine treatment despite the acidic lumen pH. However, the probe was still pH sensitive, making the analysis and interpretation of the data difficult and requiring a tricky pH calibration. An important breakthrough is probably the recent identification of a new pH-resistant GFP (Shinoda et al., 2018) that will probably allow the development of new sensors suitable for acidic compartments. Table 2. Summary of measured and estimated Ca2+ concentrations at resting conditions in the different subcellular compartments of plant cells Subcellular compartment Range/estimation of resting free Ca2+ Method of measurement References Apoplast 330 µM–1 mM X-ray microanalysis Conn et al. (2011) Cytosol 50–100 nM Aequorin, Cameleon (YC3.6), R-Geco1 Knight and Knight (1995); Logan and Knight (2003); Wagner et al. (2015a); Waadt et al. (2017) Nucleus 100 nM Aequorin Van der Luit et al. (1999); Mithöfer and Mazars (2002) Mitochondria matrix 100–600 nM Aequorin, Cameleon (YC3.6), Fura-2 Zottini and Zannoni (1993); Logan and Knight (2003); Wagner et al. (2015a) Chloroplast stroma 100–200 nM Aequorin, Cameleon (YC3.6) Mehlmer et al. (2012); Nomura et al. (2012); Sello et al. (2016); Loro et al. (2016) Thylakoid lumen 500 nM Aequorin Sello et al. (2018) Amyloplast/plastid stroma 80–100 nM Aequorin, Cameleon (YC3.6) Sello et al. (2016); Loro et al. (2016) Vacuolar lumen 200 µM–50 mM X-ray microanalysis Conn and Gilliham (2010); Conn et al. (2011) Endoplasmic reticulum lumen 50–500 µM Cameleon (CRT-D4ER) Iwano et al. (2009); Bonza et al. (2013) Golgi lumen 700 nM Aequorin Ordenes et al. (2012) Peroxisome lumen 150 nM–2 µM Cameleon (D3cpv-KVK-SKL) Costa et al. (2010) Subcellular compartment Range/estimation of resting free Ca2+ Method of measurement References Apoplast 330 µM–1 mM X-ray microanalysis Conn et al. (2011) Cytosol 50–100 nM Aequorin, Cameleon (YC3.6), R-Geco1 Knight and Knight (1995); Logan and Knight (2003); Wagner et al. (2015a); Waadt et al. (2017) Nucleus 100 nM Aequorin Van der Luit et al. (1999); Mithöfer and Mazars (2002) Mitochondria matrix 100–600 nM Aequorin, Cameleon (YC3.6), Fura-2 Zottini and Zannoni (1993); Logan and Knight (2003); Wagner et al. (2015a) Chloroplast stroma 100–200 nM Aequorin, Cameleon (YC3.6) Mehlmer et al. (2012); Nomura et al. (2012); Sello et al. (2016); Loro et al. (2016) Thylakoid lumen 500 nM Aequorin Sello et al. (2018) Amyloplast/plastid stroma 80–100 nM Aequorin, Cameleon (YC3.6) Sello et al. (2016); Loro et al. (2016) Vacuolar lumen 200 µM–50 mM X-ray microanalysis Conn and Gilliham (2010); Conn et al. (2011) Endoplasmic reticulum lumen 50–500 µM Cameleon (CRT-D4ER) Iwano et al. (2009); Bonza et al. (2013) Golgi lumen 700 nM Aequorin Ordenes et al. (2012) Peroxisome lumen 150 nM–2 µM Cameleon (D3cpv-KVK-SKL) Costa et al. (2010) View Large Table 2. Summary of measured and estimated Ca2+ concentrations at resting conditions in the different subcellular compartments of plant cells Subcellular compartment Range/estimation of resting free Ca2+ Method of measurement References Apoplast 330 µM–1 mM X-ray microanalysis Conn et al. (2011) Cytosol 50–100 nM Aequorin, Cameleon (YC3.6), R-Geco1 Knight and Knight (1995); Logan and Knight (2003); Wagner et al. (2015a); Waadt et al. (2017) Nucleus 100 nM Aequorin Van der Luit et al. (1999); Mithöfer and Mazars (2002) Mitochondria matrix 100–600 nM Aequorin, Cameleon (YC3.6), Fura-2 Zottini and Zannoni (1993); Logan and Knight (2003); Wagner et al. (2015a) Chloroplast stroma 100–200 nM Aequorin, Cameleon (YC3.6) Mehlmer et al. (2012); Nomura et al. (2012); Sello et al. (2016); Loro et al. (2016) Thylakoid lumen 500 nM Aequorin Sello et al. (2018) Amyloplast/plastid stroma 80–100 nM Aequorin, Cameleon (YC3.6) Sello et al. (2016); Loro et al. (2016) Vacuolar lumen 200 µM–50 mM X-ray microanalysis Conn and Gilliham (2010); Conn et al. (2011) Endoplasmic reticulum lumen 50–500 µM Cameleon (CRT-D4ER) Iwano et al. (2009); Bonza et al. (2013) Golgi lumen 700 nM Aequorin Ordenes et al. (2012) Peroxisome lumen 150 nM–2 µM Cameleon (D3cpv-KVK-SKL) Costa et al. (2010) Subcellular compartment Range/estimation of resting free Ca2+ Method of measurement References Apoplast 330 µM–1 mM X-ray microanalysis Conn et al. (2011) Cytosol 50–100 nM Aequorin, Cameleon (YC3.6), R-Geco1 Knight and Knight (1995); Logan and Knight (2003); Wagner et al. (2015a); Waadt et al. (2017) Nucleus 100 nM Aequorin Van der Luit et al. (1999); Mithöfer and Mazars (2002) Mitochondria matrix 100–600 nM Aequorin, Cameleon (YC3.6), Fura-2 Zottini and Zannoni (1993); Logan and Knight (2003); Wagner et al. (2015a) Chloroplast stroma 100–200 nM Aequorin, Cameleon (YC3.6) Mehlmer et al. (2012); Nomura et al. (2012); Sello et al. (2016); Loro et al. (2016) Thylakoid lumen 500 nM Aequorin Sello et al. (2018) Amyloplast/plastid stroma 80–100 nM Aequorin, Cameleon (YC3.6) Sello et al. (2016); Loro et al. (2016) Vacuolar lumen 200 µM–50 mM X-ray microanalysis Conn and Gilliham (2010); Conn et al. (2011) Endoplasmic reticulum lumen 50–500 µM Cameleon (CRT-D4ER) Iwano et al. (2009); Bonza et al. (2013) Golgi lumen 700 nM Aequorin Ordenes et al. (2012) Peroxisome lumen 150 nM–2 µM Cameleon (D3cpv-KVK-SKL) Costa et al. (2010) View Large A plethora of transporters and channels are active in the tonoplast, as discovered by direct patch-clamping of this organelle (Martinoia et al., 2012; Xu et al., 2015). Many of these transport systems have been molecularly identified during the last few decades (Martinoia et al., 2012; Neuhaus and Trentmann, 2014). Ca2+ is taken up into the vacuole probably by two P-type Ca2+ pumps, such as CaM-regulated autoinhibited Ca2+-ATPases (ACAs), as well as by Ca2+/proton exchangers (CAXs) (Edel et al., 2017), which exhibit a high sequence homology to their yeast counterparts also residing on the vacuolar membrane (Hirschi et al., 1996). ACA pumps exist in at least 10 isoforms in Arabidopsis (Geisler et al., 2000). The activity of the two vacuolar ACA Ca2+ pumps AtACA4 and AtACA11 (Lee et al., 2007) has been linked to the control of a salicylic acid-dependent programmed cell death (PCD) pathway in plants (Boursiac et al., 2010). Among the six CAX members in Arabidopsis, AtCAX1–AtCAX4 have been shown to locate to vacuoles (Cheng et al., 2002; Pittman et al., 2005). Knock-out mutants of AtCAX1, that is highly expressed in leaf tissue, exhibited altered plant development, perturbed hormone sensitivities, and altered expression of an auxin-regulated promoter–reporter gene fusion (Cheng et al., 2003), while indole-3-acetic acid (IAA) inhibition of abscisic acid (ABA)-induced stomatal closure was found to be impaired in cax1, cax3, and cax1/cax3 mutants (Cho et al., 2012). Vacuolar CAX4, that shows a low expression level, plays an important role in root growth under heavy metal stress conditions (Mei et al., 2009). Interestingly, some of the CAXs transport not only Ca2+, but also heavy metals such as Mn2+ and Cd2+ (Manohar et al., 2011; Martinoia et al., 2012; Socha and Guerinot, 2014; Pittman and Hirschi, 2016). The vacuole, together with the cell wall/apoplast, is the major Ca2+ store and it is generally assumed that Ca2+ released from the vacuole provides in several cases substantial contributions for the activation of signal transduction pathways. This assumption is made based on early experiments showing that inositol-1,4,5-trisphosphate (InsP3) releases Ca2+ predominantly from the vacuole (Alexandre and Lassalles, 1990; Allen et al., 1995). However, later experiments indicated that in plants, inositol-hexakisphosphate (InsP6) plays a prominent role with respect to InsP3 in intracellular signal transduction (Lemtiri-Chlieh et al., 2003; Munnik and Nielsen, 2011). Among the channels proposed to release Ca2+ from the vacuole, the Ca2+-activated two-pore non-selective, Ca2+- and K+-permeable cation channel TPC1 (e.g. Peiter et al., 2005; Carpaneto and Gradogna, 2018), whose structure has been solved (Guo et al., 2016), acts as a tonoplast channel. Since the physiological concentration of K+ both in the cytosol and inside the vacuole (~100 mM) is much higher than the concentration of Ca2+ (Table 2), K+ permeation can be expected to be largely facilitated with respect to Ca2+ through the channel. Indeed, the ability of TPC1 to conduct Ca2+ has long been debated, but combination of the patch-clamp technique with Ca2+ detection by fluorescence finally led to the demonstration that Ca2+ is able to permeate through TPC1, even if its concentration (0.5 mM) is much lower than that of K+ (105 mM) in electrophysiological experiments (Gradogna et al., 2009; Carpaneto and Gradogna, 2018). Plants lacking TPC1 are defective in both ABA-induced repression of germination and in the response of stomata to extracellular Ca2+, demonstrating a critical role for the vacuole Ca2+ release channel in various physiological processes of plants (Peiter et al., 2005). Furthermore, the propagation of salt stress-induced long-distance Ca2+ waves as well as wounding/herbivory-triggered Ca2+ waves was found to be dependent on TPC1 in Arabidopsis (Choi et al., 2014; Kiep et al., 2015). However, other reports dismiss a prominent role for TPC1 in vacuolar Ca2+ release, assessed either by aequorin or by direct patch-clamp analyses. For example, unaltered cytosolic Ca2+ signals were recorded in intact plants either lacking or overexpressing TPC1 upon exposure to various biotic and abiotic stimuli (Ranf et al., 2008). At physiological pH and Ca2+ gradients, TPC1 was shown to conduct Ca2+ into the vacuole, suggesting that it dissipates rather than generates cytosolic Ca2+ signals (at least during external Ca2+-induced stomatal closure) (Rienmüller et al., 2010). Likewise, the findings that the fou2 mutant plants harbouring a hyperactive TPC1 channel variant (D454N) show an increased vacuolar Ca2+ content (Beyhl et al., 2009) and a decreased resting cytosolic Ca2+ level compared with the wild type argues against a Ca2+ release function of TPC1. The fou2 mutant also has a slightly lower resting cytosolic [Ca2+] compared with the wild type, and cytosolic Ca2+ increases after wounding were found to be similar in both plants (Lenglet et al., 2017). On the other hand, a recent study, carried out by using the fluorescent Ca2+ biosensor GCaMP3, highlighted a functionally relevant interplay between the plant defence co-receptor Brassinosteroid insensitive-associated kinase1 (BAK1), the PM-localized glutamate receptors GLR3.3 and GLR3.6, and TPC1 to mediate cytosolic Ca2+ elevations following biotic stress such as aphid attack (Vincent et al., 2017a). Interestingly, another study highlighted the importance of endomembrane cation fluxes in controlling the basal level of the wound-inducible defence mediator jasmonate acid, thanks to the use of the fou2 mutant of TPC1 (Lenglet et al., 2017). Thus, altogether, TPC1 is emerging as a possible regulator of cytosolic Ca2+ signals, although many questions still remain open (Hedrich et al., 2018). The readers are advised to consult excellent reviews on the state of the art and hot topics in vacuolar transport research, including those discussing the regulation of vacuolar channels by cytoplasmic/luminal factors (Hedrich, 2012; Martinoia et al., 2012; Edel et al., 2017; Francisco and Martinoia, 2018). A role for the endoplasmic reticulum in plant intracellular Ca2+ signalling? Another main intracellular Ca2+ store is the ER. Not much is known about the Ca2+ storage properties of the plant ER in contrast to the animal field, where it is well explored (Sammels et al., 2010; Raffaello et al., 2016). In animal cells, the total Ca2+ concentration in the ER is thought to be 2 mM, whereas the free Ca2+ concentration ranges between 50 μM and 500 μM (Rizzuto et al., 2009; Stael et al., 2012). The involvement of the ER in Ca2+ homeostasis and signalling in plant cells has long been underappreciated, possibly overshadowed by the prominent role commonly ascribed to the vacuole, and because of the lack, for a long time, of direct measurements of luminal [Ca2+] ([Ca2+]ER) and its potential variations during signal transduction. Functional conservation of calreticulin as the major high-capacity (15–30 mol of Ca2+ per mol of protein), low-affinity (Kd=0.5 mM) Ca2+-binding protein in the lumen of the plant ER (for reviews, see, for example, Mariani et al., 2003; Jia et al., 2009) has provided circumstantial evidence for submillimolar [Ca2+]ER. In addition to ER Ca2+ storage and modulation of Ca2+ homeostasis, calreticulin has been shown to function, together with calnexin, as a molecular chaperone for glycoprotein folding and quality control in the ER (Jin et al., 2009). Interestingly, overexpression of calreticulin was found to enhance the survival of plants grown in low Ca2+ medium (Persson et al., 2001) and to increase plant salinity tolerance (Xiang et al., 2015). The targeting of a Cameleon probe (YC4.6, with two KDs of 58 nM and 14.4 µM) (Table 1) to the ER of pollen tubes has highlighted a potential involvement of the ER in the fine regulation of the tip-focused [Ca2+]cyt gradient required for pollen tube growth (Iwano et al., 2009). Arabidopsis contains four P(IIA)-type ATPase genes, AtECA1–AtECA4, which are expressed in all major organs of Arabidopsis. ECA1 knock-out mutants grew poorly on medium with low Ca2+ or high Mn2+, indicating that ECA1-mediated uptake of these divalent cations into the ER is required for plant growth under conditions of Mn2+ toxicity or Ca2+ deficiency (Wu et al., 2002). The silencing of an ER-localized type IIB Ca2+-ATPase (ACA like) in tobacco has been found to alter intracellular Ca2+ signalling and accelerate PCD during the plant innate immune response, indicating that the Ca2+ uptake pathway into the ER functions as a regulator of PCD (Zhu et al., 2010). Ca2+ release from the ER has also been proposed to play an essential role in sieve tube occlusion via Ca2+-dependent forisome dispersion in legumes in response to burning stimuli (Furch et al., 2009; Tuteja et al., 2010). The recently reported targeting of another Cameleon variant, the CRT-D4ER (with a KD of 195 µM) (Table 1), allowed the dynamic, in vivo monitoring of ER luminal Ca2+, showing that the ER may also work as a capacitor/buffer of cytosolic Ca2+ transients (Bonza et al., 2013). In fact, cytosolic Ca2+ increases triggered by different stimuli (salt stress, external ATP, and glutamate) were followed by Ca2+ accumulation into the ER lumen, but not by release. Moreover, dynamically, the ER Ca2+ increases followed temporally the cytosolic increases, showing a slower rate of accumulation and release (Bonza et al., 2013; Corso et al., 2018). Another clue in favour of the role of the ER as a cytosolic Ca2+ capacitor is confirmed by the effect of cyclopiazonic acid (CPA) (an inhibitor of IIA Ca2+-ATPase ECA) which reduced the luminal ER Ca2+ concentration and increased that in the cytosol (Zuppini et al., 2004; Bonza et al., 2013), indicating the ECAs as fundamental players for ER Ca2+ homeostasis. Nonetheless, our recent work has demonstrated that the Arabidopsis CCX2 is localized in the ER where it is directly involved in the control of Ca2+ fluxes between the ER and the cytosol, playing a key role in the ability of plants to cope with osmotic stresses (Corso et al., 2018). Concerning Ca2+-permeable channels located in higher plant ERs, early biochemical studies have indicated the occurrence of ER Ca2+ mobilization pathways activated by voltage (Klüsener et al., 1995) and by two structurally related molecules, namely the pyridine nucleotide derivatives nicotinic acid adenine dinucleotide phosphate (NAADP) (Navazio et al., 2000) and cyclic ADP-ribose (cADPR) (Navazio et al., 2001). The molecular identity of the above voltage- and ligand-gated Ca2+-permeable channels, however, has not been unravelled yet. The ER has a unique architecture that facilitates the spatio-temporal segregation of biochemical reactions and the establishment of interorganellar communication networks. Spatially confined ER–PM microdomains are emerging as highly specialized signalling hubs both in animal systems (e.g. Son et al., 2016; Demaurex and Guido, 2017) and in plants (Bayer et al., 2017). In addition, the continuity between ER membranes and the outer nuclear membrane suggests a potential role for the ER as a Ca2+ store participating in the repetitive Ca2+ release/uptake from the nucleoplasm and perinuclear cytosol during legume symbioses (Capoen et al., 2011). CNGCs have recently been demonstrated to mediate these nuclear-associated Ca2+ oscillations induced in response to beneficial plant microbes during the nitrogen-fixing symbiosis and arbuscular mycorrhizal symbiosis (Charpentier et al., 2016). Structural and functional interactions have been demonstrated to occur between the ER membranes and stromules, dynamic stroma-filled tubules continuously extending and retracting from plastids (Schattat et al., 2011). The occurrence of specific contact sites through which the ER and plastids may exchange not only lipids, but also ions such as Ca2+, opens up the possibility of a complex and finely tuned Ca2+ regulation, involving potential ER–plastid crosstalk (Mehrshahi et al., 2013). On the other hand, the role of ER–mitochondria contact sites in shaping the cytosolic Ca2+ signalling is well documented in mammals (e.g. Rizzuto et al., 2012; Brini et al., 2018), but not in plants. We can envisage such an intimate liason also in the case of plant cells, although direct proof is missing in this case. The plant Golgi apparatus: a rather unexplored Ca2+ store The Golgi apparatus in plant cells is made of discrete stacks (formerly indicated as dictyosomes) dispersed throughout the cytoplasm and rapidly moving (several micrometres per second) along the surface of the ER (Robinson et al., 2015). In addition to essential roles in protein glycosylation and trafficking (Vitale and Galili, 2001), the plant Golgi apparatus serves as factory of polysaccharides (hemicellulose and pectins) for the cell wall matrix, whose architecture is known to be regulated by Ca2+ (Mravec et al., 2017). Moreover, the Golgi apparatus is the source for exocytotic vesicles, and it is known that exo- and endocytosis can be modulated by Ca2+ (Cucu et al., 2017). In view of the unique structural and functional features of the plant Golgi apparatus with respect to animal cells, we can expect that Ca2+ handling by this compartment may also exhibit some specificity in plant cells. Compared with the extensive information about Ca2+ handling by the Golgi in mammalian cells (for a review, see Pizzo et al., 2011), knowledge about Ca2+ homeostasis and signalling in the plant Golgi is still scarce. Free Ca2+ levels in the Golgi ([Ca2+]Golgi) were estimated to be ~0.70 μM (Table 2) (Ordenes et al., 2012), a value which is much lower than [Ca2+]Golgi measured in mammalian cells (ranging from ~250 μM in the cis-Golgi to ~130 μM in the trans-Golgi) (Pizzo et al., 2011). This suggests the existence of Ca2+-buffering systems inside the Golgi, and indeed calreticulin has been reported to be localized at the plant Golgi, in addition to the ER (Navazio et al., 2002; Nardi et al., 2006). Interestingly, transient increases in Ca2+ dynamics were observed in response to several abiotic stimuli, such as cold shock, mechanical stimulation, and hyperosmotic stress, whereas the administration of the synthetic auxin analogue 2,4-dichlorophenoxy acetic acid (2,4-D) induced a slow decrease of organellar Ca2+ (Ordenes et al., 2012). Concerning Ca2+ decoding mechanisms, two calmodulin-like proteins from Arabidopsis thaliana, AtCML4 and AtCML5, were found to be localized in vesicular structures between the Golgi and the endosomal system. Nevertheless, their C-terminal CaM domain was found to be exposed to the cytosolic surface of the vesicles, suggesting that they may sense and decode cytosolic, rather than luminal, Ca2+ signals (Ruge et al., 2016). The nature of Ca2+-transporting proteins still awaits clarification. Among the four IIA Ca2+-ATPase in Arabidopsis, AtECA3 was proposed to function in the transport of Ca2+ and Mn2+ ions into the Golgi (Mills et al., 2008). From the data so far available, it is clear that the information on the Ca2+ toolkit of the plant endomembrane system awaits further investigation of its precise molecular components and of the specific involvement of the different compartments of the plant secretory pathway as Ca2+-mobilizable stores in Ca2+-mediated signal transduction events. Chloroplasts as Ca2+ signal-shaping components in plant cells Recent studies have revealed that plant mitochondria and chloroplasts respond to biotic and abiotic stresses with specific Ca2+ signals (reviewed by McAinsh and Pittman, 2009; Rocha and Vothknecht, 2012; Nomura and Shiina, 2014; Kmiecik et al., 2016). Chloroplasts, which possess a high concentration of Ca2+, serve as important intracellular cytosolic ‘Ca2+ capacitors’ in plant cells, and they may also influence the entire cellular Ca2+ network by modulating cytosolic Ca2+ transients. Thus, they can contribute to shaping cytoplasmic Ca2+ signatures (Nomura et al., 2012; Loro et al., 2016; Sello et al., 2016). The predominant portion of the chloroplastic Ca2+ (~15 mM) is bound to the negatively charged thylakoid membranes or to Ca2+-binding proteins, keeping the resting free [Ca2+]stroma as low as 150 nM (Table 2) to avoid the precipitation of phosphates (Hochmal et al., 2015). Importantly, this concentration can be actively regulated: light-dependent depletion of cytosolic Ca2+ in the vicinity of chloroplasts has been observed in green algae, suggesting that an active Ca2+ uptake machinery is present on the envelope membranes, which is regulated by light/dark transitions and/or photosynthesis (Sai and Johnson, 2002). Specific, high-resolution tools have been exploited to monitor and quantify plastid Ca2+ dynamics (Table 1). The bioluminescent Ca2+ reporter aequorin was targeted to the chloroplast stroma, highlighting induction of Ca2+ influx into the stroma upon light to dark transition (Sai and Johnson, 2002) as well as a role for stromal Ca2+ signals in the activation of plant innate immunity (Nomura et al., 2012; Stael et al., 2015). Constructs encoding YFP–aequorin chimeras targeted to the outer and inner membrane of the chloroplast envelope, in addition to the stroma, are also available to investigate Ca2+ dynamics in these compartments (Mehlmer et al., 2012). We have recently used these plastid-targeted aequorin probes to reveal differential stimulus-specific Ca2+ responses of amyloplasts versus chloroplasts (Sello et al., 2016), suggesting that Ca2+ signalling might have specific roles during plastid development. Interestingly, using a chloroplast-targeted Cameleon probe, Ca2+ spikes could be detected in a large portion (>80%) of guard cell chloroplasts (Loro et al., 2016). The observed unique spiking pattern for each chloroplast strongly suggests that these Ca2+ signals can be modulated at the level of the single organelle (Loro et al., 2016). The reported observations support the concept that Ca2+ plays a key role in integrating internal and external stimuli at the level of individual chloroplasts. Ca2+ spikes appeared under chloroplast-autonomous control, even though the source of the Ca2+ causing the spike may be the cytosol. It has been hypothesized that opening of individual Ca2+ channels following a stimulus from within the chloroplast itself may allow influx of Ca2+ from the cytosol along the negative electrochemical gradient across the chloroplast envelope. For the inner envelope membrane, a value of approximately –110 mV has been reported (Wu et al., 1991). However, the nature of such channel(s) remains elusive (for recent reviews, see, for example, Finazzi et al., 2015; Carraretto et al., 2016; Pottosin and Shabala, 2016). Light-dependent uptake of Ca2+ into isolated chloroplast is thought to be mediated by a Ruthenium Red-sensitive uniport-type carrier in the envelope membrane and to be linked to photosynthetic electron transport via the membrane potential (Kreimer et al., 1985). Electrophysiological studies suggest the existence of voltage-dependent Ca2+ uptake activity [the fast-activating cation channel (FACC)] in the inner envelope membrane of pea chloroplasts (Pottosin et al., 2005). However, the molecular identity of FAAC remains elusive, and sensitivity to Ruthenium Red has not been investigated. Presuming that the outer membrane is permeable to Ca2+ via porin-like molecules (Szabo and Zoratti, 2014; Carraretto et al., 2016), the most promising inner envelope-located candidates include ion channels that may mediate the negative voltage-driven Ca2+ uptake across the inner envelope membrane (Heiber et al., 1995). These channel-forming proteins include the plastid-located glutamate receptors GLR3.4 (Teardo et al., 2010, 2011) and GLR3.5 (Teardo et al., 2015) and the mechanosensitive MSL2/3 channels (Haswell and Meyerowitz, 2006). A Ca2+-ATPase like protein (ACA1) (Huang et al., 1993), as well as HMA1 P-type ATPase (Ferro et al., 2010) are also candidates for mediating Ca2+ flux across the inner envelope membrane. The possible role, localization (for ACA1), and specificity of the latter two proteins are, however, highly debated (Hochmal et al., 2015). The recently identified member of the UPF0016 family, the PHOTOSYNTHESIS AFFECTED MUTANT71 (PAM71), located to the thylakoid membrane was reported to function in manganese transport in higher plants (Schneider et al., 2016). The closest homologue of PAM71, PAM71-HL, is located to the chloroplast envelope and is likely to exert the same function (Schneider et al., 2016; Eisenhut et al., 2018), as the homologues in cyanobacteria are also linked to manganese homeostasis (Gandini et al., 2017). On the other hand, the thylakoid-located PAM71 was proposed to encode a putative Ca2+/H+ antiporter with critical functions in the regulation of PSII and in chloroplast Ca2+ and pH homeostasis in Arabidopsis (Wang et al., 2016). The possibility that this protein is able to transport manganese in a Ca2+-dependent way or to transport both cations will have to be explored in a simplified, reconstituted system. A further candidate for Ca2+ transport across chloroplast membranes is represented by one of the six homologues of the Ruthenium Red-sensitive mammalian mitochondrial uniporter (MCU), which displays an ambiguous N-terminal sequence, possibly allowing targeting to both mitochondria and chloroplasts (Stael et al., 2012). However, the localization, channel activity, and the permeability for Ca2+ of this putative plastidial member of the AtMCU family have not been described up to now, in contrast to four other mitochondria-located AtMCU homologues (Wagner et al., 2015a; Carraretto et al., 2016; Teardo et al., 2017). At present, it is difficult to understand whether the FAAC might correspond to one of the above entities. In addition to Ca2+-permeable channels in chloroplasts, other regulatory cation fluxes may shape the cytosolic Ca2+ signature during stress. Stephan et al. (2016) provided evidence for involvement of two envelope-located K+/H+ antiporters, namely KEA1 and KEA2, in Ca2+-induced cytoplasmic responses during osmotic stress. In particular, the double kea1/kea2 mutant showed a reduced cytosolic Ca2+ level upon treatment with a hyperosmotic sorbitol solution, suggesting that the function of the two K+/H+ antiporters is intimately linked to Ca2+ mobilization pathways at the chloroplast membranes under these conditions. However, the exact mode of action is still unclear. In addition to the above-mentioned ion channels and transporters, several candidate Ca2+-binding proteins and Ca2+ sensors have been identified in these organelles and shown to contribute critically to Ca2+ homeostasis (Rocha and Vothknecht, 2012; Stael et al., 2012; Hochmal et al., 2015). The impact of impaired organellar Ca2+ handling for plant physiology has been convincingly illustrated in the cases of the chloroplast-localized Ca2+ sensor protein CAS, the thylakoid-located Post-Floral-specific gene 1 PPF1, and the glycosyltransferase QUASIMODO1 (QUA1) (Wang et al., 2003; Nomura et al., 2008; Petroutsos et al., 2011; Zheng et al., 2017). In addition, another Ca2+-binding protein, CP12, was shown to play an important role in the regulation of the Calvin–Benson–Bassham cycle (Rocha and Vothknecht, 2013). Studies on the thylakoid-localized Ca2+-sensing receptor CAS showed that chloroplasts modulate intracellular Ca2+ signals by controlling external Ca2+-induced cytosolic Ca2+ transients during stomatal closure (Nomura et al., 2008; Weinl et al., 2008). Indeed, mutation of the putative chloroplastic Ca2+ sensor CAS led to impaired stomatal movement and impaired plant growth, although the detailed molecular mechanism underlying CAS-related effects under various conditions has not yet been fully elucidated (Wang et al., 2012, 2016; Fu et al., 2013). Pathogen-associated molecular pattern (PAMP) signals evoked specific Ca2+ signatures in the stroma in chloroplasts, and CAS was involved in stromal Ca2+ transients (Nomura et al., 2012). CAS, and thus Ca2+, was shown to regulate chloroplast salicylic acid (SA) biosynthesis, and plants depleted of CAS failed to induce SA production in response to pathogen infection. Transcriptome analysis demonstrated that CAS allowed chloroplast-mediated transcriptional reprogramming during plant immune responses, as expression of several nuclear defence-related genes was shown to be dependent on CAS. Furthermore, the activity of mitogen-activated protein kinases (MAPK) was shown to be regulated in a CAS-dependent manner, suggesting that chloroplast-modulated Ca2+ signalling controls the MAPK pathway for the activation of critical components of the retrograde signalling chain (Guo et al., 2016; Leister et al., 2017). Thus, it is expected that chloroplasts could play pivotal roles in the Ca2+ signalling in plant cells upon different stress stimuli, as indeed indicated by recent results linking the CAS protein to chloroplast-dependent Ca2+ signalling under salt and drought stresses (Zhao et al., 2015; Zheng et al., 2017). Finally, QUA1 was also recently identified as a regulator of [Ca2+]cyt in response to drought and salt stress (Zheng et al., 2017). In addition to the chloroplast stroma, Ca2+ is required for the function of thylakoid lumen-located proteins such as the oxygen-evolving complex, suggesting that changes in free [Ca2+] are likely also to occur in the lumen. Recently, aequorin-based chimeras have been targeted to the thylakoid lumen and the stromal surface of the thylakoid membrane (Sello et al., 2018). The design of these thylakoid-specific Ca2+ indicators allowed measurement of Ca2+ concentrations inside and around thylakoids (Table 2) and to monitor dynamic Ca2+ changes in the above subchloroplast locations in response to different environmental cues. The availability of this complex toolkit of chloroplast-targeted Ca2+ reporters will pave the way for future studies on chloroplast Ca2+ homeostasis and signalling, and rapidly advance our understanding of the still enigmatic integration of these organelles in the plant Ca2+ signalling network. In summary, a systematic study linking the possible players of chloroplast Ca2+ dynamics to specific plant stress responses using envelope-, stroma-, thylakoid membrane-, and thylakoid lumen-targeted Ca2+ sensors would be of great importance to highlight further the importance of this organelle in global Ca2+ signalling within plant cells. Moreover, a promising field of investigation concerns the analysis of Ca2+ handling by non-green plastids in non-photosynthetic tissues and organs, such as the root. Indeed, the cell type-specific cytosolic Ca2+ responses of the root to environmental cues (Kiegle et al., 2000) may entail a differential contribution of root plastids. Moreover, it can be envisaged that root plastids may also play relevant roles in Ca2+ signalling during plant interactions with microorganisms of the rhizosphere, either pathogenic or beneficial. Mitochondrial Ca2+ signalling in plants Plant mitochondrial Ca2+ signalling has recently been reviewed (Stael et al., 2012; Nomura and Shiina, 2014; Carraretto et al., 2016; Wagner et al., 2016); therefore, here we prevalently focus on the missing links to understand the role(s) played by mitochondria in Ca2+ signalling processes. The emerging idea is that, similarly to animal cells, plant mitochondria can play a role in the modulation of cytosolic Ca2+ signatures, hence participating in the general intracellular Ca2+ homeostasis. The complex series of redox reactions of the mitochondrial electron transport chain (ETC) coupled to proton movement against the electrochemical gradient across the inner mitochondrial membrane (IMM) generates a proton motive force (pmf) composed of a proton gradient (ΔpH) across the IMM of ~0.9 pH units, and of an electric component (Δψ) reaching values of around –180 mV/–220 mV (Poburko et al., 2011; Szabo and Zoratti, 2014). The generated pmf is exploited to synthesize ATP, and for the import of proteins as well as of several charged substrates and cofactors that are translocated into the matrix via specialized co-transporters (Lee and Millar, 2016). Moreover, the negative matrix-side Δψ drives the import of positively charged ions, such as Ca2+, which flux into the matrix passively through channels, reaching free Ca2+ concentrations with values ranging from 100 nM to 600 nM (Table 2) (depending on the plant species and cell type; Zottini and Zannoni, 1993; Logan and Knight, 2003; Wagner et al., 2015b). In mammals, the free matrix Ca2+ has been shown to stimulate the activity of several enzymes of the Krebs cycle and the ATP synthase (Bagur and Hajnóczky, 2017). While in mammals mitochondria are essential players of Ca2+-based signalling processes, in plant cells clear-cut, unambiguous evidence demonstrating the involvement of this organelle in Ca2+ signalling processes is still lacking. The recent use of Ca2+ sensors (Rhod-2; aequorin and Cameleon) targeted to the plant mitochondrial matrix (Table 1) have allowed study in vivo of the mitochondrial Ca2+ dynamics both in resting conditions and after challenging the plant cells with different stimuli or drug treatments (Logan and Knight, 2003; Loro et al., 2012). A side by side use of transgenic plants stably expressing genetically encoded Ca2+ sensors targeted to the cytosol or mitochondria has then enabled the relationship, in terms of Ca2+ handling among these two different compartments, to be defined. An important and fundamental finding resulting from these in vivo studies was the ability of mitochondria to accumulate and release Ca2+ following cytosolic Ca2+ transients, being essentially dependent on them (Logan and Knight, 2003; Loro et al., 2012; Manzoor et al., 2012; Teardo et al., 2015; Wagner et al., 2015a). Moreover, stimuli which induce different cytosolic Ca2+ increases, in terms of dynamics and magnitudes, were also able to generate different mitochondrial Ca2+ dynamics, again confirming the existence of a strict relationship between the cytosol and mitochondria. Intriguingly, these works highlighted that mitochondria show slower dynamics of Ca2+ accumulation and release with respect to the cytoplasmic variations, strongly pointing to the possibility that plant mitochondria operate as cytosolic Ca2+ capacitors, at least locally, playing a role in the shaping of cytosolic Ca2+ signals (McAinsh and Pittman, 2009). In mammals, several reports demonstrated a role for mitochondria in cytosolic Ca2+ clearing and buffering, thus affecting and regulating Ca2+-based signalling responses (Rizzuto et al., 2012). However, indications that this mode of regulation also operates in plant cells are still lacking; therefore, new experimental strategies to demonstrate the existence, if any, of such a mechanism are needed. In this respect, the recent identification of some of the molecular components responsible for the mitochondrial Ca2+ transport across the IMM may be of help to test if the ‘mitochondria clearing hypothesis’ is valid in plants. The molecular identification of the MCU (Baughman et al., 2011; De Stefani et al., 2011) allowed important steps in the plant field also to be achieved. In Arabidopsis, AtMCU1 and AtMCU2, homologues of the mammalian MCU, were shown to localize to mitochondria and to transport Ca2+ when expressed in cell-free or heterologous systems (Tsai et al., 2016; Teardo et al., 2017). However, root mitochondria of the mcu1 knock-out (KO) Arabidopsis plants showed just a small reduction in the Ca2+ uptake rate compared with the wild type in vivo (Teardo et al., 2017), pointing to a functional redundancy, in line with the prediction of mitochondrial localization of at least five out the six MCU homologues in Arabidopsis (Stael et al., 2012). Interestingly, these isoforms appear to display tissue-specific distribution (Selles et al., 2018), possibly allowing clarification of the role of MCUs in a certain tissue using double/triple KO plants. Besides the MCUs, other possible routes for mitochondrial Ca2+ accumulation also exist in planta. Some members of the ionotropic glutamate-like receptor (GLR) family have been shown to transport Ca2+ (Vincill et al., 2012; Tapken et al., 2013; Ortiz-Ramírez et al., 2017). In addition to the predominant localization of the Arabidopsis GLR3.5 to plastids, a splicing variant is localized to mitochondria, and a glr3.5 KO mutant showed a reduction of the mitochondrial Ca2+ accumulation rate compared with the wild type (Teardo et al., 2015). A priori, GLR3.5 might work agonistically with the MCU for the accumulation of Ca2+. Although in Arabidopsis there are no reports showing mitochondrial Ca2+ dynamics in transgenic lines that overexpress MCUs, the recently described mutant lacking the mitochondrial Ca2+ uptake regulator protein (MICU) that inhibits channel activity showed an overaccumulation of mitochondrial Ca2+ (even in resting conditions) when compared with the wild type (Wagner et al., 2015a), therefore potentially mimicking the effects of MCU overexpression. Moreover, the lack of MICU accelerated the speed of mitochondrial Ca2+ accumulation in root tip cells in response to external stimuli. However, cytosolic Ca2+ dynamics assayed in the micu mutant background did not show significant differences if compared with the wild type, indicating that an increase of mitochondrial Ca2+ accumulation does not necessarily boost the cytosolic Ca2+ clearing. In summary, the study of both mitochondrial and cytosolic Ca2+ dynamics would be fundamental in plants simultaneously lacking MCU isoforms and GLR3.5 (possibly with an inducible system) to define the role of mitochondria in clearing of cytosolic Ca2+ and therefore their role for the regulation of Ca2+ signalling. In addition to MCUs and GLR3.5, plant mitochondria may have other routes for Ca2+ uptake. Three-mitochondrial adenine nucleotide/phosphate carriers (AtAPC1–AtAPC3) can transport ATP-Ca in reconstituted liposomes (Lorenz et al., 2015). However, evidence that the ATP-Ca transport in mitochondria occurs in vivo is lacking. Thus, it would be extremely interesting to study Ca2+ dynamics in the mitochondria and cytosol of apcs mutants carrying mitochondria- and cytosol-targeted Ca2+ probes. Pollen tubes or root hairs, where both Ca2+ and ATP are fundamental players for a proper growth (Winship et al., 2016), might represent especially useful systems for these studies. Indeed, a recent study highlighted the importance of MCU2 in pollen tube development even if it was not clear whether the observed phenotype was dependent or not on an altered mitochondrial or cytosolic Ca2+ homeostasis (Selles et al., 2018). It must be mentioned that the experiments presented so far were mainly carried out in Arabidopsis root tip cells and essentially designed to study fast Ca2+ dynamics in the mitochondria and cytosol. The mcu1, mcu2, micu, and glr3.5 plants showed mild phenotypes such as altered mitochondrial morphology, reduced pollen tube germination and growth in vitro, accelerated senescence, or reduced seedling root lengths that can be somewhat difficult to correlate directly with short-term signalling events. The lack of a strong phenotype in terms of mitochondrial and cytosolic Ca2+ dynamics and Ca2+-related signalling events can be explained by the lack of a true null mutant (unable to accumulate Ca2+ into mitochondria). Nevertheless, studies at specific developmental stages and in specific organs/tissues or cell types may be of help. In support of this idea, it has previously been demonstrated that the concentration of free Ca2+ in mitochondria is higher in the tip of the root hairs (500 nM) than in the shanks (200 nM), hence essentially following the cytosolic Ca2+ gradient (Wang et al., 2010). As mentioned above, growing pollen tubes and root hairs, that both have a high demand of metabolic energy and require the establishment of a defined cytosolic tip Ca2+ gradient, may represent the most suitable systems (Michard et al., 2017). Indeed, the recent demonstration that the mcu2 mutant shows a phenotype in pollen tubes supports this idea (Selles et al., 2018). It might also be interesting to understand if and how mitochondrial Ca2+ release can regulate cytosolic Ca2+ recovery. In a simplified way, we may hypothesize that the slow decrease of mitochondrial Ca2+ might delay the cytosolic Ca2+ recovery phase. The Arabidopsis genome contains two genes with homology to the mammalian LETM1, an EF-hand protein proposed to be involved in the export of Ca2+ from the mitochondria (Shao et al., 2016; Austin et al., 2017). Both Arabidopsis homologues, LETM1 and LETM2, reside in the IMM, and the double knockout mutant is not viable (Zhang et al., 2012). To date there are no data showing Ca2+ dynamics in the mitochondria or cytosol (of at least single LETM mutants), and the ion species transported by this protein are a matter of debate even in the mammalian system. Hence, it would be important to analyse the cytosol/mitochondria Ca2+ handling relationships in an Arabidopsis mutant lacking both LETMs, possibly by using an inducible silencing system to avoid embryonic lethality. In order to systematically study the role of mitochondria in the regulation of Ca2+ signalling, a forward genetic strategy for the isolation of mutants impaired in mitochondrial Ca2+ homeostasis could be of relevance. In this case, the use of molecular imaging coupled with high-throughput screenings and possibly with a relatively simple genetic system (e.g. Physcomitrella patens, Marchantia polymorpha, or Chlamydomonas reinhardtii) could provide a series of potential new candidate genes that could help to elucidate the role of mitochondria in Ca2+ signalling processes. A similar approach was pursued by Zhao et al. (2013) who screened an A. thaliana T-DNA insertion pool to identify mutants defective in salt stress-induced increases in cytosolic Ca2+. This screening pointed to Actin-Related Protein2 (Arp2) which affected not only the salt-induced cytosolic Ca2+ increases, but also mitochondria movement, mitochondrial membrane potential, and opening of the cell death-triggering permeability transition pore (PTP). An interesting observation was that the pharmacological block of the mitochondrial PTP opening prevented the cytosolic Ca2+ increase, but unfortunately the authors did not provide any direct evidence on altered mitochondrial Ca2+ dynamics. Another recent work identified the WRKY15 transcription factor as a negative regulator of salt and osmotic stress tolerance in Arabidopsis (Vanderauwera et al., 2012). Importantly, the authors revealed that WRKY15 overexpression induced an unfolded protein response which impaired the cytosolic Ca2+ homeostasis and affected the mitochondrial retrograde regulation mechanism, de facto triggering a stress hypersensitivity. Treatment with CPA that affects the activity of ECAs (see above), promoted mitochondrial responses, placing this organelle at the crossroads of ER stress and general cellular responses. A detailed description of Ca2+ dynamics in mitochondria, ER, and cytosol has not been provided, making it difficult to assign a specific role for mitochondria in the regulation of cytosolic Ca2+ under these experimental conditions. In summary, a suggested role for mitochondrial Ca2+ regulation in the salt and osmotic stress response in these latter works is of high interest and deserves further investigation. Peroxisomal Ca2+ signalling When discussing the role of organelles in Ca2+ signalling, peroxisomes also have to be taken into account, even if only a few studies addressed their involvement during the last few years. Peroxisomes are ubiquitous single-membrane-bounded organelles that fulfil essential roles in cellular metabolism. In contrast to mitochondria, peroxisomes do not have any ETC and, to the best of our knowledge, the existence of a membrane potential has not been reported. However, the peroxisomal membrane is impermeable to high molecular weight molecules (>1000 Da), and specific carriers are expressed in the organelles for the transport of different metabolites (Linka and Weber, 2010; Linka and Esser, 2012). Both mammalian and plant peroxisomes accumulate Ca2+ in the lumen in response to stimuli that trigger cytosolic Ca2+ increases (Lasorsa et al., 2008; Costa et al., 2010, 2013). The resting intraperoxisomal luminal Ca2+ concentration has been estimated to range between 150 nM and 2 µM (Table 2) (Drago et al., 2008). In mammals, stimuli which induce cytosolic Ca2+ increases are followed by slow rises in intraperoxisomal Ca2+ that do not require either ATP, membrane potential, and a H+ gradient (Drago et al., 2008). In plant cells, only two reports showed a stimulus-induced peroxisomal Ca2+ increase in guard cells and root tip cells (Costa et al., 2010; 2013). In both cases, the peroxisomal Ca2+ dynamics were like the cytosolic dynamics, reminescent of what is reported in mammalian cells. From the available results, we can summarize that peroxisomes essentially show an equilibration of the peroxisomal luminal Ca2+ with that of the cytosol and only potentially work as an additional cytosolic Ca2+ buffer. On the other hand, catalase 3 (CAT3) controls the H2O2 levels in guard cells (Zou et al., 2015), and this regulation is dependent on Ca2+ in two different ways—one mediated by CaM (Yang and Poovaiah, 2002) that operates in peroxisomes, and one mediated by CPK8 operating in the cytosol (Zou et al., 2015). Hence, a stimulus that induces both a cytosolic and peroxisomal Ca2+ increase can activate the same enzyme in different locations, via different mechanisms. Another recent observation reports that the peroxisomal Ca2+ is required, via a CaM-dependent mechanism, for protein import and for the normal functionality of peroxisomal enzymes, including antioxidant and photorespiratory enzymes, as well as for nitric oxide production (Corpas and Barroso, 2017). In conclusion, the property of peroxisomes to accumulate and release Ca2+ into and out of the lumen has a functional role in the plant cell; however, currently we lack information about the identity of possible transporters/channels involved in these fluxes. The apoplast as a main source of Ca2+ in signalling The apoplast is obviously not an intracellular organelle; however, together with the vacuole, the cell wall represents the main Ca2+ store in plants cells, with an estimated concentration of free Ca2+ ranging from 0.33 mM to 1 mM (Table 2) (Conn and Gilliham, 2010; Stael et al., 2012). Remarkably, the apoplast is considered the first plant compartment encountering environmental signals (Gao et al., 2004) and, in support of this, there are several pieces of evidence which demonstrate that the apoplast represents the primary source for the entry of Ca2+ into the cell upon the perception of a given stimulus. In fact, by chelating extracellular Ca2+, using EGTA or BAPTA, or by blocking the PM non-selective cation channels with La3+ or Gd3+, stimuli-induced cytosolic Ca2+ increases are strongly reduced if not completely abolished (Knight et al., 1996; Lamotte et al., 2004; Ali et al., 2007; Navazio et al., 2007). Despite the importance of the apoplast in the generation of cytosolic Ca2+ increases, a limited number of studies have reported in vivo direct measurements of apoplastic Ca2+. This is mainly due to the high Ca2+ concentration and the low pH of the apoplast which make Ca2+ measurements challenging, similarly to what we underlined for the vacuole. However, Gao and colleagues were able to target aequorin to the extracellular space and measure apoplastic Ca2+ dynamics in response to cold stress, revealing that they were different from the cytosolic dynamics. Remarkably, the authors also showed that the permanent washout of apoplastic Ca2+ determined a continuing aequorin signal decay, hence confirming the probe functionality (Gao et al., 2004). More recently, Wang and colleagues have instead used the Oregon Green BAPTA 488 5N dye to demonstrate that leaf cells of the cngc2 and cax1cax3 mutants overaccumulate apoplastic Ca2+ compared with the wild type, when grown in the presence of high external Ca2+ in the medium (Wang et al., 2017). Interestingly, the overaccumulation of apoplastic Ca2+ in the cax1/cax3 mutant was previously reported by Conn and co-workers performing X-ray microanalysis (Conn et al., 2011). The fact that CAX1 and CAX3 are tonoplast-localized Ca2+/H+ exchangers makes this latter observation of primary importance since it supports the existence of a potential communication between the apoplast and vacuole, which will probably deserve more attention. Conclusion and perspectives Although there are common elements in Ca2+-based signal transduction networks in all eukaryotes, unique traits of plant Ca2+ signalling derive from both structural features of the plant cell and from differences in the lifestyle and developmental programmes of plants. Genetic approaches using mutant plants defective in specific Ca2+ transporters/channels, together with pharmacological approaches using Ca2+ chelators and/or inhibitors of Ca2+ channels differentially distributed across cellular membranes, have elucidated how the different stimulus-specific cytosolic Ca2+ signatures often derive from the joint contribution of more than one source of Ca2+. Figure 2 summarizes the different channels/transporters possibly involved in Ca2+ fluxes in different intracellular membranes. Crosstalk among cellular compartments, possibly due to structurally close contacts, may also affect the ensuing global cytoplasmic Ca2+ signal. In this respect, the possible use of optical molecular tweezers (Sparkes, 2016) might be of relevance. Fig. 2. View largeDownload slide Organelle-located Ca2+-permeable channels and transporters possibly involved in Ca2+ transport across intracellular membranes. Different channels/transporters putatively involved in Ca2+ uptake/release into/from organelles and endomembranes are listed. The proteins involved are cyclic nucleotide-gated channels (CNGCs), glutamate receptor-like channels (GLRs), two-pore channels (TPCs), mechanosensitive channels (MSLs), autoinhibited Ca2+-ATPases (ACAs), ER-type Ca2+-ATPases (ECAs), P1-ATPases (HMA1), mitochondrial Ca2+ uniporter complex (MCUC), Ca2+/H+ exchangers (CAXs), and cation/Ca2+ exchangers (CCX2). See text for further details. Fig. 2. View largeDownload slide Organelle-located Ca2+-permeable channels and transporters possibly involved in Ca2+ transport across intracellular membranes. Different channels/transporters putatively involved in Ca2+ uptake/release into/from organelles and endomembranes are listed. The proteins involved are cyclic nucleotide-gated channels (CNGCs), glutamate receptor-like channels (GLRs), two-pore channels (TPCs), mechanosensitive channels (MSLs), autoinhibited Ca2+-ATPases (ACAs), ER-type Ca2+-ATPases (ECAs), P1-ATPases (HMA1), mitochondrial Ca2+ uniporter complex (MCUC), Ca2+/H+ exchangers (CAXs), and cation/Ca2+ exchangers (CCX2). See text for further details. In summary, it is clear that the combination of an increasing understanding of the molecular players and elements underlying plant Ca2+ signalling in organelles, together with newly generated detection systems for measuring organellar Ca2+ concentrations in intact plants, should provide fruitful grounds for ground-breaking discoveries. The view is emerging that, beside transporters, intracellular ion channels also contribute to fine-tuning of cytoplasmic Ca2+ dynamics. In this respect, existing proteomic data for different organelles (e.g. Prime et al., 2000) might allow the identification of further, new (putative) Ca2+ transport modules and decoders that might play a role in shaping Ca2+ homeostasis within the plant cell. One of the greatest challenges in the field is the elucidation of how influx and efflux Ca2+ transporters/channels are regulated in a concerted manner to translate specific information into a Ca2+ signature. The reported values are an estimation of Ca2+ concentrations in the different compartments based on direct measurements or deducted from the in vitro KD of the Ca2+ sensors reported in Table 1 Acknowledgements The original work by the authors has been supported over the years by grants from the Italian Ministry of University and Research (MIUR) [Progetti di Ricerca di Rilevante Interesse Nazionale (PRIN) and Fondo per gli Investimenti della Ricerca di Base (FIRB)], the Human Frontier Science Program (HFSP), and the University of Padova [Progetti di Ricerca di Ateneo (PRAT)]. We thank all lab members who contributed to the results reported in this review, and Marisa Brini, Fiorella Lo Schiavo, and Ute Vothknecht for continuous stimulating discussion on organellar Ca2+ signalling. Further, we are grateful to R. Mazzaro (Padova, Italy) for help with drawing Fig. 2. References Albrecht T , Zhao Y , Nguyen TH , Campbell RE , Johnson JD . 2015 . 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doi: 10.1093/jxb/ery154pmid: 29701811

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. 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doi: 10.1093/jxb/ery201pmid: 29800460

Abstract Soil composition largely defines the living conditions of plants and represents one of their most relevant, dynamic, and complex environmental cues. The effective concentrations of many either tolerated or essential ions and compounds in the soil usually differ from the optimum that would be most suitable for plants. In this regard, salinity—caused by excess NaCl—represents a widespread adverse growth condition, but shortage of ions such as K+, NO3−, and Fe2+ also restrains plant growth. During the past years, many components and mechanisms that function in the sensing and establishment of ion homeostasis have been identified and characterized. Here, we reflect on recent insights that extended our understanding of components and mechanisms which govern and fine-tune plant salt stress tolerance and ion homeostasis. We put special emphasis on mechanisms that allow for interconnection of the salt overly sensitive pathway with plant development and discuss newly emerging functions of Ca2+ signaling in salinity tolerance. Moreover, we review and discuss accumulating evidence for a central and unifying role for Ca2+ signaling and Ca2+-dependent protein phosphorylation in regulating sensing, uptake, transport, and storage processes of various ions. Finally, based on this cross-field inventory, we deduce emerging concepts and questions arising for future research. Calcium, CBL–CIPK, iron, metals, nitrate, nutrient sensing, potassium, salt stress, SOS Introduction Because the sessile lifestyle of plants is inevitably connected with their rooting in the ground, the actual composition of the soil and especially fluctuations of its components such as minerals, metals, and ions are vitally important for plants. In nature, but also in artificial agricultural environments, an optimal soil composition is the exception rather than the rule. Instead, deviations of many soil components from the optimum either towards a shortage or alternatively towards excess supply represent the daily challenge for plant life. Suboptimal supply of essential nutrients and too high concentrations of normally tolerated ions in any case represent stress conditions to plants. The main soil stresses that affect plants and crops in the field include soil salinity, adverse pH, insufficient water availability, nutrient depletion, and anaerobic stresses connected with flooding (Suzuki et al., 2014). A steadily growing share of agriculturally used areas but also of natural ecosystems is facing salinity stress due to high salt in soils, currently affecting 20% of the total, and 33% of irrigated agricultural lands worldwide (Munns and Tester, 2008; Cominelli et al., 2013; Schroeder et al., 2013). Due to the predominant role of NaCl, we refer within this article to the effects of high Na+, when using the terms saline or salt. However, it should be considered that high concentrations of Cl− anions are also toxic for plants (Tester and Davenport, 2003). The physiological and molecular implications of high Cl− concentrations on plants as well as their Cl− transport and evasion strategies have been most recently discussed in a comprehensive review article (Geilfus, 2018). Salt accumulation in arable soils is mainly derived from irrigation water that contains trace amounts of NaCl, and from seawater (Munns and Tester, 2008; Deinlein et al., 2014). Soils are considered saline when their electric conductivity reaches ≥4 dS m−1, which is equivalent to 40 mM NaCl (Marschner, 1995). However, some crops, such as beans, are sensitive to much lower salt concentrations (Cabot et al., 2014). In saline environments, the Na+ concentration in soils has been determined to range from ~100 mM to 2380 mM Na+ (Flowers 1985). Most crops will stall, die off, or at least not produce fruits or seeds under growth conditions exceeding 100 mM NaCl (Mass and Hoffman, 1977; Zhu, 2001; Flowers, 2004). High concentrations of Na+ can alter the basic texture of the soil, resulting in decreased soil porosity and consequently reduced soil aeration and water conductance. High salt deposition in the soil generates a zone of low water potential, making it increasingly difficult for the roots to acquire both water and nutrients. One facet of salt stress for the plants is therefore the resulting water deficit mimicking a physiological drought in combination with nutrient deficiencies (Mahajan et al., 2008). By causing osmotic stress, salt stress can inhibit the activity of many essential enzymes, cell division, and cell expansion, and can cause membrane disorganization and osmotic imbalance, which finally lead to growth inhibition (Marschner, 1995; Tuteja, 2007). Higher concentrations of Na+ also provoke the stomatal limitation of photosynthesis and excess production of reactive oxygen species (ROS) (Chaves et al., 2009; Flowers et al., 2010). Irrespective of these aspects, the sole ionic component of salinity stress, namely the toxicity of Na+, also has enormous consequences for the plant and exerts dramatic adverse effects on plant physiology including imbalances in the homeostasis of other ions such as K+ and Ca2+ (Munns and Tester, 2008; Anschütz, et al., 2014; Julkowska and Testerink, 2015). The maintenance of intracellular, but also organismic scale, ionic homeostasis is therefore fundamental to plant physiology. The versatility and diversity of implications that salt stress has on plants is reflected by the complexity and variety of sensing and adaptation mechanisms that are triggered in challenged plants. These mechanisms involve, for example, the control of cellular water and ion homeostasis in different plant tissues as well as triggering scavenging mechanisms of toxic compounds (Hasegawa et al., 2000). To mount an effective response to cope with salinity stress, land plants have developed the ability to sense and adapt to both osmotic stress and Na+ (Deinlein et al., 2014), which include Na+/H+ transporters at both the plasma membrane and tonoplast that enable efflux of Na+ ions out of the cell into the apoplastic space or to sequester them in the vacuole, respectively. Also, transcriptional reprogramming and specific synthesis of osmolytes, as well as avoidance of salt stress by directing root growth away from high salt concentrations belong to the repertoire of plant response strategies. All these aspects of plant stress adaptation have been most informatively covered in a number of recent review articles (Julkowska and Testerink, 2015; Han et al., 2017; Ismail and Horie, 2017; Köster et al., 2018; Yang and Guo, 2018). Similar to salt stress, lack of essential ions or nutrients (or in rarer cases their excess) triggers plant adaptation reactions to re-establish an optimal physiological homeostasis. These processes have been intensively studied for important nutrients and ions such as K+, Mg2+, Fe2+, NO3−, and NH4+ (Nieves-Cordones et al., 2016; Jeong et al., 2017; Tang and Luan, 2017; Tegeder and Masclaux-Daubresse, 2018). Although our knowledge about the plant perception and adaptation of such environmental cues has been steadily increasing, for many of these substances it remains unclear how they are sensed, and details of their signal transduction and adaptation reactions are only beginning to unfold. Nevertheless, an emerging common theme of these fundamental plant response reactions appears to be the occurrence of Ca2+ signals somewhere during the chain of events resulting in plant physiological adjustment. Consequently, in this review, we first focus on recent findings in the field of salt stress-related Ca2+ signaling. In this context, we discuss emerging indications for the integration of the Ca2+-triggered ‘salt overly sensitive’ (SOS) pathway with the co-ordination of developmental programs, nutritional status, and hormonal regulation to achieve plant developmental plasticity especially in stress conditions. From this, we expand our reflections towards the arising theme of Ca2+ signaling as a common and versatile mechanism shared by an increasingly appreciated number of plant nutrient and ion sensing and adaptation processes. Calcium signaling in response to salt stress Early studies on the possible involvement of Ca2+ signals in salt stress responses have already revealed strong indications that Ca2+ signaling indeed appears to be an early and prominent feature of the plant responses to this environmental cue (Lynch and Läuchli, 1988; Lynch et al., 1989). It was found that plants exhibit a rapid increase in cytosolic calcium concentration ([Ca2+]cyt) within seconds of being exposed to NaCl or mannitol (Knight et al., 1997). The similarity of Ca2+ signatures induced by either NaCl or non-ionic osmotic triggers has up to now prevented conclusive discernment of whether Na+ as an ion specifically triggers a Ca2+ response independently of its osmotic impact. Also, the still unknown identity of functional salt stress receptors up to now impedes the advancement of our understanding of these most important early steps in salt stress signaling (Köster et al., 2018). Several studies observed that the salt-induced Ca2+ rise originated within the roots (Kiegle et al., 2000; Tracy et al., 2008). Moreover, more detailed studies on the location and dynamics of the initial Ca2+ signals already suggested cell specificity and tissue heterogenicity of this response. For example, Kiegle and colleagues observed a most pronounced increase in cytoplasmic Ca2+ concentration in endodermal cells throughout the root in response to acute salt stress (Kiegle et al., 2000). Independently, it was found that an initial Ca2+ signal after salt stress was formed close to the root apex and then appeared to disperse to basal parts of this organ (Moore et al., 2002). More recently, the application of advanced Ca2+ reporter proteins has provided further important details about the Ca2+ signals that are triggered in response to NaCl application. Mazars and colleagues observed that exposure of roots to NaCl was sufficient to trigger systemic, wave-like Ca2+ elevations in plant leaves (Xiong et al., 2014). Moreover, targeted local application of NaCl to distinct parts of Arabidopsis roots was found to trigger an initial locally restricted Ca2+ signal within a few seconds that subsequently expanded through the root and whole plant in a wave-like manner (Choi et al., 2014). These important studies already suggest that Ca2+ signaling during salt stress represents a rather complex phenomenon and that its function and consequences are not restricted to the single-cell level. Moreover, these studies also provided initial evidence that Ca2+ signals fulfill a central role in orchestrating the whole-plant response to soil-borne signals that are formed by salt stress. Subsequently, it was reported that the tonoplast-localized cation channel TPC1 and the plasma membrane-localized NADPH oxidase RBOHD crucially contribute to the appropriate formation of salt-induced long-distance signals (Choi et al., 2014; Evans et al., 2016). These findings reinforce the emerging complexity of these processes, since they indicate the involvement of multiple cellular compartments and the contribution of several signaling systems to the full manifestation of Ca2+ signaling in response to salt stress. However, as important as the formation of Ca2+ signals for the implementation of appropriate physiological responses is their decoding and translation into downstream reactions. Converting Ca2+ signals into salt tolerance: the SOS pathway Our current knowledge on central processes and components of plant salt tolerance is to a large extent due to an elegant genetic screen for SOS mutants that identified (and allowed characterization of) central components of this pathway. Despite the non-biased character of this screen, the very first gene to be identified serendipitously turned out to be that encoding the Ca2+-binding protein SOS3 that most probably functions to decode the above-described Ca2+ signals (Liu and Zhu, 1997). Subsequently, SOS3 was found to be the fourth member (designated as CBL4) of the family of calcineurin B-like proteins (CBLs) which function as Ca2+ sensor proteins in many fundamental processes in plants (Kudla et al., 1999; Kolukisaoglu et al., 2004). Subsequent research established that SOS3 interacts with and activates the CBL-interacting protein kinase (CIPK) SOS2/CIPK24 (Halfter et al., 2000; Liu et al., 2000). The resulting Ca2+ sensor–kinase complex then phosphorylates and activates the Na+/H+ antiporter SOS1, which functions in Na+ extrusion and long-distance Na+ transport in plants (Shi et al., 2000, 2002). The SOS pathway, consisting of a Ca2+ sensor, a protein kinase, and a substrate, represents the first completely identified CBL–CIPK pathway for maintaining ion homeostasis in plant cells (Qiu et al., 2002) (Fig. 1). Although, the formation and activation of SOS3/CBL4–SOS2/CIPK24 complexes in vitro was not promoted by increased Ca2+ concentrations, it is generally accepted that the SOS pathway lies downstream of the salt-induced root [Ca2+]cyt increases (Yang and Guo, 2018). Fig. 1. View largeDownload slide Ca2+ signaling crucially functions in various facets of plant salt stress responses. The receptor-like kinase FERONIA (FER) is required for single cell-specific Ca2+ signaling that mediates cell wall integrity control during late salt stress responses, which are linked to re-establishment of growth. The recognition of Na+-induced cell wall softening probably involves physical interaction between the extracellular domain of FER, and pectin- and FER-dependent signaling elicits cell-specific Ca2+ signals within this process. The molecular nature of the receptor(s) for both the ionic (Na+) and the osmotic (π) component of salt stress, as well as the identity of the downstream Ca2+ channel(s) responsible for the well-documented cellular Ca2+ influx upon salt stress remain to be uncovered. Na+ extrusion from the cytosol is achieved by the activity of the Na+/H+ antiporter SOS1. Upon salt stress, the transporter is phosphorylated and thereby activated by the kinase SOS2/CIPK24, which is in turn regulated by the interacting Ca2+ sensor protein SOS3/CBL4. Besides CBL4, other members of the CBL protein family can potentially form functional complexes with SOS2/CIPK24, such as CBL10 and CBL1. Along with activation by CBL Ca2+ sensors, additional CIPK24 activity modulators have been recently identified. The geminivirus Rep-interacting kinases (GRIKs) 1 and 2, which both play critical roles in sugar/energy signaling by phosphorylating SnRK1-type kinases, were identified also to phosphorylate CIPK24 within its activation loop at Thr168. This phosphorylation is required for full SOS2 activation and accurate responses during salt stress. In the absence of salt stress, CIPK24 is kept in an inactive state by both the flowering time regulator GIGANTEA and the 14-3-3 proteins λ/κ. Whether these inhibitory interactions interfere with CBL binding to CIPK24, or if they represent CBL-independent mechanisms, remains to be addressed. Within the aerial parts of the plant, upon salt stress Na+ sequestration into the vacuole is a strategy to retain cytoplasmic Na+ concentrations in a tolerable range. Here, CBL10/CIPK24 complexes probably activate a still unknown mechanism of Na+ transport into the vacuole. One of the toxic effects of Na+ within the cells is the perturbance of the Na+/K+ ratio. To counteract this effect triggered by Na+ influx into cells during salt stress, the import of K+ ions can be enhanced to achieve adjustment of the Na+/K+ ratio by Ca2+ elevations through activation of the plasma membrane-localized K+ transporter AKT1 by CBL1/9—CIPK23 complexes. The regulation of the pH gradient across the plasma membrane is crucial for salt tolerance responses, as this gradient represents the driving force for the SOS1-mediated Na+ extrusion from the cytosol. In the absence of salt stress, the AHA2 proton pump is negatively regulated by CBL2 and/or CIPK11 to prevent excess ATP consumption. During the onset of stress, this negative regulation is released. During endodermal differentiation, a ring-like, lignified tertiary cell wall deposition called the Casparian strip is formed to block the apoplastic and the coupled trans-cellular pathway into the stele to prevent excess uptake of toxic elements such as Na+ into, as well as leakage of valuable metabolites from, the vasculature. For this barrier formation, precisely localized activity of the NADPH oxidase RBOHF is required to produce H2O2 needed for lignin cross-linking by peroxidases. RBOHF is known to be directly regulated by Ca2+ binding and phosphorylation by CBL/CIPK26 complexes. rbohF mutants with disturbed Casparian strip formation display a specific transpiration-dependent salt stress phenotype. Fig. 1. View largeDownload slide Ca2+ signaling crucially functions in various facets of plant salt stress responses. The receptor-like kinase FERONIA (FER) is required for single cell-specific Ca2+ signaling that mediates cell wall integrity control during late salt stress responses, which are linked to re-establishment of growth. The recognition of Na+-induced cell wall softening probably involves physical interaction between the extracellular domain of FER, and pectin- and FER-dependent signaling elicits cell-specific Ca2+ signals within this process. The molecular nature of the receptor(s) for both the ionic (Na+) and the osmotic (π) component of salt stress, as well as the identity of the downstream Ca2+ channel(s) responsible for the well-documented cellular Ca2+ influx upon salt stress remain to be uncovered. Na+ extrusion from the cytosol is achieved by the activity of the Na+/H+ antiporter SOS1. Upon salt stress, the transporter is phosphorylated and thereby activated by the kinase SOS2/CIPK24, which is in turn regulated by the interacting Ca2+ sensor protein SOS3/CBL4. Besides CBL4, other members of the CBL protein family can potentially form functional complexes with SOS2/CIPK24, such as CBL10 and CBL1. Along with activation by CBL Ca2+ sensors, additional CIPK24 activity modulators have been recently identified. The geminivirus Rep-interacting kinases (GRIKs) 1 and 2, which both play critical roles in sugar/energy signaling by phosphorylating SnRK1-type kinases, were identified also to phosphorylate CIPK24 within its activation loop at Thr168. This phosphorylation is required for full SOS2 activation and accurate responses during salt stress. In the absence of salt stress, CIPK24 is kept in an inactive state by both the flowering time regulator GIGANTEA and the 14-3-3 proteins λ/κ. Whether these inhibitory interactions interfere with CBL binding to CIPK24, or if they represent CBL-independent mechanisms, remains to be addressed. Within the aerial parts of the plant, upon salt stress Na+ sequestration into the vacuole is a strategy to retain cytoplasmic Na+ concentrations in a tolerable range. Here, CBL10/CIPK24 complexes probably activate a still unknown mechanism of Na+ transport into the vacuole. One of the toxic effects of Na+ within the cells is the perturbance of the Na+/K+ ratio. To counteract this effect triggered by Na+ influx into cells during salt stress, the import of K+ ions can be enhanced to achieve adjustment of the Na+/K+ ratio by Ca2+ elevations through activation of the plasma membrane-localized K+ transporter AKT1 by CBL1/9—CIPK23 complexes. The regulation of the pH gradient across the plasma membrane is crucial for salt tolerance responses, as this gradient represents the driving force for the SOS1-mediated Na+ extrusion from the cytosol. In the absence of salt stress, the AHA2 proton pump is negatively regulated by CBL2 and/or CIPK11 to prevent excess ATP consumption. During the onset of stress, this negative regulation is released. During endodermal differentiation, a ring-like, lignified tertiary cell wall deposition called the Casparian strip is formed to block the apoplastic and the coupled trans-cellular pathway into the stele to prevent excess uptake of toxic elements such as Na+ into, as well as leakage of valuable metabolites from, the vasculature. For this barrier formation, precisely localized activity of the NADPH oxidase RBOHF is required to produce H2O2 needed for lignin cross-linking by peroxidases. RBOHF is known to be directly regulated by Ca2+ binding and phosphorylation by CBL/CIPK26 complexes. rbohF mutants with disturbed Casparian strip formation display a specific transpiration-dependent salt stress phenotype. The SOS pathway appears to be conserved in every plant species that has been studied in this regard, and experimental evidence for its functional conservation has been obtained not only for higher plants, but also for mosses such as Physcomitrella patens (Benito and Rodríguez-Navarro, 2003; Fraile-Escanciano et al., 2010). Strong evidence for such functional conservation has, for example, been obtained by heterologous complementation of Arabidopsis sos mutants with respective SOS cDNAs from different species, including, for example, the crops maize and rice (Martínez-Atienza et al., 2007; Wang et al., 2007; Olías et al., 2009). Comparative evolutionary studies suggest that SOS3/CBL4 and SOS2/CIPK24 may even constitute the most basal representatives of their respective gene families with single homologs already present in green algae, suggesting that a precursor of this pathway and its regulation by Ca2+ may have been already functional very early in plant evolution (Edel and Kudla, 2015). Recent research has further corroborated the central role of the kinase SOS2 in salt tolerance, but has also extended the complexity of the Ca2+ sensing components that contribute to the SOS pathway. The Ca2+ sensor CBL10/SCaBP8 has also been found to form complexes with SOS2. In contrast to the root-expressed CBL4, CBL10 is predominantly expressed in the aerial tissues of Arabidopsis. Here, CBL10–SOS2 complexes can again target SOS1 and in this way contribute to salt extrusion over the plasma membrane (Quan et al., 2007). Moreover, CBL10–SOS2 complexes are also localized at the tonoplast membrane and appear to activate an as yet unknown transport protein for Na+ sequestration in the vacuole (Fig. 1). In line with this latter function, cbl10 mutants accumulate significantly less Na+ during salt stress despite their strong salt sensitivity (Kim et al., 2007). Most recently, analyses of a tomato T-DNA insertion mutant in combination with complementation of an Arabidopsis cbl10 mutant with a tomato CBL10 cDNA provided strong evidence for a conserved function of tomato CBL10 in salt tolerance by regulating Na+ fluxes in the vacuole (Egea et al., 2018). Also, in poplar (Populus trichocarpa), which expresses two CBL10 homologs, both isoforms have been localized at the tonoplast and have been shown to function in shoot salt tolerance via interaction with P. trichocarpa SOS2 (Tang et al., 2014). While CBL4 function has so far only been specifically associated with salt tolerance, CBL10 appears to be a multifunctional Ca2+ sensor. In tomato, CBL10 together with CIPK6 were found to interact with RBOHB at the plasma membrane and to contribute to the regulation of ROS generation during effector-triggered immunity (de la Torre et al., 2013). Moreover, in rice, variations in the OsCBL10 promoter have been reported crucially to determine the flooding tolerance of the respective rice cultivars by affecting OsCIPK15 protein accumulation (Ye et al., 2018). These cases already clearly illustrate the integrative capacity of Ca2+ signaling to interconnect salt stress responses with additional physiological processes at the level of Ca2+ sensor function. Finally, in Arabidopsis, it was found that CBL10 can also interact with the K+ channel Arabidopsis K+ Transporter 1 (AKT1) and, in this case, counteract effective accumulation of AKT1 at the plasma membrane to regulate K+ uptake negatively (potentially in the absence of salt stress) (Ren et al., 2013; Yang and Guo, 2018). Such a function would be in line with the importance of maintaining an optimal K+/Na+ ratio in plants for sustaining salt tolerance and may also explain the salt-sensitive phenotype of cbl1 mutants that has been reported previously (Albrecht et al., 2003; Cheong et al., 2003; D’Angelo et al., 2006). The Ca2+ sensor CBL1 constitutes a major activator of the K+ uptake channel AKT1, and the impairment of AKT1 activation in cbl1 mutants may underlie their salt-sensitive phenotype (Cheong et al., 2003; Xu et al., 2006) (Fig. 1). In the future, it will be most interesting to address whether other CBL proteins or alternative Ca2+ sensors contribute to the fine-tuning or functional flexibility of the core SOS pathway. Integrating the SOS pathway with a plant’s daily routine Salt stress can hit plants by chance at any given time and under any given physiological condition. Consequently, the ability for a prompt and appropriate activation of the SOS pathway (in particular) and its co-ordination with either ongoing developmental processes, physiological programs, or other stress responses is crucial for a plant’s fitness. Recent research has provided first important insights into such interconnections. One emerging principle of SOS2 regulation and co-ordination appears to be phosphorylation by other kinases (and of course the corresponding dephosphorylation). In this regard, it was found that SOS2 directly interacts with the protein phosphatase ABI2, which prominently functions in ABA signaling (Ohta et al., 2003). Unfortunately, the physiological and mechanistic consequences of this interaction still await further elucidation. However, endodermis-specific expression of the abscisic acid (ABA)-insensitive 1-1 mutant protein (ABI1-1), a dominant-negative version of the ABA-regulated phosphatase ABI1, which is most closely related to ABI2, negatively affected plant salt responses, underscoring the important role of ABA signaling and its likely interconnection with Ca2+ signaling in plant salt tolerance (Duan et al., 2013; Dinneny 2015). More recently, Yan Guo and colleagues identified and investigated the consequences of phosphorylation at amino acid Ser294, which is located in the junction domain of SOS2 (Zhou et al., 2014). In non-stressed conditions, this residue is phosphorylated and allows for interaction with the 14-3-3 proteins λ and κ. This binding of 14-3-3 proteins to SOS2 renders this kinase inactive and thereby maintains the SOS pathway in a dormant status (Fig. 1). Salt stress probably triggers dephosphorylation of S294 (by an as yet unknown phosphatase), resulting in 14-3-3 dissociation from SOS2 and thereby allowing for activation of the SOS pathway. Moreover, salt stress has been found to accelerate 26S–proteasome-mediated degradation of both 14-3-3 proteins (Tan et al., 2016). These conclusions were further corroborated by genetic analyses. A non-phosphorylatable S294A version of SOS2 failed to complement the salt-sensitive phenotype of sos2 mutants. Also, 14-3-3 λ/κ double mutants displayed an enhanced salt tolerance when compared with the wild type, a phenotype in line with their negative regulatory role in this pathway. This mechanism potentially ensures rapid SOS pathway activation immediately after the onset of the stress, as well as pathway stabilization. The dephosphorylation of SOS2 instantly turns on the pathway, while simultaneous degradation of 14-3-3s facilitates its continuous function. Another trans-phosphorylation site, Thr168, was found to interconnect the SOS pathway with the energy status of the plant (Barajas-Lopez et al., 2018). This residue was determined to be phosphorylated by the kinases GRIK1 and 2 which previously have been reported to activate SnRK1s that function centrally in metabolic regulation in eukaryotes. GRIK phosphorylation appeared to enhance SOS2 activity and, accordingly, mutation of Thr168 prevented complementation of salt sensitivity of a sos2 mutant. Moreover, grik1/2 double mutants were sensitive to Na+ stress. Reconstitution analysis of the SOS pathway in an appropriate yeast mutant background also indicated that GRIK co-expression increased the capacity of SOS proteins to confer salt tolerance. Considering the dual role of GRIKs, these data identify a mechanism that may allow for a co-ordinated modulation of the plant sugar-sensing/energy status via SnRK1 phosphorylation and salt stress tolerance via SOS2 phosphorylation. Intriguing insights into how a plant’s salt tolerance can be directly interconnected with the regulation of fundamental developmental programs such as plant flowering were enabled by the finding that the well-known circadian clock, flowering time, and light signaling regulator GIGANTEA (GI) directly interacts with SOS2 (Kim et al., 2013). Interaction of SOS2 with GI appears to interfere with SOS2 function in the SOS pathway and cages this kinase in the nucleus and cytoplasm (Fig. 1). Salt stress triggers GI degradation and thereby releases SOS2, which most probably allows for SOS3 interaction and, in this way, enhances its function in the salt stress pathway. This process of SOS2 caging and release is reflected in the corresponding phenotypes. GI-overexpressing plants are more salt sensitive than wild-type plants, whereas GI mutants are markedly salt tolerant. Together these findings identify GI as a transitory modulator of the salt stress response that enables integration of stress responses with flowering time regulation and provides a mechanistic explanation for the long-observed impact of salt stress on flowering time regulation in plants. Multifaceted roles of Ca2+ in salt stress responses Despite the obvious central role of Ca2+ signaling in activating the core SOS pathway, recent findings suggest that the function of this second messenger in salt stress responses may be even more versatile than so far appreciated. In this regard, accumulating evidence points to the involvement of a diverse array of Ca2+ sensor proteins in the various aspects of salinity tolerance. For example, CML9 [a member of the calmodulin-like (CML) gene family] was found to be up-regulated during salt stress, and cml9 loss-of-function mutants displayed hypersensitivity in germination assays on medium containing either NaCl or ABA, whereas adult cml9 plants show enhanced tolerance towards irrigation with salt water (Magnan et al., 2008). While transcriptomic analysis revealed altered expression of several stress marker genes in cml9 mutants, the target proteins of CML9 and the exact nature of the regulated process(es) remain to be uncovered. Moreover, the Ca2+-dependent protein kinase 3 (CDPK3, designated as CPK3 in Arabidopsis) was found to interact with the vacuolar two-pore K+ channel 1 (TPK1) in planta and to phosphorylate it in a Ca2+-dependent manner in vitro (Latz et al., 2013). As this phosphorylation is specifically induced in planta upon salt stress and as both tpk1 and cpk3 loss-of-function mutants show salt-sensitive phenotypes, this study points to a function of CPKs in the regulation of the K+/Na+ ratio during saline conditions (Latz et al., 2013). Another example in this regard is the function of the NADPH oxidase RBOHF in transpiration-dependent salt stress signaling (Jiang et al., 2012; Köster et al., 2018). The genomes of higher plants encode NADPH oxidases as gene families of considerable size with, for example, 10 members in Arabidopsis. In plants, these proteins are usually referred to as ‘respiratory burst oxidase homolog’ (RBOH) proteins, with alphabetic nomenclature. These proteins are localized at the plasma membrane and have been reported to regulate various biological processes including pathogen responses, plant development, and abiotic stress tolerance by their ability for a fine-tuned production of ROS as second messengers. (Sagi and Fluhr, 2006; Miller et al., 2009; Steinhorst and Kudla, 2013) A common feature specific to plant RBOH proteins, that directly links Ca2+ signaling to ROS signaling, is the occurrence of two Ca2+-binding EF-hands in their N-terminal, cytoplasmic domain. These elements of Ca2+ regulation are also present in RBOHF, which has been shown to be strongly expressed in the endodermal cell layer and to play a crucial role in the formation of the Casparian strip, which functions as a diffusion barrier, preventing both uncontrolled access of solutes into the vasculature and unwanted leakage out of the stele (Jiang et al., 2012; Lee et al., 2013). This function in plant development coincides with a second striking phenotype in that rbohF mutants are hypersensitive to salt stress when they are grown on soil, but indistinguishable from the wild type in their salt tolerance when grown in vitro on agar plates. The reason for this phenotypic difference lies in the transpiration dependence of enhanced Na+ accumulation in the stele, the xylem sap, and shoot tissues, that is causing this mutant phenotype (Jiang et al., 2012). Although the available data suggest that the transport and accumulation of multiple ions appear to be affected in mutants defective in endodermal barrier formation, the supposed agronomical relevance of this transpiration-dependent salt tolerance pathway argues for a consideration of this process in the context of plant salt tolerance (Pfister et al., 2014; Doblas et al., 2017; Köster et al., 2018). Currently, it cannot be fully distinguished whether the morphological alterations in rbohF mutants or the modulation of ROS generation (or even both aspects together) are causative for the resulting salt sensitivity. Independent studies identified RBOHF as a target of Ca2+-activated CBL1–CIPK26 complexes (Drerup et al., 2013). Unfortunately, a potential role for CBLs or CIPKs in general, or more specifically for CBL1 and CIPK26, in transpiration-dependent salt tolerance has not been addressed. Nevertheless, the dual indication for a regulatory role for Ca2+ in ROS production (via Ca2+ binding to the EF-hands and via Ca2+-dependent phosphorylation) by RBOHF leads to a tempting hypothesis that this second messenger plays a considerable role in endodermal differentiation and transpiration-dependent salt tolerance. This facet of Ca2+ signaling clearly deserves more attention in the future. Establishment of salt tolerance not only requires proper adjustment to the acute stress, but also ‘damage surveillance’ and implementation of growth recovery. One of the primary targets of toxic Na+ ions is the cell wall. Novel findings have linked Ca2+ signaling to these aspects of salt tolerance. Feng and colleagues investigated the contribution of the FERONIA (FER) receptor kinase to salt stress tolerance in Arabidopsis (Feng et al., 2018). Quite intriguingly, they identified a novel class of Ca2+ signals that occurred as late-stage, stress-induced, local transients mostly in the early elongation zone for up to 15 h. These transients were localized to individual cells, persisted for <1 min, and were spatially and temporally correlated with growth recovery of the root. Importantly, these transients were strongly reduced in frequency in fer mutants, which correlated with their inability to recover growth and the frequent occurrence of salt stress-induced cell bursts in this mutant. Altogether, this study identified FER-dependent Ca2+ signaling as a consequence of salinity-induced softening of the cell wall and provided evidence for a FER-dependent process of cell wall integrity restoration and root growth. In this context, the study also uncovered a novel extracellular toxicity of salinity and again highlighted that salt tolerance is manifested not only at the cellular level, but also on the level of tissue and organ integrity. A co-ordinating role for Ca2+ signaling in plant nutrition Any snapshot depicting the distribution of nutrients and essential ions within a plant would reveal spatially defined concentration differences that record the physiological situation of the plant, and in any case the accumulation of nutrients and ions would highly exceed their concentration in the soil. This situation defines nutrient and ion transport processes as central for plant biology. Consequently, the identification and characterization of such transporters has been a central subject of plant research for many years. However, during the past decade, the regulation of such transporter and channel proteins has taken center stage of research. One emerging outcome of these investigations is that Ca2+-regulated phosphorylation (often in combination with phosphorylation/dephosphorylation processes that are triggered by hormones or initiated by receptors such as kinases) appears to be a common theme of transport regulation (Osakabe et al., 2014; Kudla et al., 2018). Ca2+-dependent phosphorylation can be brought about by Ca2+-dependent kinases (CDPKs designated as CPKs in Arabidopsis) or by a Ca2+ decoding network that is formed by CBLs (with 10 members in Arabidopsis), which interact with CIPKs (with 26 members in Arabidopsis) (Kolukisaoglu et al., 2004; Weinl and Kudla, 2009; Wilkins et al., 2016). As described above CBL–CIPK complexes were first implemented in salt tolerance. However, more and more research reveals that they appear to be involved in the regulation of nutrient uptake and ion transport of any molecule that has been studied intensively enough. Quite remarkably, out of the 26 CIPKs, one particular kinase, namely CIPK23, appears to function as a kind of central nutrient regulator, since it functions in a remarkably large number of nutrient-related processes (Fig. 2). This situation will be discussed in more detail below. Fig. 2. View largeDownload slide Arabidopsis CIPK23 represents a general regulator of ion homeostasis and nutrient uptake. The transport protein IRT1 is the major uptake facilitator for iron in Arabidopsis. Under low iron conditions, the transporter’s expression is up-regulated, and it accumulates at the plasma membrane. In addition to iron, other metals Zn, Mn, and Co can also enter the root through IRT1. Since overaccumulation of the latter ions is highly toxic, the accumulation and stability of IRT1 in the plasma membrane is tightly regulated on the post-transcriptional level. In the presence of low concentrations of non-iron metals (+ METALS), the cytoplasmic loop of IRT1 is monoubiquitinated (U) at two lysine residues. This modification initiates endocytosis of IRT1 to reduce the pool of active metal transporters at the plasma membrane. De-ubiquitination (DUB) of IRT1 allows recycling of those endosomes back to the membrane. Upon higher (even more toxic) non-iron metal concentrations (+++ METALS), these metals, especially Mn, directly bind to a histidine-rich stretch within the cytoplasmic loop. Subsequent recruitment of the kinase CIPK23 and CIPK23-mediated phosphorylation within the cytoplasmic loop of IRT1 initiates polyubiquitination. Dual post-transcriptional modification by phosphorylation and polyubiquitination triggers subsequent sorting of IRT1 towards late endosomes and degradation within the vacuole. Whether CBL1 and/or CBL9 have any influence on this CIPK23-regulated process remains to be explored. Both the Arabidopsis high affinity K+ uptake transporter HAK5 and the low affinity channel AKT1 are activated by complexes consisting of the kinase CIPK23 and interacting CBL1/9 Ca2+ sensors, which are probably activated by Ca2+ signals triggered during K+-deficient conditions. Besides the macronutrient K+, the uptake of nitrogen is also regulated by CBL1/9–CIPK23 complexes. The nitrate transporter NRT1.1/NPF6.3/CHL1 is switched from low affinity mode to high affinity mode by phosphorylation mediated by the CBL1/9–CIPK23 complexes, enabling nitrate uptake at low external concentrations of this nutrient. The ammonium transporters AMT1.1 and AMT1.2 were shown to be negatively regulated by the same sensor–kinase complexes. Regulation of Mg2+ homeostasis under excess concentrations of these ions is impaired in mutants lacking the tonoplast-localized Ca2+ sensors CBL2 and CBL3, as well as in cipk3/9/23/26 quadruple mutants. Note that the targets of the CBL2/3–CIPK complexes still await identification. Besides SnRK2 kinases and CPKs, CBL1/9–CIPK23 complexes represent regulatory elements that can activate the anion channel SLAC1, which represents an essential component of stomatal closure. Together, these findings strongly support the conclusion that CIPK23 represents a central integration and co-ordination hub of plant ion homeostasis. Fig. 2. View largeDownload slide Arabidopsis CIPK23 represents a general regulator of ion homeostasis and nutrient uptake. The transport protein IRT1 is the major uptake facilitator for iron in Arabidopsis. Under low iron conditions, the transporter’s expression is up-regulated, and it accumulates at the plasma membrane. In addition to iron, other metals Zn, Mn, and Co can also enter the root through IRT1. Since overaccumulation of the latter ions is highly toxic, the accumulation and stability of IRT1 in the plasma membrane is tightly regulated on the post-transcriptional level. In the presence of low concentrations of non-iron metals (+ METALS), the cytoplasmic loop of IRT1 is monoubiquitinated (U) at two lysine residues. This modification initiates endocytosis of IRT1 to reduce the pool of active metal transporters at the plasma membrane. De-ubiquitination (DUB) of IRT1 allows recycling of those endosomes back to the membrane. Upon higher (even more toxic) non-iron metal concentrations (+++ METALS), these metals, especially Mn, directly bind to a histidine-rich stretch within the cytoplasmic loop. Subsequent recruitment of the kinase CIPK23 and CIPK23-mediated phosphorylation within the cytoplasmic loop of IRT1 initiates polyubiquitination. Dual post-transcriptional modification by phosphorylation and polyubiquitination triggers subsequent sorting of IRT1 towards late endosomes and degradation within the vacuole. Whether CBL1 and/or CBL9 have any influence on this CIPK23-regulated process remains to be explored. Both the Arabidopsis high affinity K+ uptake transporter HAK5 and the low affinity channel AKT1 are activated by complexes consisting of the kinase CIPK23 and interacting CBL1/9 Ca2+ sensors, which are probably activated by Ca2+ signals triggered during K+-deficient conditions. Besides the macronutrient K+, the uptake of nitrogen is also regulated by CBL1/9–CIPK23 complexes. The nitrate transporter NRT1.1/NPF6.3/CHL1 is switched from low affinity mode to high affinity mode by phosphorylation mediated by the CBL1/9–CIPK23 complexes, enabling nitrate uptake at low external concentrations of this nutrient. The ammonium transporters AMT1.1 and AMT1.2 were shown to be negatively regulated by the same sensor–kinase complexes. Regulation of Mg2+ homeostasis under excess concentrations of these ions is impaired in mutants lacking the tonoplast-localized Ca2+ sensors CBL2 and CBL3, as well as in cipk3/9/23/26 quadruple mutants. Note that the targets of the CBL2/3–CIPK complexes still await identification. Besides SnRK2 kinases and CPKs, CBL1/9–CIPK23 complexes represent regulatory elements that can activate the anion channel SLAC1, which represents an essential component of stomatal closure. Together, these findings strongly support the conclusion that CIPK23 represents a central integration and co-ordination hub of plant ion homeostasis. Plant potassium homeostasis K+ represents the most abundant inorganic ion in plants which accumulates at cytoplasmic concentrations of ~100 mM, while the abundance of K+ in the soil is usually within the micromolar range (Wang and Wu, 2013). When challenged with salt stress, plants attempt to maintain a high K+ to Na+ ratio in the cytosol. They do this by regulating the expression and activity of K+ and Na+ transporters and H+ pumps that generate the driving force for transport. Plants must maintain cytosolic K+ at ~80 mM for optimal growth even under adverse conditions (Shabala and Pottosin, 2014). It has been long known that plasma membrane hyperpolarization represents a fast response to K+ deprivation (Demidchik et al., 2002). However, despite the reported Ca2+ -dependent regulation of K+ channels >10 years ago, only recently has the occurrence of specific Ca2+ signals in defined regions of the roots in response to K+ depletion been unambiguously detected (Xu et al., 2006; Behera et al., 2017). Central for plant supply with K+, especially under limiting conditions, are the K+ channel AKT1 and the Arabidopsis thaliana High Affinity K+ transporter 5 (HAK5). A breakthrough study in 2006 identified AKT1 as subject to regulation by CBL1/9–CIPK23 complexes (Xu et al., 2006). Phosphorylation of AKT1 by this Ca2+ sensor–kinase complex was found to be absolutely essential for channel activity, and later studies identified the protein phosphatase 2C (PP2C)-type phosphatase AIP1 as counteracting this activation (Xu et al., 2006; Cheong et al., 2007; Lan et al., 2011). This regulation of K+ uptake is recapitulated in rice roots, where OsCBL1–CIPK23 complexes activate OsAKT1 (Li et al., 2014). Low K+ conditions also induce the expression of HAK5. HAK5 encodes a high-affinity K+ transporter, which facilitates K+ uptake at low external concentrations that are thermodynamically unfavorable for channel-mediated influx (Nieves-Cordones et al., 2016). Quite remarkably, it was recently found that HAK5 activity in Arabidopsis roots is also positively regulated by CIPK23-mediated phosphorylation in conjunction with CBL1 and CBL9 (Ragel et al., 2015). This simultaneous regulation of both K+ uptake components by the very same kinase obviously provides the opportunity for co-ordinating these processes. However, details of the underlying mechanisms are still awaiting further elucidation. Moreover, it will be most interesting to clarify how alternative activation of either AKT1 or HAK5, which may be favorable under specific conditions, could be brought about by one and the same kinase. Uptake and homeostasis of nitrogen-containing molecules Like K+, nitrogen also represents an essential component of plant nutrition. Most plants can obtain this nutrient mainly in the form of inorganic compounds such as NO3− and NH4+. The inter-relationships of NO3− and NH4+ in plant nutrition are manifold and complex (Xuan et al., 2017). Simply formulated, whenever possible, a plant would prefer to obtain NO3−, instead of NH4+. In contrast to K+, where concentration decreases have been found to induce Ca2+ signals in roots, for NO3− it has been observed that an increase in concentration triggers Ca2+ signals (Riveras et al., 2015; Liu et al., 2017). NO3− uptake and distribution within plants are facilitated by a complex family of NO3− transporters. One important transporter that mediates NO3− uptake in plants is NRT1.1/NPF6.3. Remarkably, this protein appears not only to function in NO3− transport, but simultaneously exerts a function as a receptor (transceptor) for this ion (Ho et al., 2009; Noguero and Lacombe, 2016). Ca2+-dependent phosphorylation has been reported to regulate the NO3− uptake activity negatively and to affect the sensing function of NPF6.3. Subsequent structure–function analysis of the CIPK23 target residue Thr101 revealed that NPF6.3 activates several distinct signaling responses and that the phosphorylated and non-phosphorylated forms at Thr101 have distinct signaling functions (Léran et al., 2015). Moreover, the Ca2+ sensor CBL1 and the PP2C ABI2 were identified as additional components regulating NPF6.3, which is inhibited by the stress response hormone ABA. ABI2-mediated dephosphorylation appeared to enhance NPF6.3-dependent NO3− transport sensing and signaling (Léran et al., 2015). Consequently, these results not only highlight the general function of CIPK23 in nutrient homeostasis, but they also suggest that co-regulation of ion transport by Ca2+ and ABA may functionally link stress-regulated control of growth with energy-expensive nitrate utilization. Intriguingly, CIPK23 was found to bring about negative regulation of NH4+ uptake (again in combination with CBL1 and CBL9) by negatively regulating the ammonium transporters AMT1.1 and AMT1.2 (Straub et al., 2017). In addition to providing the obvious opportunity to regulate NO3− and NH4+ influx antagonistically, this finding, however, raises the difficult question of how one pair of Ca2+ sensors (CBL1 and CBL9) can bring about the specificity in decoding Ca2+ signals that allows the execution of such divergent response reactions. Quite remarkably, the other major family of Ca2+ signal decoding kinases—the CDPKs—has also been found to function crucially in regulating NO3− signaling and primary NO3− responses. Extensive mutant studies involving cpk triple mutants together with the elegant engineering of chemical switchable cpk mutant versions identified CPK10, CPK30, and CPK32 as central components of nitrate signaling (Liu et al., 2017). These kinases phosphorylated conserved NIN-like protein (NLP) transcription factors to adjust specifically the expression of downstream genes for NO3− assimilation and metabolism. Together with the work on Ca2+-regulated NO3− transport, these findings underscore the conclusion that Ca2+ signaling networks integrate many processes of plant physiology to adjust plant growth and development appropriately to nutrient availability, thereby ensuring the developmental plasticity of plants. Magnesium ion homeostasis Mg2+ is also an essential ion for plants; it accumulates in relatively high cytoplasmic concentrations and regulates a multitude of cellular processes. However, excess Mg2+, under specific conditions, such as, for example, in serpentine soils, can also be toxic to plants, underscoring the importance of a well-balanced Mg2+ homeostasis for appropriate plant development. Consequently, as for other ions, an elaborate network of Mg2+ transporters mediates uptake, distribution, and subcellular sequestration of Mg2+ in plants (Schmitz et al., 2013; Tang and Luan, 2017). In contrast to the situation with other ions, our knowledge about a potential role for Ca2+ signaling or more specifically Ca2+-dependent phosphorylation in regulating Mg2+ uptake is still very limited. However, recently, important insights into the role of Ca2+ in regulating Mg2+ sequestration have been gained. Here again, a crucial function of the CBL–CIPK signaling network in vacuole-mediated detoxification of high external Mg2+ was uncovered (Tang et al., 2015). Analysis of cbl2/3 double mutants revealed that these Ca2+ sensors are regulating vacuole-mediated Mg2+ ion homeostasis in cells (Tang et al., 2015). The cbl2/3 double mutant was hypersensitive to high concentrations of external Mg2+, and ionic profile analyses also showed a reduced amount of Mg2+ accumulation in cbl2/3 double mutant plants. Tang and colleagues found that the kinases CIPK3/9/23/26 physically interacted with CBL2 and CBL3 at the tonoplast, that cipk3/9/23/26 quadruple mutants displayed severe hypersensitivity towards excess Mg2+, and that these mutants exhibited a similar ionic profile to cbl2/3 mutants (Tang et al., 2015). These results strongly suggest that CIPK3/9/23/26 work together with CBL2/3 at the tonoplast to alleviate the toxic effects of external high Mg2+ concentrations via vacuolar sequestration. An independent study by K. Shinozaki and co-workers not only simultaneously identified the very same CIPKs as crucial components of Mg2+ homeostasis but also provided important hints that interconnect the regulation of Mg2+ distribution with the implementation of stress tolerance in plants (Mogami et al., 2015). These authors did not only find that ABA synthesis-deficient mutants or ABA signaling mutants of SnRK2s displayed sensitivity to high external Mg2+ concentrations. Most importantly, this study also provided biochemical and genetic evidence for a direct interaction between SnRK2-type and CIPK-type kinases. Consequently, these insights again uncover molecular and genetic interconnections between abiotic stress tolerance and ion accumulation, and in this way (considering the results from research in the nitrate field) point to the fundamental importance of this signaling integration for plant biology. Ca2+ signaling in metal uptake regulation Many less abundant ions such as, for example, metals fulfill essential functions in plants including as cofactors. Among these, iron is of outstanding importance to plants due to its ability to change redox states and its central role in photosynthesis and respiration (Brumbarova et al., 2015). Although abundant in nature, iron is often poorly available for plants, and diverse mechanisms for optimizing plant supply of iron have evolved. Also in this field, increasing evidence indicates that not only transcriptional regulation but also post-transcriptional regulation of iron transport processes is of crucial importance. In this regard, most recent findings have shed new light on the regulation of IRT1. IRT1 is a broad-spectrum transporter driving the uptake of iron into plants, but also of non-iron essential heavy metals such as manganese, zinc, and cobalt (Vert et al., 2002). A recent study by Dubeaux and colleagues has uncovered that IRT1 not only acts as an iron transporter, but also functions as a receptor (transceptor) in a rather sophisticated manner. For its latter function, IRT1 appears to sense directly the excess of non-iron metals in the cytoplasm, and this sensing appears to regulate its own degradation. Direct metal binding to a histidine-rich region in IRT1 triggers interaction with and phosphorylation by the Ca2+-activated kinase CIPK23 (Dubeaux et al., 2018) (Fig. 2). Although this work did not address the potential role of CBL proteins in this process, it is tempting to conclude that this mechanism indicates Ca2+ control of metal homeostasis in plants. Phosphorylation of IRT1 facilitates the recruitment of an E3 ligase, and phosphorylation and lysine polyubiquitination jointly drive endocytosis and vacuolar degradation of IRT1. In this way, IRT1 directly senses metal concentration and integrates this information for optimized iron uptake. This study not only provides new insights into the potential complexity of how Ca2+ can impact on iron homeostasis regulation, but it also raises the intriguing question of whether such an intracellular sensing mechanism of plants will be specific for IRT1 or instead may be considered as an aspect which may also be relevant for other plant transceptor proteins. Last but not least: Ca2+-controlled anion fluxes for ensuring plant nutrition Although gaseous, the nutrient function of CO2 for plants is as important as the above discussed ions. Opposite to the previous cases, CO2 uptake via the roots can be neglected and here specific pores in leaves that are designated as stomata, which are formed and regulated by the aperture of guard cells, represent the dominant uptake pathway (Hetherington, 2001; Jezek and Blatt, 2017). Nevertheless, as in roots, Ca2+ signaling (and its interactions with ABA signaling) also represents a reoccurring theme in this case. Centrally functioning in regulating guard cell aperture is the anion-conducting ion channel SLAC1 (Negi et al., 2008; Vahisalu et al., 2008; Geiger et al., 2009). This channel has become a model case for studying and illustrating the complexity of processes that converge on the regulation of ion conductance (Jeworutzki et al., 2010; Kim et al., 2010; Krol et al., 2010; Hedrich and Geiger, 2017). In general, multiple kinases from at least three distinct families and PP2C phosphatases convey the fine-tuning of this channel (Fig. 2). Again, CDPKs, CBL–CIPKs, and SnRK2s appear to form an alliance counteracting the function of ABA-inhibited phosphatases. Details of this regulation have been worked out in many very informative original publications (Geiger et al., 2009, 2010; Lee et al., 2009; Maierhofer et al., 2014; Yu et al., 2012; Brandt et al., 2012, 2015), and have been recently reviewed (Munemasa et al., 2015; Edel and Kudla, 2015). Therefore, in the context of this article, we are not elaborating on these details again. Instead we draw the reader’s attention to the most complex regulation of SLAC1 with the intention to underscore the broad biological significance of Ca2+ signaling and Ca2+-dependent phosphorylation for plant biology. Obviously, this regulatory principle has not remained restricted to the regulation of ion transport and homeostasis, but also governs other distinct biological processes. By elucidating commonalities which are shared on the level of the biological challenges to the plant and simultaneously deciphering similarities and differences in their mechanistic solution, we may advance in our quest for a comprehensive appreciation of the grace and sophistication of plants. Acknowledgements JK is grateful for support from the Deutsche Forschungsgemeinschaft (FOR964 and KU 931/14) and by a Distinguished Scientist Fellowship Program, King Saud University, Saudi Arabia. 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