Accepted Manuscript
Ion channels in plant bioenergetic organelles chloroplast and mitochondria: from
molecular identification to function
Luca Carraretto, Enrico Teardo, Vanessa Checchetto, Giovanni Finazzi, Nobuyuki
Uozumi, Ildiko Szabo
PII:
DOI:
Reference:
S1674-2052(15)00457-8
10.1016/j.molp.2015.12.004
MOLP 223
To appear in: MOLECULAR PLANT
Accepted Date: 1 December 2015
Please cite this article as: Carraretto L., Teardo E., Checchetto V., Finazzi G., Uozumi N., and Szabo
I. (2016). Ion channels in plant bioenergetic organelles chloroplast and mitochondria: from molecular
identification to function. Mol. Plant. doi: 10.1016/j.molp.2015.12.004.
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ACCEPTED MANUSCRIPT
Ion channels in plant bioenergetic organelles chloroplast and
mitochondria: from molecular identification to function
Uozumi4 and Ildiko Szabo1,2
1 Department of Biology, University of Padova, 35121, Italy
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2 CNR Institute of Neuroscience, University of Padova, 35121 Italy
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Luca Carraretto1, Enrico Teardo1,2, Vanessa Checchetto1, Giovanni Finazzi3, Nobuyuki
3 UMR 5168 Laboratoire de Physiologie Cellulaire Végétale (LPCV) CNRS/ UJF / INRA / CEA,
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Institut de Recherches en Technologies et Sciences pour le Vivant (iRTSV), CEA Grenoble, 38054
Grenoble, France
4 Department of Biomolecular Engineering, Graduate School of Engineering, Tohoku University
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Sendai, 980-8579, Japan
Correspondence should be sent to:
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Ildiko Szabo ([email protected]),Giovanni Finazzi ([email protected]), Nobuyuki Uozumi
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([email protected])
Keywords: ion channels, chloroplasts, mitochondria, cyanobacteria, endosymbiosis, plant
physiology
Abstract
Recent technical advances in electrophysiological measurements, organelle-targeted fluorescence
imaging and organelle proteomics have pushed a step forward in the research of ion transport in the
case of the plant bioenergetic organelles chloroplasts and mitochondria, leading to the molecular
identification and functional characterization of several ion transport systems during the last years.
Here we focus on channels which mediate relatively high-rate ion and water flux and summarize the
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current knowledge in this field, focusing on targeting mechanisms, proteomics, electrophysiology
and physiological function. In addition, since chloroplasts evolved from a cyanobacterial ancestor,
we give an overview of the information available about cyanobacterial ion channels and discuss
evolutionary origin of chloroplast channels. The recent molecular identification of some of these ion
channels allowed to study their physiological functions using genetically modified Arabidopsis
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plants and cyanobacteria. The view is emerging that alteration of chloroplast and mitochondrial ion
homeostasis leads to organelle dysfunction which in turn significantly affects energy metabolism of
the whole organism. Clear-cut identification of genes encoding for channels in these organelles
however remains a major challenge in this rapidly developing field. Multiple strategies including
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bioinformatics, cell biology, electrophysiology, use of organelle-targeted ion–sensitive probes,
genetics and identification of signals eliciting specific ion fluxes across organelle membranes
should provide in the future a better understanding of the physiological role of organellar channels
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and of their contribution to signaling pathways in plants.
INTRODUCTION
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Although over 80% of transport processes occur inside the cells, the mechanisms of ion flux
across intracellular membranes are difficult to investigate and remain poorly understood, except for
a few cases (e.g. plant vacuoles, (Hedrich 2012; Martinoia et al. 2012; Xu et al. 2015)). Plastids are
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the hallmark organelles of all photosynthetic eukaryotes and are the site of major anabolic
pathways. Hence, a plethora of metabolite and ion transport systems are found in plastids. Excellent
reviews deal with metabolite/ion transporters (Weber et al. 2005; Linka and Weber 2010;
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Facchinelli and Weber 2011; Rolland et al. 2012; Finazzi et al. 2015; Oh and Hwang 2015; Pfeil et
al. 2014), and therefore we will focus here on ion channels of plastids. As to mitochondria, similarly
to the mammalian organelle, the existence of ion-conducting pathways has earlier been
demonstrated by classical bioenergetic studies (for reviews see e.g. (Trono et al. 2014; Pastore et al.
2013; Millar et al. 2011; Laloi 1999; Vianello et al. 2012)) but the identity of the proteins
responsible for these activities is still largely unknown. Both organelles in plant cells serve as the
major intracellular hubs, which supply energy to the cell through the activity of membrane localized
respiratory and photosynthetic chains. To fulfil this role, they require extensive exchange of ion and
solutes also. In particularly, ion transport systems are closely related with the electrochemical
potential formation. Taking into consideration that the mitochondria and chloroplasts are of
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endosymbiontic origin (Qiu et al. 2013; Schleiff and Becker 2011; Duy et al. 2007), we address the
question of the evolutionary conservation of ion channels in relation to the possibility of exploiting
this information for identification of channels in bioenergetics organelles, mainly chloroplasts.
Knowledge about the ion channels in bacteria, like cyanobacteria, is of importance also in relation
to the structure-function of ion channels in chloroplasts and mitochondria.
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Various classes of membrane transport system are classified into channels and transporters,
the formers showing high solute permeability per second. Direct measurement of the ion currents
by electrophysiological recording approach enables to evaluate their function. Presumably a large
number of the genes encoding ion channels expressed in ancestor bacteria might be transmitted into
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the genome in the nucleus of plant cells until they have stabilized in the host cells as the
mitochondria and chloroplasts. Indeed, one of the major challenges in the field is to understand
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which of the nucleus-encoded channel-forming proteins are targeted to these organelles, since this is
a prerequisite to understand and prove their physiological roles at the level of whole organisms
using genetic tools. The sorting process and membrane trafficking of the ion channels into the
organelles from the cytosol also involves a coordinated control.
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CHLOROPLAST ION CHANNELS
Electrophysiology
Chloroplast channel activities of both the outer and inner envelope membranes (OE and IE)
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and thylakoid membranes (TM have been investigated during the last decades either by the
technically challenging direct patch-clamping of chloroplast envelope membranes and of isolated
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swollen thylakoid membranes (Pottosin 1992, 1993; Pottosin et al. 2005a; Pottosin and
Schonknecht 1996, 1995; Hinnah and Wagner 1998; van den Wijngaard and Vredenberg 1997;
Schonknecht et al. 1988) or, in most cases by incorporating membrane vesicles or purified
native/recombinant proteins into planar lipid bilayers (see Table I for references). Table I
summarizes the main characteristics of the ion channels recorded from the three membranes. The
chloroplast stroma contains approximately 150 mM K+, 50 mM Cl- and 5 mM Mg2+ as main ions
(Neuhaus and Wagner 2000). Several potassium-, and divalent cation-selective as well as anionic
ion channels have been recorded from all three chloroplast membranes with different conductance
and characteristics. Unfortunately, pharmacological characterization of these channels is rather
limited and this fact, together with the huge technical difficulty of patch clamping chloroplasts from
the completely sequenced Arabidopsis model plants either the wild type (WT) or lacking a putative
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chloroplast channel, renders difficult forwarding hypothesis about the proteins responsible for most
activities. The question also arises whether all observed activities listed in Table I. correspond to
distinct channel proteins. This is most probably not the case- different studies were performed under
different experimental conditions, making a direct comparison rather difficult. Furthermore,
although in several cases evidence has been obtained for the localization of a given putative channel
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protein in chloroplasts, direct assessment of its activity is either missing, or is studied only with
recombinant proteins in a non-physiologically relevant context (i.e. in non-native membranes) (see
Table I). Overall, it appears that while electrophysiology is essential to characterize chloroplast
channels, this technique alone is not sufficient to link unequivocally channel activity and the
previously observed physiological variations, like e.g. calcium uptake into chloroplasts (Stael et al.
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2012), increase of Mg2+ concentration in stroma during photosynthesis (Hind et al. 1974; Bulychev
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and Vredenberg 1976) and thylakoid membrane permeability changes.
Bioinformatics and proteomics: toward molecular identification
Beside the electrophysiological characterization, complementary methods may help to
understand what proteins are responsible for the observed activities. All ion channels and
transporters of chloroplasts and mitochondria are encoded by the nuclear genome, and the protein
products are imported into the organelles during biogenesis after their synthesis in the cytoplasm.
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Today, several bioinformatic algorithms exist for prediction of chloroplast or mitochondria
localization of nucleus-encoded proteins, including the most widely used prediction tools for subcellular localization, namely TargetP (Emanuelsson et al. 2000), MultiLoc2 (Blum et al. 2009) and
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WoLF PSORT (Horton et al. 2007) and ChloroP (Emanuelsson et al. 1999). Table II reports all
authentic (with proven channel function) or putative ion channels of Arabidopsis (Ward et al. 2009)
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predicted to be located to chloroplasts.
Most of these algorithms rely on the analysis of the amino acid composition of the Nterminal sequence, to detect a putative organelle targeting sequence. However alternative splicing
might lead to products with different N-terminal sequences, thus to targeting of the same protein to
different compartments (for reviews see e.g. (Karniely and Pines 2005; Yogev and Pines 2011). In
the case of mammalian system, several mitochondrial channels display dual/multiple localization
(e.g. (Szabo and Zoratti 2014; von Charpuis et al. 2015; Balderas et al. 2015). For example, the bigconductance calcium-dependent potassium channel (BKCa) has been demonstrated to be targeted to
the mitochondria or the plasma membrane based on alternative splicing (Singh et al. 2013). In
plants, relatively few examples of dually localized ion channel proteins are known so far e.g.
(Michaud et al. 2014; Elter et al. 2007; Teardo et al. 2011; Robert et al. 2012). We have recently
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demonstrated that alternative splicing-dependent mechanism accounts for dual organellar
localization of the putative channel GLR3.5 of the glutamate-receptor family protein, namely to
mitochondria and to chloroplasts (Teardo et al. 2015), suggesting that the same mechanisms leading
to dual/multiple location are present in animals and plants. Besides the possibility of alternative
splicing, other possible difficulties must be considered when trying to predict protein targeting to
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the organelles based on prediction tools. In multipass transmembrane proteins, like channels, the
targeting sequence might be internal. Furthermore and most importantly, protein-import pathways
different from the classical one via the translocon of the outer and inner membranes of the
chloroplasts (TOC/TIC) complex (Strittmatter et al. 2010; Paila et al. 2015; Rolland et al. 2012)
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operate. Indeed, a largely uncharacterized system mediates the targeting of proteins that lack
cleavable targeting signals to the chloroplast (Paila et al. 2015) and an unexpected vesicular
transport between the secretory pathway and chloroplast has been discovered in Euglena (Slavikova
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et al. 2005) as well as in higher plants (Villarejo et al. 2005). This latter pathway targets the proteins
first to the Sec translocon at the ER and subsequently exploits vesicle trafficking through the
endomembrane system for the delivery of the proteins to the chloroplast (Nanjo et al. 2006) (Paila
et al. 2015). It is however still unclear how frequently and whether these “non-canonical” pathways
which do not necessarily require N-terminal targeting sequence are used for channel proteins.
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Beside the vesicular system via the endomembrane system, vesicular transport has been proposed to
take place also within the chloroplasts, from the envelope membrane to the thylakoid via the COPIIlike system (Khan et al. 2013; Karim and Aronsson 2014). Interestingly, diacidic motifs at the Cterminus, which is typical of cargo proteins that are transported by the cytosolic vesicle transport
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system (Khan et al. 2013) have been identified in TPK3, a thylakoid-located member of the twopore potassium channel family, which plays an essential role in ion homeostasis in the chloroplast
(see below). Finally, a new “mRNA-based” mechanism of protein targeting to the organelles is
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emerging. This process could play an essential role for protein targeting to the endoplasmic
reticulum, mitochondria and chloroplasts (Weis et al. 2013). In this scenario, cis-acting elements in
the mRNA determine their subcellular localization, and mRNA become translated only once they
reached the correct location. Thus, given the complexity of the targeting mechanism to organelles,
we certainly foresee a considerable increase in the number of organelle-targeted ion channels during
the next decade. In addition, proteins annotated with unknown function might turn out to mediate
ion transport.
It is important to stress that, once hypothesized by bioinformatic analysis, the subcellular
localization of a given protein has to be proven, ideally by biochemical evidence (using specific
antibodies), or alternatively, by studying targeting in vivo, using proteins fused with fluorescent
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proteins. This latter method, which can be exploited also for the determination of the cleavage site
or the targeting peptide (Candat et al. 2013), however does not lack possible pitfalls: GFP, which is
most often used for this purpose, is rather large and might impact targeting (especially if fused to
the N-terminal part) and assembly of subunits (especially when fused to the C-termini) which is
necessary for the channel function in the case of multimeric forms. Furthermore, the generally
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strong overexpression due to strong promoters (e.g. CaMV 35S promoter) might eventually lead to
mis-targeting. Finally, expression of chloroplast-located proteins in cells lacking mature
chloroplasts or in heterologous systems (e.g. Arabidopsis proteins in tobacco cells) may yield
misleading results. Therefore, definitive and reliable protein localization studies require
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complementary strategies, also including biochemical studies. Among the different biochemical
approaches proteomic studies of purified organellar membranes e.g. (Simm et al. 2013; Salvi et al.
2011; Ferro et al. 2010; Peltier et al. 2004), and suborganelle fractions (Tomizioli et al. 2014; Yin et
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al. 2015) have been instrumental for the localization of proteins in the cell.
Mass spectrometry (MS) is a valid alternative for channel protein localization, even though,
despite technical improvement regarding sensitivity of detection, only few organelle-located
channel proteins have been identified so far in plants (Teardo et al. 2005; Hoogenboom et al. 2007;
Chang et al. 2009; Yin et al. 2015). This is likely due to very low abundance. Instead, numerous
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transporters of chloroplasts and mitochondria have been identified by MS (for reviews see e.g.
(Finazzi et al. 2015; Millar et al. 2005). In the case of the mammalian system, assembly of an
experimentally validated (by subtractive proteomics and GFP-tagged protein localization) database,
named MitoCarta (Pagliarini et al. 2008) has revolutionized the field leading to the discovery of
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new ion channels and ion/metabolite transporters (e.g. (Baughman 2011; De Stefani et al. 2011;
Herzig et al. 2012). Similarly, these approaches will expectedly lead to the identification of further
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ion channels/transporters in plant chloroplasts and mitochondria as well. In addition, the use of a
novel MS strategy in living cells, successfully applied to mammalian organelles (e.g. (Rhee et al.
2013), might be useful also in plants in order to reduce ambiguities of localization due to
contamination by other compartments and to multiple localization within the cell (Drissi et al.
2013). This method relies on a genetically organelle-targeted enzyme that modifies only nearby
proteins (e.g. by biotinylation), which can be subsequently purified and identified by MS.
Chloroplast ion channels versus cyanobacterial channels
An ancestral photoautotrophic prokaryote related to cyanobacteria is considered as the
ancestor of chloroplasts in plants and algae via an endosymbiotic event (Keeling 2013; Martin et al.
1998). After this organism was engulfed by the host cells, a large set of genes essential to plastid
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function have been transferred from the ancestral plastid genome to the nucleus e.g. (Leister 2003),
leading to the necessity of a protein targeting machinery into the plastids. Genes encoding for ion
channels and transporters of bioenergetics organelles are all encoded by the nucleus. Although
phylogenetic profiling is useful in finding nuclear-encoded chloroplast proteins of endosymbiont
origin, the physiological functions of orthologous genes may be different in chloroplasts and
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cyanobacteria (Ishikawa et al. 2009) and vice-versa, the same task might be fulfilled by unrelated
proteins (Bayer et al. 2014). Nonetheless, experimental evidence indicates that for example some
of the protein and solute/ion conducting machineries share common structure and function (Bolter
et al. 1998; Day et al. 2014; Balsera et al. 2009b). As to ion channels, many of the channel-forming
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proteins predicted to be located to chloroplast show homology with one or more cyanobacterial
protein with presumed or proven ion channel function (see Table III) and e.g. (Hamamoto and
Uozumi 2014)). For example, the glutamate receptor GLUR3.4 of Arabidopsis displays
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considerable amino acid sequence similarity to GluR0 of the completely sequenced model organism
Synechocystis (Teardo et al. 2011; Chiu et al. 2002). GluR0 (slr1257) represents a very interesting
evolutionary link between K+ channels and glutamate receptors, since it has a typical conserved
selectivity filter sequence of potassium channels, on the other hand it is gated by glutamate (Chen et
al. 1999). Indeed, many K+ channels in the Arabidopsis proteome share homology with this
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“ancient” channel protein (Schwacke et al. 2003). The mechanosensitive channels MSL2/3 of the
envelope membrane (for recent review see (Hamilton et al. 2015)) represent another example, being
similar to bacterial MscS. Instead, the Synechocystis thylakoid potassium channel SynK (Zanetti et
al. 2010) and the thylakoid two-pore potassium channel AtTPK3 (Zanetti et al. 2010; Carraretto et
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al. 2013) do not seem to share common origin (Pfeil et al. 2014), while they conserved the function
of regulating photosynthesis (see below). A cyanobacterial origin was expected for the chloride
channel family members ClC-e and ClC-f, shown to be located to thylakoids (Teardo et al. 2005;
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Marmagne et al. 2007) and to chloroplast (Teardo et al. 2005) or Golgi (Marmagne et al. 2007),
respectively. However, phylogenetic analysis did not confirm this hypothesis (Pfeil et al. 2014). In
addition, in the absence of clear comparative functional studies, it is difficult to establish if the
Synechocystis ClC and GluR0 and the chloroplast ClCe/f and GLR3.4 share the same function in
the two type of organisms: although the cyanobacterial ClC (sll0855) has been shown to work as an
electrogenic H+/Cl- antiporter with stoichiometry of 2/1 (Jayaram et al. 2011), AtClC-e has been
linked to nitrate transport (Marmagne et al. 2007). Both ClC-e and ClC-f are predicted to function
as channels rather than transporters based on their aminoacid sequences, in particular by
substitution of a conserved Glu, implicated in proton/chloride antiporter activity (Zifarelli and
Pusch 2010). Interestingly, the expression of a probable chloride channel (sll1864) increased four
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times upon inhibition of photosystem II function (Hihara et al. 2003), but no other information is
available about this protein. For GluR0, its function has not been studied in cyanobacteria to our
knowledge, while for the two chloroplast-located GLR members, AtGLR3.4 and AtGLR3.5, a
clear-cut evidence for their channel-forming ability and selectivity has still to be obtained.
Expression of AtGLR3.4 in mammalian cells results in calcium-permeable activity (Vincill et al.
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2012) and inner envelope vesicle purified from spinach chloroplasts containing members of family
3 of AtGLRs exhibited a calcium-permeable activity which was sensitive to GLR inhibitors (Teardo
et al. 2010); thus selectivity as well as functional impact of GluR0 and of Arabidopsis homolog
GLRs in the respective organisms might well be different.
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Unfortunately, even though an excellent and near-complete list of putative potassium
channels of different families has been published already ten years ago in a comprehensive review
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about prokaryotic potassium channels (Kuo et al. 2005) including those of cyanobacteria, only very
few studies addressed electrophysiological characterization and/or physiological function of these
channels. Collaborative work between our groups has led to the identification and characterization
in a heterologous expression system of SynK, a six-membrane spanning voltage-dependent
potassium channel (Zanetti et al. 2010). In vivo functional studies showed later that this channel
plays a crucial role in regulating photosynthetic activity and light responses (Checchetto et al. 2012)
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(see more details below). Checchetto et al. have also characterized SynCaK, a calcium-dependent
potassium channel whose lack leads to increased resistance to Zinc, present in the environment as
pollutant (Checchetto et al. 2013a). An additional potassium channel, KirBac 6.1 (slr5078) was
found to be important for Synechocystis growth (Paynter et al. 2010). The protein encoded by
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slr1270, which reside in the outer membrane of cyanobacteria has recently been shown to function
as a potassium-selective ion channel (Agarwal et al. 2014). In addition, a proton-gated pentameric
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ligand-gated cation channel from Gloeobacter violaceus (GLIC) was biophysically characterized,
revealing a single channel conductance of solely 8 pS (Velisetty and Chakrapani 2012; Bocquet et
al. 2009).
Another set of characterized cyanobacterial channels are members of the mechanosensitive
channel superfamily. The mechanosensitive large conductance non-specific channel MscL of
cyanobacteria (slr0875) is implicated in osmotic down-shock response (Nanatani et al. 2013) and in
cold and heat stress (Bachin et al. 2015). Interestingly, MscL has been proposed to allow calcium
efflux from cyanobacteria upon temperature stress (Nazarenko et al. 2003). The peak of the mscL
expression at the beginning of subjective night suggested that MscL might respond to stretchactivation of the plasma membrane due to the decomposition of photosynthetic materials produced
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in day time (Nanatani et al. 2013; Nanatani et al. 2015). A member of the cyclic nucleotide gated
(CNG) channels (part of the mechanosensitive channel of small conductance (MscS) superfamily
but universally considered as non-mechanosensitive channels) from Synechocystis has been proved
to respond to mechanical stress (Malcolm et al. 2012).
Water flux across the membranes is dependent on the intra- or extra-cellular osmotic
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changes due to the concentrations of solutes. Since the first identification of water channels,
aquaporins, in human cells and Arabidopsis thaliana (Preston et al. 1992; Maurel et al. 1993), the
physiological importance of the specialized water-conducting pore has been established. Plant cells
contain more than 30 copies in their genome (Maurel et al. 2008). Some of aquaporin homologs are
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localized in organelles (for recent review see (Beebo et al. 2013)); the tobacco aquaporin, NtAQP1
mediates CO2 transport in the plasma membrane and in the inner chloroplast membrane (Uehlein et
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al. 2008; Kaldenhoff et al. 2014). It has to be mentioned however that further work would be useful
to exclude with certainty that the presence of some of the AQPs in the organellar membranes is due
to contamination. Synechocystis PCC 6803 possesses a single gene encoding an aquaporin, aqpZ
(slr2057) (Fujimori et al. 2005). The AqpZ facilitated water flux across the plasma membrane (Akai
et al. 2011; Pisareva et al. 2011). The AqpZ functions in response to the osmolarity oscillations
(Shapiguzov et al. 2005; Azad et al. 2011; Akai et al. 2012) and the aqpZ mutant was sensitive to
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glucose containing in the medium, indicating that AqpZ is needed for photomixotrophic growth
(Akai et al. 2011; Ozaki et al. 2007). The expression of aqpZ was regulated by an intrinsic
biological clock, and the peak of the expression was corresponding to the early subjected night.
the carbon fixation.
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This might be coordinated with the circadian rhythm of the daily glucose metabolism supplied by
Thus, homologs of at least some of the above channels seem to be functionally present in
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chloroplasts (see Table II) and may help our understanding of the molecular identity as well as of
the function of the organelle-located channels. It also has to be mentioned, that in the case of
cyanobacterial GLIC, GluR0 and ClC, resolution of their crystal structure greatly helped
understanding of the gating mechanisms of homolog channels in higher organisms (Hilf and Dutzler
2009; Bocquet et al. 2009; Jayaram et al. 2011; Mayer et al. 2001).
In summary, combination of multiple strategies might help to discover what proteins give
rise to channel activities in the chloroplast membranes. But why is it so important to clarify this
issue? Without knowing the molecular identity of chloroplast-localized channels, final genetic proof
demonstrating their function can’t be obtained, even though important cellular and physiological
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processes seems to be regulated by these channels. Here below we focus on three aspects in which
chloroplast channels are emerging as important players.
Emerging roles of ion channels in chloroplasts and cyanobacteria: regulation of
photosynthesis
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Photosynthesis, the most important bioenergetic process on Earth, takes place in thylakoid
membranes of chloroplasts and cyanobacteria, and converts light energy into chemical energy,
ultimately supplying ATP and NADPH for autotrophic growth. Photosynthesis leads to the
generation of a proton motive force (pmf), which comprises a proton gradient (the ∆pH) and an
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electric field (the ∆Ψ). The ∆pH is generated by chloroplast lumen acidification following
plastoquinol (PQH2) oxidation by the cytochrome b6f complex and water oxidation by Photosystem
II (PSII). In parallel, H+ uptake for plastoquinone (PQ) reduction by PSII and the cyt b6f, as well as
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for NADP reduction in the stroma, makes the compartment basic in the light. The ∆Ψ is produced
by charge separation in the two photosystems (PSII and PSI) and by the activity of the cytochrome
b6f complex, the catalytic cycle of which (the so called “modified Q cycle”(Crofts et al. 1983), leads
to electron flow across the transmembrane part of this complex. The ∆Ψ is initially a localized
dipole field (positive on the luminal and negative on the stromal one), but it is rapidly delocalized
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(time of delocalization 0.1 µs ca. (Witt 1979) by ion redistribution within the hydrophilic
compartments of the chloroplast (the stroma and the lumen). Conversely ion flux between these two
compartments (i.e. across the thylakoid membranes) leads to the relaxation of the pmf. This mostly
stems from H+ translocation by the ATP synthase-ATPase CF0_Fi complex (Joliot and Delosme
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1974). This enzyme translocates H+ and thus modifies the ∆Ψ and ∆pH at the same time. CF0-Fi
activity can modulate the size, but cannot change the relative composition of the pmf. Therefore, the
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existence and implication of other ion channels/pumps in the regulation of the pmf via ion
exchanges was conceived since the formulation of the chemiosmotic theory to explain the finding
that, although the pmf is generated by a similar process in chloroplasts and mitochondria, it is
mostly composed of a ∆pH in the photosynthetic organelle and by a ∆Ψ in the respiratory one (see
e.g. review by (Bernardi 1999; Finazzi et al. 2015). Indeed, ion channels could modify the ∆Ψ/∆pH
ratio, by varying the electric field without affecting the proton gradient.
Cl- and/or K+ channels were proposed in the past as possible candidates for such a role
(Tester and Blatt 1989; Schonknecht et al. 1988). The latter possibility was recently confirmed by
the finding that some members of the TPK channel (Carraretto et al. 2013) and of the KEA K+/H+
antiporter (Kunz et al. 2014; Armbruster et al. 2014), families modulate photosynthesis. In
particular TPK3 from Arabidopsis thaliana (Carraretto et al. 2013) and its cyanobacterial
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counterpart (SynK, (Checchetto et al. 2012)), mediate K+ fluxes in the light, thereby controlling the
∆Ψ and therefore the pmf composition. Activity of the recombinant AtTPK3 channel is regulated by
protons and calcium (Carraretto et al. 2013). Silenced plants lacking AtTPK3 grown at 40 µmol
photons m–2 s–1 showed no difference with respect to WT plants, but when the light intensity was
increased to 90 µmol photons m–2 s–1, they exhibited a decreased rosette size and an increased
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anthocyan content (Figure 1a). This phenotype reflects the observed higher ∆Ψ and lower ∆pH due
to the lack of counterbalancing flux of positively charged potassium ions. This altered pmf
partitioning
results in reduced CO2 assimilation, reduced growth and also in a deficient
nonphotochemical dissipation (NPQ) of excess absorbed light, since NPQ onset is controlled by
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∆pH (see below). Conversely, K+/H+ exchangers belonging to the KEA family balance the ∆pH
and ∆Ψ during transient light shifts and in the dark, when the K+ and H+ gradients have to recover
to the situation preceding illumination. The case of both AtTPK3 and AtKEA transporters illustrate
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that in accordance with their chloroplast localization, their lack results in alteration of organelle
morphology which leads to an associated phenotype detectable at the whole-organism level.
Whether the observed chloride channel activity corresponds to ClC-e has still to be established, but
Arabidopsis plants lacking ClC-e do not display significant changes in in photosynthetic electron
flux (Marmagne et al. 2007).
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What are the consequences of the ion regulation of the pmf on photosynthesis? While ATP
synthesis can be equally fueled by the ∆pH and ∆Ψ, the rate of photosynthetic electron flow mostly
depends on the pH gradient. Indeed, the slowest step of electron transport (oxidation of PQH2 by the
cyt b6f complex) is inhibited by an acid luminal pH, leading to the so called “photosynthetic
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control”. In the light, changes in the ∆Ψ/∆pH ratio (even under constant pmf conditions) could
therefore trigger changes in the rate of electron flow, without any modification in the rate of the
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CF0-Fi enzyme. This could affect the balance between NADPH (which depends on photosynthetic
electron flow) and ATP (which only depends on CF0-Fi turnover) ratio, ultimately modifying carbon
assimilation. These effects, and their consequences on light utilization and growth were
experimentally verified in cyanobacterial and Arabidopsis mutants. Electron flow turned out to be
faster in the absence of potassium channels (i. e. a higher ∆Ψ and lower ∆pH) using time resolved
redox spectroscopy of the b6f complex (Checchetto et al. 2012; Checchetto et al. 2013b).
Consequences on ATP synthesis were also assessed, measuring membrane permeability with the
electrochromic shift of photosynthetic pigments, i.e. a modification of their absorption spectrum
caused by changes in the transthylakoid electric field (Witt 1979). This approach was however only
possible in plants (Carraretto et al. 2013), as in cyanobacteria the light harvesting pigments are not
embedded in the thylakoid membranes but rather contained in protein complexes exposed to the
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stroma. Therefore these pigments are no longer sensitive to changes in the ∆Ψ. Overall, a reduced
photosynthesis was evaluated, as shown by chlorophyll fluorescence-based assessments of
photosynthesis (Checchetto et al. 2012; Carraretto et al. 2013).
Besides consequences on electron flow and carbon assimilation, which are common to
cyanobacteria and plants, a modified composition of the pmf turned out to diminish light
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acclimation capacity in plants, via an impaired NPQ response (Carraretto et al. 2013; Armbruster et
al. 2014). The NPQ acronym (Non Photochemical Quenching of absorbed energy) refers to the
increased thermal dissipation of light energy, which is observed when its intensity overcomes the
utilization capacity of photosynthesis. NPQ is observed in both cyanobacteria and plants, but it is
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only in plants and algae that this process is directly controlled by the pmf, via ∆pH induced
conformational changes in specific protein effectors modulating thermal dissipation (Niyogi and
Truong 2013). This additional regulation of photosynthesis by proton and ion fluxes, which is
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specific of photosynthetic eukaryotes, likely triggered an altered thylakoid membrane structure,
which was observed in A. thaliana TPK3 mutants (see Figure 1b) (but not in Synechocystis lines
lacking SynK), likely reflecting the larger complexity of light harvesting regulation in eukaryotic
photosynthesis. Cation concentration might affect thylakoid structure also by counteracting negative
charges as consequence of (reversible) phosphorylation, which leads to repulsion and
destabilization of thylakoid structure (Fristedt et al. 2009). In addition, during illumination, due to
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H+ pumping by the ATPase of the inner envelope membrane, the membrane potential across this
membrane reaches ~ 100 mV (inside negative). This H+ extrusion is balanced by K+ influx across
cation selective channels of unknown identity of the IE (Heiber et al. 1995).
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Overall, these new results have opened a novel, exciting field of investigation: the molecular
regulation of photosynthesis by ion homeostasis. To complete these studies several tasks remain to
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be achieved. Besides chloride and potassium ions, other cations including magnesium, manganese
and calcium could have major roles in regulating (directly or indirectly), photosynthetic efficiency.
Indeed magnesium is also a well-established candidate for counterbalancing proton entry into the
lumen (Hind et al. 1974) and changes in stromal Mg2+ concentration are known to affect chloroplast
photosynthetic capacity by controlling H+ movement across the envelope and activation of RuBisCo
(Berkowitz and Wu 1993). Mn and calcium are necessary for the correct function of the oxygen
evolving complex (Boussac et al. 1989; Kessler 1955). Calcium plays a crucial role also in the
regulation of Calvin-cycle enzymes and recent data highlight its role in the control of cyclic
electron flow (CEF) as well (Terashima et al. 2012; Hochmal et al. 2015). Thus, research should
focus first on the identification of the molecular players involved in the flux of these ions (Nouet et
al. 2011). Moreover, the pathways allowing their entry to the correct site of action within the
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chloroplasts should be addressed, to complement their biochemical and electrophysiology studies in
vitro. Finally in vivo studied are needed to establish the link between role of each ion in
photosynthesis. Further studies should clarify to what extent the different ion channels/transporters
are involved in the regulation of pmf by comparing single and double/triple mutants and whether
their relative contribution depends on specific environmental conditions (e.g. continuous or
Emerging roles of ion channels in chloroplasts: calcium signaling
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intermittent light stress).
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Although Ca2+ is a second messenger involved in physiological and stress responses of
plants (e.g. (Choi et al. 2014; Stephan and Schroeder 2014; Michal Johnson et al. 2014; Bose et al.
2011)), little attention has been paid to the potential roles, except for the regulation of
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photosynthesis, of chloroplasts in cellular Ca2+ signaling of plant cells (Nomura and Shiina 2014;
Stael et al. 2012). Recent studies have revealed that plant mitochondria and chloroplasts respond to
biotic and abiotic stresses with specific Ca2+ signals (McAinsh and Pittman 2009; Rocha and
Vothknecht 2012; Stael et al. 2012) and it appears that organellar Ca2+ signaling, similarly to
mammals (Rizzuto et al. 2012), might be more important to plant cell functions than previously
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thought. In chloroplasts, the impact of impaired organellar Ca2+ handling on plant physiology has
been suggested by CAS and PPF1 ((Nomura et al. 2008) (Wang et al. 2003) (Petroutsos et al. 2011;
Huang et al. 2012)). The mutation of the putative chloroplastic Ca2+ sensor CAS or of the
transporter PPF1 led to impaired stomatal movement and impaired plant growth. Today, it is known
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that chloroplasts have a significant buffering capacity for Ca2+ ions, resulting in an extremely low
(150 nM) free (Aldakkak et al.) [Ca2+] under resting conditions. Importantly, this concentration can
be actively regulated: light-dependent depletion of cytosolic Ca2+ in the vicinity of chloroplasts has
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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 (Dodd et al.
2010; Sai and Johnson 2002). Interestingly, the bacterial pathogen-associated molecular pattern
(PAMP) inducers flagellin peptide and chitin both induced a rapid Ca2+ transient in the cytoplasm,
followed by a long-lasting increase in the stromal free Ca2+ level (Nomura et al. 2012), suggesting
that chloroplast calcium fluxes influence plant immunity and plant stress response.
However, despite intensive research during the last decade, the fundaments of chloroplast
Ca2+ dynamics remain unanswered. In order to fully understand the impact of Ca2+ dynamics on
plant physiology, identification of the molecular players, including calcium-transporting entities, is
of utmost importance. Electrophysiological studies suggested the existence of voltage-dependent
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Ca2+ uptake activity in the inner envelope membrane of pea (Pisum sativum) chloroplasts (Pottosin
et al. 2005a) (see Table I). For the calcium import across the inner envelope membrane (presuming
that the outer membrane is freely permeable to calcium due to the presence of porins; but see
(Bolter and Soll 2001), the most promising candidates include GLR3.4 (work is under way in our
laboratories to clarify this point) and MSL2/3 channels (see above), a Ca-ATP-ase like protein
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(ACA1)(Huang et al. 1993) as well as HMA1 P-type ATP-ase, shown be located in the envelope
membrane (Ferro et al. 2010). The nature of metal ion passing the membrane through this latter
transporter is however still debated (for recent review see (Hochmal et al. 2015). Both GLR and
MSL orthologs are present in cyanobacteria (see above), but a Ca2+-ATPase seems to be responsible
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for the ability to take up calcium in these prokaryotes (Berkelman et al. 1994). A further possibility
is represented by one the six homologs of the mammalian mitochondrial uniporter (MCU), which
displays an ambiguous N-terminal sequence, possibly allowing targeting to both mitochondria and
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chloroplasts (Stael et al. 2012). However, neither the localization, nor channel activity and the
permeability for calcium of members of the AtMCU family have been described up to now (except
for one member, found in mitochondria by proteomic approach (Wagner et al. 2015)). At present, it
is difficult to understand whether the fast-activating cation channel (FACC), recorded directly in
native pea chloroplast envelope membrane (Pottosin et al. 2005a), might correspond to one of these
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entities. Since the described channel has a large conductance (from 40 to 200 pS in 250 mM KCl), it
is unlikely to arise from ATP-ase action. The channel is rather unselective, therefore the calcium
uniporter, which at least in mammals has a strict selectivity and a tiny single channel conductance
(6 pS in 100 mM calcium) (Kirichok et al. 2004; De Stefani et al. 2011), can likely be excluded.
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Whether GLR or MSL gives rise to this activity might be addressed in future studies either by
comparing the electrophysiological activity of envelope vesicles obtained from WT and mutant
plants lacking the respective channels, or by pharmacological inhibition of FACC using GLR
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inhibitors, previously shown to affect photosynthetic efficiency (Teardo et al. 2011).
Since calcium is required for the function of the oxygen evolving complex located on the
luminal side of the thylakoid membrane, calcium must cross this membrane as well. Calcium
concentration in the lumen is estimated to be rather high (total of 15 mM) (Loro et al. 2012b),
indeed the thylakoid lumen has long been postulated as the chloroplast calcium store releasing
calcium to the stroma. Based on measurements of ion fluxes, the existence of a thylakoid localized
Ca2+/H+ antiporter was hypothesized more than ten years ago (Ettinger et al. 1999). The authors
provided evidence that both light and ATP hydrolysis are coupled to Ca2+ transport through the
formation of a trans-thylakoid pH gradient and the activity of the antiporter facilitates the light14
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dependent uptake of Ca2+ by chloroplasts and by the lumen, leading to reduced stromal Ca2+ levels.
The identity of this transporter still has to be understood. In cyanobacteria, a Ca2+/H+ antiporter
SynCax has been shown to locate to the plasma membrane and impact on salt tolerance (Waditee et
al. 2004). Although in Arabidopsis several Cax transporters show clear predicted plastidial targeting
(AtCax1,AtCax3, AtCax4), all of them are recognized as vacuole transporters (Cho et al. 2012; Mei
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et al. 2009). Post-Floral-specific gene 1 (PPF1) is another candidate for mediating calcium flux
across the thylakoid membrane since it gives rise to calcium current when expressed in a human cell
line and its expression level in Arabidopsis correlates with the calcium storage capacity of
chloroplasts (Wang et al. 2003). PPF1 shows 75% identity with Alb3, a thylakoid protein that is
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part of the YidC/Alb3/Oxa1 protein family required for membrane insertion of some thylakoid
proteins (Hennon et al. 2015). Recent structural studies revealed that YidC does not form a
transmembrane channel (Kumazaki et al. 2014). In light of this discovery, it seems worthwhile to
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revisit the channel-forming ability of PPF1 in a heterologous system other than mammalian cell
and/or in an in vitro transcription/translation system. This latter system has the advantage of
avoiding contamination by membrane proteins and avoiding appearance of channel activity due to
eventual heteromerization with endogenous channel subunits. Indeed, this system has recently been
used with success to prove channel activity of various proteins displaying biophysical/structural
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characteristics similar to those found for the same proteins expressed in e.g. E. coli or P. pastoris
(Braun et al. 2014; Deniaud et al. 2010; De Stefani et al. 2011).
In summary, to provide further insight into the physiological relevance of organellar Ca2+
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signalling, it will be crucial to identify the proteins that are responsible for calcium transport, i.e.
channels and transporters. It is clear, that the combination of the molecular players and the elicitors
of calcium signalling in organelles together with newly generated detection systems for measuring
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chloroplast (and possibly sub-chloroplast) Ca2+ concentrations in intact plants should provide
fruitful grounds for further discoveries. In this respect, the construction of the chloroplast-targeted
genetically encoded aequorin-based calcium probes represents a milestone (Mehlmer et al. 2012).
Emerging roles of ion channels in chloroplasts: plastid division
Plastid division is a fundamental aspect of the physiology of plant cells (Osteryoung and
Pyke 2014). Today, it is well-recognized that the machinery required for this process shares some
similarity to the fission-fusion machinery of bacteria as well as plant and mammalian mitochondria
(Jarvis and Lopez-Juez 2013). In this latter system, beside directly affecting mitochondrial
metabolism, defects in the fission-fusion machinery have been linked to severe pathologies
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(Kasahara and Scorrano 2014). Plastid division consist of the assembly of the division machinery at
the division site, the constriction of envelope membranes, membrane fusion and then separation of
the two new organelles (Aldridge et al. 2005). The division site in chloroplasts is established by the
FtsZ ring and the external dynamin-like ARC5/DRP5B ring, which are connected to each other via
ARC6, PARC6, PDV1, and PDV2. The evolutionarily conserved components of the Min system
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confines FtsZ-ring formation to the plastid midpoint, thereby controlling the position of the division
machinery. The two well-studied mechanosensitive channels of the inner envelop membrane, MSL2 and MSL3, not only function to relieve plastid osmotic stress (Wilson et al. 2014) but also colocalize with the plastid division protein AtMinE and as a result, plants lacking both channels have
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fewer and larger chloroplasts than the wild type. The authors suggest that these channels might
directly influence division site selection, may cause altered stromal ion homeostasis and/or impede
the constriction of the FtsZ ring (Hamilton et al. 2015). Whether and how exactly the lack of these
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two channels affects chloroplast metabolism and photosynthetic efficiency is an interesting question
that awaits answer. Independently, the use of chloroplasts isolated from plastid division-deficient
Arabidopsis mutants might, in principle, represent a solution to the so-far unresolved technical
difficulty of performing patch clamp on these tiny organelles. However, osmotic imbalance across
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the envelope membranes might turn out an issue to be resolved in order to achieve seal formation.
MITOCHONDRIAL ION CHANNELS
Electrophysiology and molecular players
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Mitochondria are crucial bioenergetic organelles for providing the ATP and metabolites
required to sustain cell growth. These organelles have also a specialized metabolic role in plants,
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since they participate in photorespiration and thus impact on photosynthesis (Linka and Weber
2005; Araujo et al. 2014). Our knowledge about plant mitochondrial ion channels is rather restricted
when compared to the animal field (Szabo and Zoratti 2014) and also with respect to chloroplast
channels, except for the mitochondrial voltage-dependent anion channel (VDAC) (for review see
e.g.(Takahashi and Tateda 2013; Homble et al. 2012) and the channels formed by components of
the protein import machinery (Schleiff and Becker 2011; Carrie et al. 2010). Indeed only very few
electrophysiological studies exist on channels other than VDAC, using mainly the lipid bilayer
system employing membrane vesicles.
Table IV. summarizes channel
the channel activities
observed in different species. These studies employ species other than Arabidopsis (Jarmuszkiewicz
et al. 2010; Koszela-Piotrowska et al. 2009; Matkovic et al. 2011), even though protocols yielding
highly purified mitochondria from this model plant exist (e.g. (Sweetlove et al. 2007; Konig et al.
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2014; Millar et al. 2005; Lee et al. 2013)). For example, in potato inner membrane vesicles,
activities displaying different conductance and kinetic behavior have been recorded. These currents
were ascribed to an ATP-regulated potassium channel (mitoKATP channel), to a large-conductance
Ca2+-insensitive iberiotoxin-sensitive potassium channel, to a (DIDS) (4,4′-diisothiocyanostilbene2,2′-disulfonic acid) -sensitive chloride channel (Matkovic et al. 2011) and to a large conductance,
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calcium-activated, iberiotoxin-sensitive potassium channel (Koszela-Piotrowska et al. 2009). For
this latter channel the authors provided evidence that as expected, activation of the channel by Ca2+
and the activator NS1619 stimulated resting respiratory rate and caused partial mitochondrial
membrane depolarization (due to entry of positively charged cations into the matrix) while the
opposite effects can be observed upon inhibition by iberiotoxin. In the animal system it is well-
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established that ion channels in the mitochondrial inner membrane with a ∆Ψ of approximately 180 mV (negative inside) can drastically influence mitochondrial bioenergetic efficiency by altering
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∆Ψ and production of reactive oxygen species, and a as consequence, cell function (Szabo and
Zoratti 2014; Bernardi 1999; Szewczyk et al. 2009; Szewczyk et al. 2006). Likely the same applies
also to plant cells. Interestingly, in mammalian mitochondria, data suggesting a structural and
functional coupling between the large-conductance calcium activated BKCa potassium channel and
respiratory chain complexes has been obtained (Bednarczyk et al. 2013). Whether this is the case
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also for plant mitochondria is still unknown, due to the technical difficulty of performing
electrophysiological measurements directly on mitochondrial membranes and to uncertain protein
composition of the observed channel.
To our knowledge, only one study deals with patch clamping of mitoplasts (swollen
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mitochondria devoid of the outer membrane) obtained from wheat germ (De Marchi et al. 2010)
where the success rate of obtaining high-resistance seals was less than 5%. Arabidopsis
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mitochondria are far too small for patch clamping (unpublished observation), which prevents taking
advantage of genetic tools. The above study identified
current carried by an ATP-sensitive
potassium channel, in accordance with previous bioenergetic experiments indicating the presence
of such activity in wheat mitochondria (Pastore et al. 1999; Trono et al. 2014).
Regarding targeting predictions, the same issues discussed for chloroplasts apply to
mitochondria. Table V. gives a list of the predicted mitochondria-located proteins among the
known Arabidopsis channels. Among the proteins identified by proteomics, beside VDAC, other
channel-forming proteins are rarely found (Qin et al. 2009). Nonetheless, at least some hypothesis
has been put forward what proteins mediate the activities observed either by electrophysiology or
by classical bioenergetics. For example, the CLCNt chloride channel of tobacco cells has been
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located to the IMM by biochemical tools (Lurin et al. 2000) and ClC from Zea mays seems to
account for plant inner membrane anion channel (PIMAC) activity observed by classical
bioenergetics (Tampieri et al. 2011). For the plant BK channel, based on biochemical indication,
the authors proposed that an AtKC1-like channel might be involved (Koszela-Piotrowska et al.
2009). AtKC1 is however a silent shaker-type subunit (Reintanz et al. 2002), therefore another, so
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far unidentified subunit must be also involved in the formation of plant BK. And what is the
molecular composition of the well-studied mitochondrial ATP-dependent K+ channel (KATP)
(Pastore et al. 2013; Petrussa et al. 2008)? This is an important question that awaits answer, even in
the mammalian system where several different possibilities have been proposed (for recent review
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see e.g. (Szabo and Zoratti 2014)). However, none of the proposed proteins (Kir6.1, Kir6.2,
ROMK) have plant counterparts, except the ABC transporters of mitochondrion AtATM1-3, which
show a high degree of sequence homology with the sulfonylurea receptors SUR1 and SUR2A (De
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Marchi et al. 2010).
Hot topics: composition of the mitochondrial permeability transition pore in plants
The permeability transition pore (PTP) is a channel giving rise to permeability increase of
the inner mitochondrial membrane under specific pathophysiological conditions and has been
characterized mainly in mammalian cells (Bernardi et al. 2015). PTP can be activated by different
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insults including high matrix calcium concentration and oxidative stress, leading to swelling of
mitochondria and dissipation of energy (Zoratti and Szabo 1995). Plant mitochondria can also
undergo permeability transition (Petrussa et al. 2004; Vianello et al. 2012; Arpagaus et al. 2002)
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and plant PTP mediates DNA import into mitochondria (Koulintchenko et al. 2003) but is also
involved in nitric oxide-induced cell death (Saviani et al. 2002).
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Over the last 50 years, several different hypotheses envisioned VDAC, the adenine
nucleotide carrier, the benzodiazepine receptor, cyclophilin D and other proteins as possible
components of the mammalian PTP. The recent discovery that under conditions of oxidative stress
in the presence of calcium dimers of the F-ATP synthase can be turned into a channel whose
electrophysiological properties match those of PTP, opened a completely new perspective to the
field (Giorgio et al. 2013). This aspect seems to be conserved in different species like Drosophila
and yeast (Bernardi et al. 2015), but whether the F-ATP synthase forms the PTP in plants as well,
still has to be clarified. In this respect, it is interesting to note that the thylakoid membrane, which
also contains this ATP-producing machinery, harbors a high-conductance channel resembling PTP
(Hinnah and Wagner 1998). What could be the physiological role of such channel in the thylakoids?
May this channel open only upon strong oxidative stress and increased calcium concentration in
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chloroplasts? And if yes, what would be the consequence for the thylakoid ultrastructure? Can we
exploit knock-out Arabidopsis plants lacking different subunits of the F-ATP synthase to obtain
genetic proof in favor of the above hypothesis?
In addition, it has to be mentioned that a recent work proposes that mitochondrial spastic
paraplegia 7 (SPG7), a nuclear-encoded mitochondrial metalloprotease (m-AAA) which interacts
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with Cyclophilin D, VDAC1 and with a paraplegin-like protein and has homologs in plants, is
essential for the PTP complex formation ((Shanmughapriya et al. 2015), but see (Bernardi and Forte
2015)), rather than the ATP synthase.
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Thus, all these open questions point to an exciting future in which a combination of genetics,
electrophysiology, bioenergetics and plant physiology will hopefully provide answers.
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Hot topics: mitochondrial ion channels and calcium signaling
Plant mitochondria have been known to contain an uptake system for calcium for five
decades (Hanson et al. 1965), the functional properties of which have been thoroughly characterized
(Dieter and Marme 1980; Zottini and Zannoni 1993). A molecular basis of the regulation of calcium
import into plant mitochondria has been lacking, even though data obtained in vivo using calcium
sensors such as aequorin probe and probes of the Cameleon family revealed differential Ca2+
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dynamics between the cytosol and mitochondria (Loro et al. 2012a; Logan and Knight 2003; Sai
and Johnson 2002; Mehlmer et al. 2012; Loro et al. 2013). In the mammalian system, following the
discovery and initial characterization of MCU (De Stefani et al. 2011; Baughman 2011) and the
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regulator MICU1 (Perocchi et al. 2010), several additional proteins such as EMRE, MCUR1 and
MCUb (a dominant negative MCU isoform) as well as MICU2 and 3 were reported to be essential
components and/or regulators of the mammalian MCU complex (MCUC; for recent reviews see e.g.
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(De Stefani et al. 2015; Foskett and Philipson 2015).
In the Arabidopsis thaliana genome six genes are present which can be identified as putative
MCU channel proteins, since they are homologues of human MCU counterparts sharing the
transmembrane domains, the pore-loop domain and the conserved DVME (Asp-Val-Met-Glu)
signature sequence (Stael et al. 2012). Five out of the six Arabidopsis MCU isoforms are predicted
to localize to the mitochondria (Schwacke et al. 2003), but our recent study identified so far only
one isoform, At1g57610 in mitochondria by proteomics (Wagner et al. 2015). Figure 2. shows
mitochondrial localization of another isoform, At2g23790, when expressed in Arabidopsis using
endogenous promoter (Teardo, Carraretto et al, unpublished). Thus, these results suggest that
bioinformatic prediction regarding localization of the MCU homologs are correct also for the other
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isoforms, even though experimental proof is still lacking. Beside the MCU isoforms, other
homologues of mammalian MCUC components in plants include two homologues of MCUR1 and
one of MICU1. However, a convincing case has recently been made for MCUR1 to act as a
cytochrome oxidase assembly factor rather than a direct and essential component/regulator of
MCUC (Paupe et al. 2015). Instead, homologs of EMRE are not present. Indeed, EMRE is required
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for calcium uptake only in the case of mammalian MCU, but not of MCU derived from fungi
(Kovacs-Bogdan et al. 2014). Thus, the plant homologues of MCU and MICU1 represent good
candidates for an involvement in mitochondrial calcium homeostasis, regulation and signaling, but
formal proof for the ability of AtMCU proteins to form calcium-selective channels is still lacking.
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As to the role of AtMICU1, recent data proves that this protein is indeed able to bind calcium and
plays an active role in the regulation of mitochondrial calcium uptake since its lack increases both
steady-state calcium level and stimulus-induced calcium uptake into mitochondria (Wagner et al.
mammalian system (Patron et al. 2014).
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2015), suggesting that AtMICU1 has a function similar to that observed for MICU2 in the
In addition, to members of the putative AtMCU family, the mitochondria-targeted splicing
isoform of AtGLR3.5 might contribute to maintenance of the mitochondrial calcium homeostasis.
Because the previously studied members of the plant iGLRs of family 3 are permeable to Ca2+,
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Teardo et al. measured calcium dynamics with the genetically encoded calcium probe Cameleon
targeted to mitochondria in WT and KO plants and found a significant difference in 4-week-old
plants challenged with wounding (Teardo et al. 2015). In accordance with the mitochondrial
localization of one isoform of this putative glutamate receptor, plants lacking AtGLR3.5 displayed
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an altered mitochondrial ultrastructure (Figure 3), in particular a part of the mitochondria displayed
enlargement and partial loss of cristae. Knock-out plants underwent anticipated senescence. The
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exact link between the reduced mitochondrial calcium uptake, the mitochondrial swelling and the
anticipated onset of senescence in the mutant plants has still to be understood, but in any case this
work provides the first indication in favor of the importance of mitochondrial calcium homeostasis
for plant physiology.
In summary, we foresee a stimulating period in this field, which has to face challenges
related to redundancy of the predicted calcium transport systems into mitochondria. Therefore, it
will be important to understand whether the different proteins cooperate and/or ensure a tissue- and
stimulus specific change in calcium dynamics.
FUTURE PERSPECTIVES
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Ion channels greatly contribute to maintain the daily active organelles, chloroplast and
mitochondria. The function of the ion channels must play a central role in the energy conversion
and electrochemical potential preservation through the light perception and carbon metabolism
across the organelle membranes. Since the endosymbiontic event, the mechanisms regulating
organelle ion channels have likely been developed to coordinate biological events of the whole cell.
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Information gained in cyanobacteria about structure/function of their easily accessible PM-located
ion channels will likely be useful for discovering the function of chloroplast channels.
Hopefully, technical improvement in proteomic techniques (increase of sensibility,
preparation of highly purified membranes) will help us to identify more and more organelle-located
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channel proteins. Once the localization of a given protein is validated also in intact cells (e.g. using
tagged proteins), its channel function can be proved either by exploiting the innovative in vitro
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translation/transcription system for expression and reconstitution of active membrane proteins
(Berrier et al. 2004; Ezure et al. 2014) or by using purified membrane vesicles. Application of the
sophisticated electrophysiological patch clamp technique to native membrane of isolated
Arabidopsis organelles would be of great advantage, but for the moment this task seems very
difficult to reach. In any case, independently of the channel activity, an indication for the in vivo
function and physiological role of the putative channel might be obtained by comparison of WT and
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mutant organisms lacking a given protein that likely forms the channel. In this respect, design of
ion-specific probes, which can be targeted to organelles are really helpful (see e.g. (Mehlmer et al.
2012; Hong-Hermesdorf et al. 2015; Loro et al. 2012b). The above approach however might lead to
misleading results, since function of a missing channel can be overtaken by e.g. another member of
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the same family. This problem, arising from redundancy of proteins with structural similarities and
likely fulfilling the same task, does not have to be under-estimated. Emerging techniques allowing
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multiple genetic manipulation (e.g. CRISPR-Cas9) in diverse organisms including plants might be
of great help in this respect. Finally, elucidation of the regulation of
plant chloroplast and
mitochondrial ion channels by post-translational modification like phosphorylation also represents a
major challenge for the future. Understanding the role, if any, of ion channels in the energetic
coupling between chloroplasts and mitochondria (see e.g. (Bailleul et al. 2015) would be also an
interesting topic for future studies.
In conclusion, a combination of diversified approach will certainly lead to a considerable
increase in our knowledge of the mechanisms of organelle function mediated by the ion channels.
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ACKNOWLEDGEMENTS
35
ACCEPTED MANUSCRIPT
The authors are grateful to all colleagues in the respective laboratories for contributing to research
and to the following funding agencies: Italian Ministry (PRIN 2010CSJX4F to I.S.), Padova
University Project (to I.S.), Japan Society for the Promotion of Science (grants 24246045,
24580100, 24658090, and 25292055 to N.U), Human Frontiers Science Program (HFSP0052 to I.S.
and G.F.), Agence Nationale de la Recherche (ANR-12-BIME-0005, DiaDomOil; ANR-
RI
PT
8NT09567009, Phytadapt, theMarie Curie Initial Training Network Accliphot (FP7-PEPOPLE2012-ITN; 316427 to G.F.).
FIGURE LEGENDS AND TABLES
SC
Figure 1. Lack of the thylakoid-located potassium channel AtTPK3 induces photosensitivity
and alteration of thylakoid ultrastructure. Representative features of wt and TPK3-silenced
plants, grown under a light intensity of 40 or 90 mmol photons m–2 s–1. A)
grown at the indicated light intensities. Note the
M
AN
U
(WT and plants silenced for AtTPK3)
Arabidopsis plants
photosensitive phenotype and the reduced rosette size in the silenced plants at the higher light
intensity, which is optimal for growth of WT plants.
B)
Abnormal, disorganized thylakoid
ultrastructure in 6-week-old plants grown at the higher light intensity was observed only in the
silenced lines, as assessed by TEM analysis (arrows). For further details see the text and (Carraretto
TE
D
et al. 2013).
Figure 2. An Arabidopsis isoform of the mammalian MCU (mitochondrial calcium uniporter)
is efficiently targeted to mitochondria in Arabidopsis plants. A) At2g23790 full locus was
expressed in fusion with EGFP in Arabidopsis leaves. Ar laser (488 nm) was used to excite GFP,
EP
fluorescence signal was recovered from 505 to 525 nm for GFP, from 680 to 720 nm for
chlorophyll autofluorescence. Images were collected using a Leica TCS SP5 II confocal system
AC
C
mounted on a Leica DMI6000 inverted microscope and LAS AF software (Leica Microsystems).
“Green” organelles prevalently localize to organelles resembling mitochondria regarding both size
(see B) and motility (please see supplementary movies). Partial localization to chloroplasts (blue
color; detection due to chlorophyll autofluorescence) can be observed in both representative images.
B) Confocal microscopy image from Arabidopsis plants stably expressing GFP fused to the
presequence of the mitochondrial β-subunit of the F1-ATP synthase (see (Zottini et al. 2008; Duby
et al. 2001)) identify mitochondria. Comparison suggests that the organelles identified in images of
A) are mitochondria.
Figure 3. Lack of the mitochondria-located isoform of glutamate receptor AtGLR3.5 affects
mitochondrial morphology. TEM images from leaves of WT and KO 6-week-old plants showing
36
ACCEPTED MANUSCRIPT
swollen mitochondria with decreased electron-density and disorganization towards the center of
their matrix in the absence of AtGLR3.5. Changes in ultrastructure are less evident in 3-week old
mutant plants with respect to WT (not shown) Since expression of AtGLR3.5 increases with aging,
the function of this putative calcium-permeable channel for mitochondrial physiology seems to be
prevalent in a more advanced state of development and/or under stress conditions like those linked
RI
PT
to aging or wounding. For further details see text and (Teardo et al. 2015).
Table 1. Channel activities observed in chloroplast membranes. The table shows the main
SC
characteristics of the channels recorded in OE (outer envelope) and IE (inner envelope) membranes
as well as of the channels residing in chloroplasts membranes, which were expressed in
recombinant form. OEP16 and OEP24 which are ) specific for the transport of amino acids and
M
AN
U
amines and of triose phosphates, ATP, inorganic phosphate (Pi), dicarboxylic acid, and charged
amino acids, respectively are not listed (for review see (Neuhaus and Wagner 2000). * genetic proof
has been obtained in favor of the proposed function.
Localizati
Maximal
Selectivity/
on
conductance
Voltage-
Pharmacology
TE
D
Name
Method
of
Proposed
Reference
detection
function
Recombinant
Solute
(Goetze
exchange
al. 2015)
Protein import
(Goetze
dependence
CHANNELS IN THE ENVELOPE MEMBRANES
Cationic channels
OEP23
OE
466 pS in 250
Cationic,
Reduction
maximal open
open probability
Oep23 of pea in
probability at
by
BLM
0 mV
spermine
500 pS in 250
Cationic,
Blocked by 2
Recombinant
mM KCl
maximal open
mM
Oep37 of pea in
probability at
sensitive
0 mV
Tic32
TOC75
OE
AC
C
OEP37
EP
mM KCl
OE
10
of
mM
CuCl2,
prepeptide
to
et
et
al. 2006)
BLM
(in
µM range)
1.3 nS in 1 M
Cationic,
Voltage-
Recombinant
Protein import
(Eckart
et
KCL
maximal open
dependent block
TOC75 in BLM
*
al. 2002)
probability at
by TrOE33
by
Patch clamp of
Metabolite
(Pottosin
Konig polyanion
chloroplasts from
transport
1993, 1992)
green
across OE
0 mV
VDAC
OE
homolog
or
520 and 1016
Cationic,
contact
pS in 100 mM
closure
sites
KCl
>±50
Blocked
at
mV
37
algae
ACCEPTED MANUSCRIPT
between
(520
OE and IE
channel),
pS
Nitellopsis
at
>±30
mV
(1016
pS
obtusa
channel)
Envelope
1.14
nS
in
conducta
membrane
250/20
nce
s
Kglutamate
cation
(spinach)
mM
at
None
voltages >50
Proteoliposomes
General
(Heiber
in BLM
diffusion pore
al. 1995)
mV and <-20
mV;
IE
35 pS in 100
slightly
Cationic
mM CaCl2
permeabl
et
of OE (porinlike)
cationic
channel
Calcium-
Closed
RI
PT
Large
Blocked by 100
Purified
inner
µM DNQX
membrane
vesicles
e channel
Glutamate
(Teardo
receptor
al. 2010)
et
from
SC
spinach
chloroplast
in
BLM
IE
60 to 110 pS
Cationic
in 250 mM
KCl
Change
of
Recombinant
Protein import
(Balsera et
selectivity
in
Tic110 from pea
channel*
al. 2009a)
Regulation of
(Pottosin et
stroma
al. 2005b)
M
AN
U
Tic110
presence of 20
in BLM
mM CaCl2
FACC
Envelope
From 40 to
Cationic;
Cation
membrane
200 pS in 250
Permeable to
+
mM KCl
channel
Blocked by 100
+
K ,Na ,
2+
µM
Gd3+,
regulated by pH
2+
TE
D
Ca Mg ,
low
Patch
clamping
of
pea
chloroplasts
alkalinization
Proteoliposomes
Electrical
open
probability at
0 mV
ATP-
Envelope
100
pS
membrane
250/20
potassiu
s
KCl
m
higher
Blocked by 1
open
mM ATP (added
porbability at
to cis side) ,
negative
partially blocked
voltages
by 200 µM CsCl
160 pS in 100
Anionic,
None
mM KCl
activation
AC
C
channel
mM
Cationic;
EP
sensitive
in
in BLM
balance of H
(Heiber
+
et
al. 1995)
flux across IE
Anionic channels
Anion
OE
channel
contact
or
at
(Pottosin
chloroplasts from
1992)
sites
negative
green
between
voltages
Nitellopsis
OE and IE
VDAC-
patch clamp of
IE
like
algae
obtusa
525 pS in 150
Anionic; open
3.8 mM succinic
BLM
mM KCl
at
anhydride
isolated
IE
vesicles
from
positive
potentials
channel
Anion
Envelope
60 pS in 100
channel
membrane
mM TrisCl
increased
open
using
only
probability
spinach
Anionic
None
Path
clamp
of
proteoliposomes
(spinach)
38
Protein import
(Fuks
and
Homble
1995)
Regulation of
(Heiber
stromal pH
al. 1995)
et
ACCEPTED MANUSCRIPT
Large
Envelope
540
pS
in
conducta
membrane
250/20
nce anion
(spinach)
Kglutamate
mM
Closed
ate
None
voltages >±50
Proteoliposomes
Involved
in
in BLM
protein import
(Heiber
et
al. 1995)
mV,
Slightly
channel
selective
for
glutamate
CHANNELS IN THE THYLAKOID MEMBRANE
Unselecti
Thylakoid
620
pS
ve large-
membrane
20/100
in
Unselective
None
mM
RI
PT
Cationic /unselective channels
Patch clamp of
Correspondent
(Hinnah and
swollen
of PTP- role in
Wagner
protein import
1998)
Patch clamp of
Counterbalanc
(Hinnah and
swollen
e
Wagner
conducta
KCl
thylakoid
nce
(occasionally
pea chloroplasts
channel
1 nS)
Thylakoid
40
pS
channel
membrane
20/100
in
mM
Cationic,
None
permeable
2+
also to Mg
(occasionally
and Ca
90 pS)
channel
Thylakoid
membrane
60 pS in 100
mM KCl, 21
pS in 50 mM
2+
thylakoid
2+
Cationic,
None
permeable to
+
2+
K Mg
2+
Ca , 24 pS in
Ca
50 mM Mg2+
active
and
; more
at
of
proton
entry into the
pea chloroplasts
lumen
Patch clamp of
Counterbalanc
+
1998)
(Pottosin
swollen
e of H entry
and
thylakoid
into the lumen
Schonknech
t 1996)
membrane from
spinach
TE
D
positive
from
M
AN
U
KCl
Cation
SC
Cation
from
potential
Divalent
cation
Thylakoid
membrane
20
pS
for
2+
Ca , 35 pS
2+
for Mg
channel
Cationic,
None
Thylakoid/liposo
(Enz
permeable to
me
1993)
divalent
patch clamp
vesicle
by
C
Potassiu
m
Thylakoid
membrane
EP
cations
93 to 122 pS
in
Cationic
Inhibited by 50
1000/300
mM TEACl
mM KCl and
AC
C
channel
5 mM MgCl2
Isolated
Counterbalanc
+
thylakoid
e of H entry
membrane
into the lumen
(Tester and
Blatt 1989)
vesicles in BLM
(cis side)
Potassiu
Thylakoid
100 pS in 100
m
membrane
mM KCl
channel
TPK3
potassiu
Thylakoid
membrane
m
35 pS in 250
Cationic
None
Thylakoid/liposo
(Enz
me
1993)
vesicle
by
C
patch clamp
Cationic,
no
Activated
by
Recombinant
Counterbalanc
(Carraretto
+
et al. 2013)
mM
voltage-
100
µM
Arabidopsis
e of H influx
Kgluconate
dependence
Calcium
and
protein in BLM
into lumen *
acidic
pH,
channel
blocked
by
5
mM Barium
Anionic channels
Anion
Thylakoid
220 pS in 100
Anionic
None
39
Thylakoid/liposo
(Enz
C
ACCEPTED MANUSCRIPT
channel
membrane
mM KCl
me
vesicle
by
1993)
patch clamp
Anion
channel
Thylakoid
100 to 110 pS
membrane
-
in 114 mM Cl
Anionic; bell-
None
Patch clamp of
+
(Pottosin
shaped
swollen
e of H entry
and
voltage-
thylakoid
into the lumen
Schonknech
dependence
membrane from
(max.
Nitellopsis
open
t 1995)
obtusa
RI
PT
probability at
0 mV
Ion channels from Arabidopsis with predicted and/or proven localization to
SC
Table II.
Counterbalanc
chloroplast membranes. Consensus values are those reported at the Aramemnon site (CP
chloroplast; M mitochondria; SP secretory pathway). Only references related to either the first
M
AN
U
description of the channel or to its localization are indicated. For homology to cyanobacterial
channels blast values and percentages of identities of aminoacid sequences between the bacterial
and Arabidopsis proteins are shown. N.d.: not determined.
Cationic
Name
Predicted
channels
consensus
Observed
Homology
localization
cyanobacterial
Cationic channels
At5g46370
channel
TE
D
value
CP: 6.9
AtTPK2
Vacuole
slr1257
AtTPK3
EP
SP: 4.9
At4g18160
(two-pore
At4g32650
AC
C
potassium channel)
AtKC1
to Reference
(Voelker et al. 2006)
44
4e-05 35% identity
CP: 5.1
Thylakoid
slr1257
SP: 5.3
membrane/ vacuole
48
(3e-06),
(Voelker et al. 2006)
30%
identity/
(Carraretto
et
al.
2013)
slr0498
41
(3e-04),
33%
identity
CP:12.3
Endoplasmatic
M: 10.5
reticulum,
slr1575
when
heteromerizes
some
55
(5e-08) 29% identity
with
becomes
targeted
to
Duby
et
al.
2008)(Jeanguenin et
shaker
channels,
(Reintanz et al. 2002)
al. 2011)
the
plasmamembrane
At5g10490
AtMSL2
CP:12.1
mechanosensitive
M: 7.6
Chloroplast
glr3870
(Haswell
63
Meyerowitz 2006)
(2e-10) 24%
identity
channel
40
and
ACCEPTED MANUSCRIPT
At1g58200
CP:3.2
AtMSL3
Chloroplast
M: 6.4
glr3870
(Haswell
and
55
Meyerowitz 2006)
3e-08 21% identity
At5g12080
CP: 4.8
AtMSL10
ER and PM
Glr3870
(Veley et al. 2014)
45 (5e-05)
AtGLR3.4
CP: 4.4
Chloroplast
and
M: 4.4
plasmamembrane
SP: 15.4
At4g01010
AtCNGC13
At3g17690
AtCNGC19
all2911
(Teardo et al. 2011)
64
(Meyerhoff
9e-11 23% identity
2005)
CP: 2.7
Thylakoid
none
M: 7.8
membrane
CP: 17.0
n.d.
none
n.d.
none
M: 7.2
AtCNGC20
CP: 16.6
At2g43950
OEP37 homolog
CP:18.8
Chloroplast
outer
membrane
At5g19620
CP: 10.1
AtToc75
Chloroplast
outer
(Yin et al. 2015)
(Kugler et al. 2009)
(Kugler et al. 2009)
(Goetze et al. 2006)
alr2269
(Eckart et al. 2002)
308
M
AN
U
envelope
al.
none
SC
At3g17700
et
RI
PT
At1g05200
3e-84 35% identity
At1g06950
CP: 23.6
TIC110
Chloroplast
inner
none
(Heins et al. 2002)
None
(Maierhofer
envelope membrane
Anionic channels
At4g27970
AtSLAH2
nitrate
(putative
CP:11.4
anion
At4g10380
AtNIP5.1
putative
boric acid channel
At4g35440
TE
D
channel)
CP:6.2
Plasma membrane
SP: 6.4
CP: 10.1
AtClC-e
n.d.
EP
CP:0
Outer
M:8.5
membrane
Table III.
2014; Duby et al.
2008)
(Takano et al. 2006)
2e-33 39% identity
Thylakoid
AC
C
AtClC-f
al.
138
M: 13.9
At1g55620
glr0003
et
envelope
tll0145
(Marmagne
et
al.
244
2007) (Teardo et al.
4e-65 34% identity
2005)
tll0145
(Teardo et al. 2005)
239
1e-63 35% identity
Proteins with predicted/proven ion channel function in Synechocytsis.
Only
references where channel function has been investigated are listed. Cyanobase database
(http://genome.microbedb.jp/cyanobase/) was used in part to obtain information about putative
channel proteins.
Accession number
Molecular
function References
41
ACCEPTED MANUSCRIPT
(hypothesized or proven)
POTASSIUM CHANNELS
SynK,
slr0498
sll0993
thylakoid-located
potassium
(Zanetti et al. 2010) (Checchetto et al.
channel
2012)
KchX, potassium channel
(Berry et al. 2003; Matsuda and Uozumi
2006) (Checchetto et al. 2013a)
Kirbac6.1, similar to potassium channel
slr5078
(Paynter et al. 2010)
RI
PT
protein
sll0536
probable potassium channel protein
(Berry et al. 2003)
Slr1270
Potassium channel
(Agarwal et al. 2014)
slr1257
Glutamate receptor (Glur0), potassium
(Chen et al. 1999)
channel gated by glutamate
Mechanosensitive ion channel
CHLORIDE CHANNELS
putative voltage-gated chloride channel
sll1864
probable chloride channel protein
(Jayaram et al. 2011)
M
AN
U
sll0855
SC
probable potassium efflux system
slr1575
(Hihara et al. 2003)
-
slr0617
unknown protein putative Cl channel
(Sato et al. 2007)
apqZ,water channel protein
(Akai et al. 2011; Azad et al. 2011;
WATER CHANNELS
slr2057
Hagemann 2011)
Hypothetical protein with similarity to
ssl2749
MECHANOSENSITIVE CHANNELS
TE
D
water channel NIP4;2
MscL,
slr0875
large-conductance
(Nazarenko et al. 2003)
mechanosensitive channel
mechanosensitive ion channel homolog
slr0639
Unknown
sll0590
protein
(Kwon et al. 2010)
putative
EP
mechanosensitive ion channel
Unknown protein putative
sll1040
sll0985
slr0109
slr0510
AC
C
mechanosensitive ion channel
Mechanosensitive ion channel MscS
unknown
protein
(Malcolm et al. 2012)
putative
mechanosensitive ion channel MscS
hypothetical
protein,
putative
mechanosensitive ion channel
Table IV. Ion channel activities recorded in plant mitochondrial membranes.
Name
Localiz
ation
Conductance
Selectivity/
Pharmacology
Voltage-
42
Method
Proposed
of
function/identi
Reference
ACCEPTED MANUSCRIPT
dependence
detection
ty
Planar
metabolite
(Abrecht et
al. 2000)
Anionic channels
OM
None
3,7 or 4, 0 nS for
Large
two
conducing
lipid
transport across
state: slightly
bilayer
OE
anionic;
(BLM)
Low-
with
conductance
VDAC
state:
proteins
cationic ;
purified
Bell-shaped
from bean
voltage-
mitochon
dependence
dria
isoforms
respectively
SC
with
maximal
open
OM
VDAC3
M
AN
U
probability at
0 mV
500 pS in 300
Cationic and
AtVDAC
Metabolite
(Berrier
mM KCl
anionic open
3
transport across
al. 2015)
states;
Arabidops
None
is
close
expressed
at
in
TE
D
EP
IM
from
Tendency to
positive
et
OE
cell-
voltages,
free
open up to
system or
±60 mV
E.coli
studied in
BLM
117 pS in 50/450
Slightly
Inhibited by 200
BLM
Correspond to
(Matkovic
mM KCl
anionic;
µM DIDS
with
PIMAC?
et al. 2011)
AC
C
MitoCl channel
RI
PT
VDAC homolog
No
activity
at
negative
isolated
IM
voltage
vesicle
from
potato
Cationic channels
Large
IM
from 500 to 615
conductance BK
pS
channel
mM KCl
in
Cationic
by
BLM
Regulation
µM
with
respiratory rate
Piotrowska
isolated
and membrane
et al. 2009)
IM
potential
iberiotoxin (IC50
vesicle
mitochondria
170 nM)
from
Activated
50/450
300
Calcium,
inhibited
by
of
(Koszela-
in
potato
K(ATP)
IM
150 pS in 150
Cationic;
Inhibited by 1
43
Patch
Regulation
of
(De Marchi
ACCEPTED MANUSCRIPT
mM Kgluconate
activated by
mM ATP
negative
voltage
(in
pipette)
clamp on
respiratory rate
mitoplasts
and membrane
from
potential
wheat
mitochondria
et al. 2010)
in
germ
164 pS in 50/450
Cationic
mM KCl
Inhibited by 1
BLM
Regulation
mM ATP
with
respiratory rate
isolated
and membrane
IM
potential
vesicle
from
potato
312 pS in 50/450
Cationic,
Insensitive
mM KCl
More active
300
calcium-
at
Calcium,
insensitive
voltage
conductance
µM
BLM
with
channel
Regulation
of
respiratory rate
and membrane
IM
potential
iberiotoxin (IC50
vesicle
mitochondria
150 nM)
from
inhibited
by
et al. 2011)
in
isolated
M
AN
U
potassium
positive
to
(Matkovic
mitochondria
SC
IM
Large
of
RI
PT
IM
K(ATP)
(Matkovic
et al. 2011)
in
Table V.
TE
D
potato
Ion channels from Arabidopsis with predicted and/or proven localization to
mitochondrial membranes. Consensus values are those reported at the Aramemnon site (CP
chloroplast; M mitochondria; SP secretory pathway).
All proteins
harbouring mitochondrial
EP
targeting sequences are listed. References given for localization concerns only studies in
Access
number
AC
C
Arabidopsis unless otherwise specified. N.d.: not determined.
Name
Predicted
consensus
Observed localization
References
value
Cationic channels
At4g18290
AtKAT2
potassium
channel
AtAKT2/AKT3
M: 8.0
Plasma membrane
SP: 6.7
M: 1.3
At4g22200
Plasma membrane
potassium channel
SP: 3.9
44
(Nieves-Cordones et al.
2014)
(Very
and
Sentenac
2002) (Held et al. 2011)
ACCEPTED MANUSCRIPT
M: 5.4
AtKCO3
CP: 2.7
At5g46360
(Voelker et al. 2006;
Vacuole
Rocchetti et al. 2012)
Potassium channel
SP: 5.5
M: 5.5
endomembrane
AtAKT1
At2g26650
CP: 4.0
localization in Lilium
n.d. in Arabidopsis
longiflorum
Potassium channel
M: 5.2
nucleotide-gated
channel
SP: 1.8
AtCNGC17
At4g30360
Cyclic
M: 1.8
nucleotide-gated
channel
At4g01010
AtCNGC13
Cyclic
nucleotide-gated channel
AtCNGC16
At3g48010
Cyclic
nucleotide-gated
Cyclic
nucleotide-gated
AC
C
channel
AtCNGC18
At5g14870
Cyclic
Plasma membrane
CP: 2.4
2013a)
(Ladwig et al. 2015)
(Yin et al. 2015)
(Tunc-Ozdemir et al.
2013b)
nucleotide-gated
channel
n.d.
SP: 7.0
EP
channel
At2g46450
Thylakoid membrane
CP: 2.7
(Tunc-Ozdemir et al.
M: 7.5
nucleotide-gated
AtCNGC12
M: 7.8
TE
D
AtCNGC11
Cyclic
Plasma membrane
SP: 1.0
M: 15.1
channel
At2g46440
Plasma membrane
SC
Cyclic
CP: 0.6
M
AN
U
At1g15990
RI
PT
et al. 2015)
SP: 4.3
AtCNGC7
(Safiarian
M: 5.1
n.d.
SP: 7.0
M: 12.7
CP: 0.6
Plasma membrane
(Frietsch et al. 2007)
SP: 3.4
M: 6.2
AtGLR2.5
At5g11210
CP: 4.5
n.d.
Glutamate receptor
SP: 1.6
AtGLR3.3
M: 7.8
At1g42540
At2g32390
Glutamate receptor
SP: 21.3
AtGLR3.5
M:13.4
SP: 2.4
45
Plasma membrane
(Li et al. 2013)
Mitochondria
(Teardo et al. 2015)
ACCEPTED MANUSCRIPT
Glutamate receptor
AtMSL1
n.d.
Mechanosensitive
CP: 6.9
mitochondrial
calcium
uniporter
isoform 1
At1g09575
Putative
mitochondrial
calcium
uniporter
isoform 2
At2g23790
Putative
mitochondrial
calcium
uniporter
isoform 3
At4g36820
Putative
mitochondrial
calcium
uniporter
isoform 4
At5g42610
Putative
mitochondrial
calcium
uniporter
M: 17.6
M:20.3
M: 17.1
M: 19.2
CP: 7.5
M: 14.5
M: 4.5
channel
At5g37610
Putative
(Lee et al. 2009)
CP: 1.4
Mitochondrial outer membrane
(Robert et al. 2012)
SP: 3.0
M: 6.6
AC
C
AtVDAC6
Mitochondrial outer membrane
SP: 4.2
EP
Voltage-dependent anion
channel
n.d.
M: 4.8
AtVDAC4
At5g57490
n.d.
TE
D
Voltage-dependent anion
n.d.
CP: 4.8
Anionic channels
At3g49920
n.d.
CP:1.2
isoform 5
AtVDAC5
n.d.
CP: 1.6
RI
PT
At1g57610
Putative
SC
channel
M
AN
U
At4g00290
M: 17.5
voltage-
dependent anion channel
n.d.
SP: 13.1
AtVDAC3
At5g15090
Voltage-dependent anion
Most
M: 7.3
abundant
in
plasma
membrane
(Robert et al. 2012)
channel
AtVDAC2
At5g67500
Voltage-dependent anion
channel
At3g01280
AtVDAC1
M: 5.8
Mitochondrial outer membrane,
plasma membrane
SP: 1.6
M: 5.7
Mitochondrial outer membrane
46
(Robert et al. 2012)
(Pan et al. 2014)
ACCEPTED MANUSCRIPT
Voltage-dependent anion
SP: 0.6
channel
M:8.5
Chloroplast
CP:0
AtClC-f
outer
envelope
membrane
(Teardo et al. 2005)
RI
PT
At1g55620
Supplementary movie 1. xyt series of stable GFP-expressing mitochondria (see Figure 2B). Ar
laser (488 nm) was used to excite GFP, fluorescence signal was recovered from 505 to 525 nm. The
SC
movie (3 frames-per-second) was collected by using a Leica TCS SP5 II confocal system mounted
on a Leica DMI6000 inverted microscope and LAS AF software (Leica Microsystems).
M
AN
U
Supplementary movie 2. xyt series of MCU::GFP transformed mitochondria by using the
agroinfiltration technique (see also Figure 2A). Ar laser (488 nm) was used to excite GFP,
fluorescence signal was recovered from 505 to 525 nm for GFP, from 680 to 720 nm for
chlorophyll autofluorescence. The movie (3 frames-per-second) was collected by using a Leica TCS
SP5 II confocal system mounted on a Leica DMI6000 inverted microscope and LAS AF software
AC
C
EP
TE
D
(Leica Microsystems).
47
AC
C
EP
TE
D
M
AN
U
SC
RI
PT
ACCEPTED MANUSCRIPT
AC
C
EP
TE
D
M
AN
U
SC
RI
PT
ACCEPTED MANUSCRIPT
AC
C
EP
TE
D
M
AN
U
SC
RI
PT
ACCEPTED MANUSCRIPT