Molecular Plant
Review Article
Ion Channels in Plant Bioenergetic Organelles,
Chloroplasts and Mitochondria: From Molecular
Identification to Function
Luca Carraretto1, Enrico Teardo1,2, Vanessa Checchetto1, Giovanni Finazzi3,*,
Nobuyuki Uozumi4,* and Ildiko Szabo1,2,*
1
Department of Biology, University of Padova, Padova 35121, Italy
2
CNR Institute of Neuroscience, University of Padova, Padova 35121, Italy
3
UMR 5168 Laboratoire de Physiologie Cellulaire Végétale (LPCV) CNRS/ UJF / INRA / CEA, 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, Aobayama 6-6-07, Sendai 980-8579, Japan
*Correspondence: Giovanni Finazzi ([email protected]), Nobuyuki Uozumi ([email protected]), Ildiko Szabo ([email protected])
http://dx.doi.org/10.1016/j.molp.2015.12.004
ABSTRACT
Recent technical advances in electrophysiological measurements, organelle-targeted fluorescence imaging, and organelle proteomics have pushed the research of ion transport a step forward in the case of the
plant bioenergetic organelles, chloroplasts and mitochondria, leading to the molecular identification and
functional characterization of several ion transport systems in recent years. Here we focus on channels
that mediate relatively high-rate ion and water flux and summarize the 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 the evolutionary origin of chloroplast channels. The recent
molecular identification of some of these ion channels allowed their physiological functions to be studied
using genetically modified Arabidopsis plants and cyanobacteria. The view is emerging that alteration of
chloroplast and mitochondrial ion homeostasis leads to organelle dysfunction, which in turn significantly
affects the 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 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 a better understanding of the physiological role of organellar channels and their contribution to signaling pathways in plants in the future.
Key words: ion channels, chloroplasts, mitochondria, cyanobacteria, endosymbiosis, plant physiology
Carraretto L., Teardo E., Checchetto V., Finazzi G., Uozumi N., and Szabo I. (2016). Ion Channels in Plant
Bioenergetic Organelles, Chloroplasts and Mitochondria: From Molecular Identification to Function. Mol. Plant. 9,
371–395.
INTRODUCTION
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 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; Facchinelli and Weber, 2011; Rolland et al.,
2012; Pfeil et al., 2014; Finazzi et al., 2015; Oh and Hwang,
2015), and therefore we focus here on ion channels of plastids.
As to mitochondria, similarly to the mammalian organelle, the
existence of ion-conducting pathways has been demonstrated
previously by classical bioenergetic studies (for reviews, see
e.g., Laloi, 1999; Millar et al., 2011; Vianello et al., 2012;
Published by the Molecular Plant Shanghai Editorial Office in association with
Cell Press, an imprint of Elsevier Inc., on behalf of CSPB and IPPE, SIBS, CAS.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
371
Molecular Plant
Pastore et al., 2013; Trono et al., 2014) but the identity of the
proteins responsible for these activities is still largely unknown.
Both organelles in plant cells serve as major intracellular hubs
that supply energy to the cell through the activity of membranelocalized respiratory and photosynthetic chains. To fulfill this
role, they require extensive exchange of ions and solutes.
In particular, ion transport systems are closely related to
the formation of electrochemical potential. Taking into
consideration that the mitochondria and chloroplasts are of
endosymbiotic origin (Duy et al., 2007; Schleiff and Becker,
2011; Qiu et al., 2013), 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, such as
cyanobacteria, is of importance also in relation to the structure–
function of ion channels in chloroplasts and mitochondria.
Various classes of membrane transport system are classified into
channels and transporters, the former showing high solute
permeability per second. Direct measurement of the ion currents
by electrophysiological recording enables their function to be
evaluated. Presumably a large number of the genes encoding
ion channels expressed in ancestor bacteria might be transmitted
into the genome in the nucleus of plant cells until they have stabilized in the host cells as mitochondria and chloroplasts. Indeed,
one of the major challenges in the field is to understand 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 coordinated control.
CHLOROPLAST ION CHANNELS
Electrophysiology
Chloroplast channel activities of both the outer and inner
envelope membranes and thylakoid membranes have been
investigated during the last decades either by the technically
challenging direct patch clamping of chloroplast envelope
membranes and of isolated swollen thylakoid membranes
(Schonknecht et al., 1988; Pottosin, 1992, 1993; Pottosin and
Schonknecht, 1995, 1996; van den Wijngaard and Vredenberg,
1997; Hinnah and Wagner, 1998; Pottosin et al., 2005) or, in
most cases, by incorporating membrane vesicles or purified
native/recombinant proteins into planar lipid bilayers (see
Table 1 for references). Table 1 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 the 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 conductances 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
chloroplast channel, renders it difficult to hypothesize about the
372
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Ion Channels in Plant Bioenergetic Organelles
proteins responsible for most activities. The question also arises
whether all observed activities listed in Table 1 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 protein
in chloroplasts, direct assessment of its activity is either
missing or is studied only with recombinant proteins in nonnative membranes (see Table 1). 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, such as, e.g., calcium uptake into
chloroplasts (Stael et al., 2012), increase of Mg2+ concentration
in stroma during photosynthesis (Hind et al., 1974; Bulychev
and Vredenberg, 1976), and thylakoid membrane permeability
changes.
Bioinformatics and Proteomics: Toward Molecular
Identification
Besides 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.
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), WoLF PSORT (Horton
et al., 2007), and ChloroP (Emanuelsson et al., 1999). Table 2
reports all authentic (with proven channel function) or putative
ion channels of Arabidopsis (Ward et al., 2009) predicted to be
located in chloroplasts.
Most of these algorithms rely on the analysis of the amino acid
composition of the N-terminal 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 the 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 big-conductance 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 (Elter et al., 2007; Teardo et al., 2011; Robert
et al., 2012; Michaud et al., 2014). We have recently
demonstrated that an 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
Localization
Maximal
conductance
Selectivity/voltage
dependence
Pharmacology
Method of detection
Proposed
function
Reference
Channels in the Envelope Membranes
Cationic channels
OEP23
OE
466 pS in
250 mM KCl
Cationic, maximal open
probability at 0 mV
Reduction of open
probability by 10 mM
spermine
Recombinant Oep23 of
pea in BLM
Solute exchange
(Goetze et al., 2015)
OEP37
OE
500 pS in
250 mM KCl
Cationic, maximal open
probability at 0 mV
Blocked by 2 mM
CuCl2, sensitive to
Tic32 prepeptide
(in mM range)
Recombinant Oep37 of
pea in BLM
Protein import
(Goetze et al., 2006)
TOC75
OE
1.3 nS in
1 M KCl
Cationic, maximal open
probability at 0 mV
Voltage-dependent
block by TrOE33
Recombinant TOC75
in BLM
Protein importa
(Eckart et al., 2002)
VDAC
homolog
OE or contact 520 and
sites between 1016 pS in
OE and IE
100 mM KCl
Cationic, closure
at >±50 mV (520 pS
channel), at >±30 mV
(1016 pS channel)
Blocked by Konig
polyanion
Patch clamp of
chloroplasts from green
algae Nitellopsis obtusa
Metabolite
transport
across OE
(Pottosin, 1992, 1993)
1.14 nS in
250/20 mM
K-glutamate
Closed at voltages
>50 mV and <20 mV;
slightly cationic
None
Proteoliposomes in BLM
General diffusion (Heiber et al., 1995)
pore of OE
(porin-like)
Calcium-permeable IE
channel
35 pS in
100 mM CaCl2
Cationic
Blocked by
100 mM DNQX
Purified inner membrane
vesicles from spinach
chloroplast in BLM
Glutamate
receptor
(Teardo et al., 2010)
Tic110
IE
From 60 to
110 pS in
250 mM KCl
Cationic
Change of selectivity
in presence of
20 mM CaCl2
Recombinant Tic110 from
pea in BLM
Protein import
channela
(Balsera et al., 2009a)
FACC cation
channel
Envelope
membrane
From 40 to
200 pS in
250 mM KCl
Cationic; permeable to K+, Blocked by 100 mM
Na+, Ca2+Mg2+, low open Gd3+, regulated
probability at 0 mV
by pH
Patch clamping of pea
chloroplasts
Regulation
of stroma
alkalinization
(Pottosin et al., 2005)
ATP-sensitive
potassium channel
Envelope
membranes
100 pS in
250/20 mM KCl
Cationic; higher open
probability at negative
voltages
Blocked by 1 mM ATP Proteoliposomes in BLM
(added to cis side),
partially blocked by
200 mM CsCl
Electrical
balance
of H+ flux
across IE
(Heiber et al., 1995)
Anionic channels
Anion channel
OE or contact 160 pS in
sites between 100 mM KCl
OE and IE
Anionic, activation at
negative voltages
None
Patch clamp of chloroplasts
from green algae Nitellopsis
obtusa
VDAC-like channel
IE
Anionic; open at positive
potentials only
3.8 mM succinic
anhydride increased
open probability
BLM using isolated IE
vesicles from spinach
525 pS in
150 mM KCl
(Pottosin, 1992)
Protein import
(Fuks and Homble,
1995)
Table 1. Channel Activities Observed in Chloroplast Membranes.
373
(Continued on next page)
Molecular Plant
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Large-conductance Envelope
cation channel
membranes
(spinach)
Ion Channels in Plant Bioenergetic Organelles
Name
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Selectivity/voltage
dependence
Pharmacology
Method of detection
Proposed
function
Envelope
membrane
(spinach)
60 pS in
100 mM Tris-Cl
Anionic
None
Path clamp of
proteoliposomes
Regulation of
stromal pH
(Heiber et al., 1995)
Envelope
membrane
(spinach)
540 pS in
250/20 mM
K-glutamate
Closed ate voltages
>±50 mV, slightly
selective for glutamate
None
Proteoliposomes in BLM
Involved in
protein import
(Heiber et al., 1995)
Localization
Anion channel
Largeconductance
anion channel
Reference
Molecular Plant
374
Maximal
conductance
Name
Channels in the Thylakoid Membrane
Cationic/unselective
channels
Unselective
large-conductance
channel
Thylakoid
membrane
620 pS in
Unselective
20/100 mM
KCl (occasionally
1 nS)
None
Patch clamp of swollen
thylakoid from pea
chloroplasts
Correspondent
of PTP- role in
protein import
(Hinnah and Wagner,
1998)
Cation channel
Thylakoid
membrane
40 pS in
20/100 mM KCl
(occasionally
90 pS)
Cationic, permeable also
to Mg2+ and Ca2+
None
Patch clamp of swollen
thylakoid from pea
chloroplasts
Counterbalance
of proton entry
into the lumen
(Hinnah and Wagner,
1998)
Cation channel
Thylakoid
membrane
60 pS in
100 mM KCl,
21 pS in
50 mM Ca2+,
24 pS in
50 mM Mg2+
Cationic, permeable to K+, None
Mg2+, and Ca2+; more
active at positive potential
Patch clamp of swollen
thylakoid membrane
from spinach
Counterbalance
of H+ entry into
the lumen
(Pottosin and
Schonknecht, 1996)
Divalent cation
channel
Thylakoid
membrane
20 pS for Ca2+,
35 pS for Mg2+
Cationic, permeable to
divalent cations
None
Thylakoid/liposome
vesicle by patch clamp
Potassium channel
Thylakoid
membrane
From 93 to
122 pS in
1000/300 mM
KCl and 5 mM
MgCl2 (cis side)
Cationic
Inhibited by 50 mM
TEACl
Isolated thylakoid
membrane vesicles
in BLM
Potassium channel
Thylakoid
membrane
100 pS in
100 mM KCl
Cationic
None
Thylakoid/liposome
vesicle by patch clamp
TPK3 potassium
channel
Thylakoid
membrane
35 pS in
250 mM
K-gluconate
Cationic, no voltage
dependence
Activated by 100 mM
calcium and acidic
pH, blocked by
5 mM barium
Recombinant
Arabidopsis
protein in BLM
Thylakoid
membrane
220 pS in
100 mM KCl
Anionic
None
Thylakoid/liposome
vesicle by patch clamp
(Enz et al., 1993)
(Tester and Blatt,
1989)
(Enz et al., 1993)
Counterbalance
of H+ influx into
lumena
(Carraretto et al.,
2013)
Anionic channels
Anion channel
Table 1. Continued
(Enz et al., 1993)
(Continued on next page)
Ion Channels in Plant Bioenergetic Organelles
Counterbalance
of H+ entry into
the lumen
Table 1. Continued
The table shows the main characteristics of the channels recorded in the outer envelope (OE) and inner envelope (IE) membranes as well as of the channels residing in chloroplast membranes, which were
expressed in recombinant form. OEP16 and OEP24 which are specific for the transport of amino acids and 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).
a
Genetic proof has been obtained in favor of the proposed function.
(Pottosin and
Schonknecht, 1995)
Counterbalance
of H+ entry into
the lumen
Patch clamp of swollen
thylakoid membrane
from Nitellopsis obtusa
None
Anionic; bell-shaped
voltage dependence
(max. open
probability at 0 mV)
Thylakoid
membrane
Anion channel
From 100
to 110 pS
in 114 mM Cl
Localization
Name
Maximal
conductance
Selectivity/voltage
dependence
Pharmacology
Method of detection
Proposed
function
Reference
Ion Channels in Plant Bioenergetic Organelles
Molecular Plant
be considered when trying to predict protein targeting to
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;
Rolland et al., 2012; Paila et al., 2015) 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 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 an N-terminal targeting sequence, are used
for channel proteins. 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 COPII-like system (Khan
et al., 2013; Karim and Aronsson, 2014). Interestingly, diacidic
motifs at the C terminus, which is typical of cargo proteins that
are transported by the cytosolic vesicle transport system (Khan
et al., 2013) have been identified in TPK3, a thylakoid-located
member of the two-pore 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 emerging. This process could play an essential
role in protein targeting to the ER, 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 have 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 proteins. However, this latter
method, which can also be exploited for the determination of
the cleavage site or the targeting peptide (Candat et al., 2013),
does not lack possible pitfalls: GFP, which is most often used
for this purpose, is rather large and might affect 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 strong overexpression due to
strong promoters (e.g., CaMV 35S promoter) might eventually
lead to mistargeting. 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 complementary strategies, also including
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
375
Molecular Plant
Ion
channels
Name
Ion Channels in Plant Bioenergetic Organelles
Predicted
consensus
value
Observed localization
Homology to
cyanobacterial
channel
Reference
Cationic channels
AT5G46370
AtTPK2
CP: 6.9
SP: 4.9
Vacuole
slr1257
44 (4e-05),
35% identity
(Voelker et al., 2006)
AT4G18160
AtTPK3 (two-pore
potassium channel)
CP: 5.1
SP: 5.3
Thylakoid
membrane/vacuole
slr1257
48 (3e-06),
30% identity/slr0498
41 (3e-04),
33% identity
(Voelker et al., 2006)
(Carraretto et al., 2013)
AT4G32650
AtKC1
CP: 12.3
M: 10.5
Endoplasmic reticulum,
when heteromerizes with
some shaker channels,
becomes targeted to the
plasma membrane
slr1575
55 (5e-08),
29% identity
(Reintanz et al., 2002;
Duby et al., 2008;
Jeanguenin et al., 2011)
AT5G10490
AtMSL2
mechanosensitive
channel
CP: 12.1
M: 7.6
Chloroplast
glr3870
63 (2e-10),
24% identity
(Haswell and Meyerowitz,
2006)
AT1G58200
AtMSL3
CP: 3.2
M: 6.4
Chloroplast
glr3870
55 (3e-08),
21% identity
(Haswell and Meyerowitz,
2006)
AT5G12080
AtMSL10
CP: 4.8
ER and PM
Glr3870
45 (5e-05)
(Veley et al., 2014)
AT1G05200
AtGLR3.4
CP: 4.4
M: 4.4
SP: 15.4
Chloroplast and plasma
membrane
all2911
64 (9e-11),
23% identity
(Teardo et al., 2011)
(Meyerhoff et al., 2005)
AT4G01010
AtCNGC13
CP: 2.7
M: 7.8
Thylakoid membrane
None
(Yin et al., 2015)
AT3G17690
AtCNGC19
CP: 17.0
M: 7.2
n.d.
None
(Kugler et al., 2009)
AT3G17700
AtCNGC20
CP: 16.6
n.d.
None
(Kugler et al., 2009)
AT2G43950
OEP37 homolog
CP: 18.8
Chloroplast outer
membrane
None
(Goetze et al., 2006)
AT5G19620
AtToc75
CP: 10.1
Chloroplast outer
envelope
alr2269
308 (3e-84),
35% identity
(Eckart et al., 2002)
AT1G06950
TIC110
CP: 23.6
Chloroplast inner
envelope membrane
None
(Heins et al., 2002)
Anionic channels
AT4G27970
AtSLAH2 (putative
nitrate anion channel)
CP: 11.4
n.d.
None
(Duby et al., 2008;
Maierhofer et al., 2014)
AT4G10380
AtNIP5.1 putative boric
acid channel
CP: 6.2
SP: 6.4
Plasma membrane
glr0003
138 (2e-33),
39% identity
(Takano et al., 2006)
AT4G35440
AtClC-e
CP: 10.1
M: 13.9
Thylakoid
tll0145
244 (4e-65),
34% identity
(Marmagne et al., 2007)
(Teardo et al., 2005)
AT1G55620
AtClC-f
CP: 0
M: 8.5
Outer envelope membrane
tll0145
239 (1e-63),
35% identity
(Teardo et al., 2005)
Table 2. Ion Channels from Arabidopsis with Predicted and/or Proven Localization to 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 description of the channel or to its localization are indicated. For homology to cyanobacterial channels, blast values and percentages of identities
of amino acid sequences between the bacterial and Arabidopsis proteins are shown. n.d., not determined.
376
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Ion Channels in Plant Bioenergetic Organelles
biochemical studies. Among the different biochemical approaches, proteomic studies of purified organellar membranes
(Peltier et al., 2004; Ferro et al., 2010; Salvi et al., 2011; Simm
et al., 2013) and suborganelle fractions (Tomizioli et al., 2014;
Yin et 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 a 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 transporters of chloroplasts and mitochondria have
been identified by MS (for reviews, see e.g., Millar et al., 2005;
Finazzi et al., 2015). 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 new ion channels and ion/
metabolite transporters (e.g., Baughman, 2011; De Stefani
et al., 2011; Herzig et al., 2012). Similarly, these approaches are
expected to lead to the identification of further 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 also be useful 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 organelletargeted 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 (Martin et al., 1998; Keeling,
2013). After this organism was engulfed by the host cells, a
large set of genes essential to plastid function have been
transferred from the ancestral plastid genome to the nucleus
(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 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/ionconducting machineries share common structure and function
(Bolter et al., 1998; Balsera et al., 2009b; Day et al., 2014). As
to ion channels, many of the channel-forming proteins predicted
to be located to chloroplast show homology with one or more
cyanobacterial protein with presumed or proven ion channel
function (see Table 3 and e.g., Hamamoto and Uozumi, 2014).
For example, the glutamate receptor GLUR3.4 of Arabidopsis
displays considerable amino acid sequence similarity to GluR0
Molecular Plant
of the completely sequenced model organism Synechocystis
(Chiu et al., 2002; Teardo et al., 2011). 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 ‘‘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. The Synechocystis thylakoid potassium channel SynK
(Zanetti et al., 2010) and the thylakoid two-pore potassium channel AtTPK3 (Zanetti et al., 2010; Carraretto et al., 2013) do not
seem to share a common origin (Pfeil et al., 2014), while they
conserve 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; 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 ClC-e/-f and GLR3.4 share the same function in the
two types 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 amino acid 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 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, 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., 2012) and inner envelope vesicles purified from spinach
chloroplasts containing members of family 3 of AtGLRs
exhibited a calcium-permeable activity that 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.
Unfortunately, even though an excellent and near-complete list
of putative potassium channels of different families has been
published already 10 years ago in a comprehensive review about prokaryotic potassium channels (Kuo et al., 2005), including
those of cyanobacteria, only very few studies have 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-membranespanning 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
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
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Accession number
Ion Channels in Plant Bioenergetic Organelles
Molecular function (hypothesized or proven)
References
slr0498
SynK, thylakoid-located potassium channel
(Zanetti et al., 2010; Checchetto et al., 2012)
sll0993
KchX, potassium channel
(Berry et al., 2003; Matsuda and Uozumi, 2006;
Checchetto et al., 2013a)
Potassium channels
slr5078
Kirbac6.1, similar to potassium channel protein
(Paynter et al., 2010)
sll0536
Probable potassium channel protein
(Berry et al., 2003)
Slr1270
Potassium channel
(Agarwal et al., 2014)
slr1257
Glutamate receptor (Glur0), potassium channel gated
by glutamate
(Chen et al., 1999)
slr1575
Probable potassium efflux system mechanosensitive
ion channel
Chloride channels
sll0855
Putative voltage-gated chloride channel
(Jayaram et al., 2011)
sll1864
Probable chloride channel protein
(Hihara et al., 2003)
slr0617
Unknown protein putative Cl channel
(Sato et al., 2007)
slr2057
apqZ, water channel protein
(Akai et al., 2011; Azad et al., 2011;
Hagemann, 2011)
ssl2749
Hypothetical protein with similarity to water channel NIP4; 2
Water channels
Mechanosensitive channels
slr0875
MscL, large-conductance mechanosensitive channel
(Nanatani et al., 2013; Nazarenko et al., 2003)
(Kwon et al., 2010)
slr0639
Mechanosensitive ion channel homolog
sll0590
Unknown protein putative mechanosensitive ion channel
sll1040
Unknown protein putative mechanosensitive ion channel
sll0985
Mechanosensitive ion channel MscS
slr0109
Unknown protein putative mechanosensitive ion channel MscS
slr0510
Hypothetical protein, putative mechanosensitive ion channel
(Malcolm et al., 2012)
Table 3. 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.
light responses (Checchetto et al., 2012) (see more details
below). Checchetto et al. (2013a) have also characterized
SynCaK, a calcium-dependent potassium channel the lack of
which leads to increased resistance to zinc, present in the environment as a pollutant. An additional potassium channel, KirBac
6.1 (slr5078), was found to be important for Synechocystis growth
(Paynter et al., 2010). The protein encoded by slr1270, which
resides 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
ligand-gated cation channel from Gloeobacter violaceus (GLIC)
was biophysically characterized, revealing a single channel
conductance of solely 8 pS (Bocquet et al., 2009; Velisetty and
Chakrapani, 2012).
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
378
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
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 stretch activation of the plasma
membrane due to the decomposition of photosynthetic
materials produced in day time (Nanatani et al., 2013, 2015). A
member of the cyclic nucleotide-gated (CNG) channels (part of
the mechanosensitive channels of the small conductance
(MscS) superfamily but universally considered as a nonmechanosensitive 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 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 aquaporin homologs are localized in organelles (for
a recent review, see Beebo et al., 2013); the tobacco aquaporin,
Ion Channels in Plant Bioenergetic Organelles
Molecular Plant
Emerging Roles of Ion Channels in Chloroplasts and
Cyanobacteria: Regulation of Photosynthesis
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 m2 s1.
(A) Arabidopsis plants (WT and plants silenced for AtTPK3) grown at the
indicated light intensities. Note the 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 transmission electron microscopy analysis (arrows). For
further details see the text and Carraretto et al. (2013).
NtAQP1 mediates CO2 transport in the plasma membrane and in
the inner chloroplast membrane (Uehlein et 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 glucose in the medium, indicating that
AqpZ is needed for photomixotrophic growth (Ozaki et al., 2007;
Akai et al., 2011). The expression of aqpZ was regulated by an
intrinsic biological clock, and the peak of the expression
corresponded to the early subjected night. This might be
coordinated with the circadian rhythm of the daily glucose
metabolism supplied by carbon fixation.
Thus, homologs of at least some of the above channels seem to
be functionally present in chloroplasts (see Table 2) 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 (Mayer et al., 2001; Bocquet et al., 2009; Hilf and
Dutzler, 2009; Jayaram et al., 2011).
In summary, a 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 chloroplastlocalized channels, final genetic proof demonstrating their function cannot be obtained, even though important cellular and
physiological processes seems to be regulated by these channels. Here, we focus on three aspects in which chloroplast channels are emerging as important players.
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 (DpH) and an electric
field (DJ). DpH 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 cyt b6f, as
well as for NADP reduction in the stroma, makes the compartment basic in the light. DJ 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. DJ is initially
a localized dipole field (positive on the luminal and negative
on the stromal), but it is rapidly delocalized (time of
delocalization ca. 0.1 ms (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
relaxation of the pmf. This mostly stems from H+ translocation
by the ATP synthase-ATPase CF0-Fi complex (Joliot and
Delosme, 1974). This enzyme translocates H+ and thus
modifies DJ and DpH at the same time. CF0-Fi activity can
modulate the size but cannot change the relative composition
of the pmf. Therefore, the 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 DpH in the photosynthetic organelle and
by a DJ in the respiratory organelle (see, e.g., review by
Bernardi, 1999; Finazzi et al., 2015). Indeed, ion channels could
modify the DJ/DpH 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 (Schonknecht et al., 1988; Tester
and Blatt, 1989). 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
(Armbruster et al., 2014; Kunz et al., 2014) families modulate
photosynthesis. In particular, TPK3 from Arabidopsis thaliana
(Carraretto et al., 2013) and its cyanobacterial counterpart
(SynK; Checchetto et al., 2012), mediate K+ fluxes in the light,
thereby controlling DJ 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 mmol photons m2 s1 showed no
difference with respect to WT plants, but when the light
intensity was increased to 90 mmol photons m2 s1, they
exhibited a decreased rosette size and an increased anthocyan
content (Figure 1A). This phenotype reflects the observed
higher DJ and lower DpH due to the lack of counterbalancing
flux of positively charged potassium ions. This altered pmf
partitioning results in reduced CO2 assimilation, reduced
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
379
Molecular Plant
growth, and in a deficient nonphotochemical dissipation (NPQ)
of excess absorbed light, since NPQ onset is controlled by DpH
(see below). Conversely, K+/H+ exchangers belonging to the
KEA family balance the DpH and DJ 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 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 photosynthetic electron flux (Marmagne
et al., 2007).
What are the consequences of the ion regulation of the pmf on
photosynthesis? While ATP synthesis can be equally fueled by
DpH and DJ, 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
control. In the light, changes in the DJ/DpH ratio (even under
constant pmf conditions) could therefore trigger changes in the
rate of electron flow, without any modification in the rate of the
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 DJ
and lower DpH) using time-resolved redox spectroscopy of the
b6f complex (Checchetto et al., 2012, 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 stroma.
Therefore these pigments are no longer sensitive to changes
in DJ. 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 acclimation
capacity in plants via an impaired NPQ response (Carraretto
et al., 2013; Armbruster et al., 2014). The NPQ acronym (nonphotochemical 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 only in plants and algae that this process is
directly controlled by the pmf via DpH-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 specific to
photosynthetic eukaryotes, likely triggered an altered thylakoid
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Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Ion Channels in Plant Bioenergetic Organelles
membrane structure, which was observed in A. thaliana TPK3
mutants (see Figure 1B) (but not in Synechocystis lines lacking
SynK), possibly reflecting the larger complexity of lightharvesting regulation in eukaryotic photosynthesis. Cation
concentration might also affect thylakoid structure by
counteracting negative charges as consequence of (reversible)
phosphorylation, which leads to repulsion and destabilization of
the thylakoid structure (Fristedt et al., 2009). In addition, during
illumination, due to 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 inner envelope (Heiber et al., 1995).
Overall, these new results have opened a novel, exciting
field of investigation: the molecular regulation of photosynthesis
by ion homeostasis. Several tasks remain to be achieved to
complete these studies. 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 oxygenevolving complex (Kessler, 1955; Boussac et al., 1989).
Calcium also plays a crucial role in the regulation of Calvincycle enzymes and recent data highlight its role in the control of
cyclic electron flow as well (Terashima et al., 2012; Hochmal
et al., 2015). Thus, research should focus first on 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 chloroplasts
should be addressed to complement their biochemical and
electrophysiology studies in vitro. Finally, in vivo studies are
needed to establish the link between the 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
intermittent light stress).
Emerging Roles of Ion Channels in Chloroplasts:
Calcium Signaling
Although Ca2+ is a second messenger involved in physiological
and stress responses of plants (e.g., Bose et al., 2011; Choi
et al., 2014; Stephan and Schroeder, 2014; Michal Johnson
et al., 2014), little attention has been paid to the potential roles,
except for the regulation of photosynthesis, of chloroplasts in
cellular Ca2+ signaling of plant cells (Stael et al., 2012; Nomura
and Shiina, 2014). 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 thought. In chloroplasts, the impact of
Ion Channels in Plant Bioenergetic Organelles
impaired organellar Ca2+ handling on plant physiology has been
suggested by CAS and PPF1 (Wang et al., 2003; Nomura et al.,
2008; 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 that chloroplasts
have a significant buffering capacity for Ca2+ ions, resulting in
extremely low (150 nM) free [Ca2+] (Aldakkak et al., 2010) under
resting conditions. 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; Dodd et al.,
2010). Interestingly, the bacterial pathogen-associated molecular
pattern 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 voltagedependent Ca2+ uptake activity in the inner envelope membrane
of pea (Pisum sativum) chloroplasts (Pottosin et al., 2005) (see
Table 1). 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-ATPase like
protein (ACA1) (Huang et al., 1993), as well as HMA1 P-type
ATPase, shown be located in the envelope membrane (Ferro
et al., 2010). The nature of the 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 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 chloroplasts (Stael et al., 2012).
However, the localization, channel activity, and the permeability
for calcium of members of the AtMCU family have not been
described up to now (except for one member, found in
mitochondria by the proteomic approach; Wagner et al., 2015).
At present, it is difficult to understand whether the fastactivating cation channel (FACC), recorded directly in native
pea chloroplast envelope membrane (Pottosin et al., 2005),
might correspond to one of these entities. Since the described
channel has a large conductance (from 40 to 200 pS in 250 mM
KCl), it is unlikely to arise from ATPase 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. Whether
Molecular Plant
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 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., 2012), 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 10 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 light-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 affect 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 (Mei et al.,
2009; Cho et al., 2012). 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 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 revisit
the channel-forming ability of PPF1 in a heterologous system
other than mammalian cells and/or in an in vitro transcription/
translation system. This latter tool has the advantage of avoiding
contamination by membrane proteins and avoiding the 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 characteristics similar to
those found for the same proteins expressed in, for example,
Escherichia coli or Pichia pastoris (Deniaud et al., 2010; De
Stefani et al., 2011; Braun et al., 2014).
In summary, to provide further insight into the physiological relevance of organellar Ca2+ signaling, 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 signaling in organelles
together with newly generated detection systems for measuring
chloroplast (and possibly sub-chloroplast) Ca2+ concentrations
in intact plants should provide fruitful grounds for further discoveries. In this respect, the construction of chloroplast-targeted
genetically encoded aequorin-based calcium probes represents
a milestone (Mehlmer et al., 2012).
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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 LopezJuez, 2013). In this latter system, beside directly affecting
mitochondrial metabolism, defects in the fission-fusion machinery have been linked to severe pathologies (Kasahara and
Scorrano, 2014). Plastid division consists 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 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, MSL-2 and
MSL3, not only function to relieve plastid osmotic stress
(Wilson et al., 2014) but also co-localize with the plastid division
protein AtMinE and, as a result, plants lacking both channels
have fewer and larger chloroplasts than the WT. 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 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 the
envelope membranes might be an issue to be resolved in order
to achieve seal formation.
MITOCHONDRIAL ION CHANNELS
Electrophysiology and Molecular Players
Mitochondria are crucial bioenergetic organelles for providing
ATP and the metabolites required to sustain cell growth. These
organelles also have a specialized metabolic role in plants, 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 with the animal field (Szabo and
Zoratti, 2014) and with respect to chloroplast channels, except
for the mitochondrial voltage-dependent anion channel (VDAC)
(for review, see e.g., Homble et al., 2012; Takahashi and
Tateda, 2013) and the channels formed by components of
the protein import machinery (Carrie et al., 2010; Schleiff
and Becker, 2011). Indeed, only very few electrophysiological
studies exist on channels other than VDAC, using mainly
the lipid bilayer system employing membrane vesicles.
Table 4 summarizes the channel activities observed in different
species. These studies employ species other than Arabidopsis
(Koszela-Piotrowska et al., 2009; Jarmuszkiewicz et al., 2010;
Matkovic et al., 2011), even though protocols yielding highly
382
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Ion Channels in Plant Bioenergetic Organelles
purified mitochondria from this model plant exist (e.g., Millar
et al., 2005; Sweetlove et al., 2007; Lee et al., 2013; Konig
et al., 2014). For example, in potato inner membrane vesicles,
activities displaying different conductance and kinetic behavior
have been recorded. These currents were ascribed to an ATPregulated potassium channel (mitoKATP channel), to a largeconductance Ca2+-insensitive iberiotoxin-sensitive potassium
channel, to a (DIDS) (4,40 -diisothiocyanostilbene-2,20 -disulfonic
acid)-sensitive chloride channel (Matkovic et al., 2011), and to
a large-conductance, 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 established that ion channels
in the mitochondrial inner membrane with a DJ of
approximately 180 mV (negative inside) can drastically
influence mitochondrial bioenergetic efficiency by altering DJ
and the production of reactive oxygen species, and as a
consequence, cell function (Bernardi, 1999; Szewczyk et al.,
2006, 2009; Szabo and Zoratti, 2014). 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 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 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 mitochondria are far too small for patch clamping (Szabo
et al., unpublished results), 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 5 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
hypotheses have been put forward on what proteins mediate
the activities observed either by electrophysiology or by
classical bioenergetics. For example, the CLCNt chloride
channel of tobacco cells has been 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
indications, the authors proposed that an AtKC1-like channel
might be involved (Koszela-Piotrowska et al., 2009). AtKC1 is
Localization
Conductance
VDAC homolog
OM
3, 7, or 4, 0 nS for two
isoforms, respectively
VDAC3
OM
MitoCl channel
Selectivity/voltage
dependence
Proposed
function/identity
Pharmacology
Method of detection
Reference
Large conductance state:
slightly anionic
Low-conductance state:
cationic
Bell-shaped voltage
dependence with
maximal open probability
at 0 mV
None
Planar lipid bilayer
(BLM) with VDAC
proteins purified from
bean mitochondria
Metabolite transport
across OE
(Abrecht et al., 2000)
500 pS in 300 mM KCl
Cationic and anionic
open states; tendency to
close at positive voltages,
open up to ±60 mV
None
AtVDAC3 from
Arabidopsis expressed
in a cell-free system or
E. coli studied in BLM
Metabolite transport
across the OE
(Berrier et al., 2015)
IM
117 pS in 50/450 mM KCl
Slightly anionic; no activity
at negative voltage
Inhibited by
200 mM DIDS
BLM with isolated
IM vesicle from potato
Correspond to
PIMAC?
(Matkovic et al.,
2011)
Large-conductance
BK channel
IM
From 500 to 615 pS in
50/450 mM KCl
Cationic
Activated by
300 mM calcium,
inhibited by
iberiotoxin
(IC50 170 nM)
BLM with isolated
IM vesicle from potato
Regulation of
respiratory rate and
membrane potential
in mitochondria
(Koszela-Piotrowska
et al., 2009)
K(ATP)
IM
150 pS in 150 mM
K-gluconate
Cationic; activated by
negative voltage
(in pipette)
Inhibited by
1 mM ATP
Patch clamp on
mitoplasts from
wheat germ
Regulation of
respiratory rate
and membrane
potential in
mitochondria
(De Marchi et al.,
2010)
K(ATP)
IM
164 pS in 50/450 mM KCl
Cationic
Inhibited by
1 mM ATP
BLM with isolated
IM vesicle from potato
Regulation of
respiratory rate
and membrane
potential in
mitochondria
(Matkovic et al.,
2011)
Large-conductance
calcium-insensitive
potassium channel
IM
312 pS in 50/450 mM KCl
Cationic, more active at
positive voltage
Insensitive to
300 mM calcium,
inhibited by
iberiotoxin
(IC50 150 nM)
BLM with isolated
IM vesicle from potato
Regulation of
respiratory rate
and membrane
potential in
mitochondria
(Matkovic et al.,
2011)
Anionic channels
Ion Channels in Plant Bioenergetic Organelles
Name
Cationic channels
383
Molecular Plant
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Table 4. Ion Channel Activities Recorded in Plant Mitochondrial Membranes.
Molecular Plant
Access
number
Name
Ion Channels in Plant Bioenergetic Organelles
Predicted
consensus
value
Observed localization
References
Cationic channels
AT4G18290
AtKAT2
Potassium channel
M: 8.0
SP: 6.7
Plasma membrane
(Nieves-Cordones et al., 2014)
AT4G22200
AtAKT2/AKT3
Potassium channel
M: 1.3
SP: 3.9
Plasma membrane
(Very and Sentenac, 2002)
(Held et al., 2011)
AT5G46360
AtKCO3
Potassium channel
M: 5.4
CP: 2.7
SP: 5.5
Vacuole
(Voelker et al., 2006;
Rocchetti et al., 2012)
AT2G26650
AtAKT1
Potassium channel
M: 5.5
CP: 4.0
SP: 4.3
n.d. in Arabidopsis;
endomembrane localization
in Lilium longiflorum
(Safiarian et al., 2015)
AT1G15990
AtCNGC7
Cyclic nucleotide-gated channel
M: 5.2
CP: 0.6
SP: 1.8
Plasma membrane
(Tunc-Ozdemir et al., 2013a)
AT4G30360
AtCNGC17
Cyclic nucleotide-gated channel
M: 1.8
SP: 1.0
Plasma membrane
(Ladwig et al., 2015)
AT4G01010
AtCNGC13
Cyclic nucleotide-gated channel
M: 7.8
CP: 2.7
Thylakoid membrane
(Yin et al., 2015)
AT3G48010
AtCNGC16
Cyclic nucleotide-gated channel
M: 15.1
CP: 2.4
Plasma membrane
(Tunc-Ozdemir et al., 2013b)
AT2G46440
AtCNGC11
Cyclic nucleotide-gated channel
M: 7.5
SP: 7.0
n.d.
AT2G46450
AtCNGC12
Cyclic nucleotide-gated channel
M: 5.1
SP: 7.0
n.d.
AT5G14870
AtCNGC18
Cyclic nucleotide-gated channel
M: 12.7
CP: 0.6
SP: 3.4
Plasma membrane
AT5G11210
AtGLR2.5
Glutamate receptor
M: 6.2
CP: 4.5
SP: 1.6
n.d.
AT1G42540
AtGLR3.3
Glutamate receptor
M: 7.8
SP: 21.3
Plasma membrane
(Li et al., 2013)
AT2G32390
AtGLR3.5
Glutamate receptor
M:13.4
SP: 2.4
Mitochondria
(Teardo et al., 2015)
AT4G00290
AtMSL1
Mechanosensitive channel
M: 17.5
CP: 6.9
n.d.
AT1G57610
Putative mitochondrial calcium
uniporter isoform 1
M: 17.6
CP: 1.6
n.d.
AT1G09575
Putative mitochondrial calcium
uniporter isoform 2
M:20.3
CP:1.2
n.d.
AT2G23790
Putative mitochondrial calcium
uniporter isoform 3
M: 17.1
CP: 4.8
n.d.
AT4G36820
Putative mitochondrial calcium
uniporter isoform 4
M: 19.2
CP: 7.5
n.d.
AT5G42610
Putative mitochondrial calcium
uniporter isoform 5
M: 14.5
n.d.
M: 4.5
SP: 4.2
Mitochondrial outer
membrane
(Frietsch et al., 2007)
Anionic channels
AT3G49920
AtVDAC5
Voltage-dependent anion channel
(Lee et al., 2009)
Table 5. Ion Channels from Arabidopsis with Predicted and/or Proven Localization to Mitochondrial Membranes.
(Continued on next page)
384
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Molecular Plant
Ion Channels in Plant Bioenergetic Organelles
Access
number
Name
Predicted
consensus
value
Observed localization
References
AT5G57490
AtVDAC4
Voltage-dependent anion channel
M: 4.8
CP: 1.4
SP: 3.0
Mitochondrial outer
membrane
(Robert et al., 2012)
AT5G37610
AtVDAC6
Putative voltage-dependent anion channel
M: 6.6
SP: 13.1
n.d.
AT5G15090
AtVDAC3
Voltage-dependent anion channel
M: 7.3
Most abundant in plasma
membrane
(Robert et al., 2012)
AT5G67500
AtVDAC2
Voltage-dependent anion channel
M: 5.8
SP: 1.6
Mitochondrial outer
membrane, plasma
membrane
(Robert et al., 2012)
AT3G01280
AtVDAC1
Voltage-dependent anion channel
M: 5.7
SP: 0.6
Mitochondrial outer
membrane
(Pan et al., 2014)
AT1G55620
AtClC-f
M: 8.5
CP: 0
Chloroplast outer
envelope membrane
(Teardo et al., 2005)
Table 5. Continued
Consensus values are those reported at the Aramemnon site (CP, chloroplast; M, mitochondria; SP, secretory pathway). All proteins harboring
mitochondrial targeting sequences are listed. References given for localization concern only studies in Arabidopsis unless otherwise specified. n.d.,
not determined.
however a silent shaker-type subunit (Reintanz et al., 2002),
therefore another so 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) (Petrussa et al., 2008; Pastore et al., 2013)?
This is an important question that awaits answer, even in the
mammalian system for which several different possibilities have
been proposed (for recent review, 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
AtATM1-3 of mitochondrion, which show a high degree of
sequence homology with the sulfonylurea receptors SUR1 and
SUR2A (De 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
an increase in permeability 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 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 (Arpagaus et al., 2002; Petrussa et al., 2004; Vianello
et al., 2012) 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).
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 such as 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 chloroplasts? And if yes, what would be the
consequence for the thylakoid ultrastructure? Can we exploit
knockout 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 nuclearencoded mitochondrial metalloprotease (m-AAA) that interacts
with cyclophilin D, VDAC1, and with a paraplegin-like protein
and has homologs in plants, is essential for PTP complex formation ((Shanmughapriya et al., 2015; but see Bernardi and Forte,
2015), rather than the ATP synthase.
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.
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
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
385
Molecular Plant
Ion Channels in Plant Bioenergetic Organelles
Figure 2. An Arabidopsis Isoform of the
Mammalian Mitochondrial Calcium Uniporter Is Efficiently Targeted to Mitochondria in Arabidopsis Plants.
(A) At2g23790 full locus was expressed in fusion
with EGFP in Arabidopsis leaves. An Ar laser
(488 nm) was used to excite GFP; the fluorescence
signal was recovered from 505 to 525 nm for GFP
and from 680 to 720 nm for chlorophyll autofluorescence. Images were collected using a
Leica TCS SP5 II confocal system 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 (see Supplemental Movies 1 and 2). 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 b subunit
of the F1-ATP synthase (see Duby et al.,
2001; Zottini et al., 2008) identify mitochondria.
Comparison suggests that the organelles
identified in the images in (A) are mitochondria.
obtained in vivo using calcium sensors such as aequorin probe
and probes of the Cameleon family revealed differential Ca2+
dynamics between the cytosol and mitochondria (Sai and
Johnson, 2002; Logan and Knight, 2003; Mehlmer et al., 2012;
Loro et al., 2012, 2013). In the mammalian system, following the
discovery and initial characterization of MCU (Baughman, 2011;
De Stefani et al., 2011) and the 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 MICU3 were reported to be essential
components and/or regulators of the mammalian MCU
complex (MCUC; for recent reviews, see e.g., De Stefani et al.,
2015; Foskett and Philipson, 2015).
In Arabidopsis thaliana, six proteins that can be identified as
putative MCU channel proteins are present, since they
are homologs 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 of the six Arabidopsis MCU isoforms are predicted
to localize to the mitochondria (Schwacke et al., 2003), but
so far our recent study has identified 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 (E.T., L.C., et al., unpublished results).
Thus, these results suggest that bioinformatic prediction
386
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
regarding localization of the MCU
homologs are also correct also for the
other isoforms, even though experimental
proof is still lacking. Besides the MCU
isoforms, other homologs of mammalian
MCUC components in plants include two
homologs 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 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 homologs of
MCU and MICU1 represent good candidates for 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. As to the role of
AtMICU1, recent data prove 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 the
steady-state calcium level and stimulus-induced calcium uptake
into mitochondria (Wagner et al., 2015), suggesting that AtMICU1
has a function similar to that observed for MICU2 in the
mammalian system (Patron et al., 2014).
In addition to members of the putative AtMCU family, the
mitochondria-targeted splicing isoform of AtGLR3.5 might
contribute to maintenance of mitochondrial calcium homeostasis. Because the previously studied members of the plant
iGLRs of family 3 are permeable to Ca2+, Teardo et al. (2015)
measured calcium dynamics with the genetically encoded
calcium probe Cameleon targeted to mitochondria in WT
and knockout plants and found a significant difference in
Molecular Plant
Ion Channels in Plant Bioenergetic Organelles
Figure 3. Lack of the Mitochondria-Located Isoform of
Glutamate Receptor AtGLR3.5 Affects Mitochondrial
Morphology.
Transmission electron microscopy images from leaves of WT and
knockout 6-week-old plants showing swollen mitochondria with
decreased electron density and disorganization toward 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 to aging or wounding. For further details, see
text and Teardo et al. (2015).
4-week-old plants challenged with wounding. In accordance
with the mitochondrial localization of one isoform of this putative
glutamate receptor, plants lacking AtGLR3.5 displayed an
altered mitochondrial ultrastructure (Figure 3), in particular a
part of the mitochondria displayed enlargement and partial
loss of cristae. Knockout plants underwent anticipated
senescence. The exact link between reduced mitochondrial
calcium uptake, 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
Ion channels greatly contribute to maintain the daily active organelles, chloroplasts and mitochondria. The function of the ion
channels must play a central role in the energy conversion and
electrochemical potential preservation through light perception
and carbon metabolism across the organelle membranes. Since
the endosymbiotic event, the mechanisms regulating organelle
ion channels have likely been developed to coordinate biological
events of the whole cell. Information gained in cyanobacteria
about the structure/function of their easily accessible PMlocated ion channels will likely be useful for discovering the function of chloroplast channels.
Technical improvements in proteomic techniques (increased
sensitivity, preparation of highly purified membranes) may help
us to identify more and more organelle-located channel proteins.
Once the localization of a given protein is validated in intact
cells (e.g., using tagged proteins), its channel function can be
proved either by exploiting the innovative in vitro 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 achieve. 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 mutant organisms lacking a given protein that likely
forms the channel. In this respect, design of ion-specific probes
that can be targeted to organelles are really helpful (see e.g.,
Mehlmer et al., 2012; Loro et al., 2012; Hong-Hermesdorf et al.,
2015). However, the above approach might lead to misleading
results, since the function of a missing channel can be
overtaken by, for example, another member of 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 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 such as
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 also be an interesting topic for future
studies. A combined diversified approach will certainly lead to a
considerable increase in our knowledge of the mechanisms of
organelle function mediated by ion channels.
SUPPLEMENTAL INFORMATION
Supplemental Information is available at Molecular Plant Online.
FUNDING
The authors are grateful 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 15H02226, 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-8NT09567009, Phytadapt, the Marie Curie Initial
Training Network Accliphot (FP7-PEPOPLE-2012-ITN; 316427 to G.F.)
ACKNOWLEDGMENTS
The authors are grateful to all colleagues in the respective laboratories for
contributing to this research. No conflict of interest declared.
Received: September 1, 2015
Revised: November 22, 2015
Accepted: December 1, 2015
Published: January 2, 2016
REFERENCES
Abrecht, H., Wattiez, R., Ruysschaert, J.M., and Homble, F. (2000).
Purification and characterization of two voltage-dependent anion
channel isoforms from plant seeds. Plant Physiol. 124:1181–1190.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
387
Molecular Plant
Agarwal, R., Zakharov, S., Hasan, S.S., Ryan, C.M., Whitelegge, J.P.,
and Cramer, W.A. (2014). Structure-function of cyanobacterial
outer-membrane protein, Slr1270: homolog of Escherichia coli drug
export/colicin import protein, TolC. FEBS Lett. 588:3793–3801.
Akai, M., Onai, K., Kusano, M., Sato, M., Redestig, H., Toyooka, K.,
Morishita, M., Miyake, H., Hazama, A., Checchetto, V., et al. (2011).
Plasma membrane aquaporin AqpZ protein is essential for glucose
metabolism during photomixotrophic growth of Synechocystis sp.
PCC 6803. J. Biol. Chem. 286:25224–25235.
Akai, M., Onai, K., Morishita, M., Mino, H., Shijuku, T., Maruyama, H.,
Arai, F., Itoh, S., Hazama, A., Checchetto, V., et al. (2012). Aquaporin
AqpZ is involved in cell volume regulation and sensitivity to osmotic
stress in Synechocystis sp. strain PCC 6803. J. Bacteriol. 194:6828–
6836.
Aldakkak, M., Stowe, D.F., Cheng, Q., Kwok, W.M., and Camara, A.K.
(2010). Mitochondrial matrix K+ flux independent of large-conductance
Ca2+-activated K+ channel opening. Am. J. Physiol. Cell Physiol.
298:C530–C541.
Aldridge, C., Maple, J., and Moller, S.G. (2005). The molecular biology of
plastid division in higher plants. J. Exp. Bot. 56:1061–1077.
Araujo, W.L., Nunes-Nesi, A., and Fernie, A.R. (2014). On the role of
plant mitochondrial metabolism and its impact on photosynthesis in
both optimal and sub-optimal growth conditions. Photosynth. Res.
119:141–156.
Armbruster, U., Carrillo, L.R., Venema, K., Pavlovic, L., Schmidtmann,
E., Kornfeld, A., Jahns, P., Berry, J.A., Kramer, D.M., and Jonikas,
M.C. (2014). Ion antiport accelerates photosynthetic acclimation in
fluctuating light environments. Nat. Commun. 5:5439.
Arpagaus, S., Rawyler, A., and Braendle, R. (2002). Occurrence and
characteristics of the mitochondrial permeability transition in plants.
J. Biol. Chem. 277:1780–1787.
Azad, A.K., Sato, R., Ohtani, K., Sawa, Y., Ishikawa, T., and Shibata, H.
(2011). Functional characterization and hyperosmotic regulation of
aquaporin in Synechocystis sp. PCC 6803. Plant Sci. 180:375–382.
Bachin, D., Nazarenko, L.V., Mironov, K.S., Pisareva, T.,
Allakhverdiev, S.I., and Los, D.A. (2015). Mechanosensitive ion
channel MscL controls ionic fluxes during cold and heat stress in
Synechocystis. FEMS Microbiol. Lett. 362:fnv090.
Bailleul, B., Berne, N., Murik, O., Petroutsos, D., Prihoda, J., Tanaka,
A., Villanova, V., Bligny, R., Flori, S., Falconet, D., et al. (2015).
Energetic coupling between plastids and mitochondria drives CO2
assimilation in diatoms. Nature 524:366–369.
Balderas, E., Zhang, J., Stefani, E., and Toro, L. (2015). Mitochondrial
BKCa channel. Front. Physiol. 6:104.
Balsera, M., Goetze, T.A., Kovacs-Bogdan, E., Schurmann, P.,
Wagner, R., Buchanan, B.B., Soll, J., and Bolter, B. (2009a).
Characterization of Tic110, a channel-forming protein at the inner
envelope membrane of chloroplasts, unveils a response to Ca(2+)
and a stromal regulatory disulfide bridge. J. Biol. Chem. 284:2603–
2616.
Balsera, M., Soll, J., and Bolter, B. (2009b). Protein import machineries in
endosymbiotic organelles. Cell Mol. Life Sci. 66:1903–1923.
Baughman, J.M. (2011). Integrative genomics identifies MCU as an
essential component of the mitochondrial calcium uniporter. Nature
476:341–345.
Ion Channels in Plant Bioenergetic Organelles
functional coupling of the mitochondrial BK channel to the
respiratory chain. PLoS One 8:e68125.
Beebo, A., Mathai, J.C., Schoefs, B., and Spetea, C. (2013).
Assessment of the requirement for aquaporins in the thylakoid
membrane of plant chloroplasts to sustain photosynthetic water
oxidation. FEBS Lett. 587:2083–2089.
Berkelman, T., Garret-Engele, P., and Hoffman, N.E. (1994). The
pacL gene of Synechococcus sp. strain PCC 7942 encodes a
Ca(2+)-transporting ATPase. J. Bacteriol. 176:4430–4436.
Berkowitz, G.A., and Wu, W. (1993). Magnesium, potassium flux and
photosynthesis. Magnes. Res. 6:257–265.
Bernardi, P. (1999). Mitochondrial transport of cations: channels,
exchangers, and permeability transition. Physiol. Rev. 79:1127–1155.
Bernardi, P., and Forte, M. (2015). Commentary: SPG7 is an essential
and conserved component of the mitochondrial permeability
transition pore. Front. Physiol. 6:320.
Bernardi, P., Rasola, A., Forte, M., and Lippe, G. (2015). The
mitochondrial permeability transition pore: channel formation by
F-ATP synthase, integration in signal transduction, and role in
pathophysiology. Physiol. Rev. 95:1111–1155.
Berrier, C., Park, K.H., Abes, S., Bibonne, A., Betton, J.M., and Ghazi,
A. (2004). Cell-free synthesis of a functional ion channel in the absence
of a membrane and in the presence of detergent. Biochemistry
43:12585–12591.
Berrier, C., Peyronnet, R., Betton, J.M., Ephritikhine, G., BarbierBrygoo, H., Frachisse, J.M., and Ghazi, A. (2015). Channel
characteristics of VDAC-3 from Arabidopsis thaliana. Biochem.
Biophys. Res. Commun. 459:24–28.
Berry, S., Esper, B., Karandashova, I., Teuber, M., Elanskaya, I.,
Rogner, M., and Hagemann, M. (2003). Potassium uptake in the
unicellular cyanobacterium Synechocystis sp. strain PCC 6803
mainly depends on a Ktr-like system encoded by slr1509 (ntpJ).
FEBS Lett. 548:53–58.
Blum, T., Briesemeister, S., and Kohlbacher, O. (2009). MultiLoc2:
integrating phylogeny and Gene Ontology terms improves subcellular
protein localization prediction. BMC Bioinformatics 10:274.
Bocquet, N., Nury, H., Baaden, M., Le Poupon, C., Changeux, J.P.,
Delarue, M., and Corringer, P.J. (2009). X-ray structure of
a pentameric ligand-gated ion channel in an apparently open
conformation. Nature 457:111–114.
Bolter, B., and Soll, J. (2001). Ion channels in the outer membranes
of chloroplasts and mitochondria: open doors or regulated gates?
EMBO J. 20:935–940.
Bolter, B., Soll, J., Schulz, A., Hinnah, S., and Wagner, R. (1998). Origin
of a chloroplast protein importer. Proc. Natl. Acad. Sci. USA 95:15831–
15836.
Bose, J., Pottosin, I.I., Shabala, S.S., Palmgren, M.G., and Shabala, S.
(2011). Calcium efflux systems in stress signaling and adaptation in
plants. Front. Plant Sci. 2:85.
Boussac, A., Zimmermann, J.L., and Rutherford, A.W. (1989). EPR
signals from modified charge accumulation states of the oxygen
evolving enzyme in Ca2+-deficient photosystem II. Biochemistry
28:8984–8989.
Bayer, R.G., Köstler, T., Jain, A., Stael, S., Ebersberger, I., and Teige,
M. (2014). Higher plant proteins of cyanobacterial origin: are they or are
they not preferentially targeted to chloroplasts? Mol. Plant 7:1797–
1800.
Braun, C.J., Lachnit, C., Becker, P., Henkes, L.M., Arrigoni, C., Kast,
S.M., Moroni, A., Thiel, G., and Schroeder, I. (2014). Viral
potassium channels as a robust model system for studies of
membrane-protein interaction. Biochim. Biophys. Acta 1838:1096–
1103.
Bednarczyk, P., Wieckowski, M.R., Broszkiewicz, M., Skowronek, K.,
Siemen, D., and Szewczyk, A. (2013). Putative structural and
Bulychev, A.A., and Vredenberg, W.J. (1976). Effect of ionophores
A23187 and nigericin on the light-induced redistribution of Mg2+,
388
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Ion Channels in Plant Bioenergetic Organelles
K+ and H+ across the thylakoid membrane. Biochim. Biophys. Acta
449:48–58.
Candat, A., Poupart, P., Andrieu, J.P., Chevrollier, A., Reynier, P.,
Rogniaux, H., Avelange-Macherel, M.H., and Macherel, D. (2013).
Experimental determination of organelle targeting-peptide cleavage
sites using transient expression of green fluorescent protein
translational fusions. Anal. Biochem. 434:44–51.
Carraretto, L., Formentin, E., Teardo, E., Checchetto, V., Tomizioli, M.,
Morosinotto, T., Giacometti, G.M., Finazzi, G., and Szabo, I. (2013).
A thylakoid-located two-pore K+ channel controls photosynthetic light
utilization in plants. Science 342:114–118.
Carrie, C., Murcha, M.W., and Whelan, J. (2010). An in silico analysis of
the mitochondrial protein import apparatus of plants. BMC Plant Biol.
10:249.
Chang, I.F., Curran, A., Woolsey, R., Quilici, D., Cushman, J.C., Mittler,
R., Harmon, A., and Harper, J.F. (2009). Proteomic profiling of tandem
affinity purified 14-3-3 protein complexes in Arabidopsis thaliana.
Proteomics 9:2967–2985.
Checchetto, V., Segalla, A., Allorent, G., La Rocca, N., Leanza, L.,
Giacometti, G.M., Uozumi, N., Finazzi, G., Bergantino, E., and
Szabo, I. (2012). Thylakoid potassium channel is required for efficient
photosynthesis in cyanobacteria. Proc. Natl. Acad. Sci. USA
109:11043–11048.
Checchetto, V., Formentin, E., Carraretto, L., Segalla, A., Giacometti,
G.M., Szabo, I., and Bergantino, E. (2013a). Functional
characterization and determination of the physiological role of a
calcium-dependent potassium channel from cyanobacteria. Plant
Physiol. 162:953–964.
Checchetto, V., Teardo, E., Carraretto, L., Formentin, E., Bergantino,
E., Giacometti, G.M., and Szabo, I. (2013b). Regulation of
photosynthesis by ion channels in cyanobacteria and higher plants.
Biophys. Chem. 182:51–57.
Chen, G.Q., Cui, C., Mayer, M.L., and Gouaux, E. (1999). Functional
characterization of a potassium-selective prokaryotic glutamate
receptor. Nature 402:817–821.
Chiu, J.C., Brenner, E.D., DeSalle, R., Nitabach, M.N., Holmes, T.C.,
and Coruzzi, G.M. (2002). Phylogenetic and expression analysis of
the glutamate-receptor-like gene family in Arabidopsis thaliana. Mol.
Biol. Evol. 19:1066–1082.
Cho, D., Villiers, F., Kroniewicz, L., Lee, S., Seo, Y.J., Hirschi, K.D.,
Leonhardt, N., and Kwak, J.M. (2012). Vacuolar CAX1 and CAX3
influence auxin transport in guard cells via regulation of apoplastic
pH. Plant Physiol. 160:1293–1302.
Choi, W.G., Toyota, M., Kim, S.H., Hilleary, R., and Gilroy, S. (2014). Salt
stress-induced Ca2+ waves are associated with rapid, long-distance
root-to-shoot signaling in plants. Proc. Natl. Acad. Sci. USA
111:6497–6502.
Crofts, A.R., Meinhardt, S.W., Jones, K.R., and Snozzi, M. (1983). The
role of the quinone pool in the cyclic electron-transfer chain of
Rhodopseudomonas sphaeroides: a modified Q-cycle mechanism.
Biochim. Biophys. Acta 723:202–218.
Day, P.M., Potter, D., and Inoue, K. (2014). Evolution and targeting of
Omp85 homologs in the chloroplast outer envelope membrane.
Front. Plant Sci. 5:535.
De Marchi, U., Checchetto, V., Zanetti, M., Teardo, E., Soccio, M.,
Formentin, E., Giacometti, G.M., Pastore, D., Zoratti, M., and
Szabo, I. (2010). ATP-sensitive cation-channel in wheat (Triticum
durum Desf.): identification and characterization of a plant
mitochondrial channel by patch-clamp. Cell Physiol. Biochem.
26:975–982.
Molecular Plant
De Stefani, D., Raffaello, A., Teardo, E., Szabo, I., and Rizzuto, R.
(2011). A forty-kilodalton protein of the inner membrane is the
mitochondrial calcium uniporter. Nature 476:336–340.
De Stefani, D., Patron, M., and Rizzuto, R. (2015). Structure and function
of the mitochondrial calcium uniporter complex. Biochim. Biophys.
Acta 1853:2006–2011.
Deniaud, A., Liguori, L., Blesneac, I., Lenormand, J.L., and PebayPeyroula, E. (2010). Crystallization of the membrane protein hVDAC1
produced in cell-free system. Biochim. Biophys. Acta 1798:1540–
1546.
Dieter, P., and Marme, D. (1980). Ca2+ transport in mitochondrial and
microsomal fractions from higher plants. Planta 150:1–8.
Dodd, A.N., Kudla, J., and Sanders, D. (2010). The language of calcium
signaling. Annu. Rev. Plant Biol. 61:593–620.
Drissi, R., Dubois, M.L., and Boisvert, F.M. (2013). Proteomics methods
for subcellular proteome analysis. FEBS J. 280:5626–5634.
Duby, G., Oufattole, M., and Boutry, M. (2001). Hydrophobic residues
within the predicted N-terminal amphiphilic alpha-helix of a plant
mitochondrial targeting presequence play a major role in in vivo
import. Plant J. 27:539–549.
Duby, G., Hosy, E., Fizames, C., Alcon, C., Costa, A., Sentenac, H., and
Thibaud, J.B. (2008). AtKC1, a conditionally targeted Shaker-type
subunit, regulates the activity of plant K+ channels. Plant J.
53:115–123.
Duy, D., Soll, J., and Philippar, K. (2007). Solute channels of the outer
membrane: from bacteria to chloroplasts. Biol. Chem. 388:879–889.
Eckart, K., Eichacker, L., Sohrt, K., Schleiff, E., Heins, L., and Soll, J.
(2002). A Toc75-like protein import channel is abundant in
chloroplasts. EMBO Rep. 3:557–562.
Elter, A., Hartel, A., Sieben, C., Hertel, B., Fischer-Schliebs, E., Luttge,
U., Moroni, A., and Thiel, G. (2007). A plant homolog of animal chloride
intracellular channels (CLICs) generates an ion conductance in
heterologous systems. J. Biol. Chem. 282:8786–8792.
Emanuelsson, O., Nielsen, H., and von Heijne, G. (1999). ChloroP,
a neural network-based method for predicting chloroplast transit
peptides and their cleavage sites. Protein Sci. 8:978–984.
Emanuelsson, O., Nielsen, H., Brunak, S., and von Heijne, G. (2000).
Predicting subcellular localization of proteins based on their
N-terminal amino acid sequence. J. Mol. Biol. 300:1005–1016.
Enz, C., Steinkamp, T., and Wagner, R. (1993). Ion channels in the
thylakoid membrane (a patch-clamp study). Biochim. Biophys. Acta
1143:67–76.
Ettinger, W.F., Clear, A.M., Fanning, K.J., and Peck, M.L. (1999).
Identification of a Ca2+/H+ antiport in the plant chloroplast thylakoid
membrane. Plant Physiol. 119:1379–1386.
Ezure, T., Nanatani, K., Sato, Y., Suzuki, S., Aizawa, K., Souma, S., Ito,
M., Hohsaka, T., von Heijine, G., Utsumi, T., et al. (2014). A cell-free
translocation system using extracts of cultured insect cells to yield
functional membrane proteins. PLoS One 9:e112874.
Facchinelli, F., and Weber, A.P. (2011). The metabolite transporters of
the plastid envelope: an update. Front. Plant Sci. 2:50.
Ferro, M., Brugiere, S., Salvi, D., Seigneurin-Berny, D., Court, M.,
Moyet, L., Ramus, C., Miras, S., Mellal, M., Le Gall, S., et al.
(2010). AT_CHLORO, a comprehensive chloroplast proteome
database with subplastidial localization and curated information on
envelope proteins. Mol. Cell. Proteomics 9:1063–1084.
Finazzi, G., Petroutsos, D., Tomizioli, M., Flori, S., Sautron, E.,
Villanova, V., Rolland, N., and Seigneurin-Berny, D. (2015).
Ions channels/transporters and chloroplast regulation. Cell Calcium
58:86–97.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
389
Molecular Plant
Ion Channels in Plant Bioenergetic Organelles
Foskett, J.K., and Philipson, B. (2015). The mitochondrial Ca(2+)
uniporter complex. J. Mol. Cell. Cardiol. 78:3–8.
cyanobacterium Synechocystis sp. strain PCC 6803. J. Bacteriol.
185:1719–1725.
Frietsch, S., Wang, Y.F., Sladek, C., Poulsen, L.R., Romanowsky,
S.M., Schroeder, J.I., and Harper, J.F. (2007). A cyclic nucleotidegated channel is essential for polarized tip growth of pollen. Proc.
Natl. Acad. Sci. USA 104:14531–14536.
Hilf, R.J., and Dutzler, R. (2009). Structure of a potentially open state
of a proton-activated pentameric ligand-gated ion channel. Nature
457:115–118.
Fristedt, R., Willig, A., Granath, P., Crevecoeur, M., Rochaix, J.D., and
Vener, A.V. (2009). Phosphorylation of photosystem II controls
functional macroscopic folding of photosynthetic membranes in
Arabidopsis. Plant Cell 21:3950–3964.
Fujimori, T., Higuchi, M., Sato, H., Aiba, H., Muramatsu, M., Hihara, Y.,
and Sonoike, K. (2005). The mutant of sll1961, which encodes a
putative transcriptional regulator, has a defect in regulation of
photosystem stoichiometry in the cyanobacterium Synechocystis sp.
PCC 6803. Plant Physiol. 139:408–416.
Fuks, B., and Homble, F. (1995). A voltage-dependent porin-like channel
in the inner envelope membrane of plant chloroplasts. J. Biol. Chem.
270:9947–9952.
Giorgio, V., von Stockum, S., Antoniel, M., Fabbro, A., Fogolari, F.,
Forte, M., Glick, G.D., Petronilli, V., Zoratti, M., Szabo, I., et al.
(2013). Dimers of mitochondrial ATP synthase form the permeability
transition pore. Proc. Natl. Acad. Sci. USA 110:5887–5892.
Goetze, T.A., Philippar, K., Ilkavets, I., Soll, J., and Wagner, R. (2006).
OEP37 is a new member of the chloroplast outer membrane ion
channels. J. Biol. Chem. 281:17989–17998.
Goetze, T.A., Patil, M., Jeshen, I., Bolter, B., Grahl, S., and Soll, J.
(2015). Oep23 forms an ion channel in the chloroplast outer
envelope. BMC Plant Biol. 15:47.
Hagemann, M. (2011). Molecular biology of cyanobacterial salt
acclimation. FEMS Microbiol. Rev. 35:87–123.
Hamamoto, S., and Uozumi, N. (2014). Organelle-localized potassium
transport systems in plants. J. Plant Physiol. 171:743–747.
Hamilton, E.S., Schlegel, A.M., and Haswell, E.S. (2015). United in
diversity: mechanosensitive ion channels in plants. Annu. Rev. Plant
Biol. 66:113–137.
Hanson, J.B., Malhotra, S.S., and Stoner, C.D. (1965). Action of calcium
on corn mitochondria. Plant Physiol. 40:1033–1040.
Haswell, E.S., and Meyerowitz, E.M. (2006). MscS-like proteins control
plastid size and shape in Arabidopsis thaliana. Curr. Biol. 16:1–11.
Hedrich, R. (2012). Ion channels in plants. Physiol. Rev. 92:1777–1811.
Heiber, T., Steinkamp, T., Hinnah, S., Schwarz, M., Flugge, U.I.,
Weber, A., and Wagner, R. (1995). Ion channels in the chloroplast
envelope membrane. Biochemistry 34:15906–15917.
Heins, L., Mehrle, A., Hemmler, R., Wagner, R., Kuchler, M., Hormann,
F., Sveshnikov, D., and Soll, J. (2002). The preprotein conducting
channel at the inner envelope membrane of plastids. EMBO J.
21:2616–2625.
Held, K., Pascaud, F., Eckert, C., Gajdanowicz, P., Hashimoto, K.,
Corratge-Faillie, C., Offenborn, J.N., Lacombe, B., Dreyer, I.,
Thibaud, J.B., et al. (2011). Calcium-dependent modulation and
plasma membrane targeting of the AKT2 potassium channel by
the CBL4/CIPK6 calcium sensor/protein kinase complex. Cell Res.
21:1116–1130.
Hind, G., Nakatani, H.Y., and Izawa, S. (1974). Light-dependent
redistribution of ions in suspensions of chloroplast thylakoid
membranes. Proc. Natl. Acad. Sci. USA 71:1484–1488.
Hinnah, S.C., and Wagner, R. (1998). Thylakoid membranes contain a
high-conductance channel. Eur. J. Biochem. 253:606–613.
Hochmal, A.K., Schulze, S., Trompelt, K., and Hippler, M. (2015).
Calcium-dependent regulation of photosynthesis. Biochim. Biophys.
Acta 1847:993–1003.
Homble, F., Krammer, E.M., and Prevost, M. (2012). Plant VDAC: facts
and speculations. Biochim. Biophys. Acta 1818:1486–1501.
Hong-Hermesdorf, A., Miethke, M., Gallaher, S.D., Kropat, J., Dodani,
S.C., Chan, J., Barupala, D., Domaille, D.W., Shirasaki, D.I., Loo,
J.A., et al. (2015). Subcellular metal imaging identifies dynamic sites
of Cu accumulation in Chlamydomonas. Nat. Chem. Biol. 11:235.
Hoogenboom, B.W., Suda, K., Engel, A., and Fotiadis, D. (2007). The
supramolecular assemblies of voltage-dependent anion channels in
the native membrane. J. Mol. Biol. 370:246–255.
Horton, P., Park, K.J., Obayashi, T., Fujita, N., Harada, H., AdamsCollier, C.J., and Nakai, K. (2007). WoLF PSORT: protein
localization predictor. Nucleic Acids Res. 35:W585–W587.
Huang, L., Berkelman, T., Franklin, A.E., and Hoffman, N.E. (1993).
Characterization of a gene encoding a Ca(2+)-ATPase-like protein in
the plastid envelope. Proc. Natl. Acad. Sci. USA 90:10066–10070.
Huang, S.S., Chen, J., Dong, X.J., Patton, J., Pei, Z.M., and Zheng, H.L.
(2012). Calcium and calcium receptor CAS promote Arabidopsis
thaliana de-etiolation. Physiol. Plant. 144:73–82.
Ishikawa, M., Fujiwara, M., Sonoike, K., and Sato, N. (2009).
Orthogenomics of photosynthetic organisms: bioinformatic and
experimental analysis of chloroplast proteins of endosymbiont origin
in Arabidopsis and their counterparts in Synechocystis. Plant Cell
Physiol. 50:773–788.
Jarmuszkiewicz, W., Matkovic, K., and Koszela-Piotrowska, I. (2010).
Potassium channels in the mitochondria of unicellular eukaryotes and
plants. FEBS Lett. 584:2057–2062.
Jarvis, P., and Lopez-Juez, E. (2013). Biogenesis and homeostasis of
chloroplasts and other plastids. Nat. Rev. Mol. Cell Biol. 14:787–802.
Jayaram, H., Robertson, J.L., Wu, F., Williams, C., and Miller, C. (2011).
Structure of a slow CLC Cl(-)/H+ antiporter from a cyanobacterium.
Biochemistry 50:788–794.
Jeanguenin, L., Alcon, C., Duby, G., Boeglin, M., Cherel, I., Gaillard, I.,
Zimmermann, S., Sentenac, H., and Very, A.A. (2011). AtKC1 is a
general modulator of Arabidopsis inward Shaker channel activity.
Plant J. 67:570–582.
Joliot, P., and Delosme, R. (1974). Flash-induced 519 nm absorption
change in green algae. Biochim. Biophys. Acta 357:267–284.
Kaldenhoff, R., Kai, L., and Uehlein, N. (2014). Aquaporins and
membrane diffusion of CO2 in living organisms. Biochim. Biophys.
Acta 1840:1592–1595.
Hennon, S.W., Soman, R., Zhu, L., and Dalbey, R.E. (2015). YidC/Alb3/
Oxa1 family of insertases. J. Biol. Chem. 290:14866–14874.
Karim, S., and Aronsson, H. (2014). The puzzle of chloroplast vesicle
transport - involvement of GTPases. Front. Plant Sci. 5:472.
Herzig, S., Raemy, E., Montessuit, S., Veuthey, J.L., Zamboni, N.,
Westermann, B., Kunji, E.R., and Martinou, J.C. (2012).
Identification and functional expression of the mitochondrial pyruvate
carrier. Science 337:93–96.
Karniely, S., and Pines, O. (2005). Single translation–dual destination:
mechanisms of dual protein targeting in eukaryotes. EMBO Rep.
6:420–425.
Hihara, Y., Sonoike, K., Kanehisa, M., and Ikeuchi, M. (2003). DNA
microarray analysis of redox-responsive genes in the genome of the
390
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Kasahara, A., and Scorrano, L. (2014). Mitochondria: from cell death
executioners to regulators of cell differentiation. Trends Cell Biol.
24:761–770.
Ion Channels in Plant Bioenergetic Organelles
Keeling, P.J. (2013). The number, speed, and impact of plastid
endosymbioses in eukaryotic evolution. Annu. Rev. Plant Biol.
64:583–607.
Kessler, E. (1955). On the role of manganese in the oxygen-evolving
system of photosynthesis. Arch. Biochem. Biophys. 59:527–529.
Khan, N.Z., Lindquist, E., and Aronsson, H. (2013). New putative
chloroplast vesicle transport components and cargo proteins
revealed using a bioinformatics approach: an Arabidopsis model.
PLoS One 8:e59898.
Kirichok, Y., Krapivinsky, G., and Clapham, D.E. (2004). The
mitochondrial calcium uniporter is a highly selective ion channel.
Nature 427:360–364.
Konig, A.C., Hartl, M., Boersema, P.J., Mann, M., and Finkemeier, I.
(2014). The mitochondrial lysine acetylome of Arabidopsis.
Mitochondrion 19:252–260.
Koszela-Piotrowska, I., Matkovic, K., Szewczyk, A., and
Jarmuszkiewicz, W. (2009). A large-conductance calciumactivated potassium channel in potato (Solanum tuberosum) tuber
mitochondria. Biochem. J. 424:307–316.
Molecular Plant
Leister, D. (2003). Chloroplast research in the genomic age. Trends
Genet. 19:47–56.
Li, F., Wang, J., Ma, C., Zhao, Y., Wang, Y., Hasi, A., and Qi, Z. (2013).
Glutamate receptor-like channel3.3 is involved in mediating
glutathione-triggered cytosolic calcium transients, transcriptional
changes, and innate immunity responses in Arabidopsis. Plant
Physiol. 162:1497–1509.
Linka, M., and Weber, A.P. (2005). Shuffling ammonia between
mitochondria and plastids during photorespiration. Trends Plant Sci.
10:461–465.
Linka, N., and Weber, A.P. (2010). Intracellular metabolite transporters in
plants. Mol. Plant 3:21–53.
Logan, D.C., and Knight, M.R. (2003). Mitochondrial and cytosolic
calcium dynamics are differentially regulated in plants. Plant Physiol.
133:21–24.
Loro, G., Drago, I., Pozzan, T., Schiavo, F.L., Zottini, M., and Costa, A.
(2012). Targeting of Cameleons to various subcellular compartments
reveals a strict cytoplasmic/mitochondrial Ca2+ handling relationship
in plant cells. Plant J. 71:1–13.
Koulintchenko, M., Konstantinov, Y., and Dietrich, A. (2003). Plant
mitochondria actively import DNA via the permeability transition pore
complex. EMBO J. 22:1245–1254.
Loro, G., Ruberti, C., Zottini, M., and Costa, A. (2013). The D3cpv
Cameleon reports Ca2+ dynamics in plant mitochondria with similar
kinetics of the YC3.6 Cameleon, but with a lower sensitivity. J.
Microsc. 249:8–12.
Kovacs-Bogdan, E., Sancak, Y., Kamer, K.J., Plovanich, M.,
Jambhekar, A., Huber, R.J., Myre, M.A., Blower, M.D., and
Mootha, V.K. (2014). Reconstitution of the mitochondrial calcium
uniporter in yeast. Proc. Natl. Acad. Sci. USA 111:8985–8990.
Lurin, C., Guclu, J., Cheniclet, C., Carde, J.P., Barbier-Brygoo, H., and
Maurel, C. (2000). CLC-Nt1, a putative chloride channel protein
of tobacco, co-localizes with mitochondrial membrane markers.
Biochem. J. 348:291–295.
Kugler, A., Kohler, B., Palme, K., Wolff, P., and Dietrich, P. (2009). Saltdependent regulation of a CNG channel subfamily in Arabidopsis. BMC
Plant Biol. 9:140.
Maierhofer, T., Lind, C., Huttl, S., Scherzer, S., Papenfuss, M., Simon,
J., Al-Rasheid, K.A., Ache, P., Rennenberg, H., Hedrich, R., et al.
(2014). A single-pore residue renders the Arabidopsis root anion
channel SLAH2 highly nitrate selective. Plant Cell 26:2554–2567.
Kumazaki, K., Chiba, S., Takemoto, M., Furukawa, A., Nishiyama, K.,
Sugano, Y., Mori, T., Dohmae, N., Hirata, K., Nakada-Nakura, Y.,
et al. (2014). Structural basis of Sec-independent membrane protein
insertion by YidC. Nature 509:516–520.
Kunz, H.H., Gierth, M., Herdean, A., Satoh-Cruz, M., Kramer, D.M.,
Spetea, C., and Schroeder, J.I. (2014). Plastidial transporters KEA1,
-2, and -3 are essential for chloroplast osmoregulation, integrity,
and pH regulation in Arabidopsis. Proc. Natl. Acad. Sci. USA
111:7480–7485.
Kuo, M.M., Haynes, W.J., Loukin, S.H., Kung, C., and Saimi, Y. (2005).
Prokaryotic K(+) channels: from crystal structures to diversity. FEMS
Microbiol. Rev. 29:961–985.
Kwon, J., Oh, J., Park, C., Cho, K., Kim, S.I., Kim, S., Lee, S., Bhak, J.,
Norling, B., and Choi, J.S. (2010). Systematic cyanobacterial
membrane proteome analysis by combining acid hydrolysis and
digestive enzymes with nano-liquid chromatography-Fourier
transform mass spectrometry. J. Chromatogr. A 1217:285–293.
Ladwig, F., Dahlke, R.I., Stuhrwohldt, N., Hartmann, J., Harter, K., and
Sauter, M. (2015). Phytosulfokine regulates growth in Arabidopsis
through a response module at the plasma membrane that
includes CYCLIC NUCLEOTIDE-GATED CHANNEL17, H+-ATPase,
and BAK1. Plant Cell 27:1718–1729.
Laloi, M. (1999). Plant mitochondrial carriers: an overview. Cell Mol. Life
Sci. 56:918–944.
Lee, S.M., Hoang, M.H., Han, H.J., Kim, H.S., Lee, K., Kim, K.E., Kim,
D.H., Lee, S.Y., and Chung, W.S. (2009). Pathogen inducible
voltage-dependent anion channel (AtVDAC) isoforms are localized to
mitochondria membrane in Arabidopsis. Mol. Cell 27:321–327.
Lee, C.P., Taylor, N.L., and Millar, A.H. (2013). Recent advances in the
composition and heterogeneity of the Arabidopsis mitochondrial
proteome. Front. Plant Sci. 4:4.
Malcolm, H.R., Heo, Y.Y., Caldwell, D.B., McConnell, J.K., Hawkins,
J.F., Guayasamin, R.C., Elmore, D.E., and Maurer, J.A. (2012). SsbCNGa: a unique member of the bacterial cyclic nucleotide gated
(bCNG) channel family that gates in response to mechanical tension.
Eur. Biophys. J. 41 (12):1003–1013.
Marmagne, A., Vinauger-Douard, M., Monachello, D., de Longevialle,
A.F., Charon, C., Allot, M., Rappaport, F., Wollman, F.A., BarbierBrygoo, H., and Ephritikhine, G. (2007). Two members of the
Arabidopsis CLC (chloride channel) family, AtCLCe and AtCLCf, are
associated with thylakoid and Golgi membranes, respectively. J.
Exp. Bot. 58:3385–3393.
Martin, W., Stoebe, B., Goremykin, V., Hapsmann, S., Hasegawa, M.,
and Kowallik, K.V. (1998). Gene transfer to the nucleus and the
evolution of chloroplasts. Nature 393:162–165.
Martinoia, E., Meyer, S., De Angeli, A., and Nagy, R. (2012). Vacuolar
transporters in their physiological context. Annu. Rev. Plant Biol.
63:183–213.
Matkovic, K., Koszela-Piotrowska, I., Jarmuszkiewicz, W., and
Szewczyk, A. (2011). Ion conductance pathways in potato tuber
(Solanum tuberosum) inner mitochondrial membrane. Biochim.
Biophys. Acta 1807:275–285.
Matsuda, N., and Uozumi, N. (2006). Ktr-mediated potassium transport,
a major pathway for potassium uptake, is coupled to a proton gradient
across the membrane in Synechocystis sp. PCC 6803. Biosci.
Biotechnol. Biochem. 70:273–275.
Maurel, C., Reizer, J., Schroeder, J.I., and Chrispeels, M.J. (1993). The
vacuolar membrane protein gamma-TIP creates water specific
channels in Xenopus oocytes. EMBO J. 12:2241–2247.
Maurel, C., Verdoucq, L., Luu, D.T., and Santoni, V. (2008). Plant
aquaporins: membrane channels with multiple integrated functions.
Annu. Rev. Plant Biol. 59:595–624.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
391
Molecular Plant
Mayer, M.L., Olson, R., and Gouaux, E. (2001). Mechanisms for ligand
binding to GluR0 ion channels: crystal structures of the glutamate
and serine complexes and a closed apo state. J. Mol. Biol.
311:815–836.
McAinsh, M.R., and Pittman, J.K. (2009). Shaping the calcium signature.
New Phytol. 181:275–294.
Mehlmer, N., Parvin, N., Hurst, C.H., Knight, M.R., Teige, M., and
Vothknecht, U.C. (2012). A toolset of aequorin expression vectors
for in planta studies of subcellular calcium concentrations in
Arabidopsis thaliana. J. Exp. Bot. 63:1751–1761.
Mei, H., Cheng, N.H., Zhao, J., Park, S., Escareno, R.A., Pittman, J.K.,
and Hirschi, K.D. (2009). Root development under metal stress in
Arabidopsis thaliana requires the H+/cation antiporter CAX4. New
Phytol. 183:95–105.
Meyerhoff, O., Muller, K., Roelfsema, M.R., Latz, A., Lacombe, B.,
Hedrich, R., Dietrich, P., and Becker, D. (2005). AtGLR3.4, a
glutamate receptor channel-like gene is sensitive to touch and cold.
Planta 222:418–427.
Michal Johnson, J., Reichelt, M., Vadassery, J., Gershenzon, J., and
Oelmuller, R. (2014). An Arabidopsis mutant impaired in intracellular
calcium elevation is sensitive to biotic and abiotic stress. BMC Plant
Biol. 14:162.
Michaud, M., Ubrig, E., Filleur, S., Erhardt, M., Ephritikhine, G.,
Marechal-Drouard, L., and Duchene, A.M. (2014). Differential
targeting of VDAC3 mRNA isoforms influences mitochondria
morphology. Proc. Natl. Acad. Sci. USA 111:8991–8996.
Millar, A.H., Heazlewood, J.L., Kristensen, B.K., Braun, H.P., and
Moller, I.M. (2005). The plant mitochondrial proteome. Trends Plant
Sci. 10:36–43.
Millar, A.H., Whelan, J., Soole, K.L., and Day, D.A. (2011). Organization
and regulation of mitochondrial respiration in plants. Annu. Rev. Plant
Biol. 62:79–104.
Nanatani, K., Shijuku, T., Akai, M., Yukutake, Y., Yasui, M.,
Hamamoto, S., Onai, K., Morishita, M., Ishiura, M., and Uozumi,
N. (2013). Characterization of the role of a mechanosensitive channel
in osmotic down shock adaptation in Synechocystis sp PCC 6803.
Channels (Austin) 7:238–242.
Nanatani, K., Shijuku, T., Takano, Y., Zulkifli, L., Yamazaki, T.,
Tominaga, A., Souma, S., Onai, K., Morishita, M., Ishiura, M.,
et al. (2015). Comparative analysis of kdp and ktr mutants
reveals distinct roles of the potassium transporters in the model
cyanobacterium Synechocystis sp. strain PCC 6803. J. Bacteriol.
197:676–687.
Nanjo, Y., Oka, H., Ikarashi, N., Kaneko, K., Kitajima, A., Mitsui, T.,
Munoz, F.J., Rodriguez-Lopez, M., Baroja-Fernandez, E., and
Pozueta-Romero, J. (2006). Rice plastidial N-glycosylated
nucleotide pyrophosphatase/phosphodiesterase is transported from
the ER-Golgi to the chloroplast through the secretory pathway. Plant
Cell 18:2582–2592.
Ion Channels in Plant Bioenergetic Organelles
Niyogi, K.K., and Truong, T.B. (2013). Evolution of flexible nonphotochemical quenching mechanisms that regulate light harvesting
in oxygenic photosynthesis. Curr. Opin. Plant Biol. 16:307–314.
Nomura, H., and Shiina, T. (2014). Calcium signaling in plant
endosymbiotic organelles: mechanism and role in physiology. Mol.
Plant 7:1094–1104.
Nomura, H., Komori, T., Kobori, M., Nakahira, Y., and Shiina, T. (2008).
Evidence for chloroplast control of external Ca2+-induced cytosolic
Ca2+ transients and stomatal closure. Plant J. 53:988–998.
Nomura, H., Komori, T., Uemura, S., Kanda, Y., Shimotani, K., Nakai,
K., Furuichi, T., Takebayashi, K., Sugimoto, T., Sano, S., et al.
(2012). Chloroplast-mediated activation of plant immune signalling in
Arabidopsis. Nat. Commun. 3:926.
Nouet, C., Motte, P., and Hanikenne, M. (2011). Chloroplastic and
mitochondrial metal homeostasis. Trends Plant Sci. 16:395–404.
Oh, Y.J., and Hwang, I. (2015). Targeting and biogenesis of transporters
and channels in chloroplast envelope membranes: unsolved
questions. Cell Calcium 58:122–130.
Osteryoung, K.W., and Pyke, K.A. (2014). Division and dynamic
morphology of plastids. Annu. Rev. Plant Biol. 65:443–472.
Ozaki, H., Ikeuchi, M., Ogawa, T., Fukuzawa, H., and Sonoike, K.
(2007). Large-scale analysis of chlorophyll fluorescence kinetics in
Synechocystis sp. PCC 6803: identification of the factors involved
in the modulation of photosystem stoichiometry. Plant Cell Physiol.
48:451–458.
Pagliarini, D.J., Calvo, S.E., Chang, B., Sheth, S.A., Vafai, S.B., Ong,
S.-E., Walford, G.A., Sugiana, C., Boneh, A., Chen, W.K., et al.
(2008). A mitochondrial protein compendium elucidates complex I
disease biology. Cell 134:112–123.
Paila, Y.D., Richardson, L.G., and Schnell, D.J. (2015). New insights into
the mechanism of chloroplast protein import and its integration with
protein quality control, organelle biogenesis and development. J.
Mol. Biol. 427:1038–1060.
Pan, X., Chen, Z., Yang, X., and Liu, G. (2014). Arabidopsis voltagedependent anion channel 1 (AtVDAC1) is required for female
development and maintenance of mitochondrial functions related to
energy-transaction. PLoS One 9:e106941.
Pastore, D., Stoppelli, M.C., Di Fonzo, N., and Passarella, S. (1999).
The existence of the K(+) channel in plant mitochondria. J. Biol.
Chem. 274:26683–26690.
Pastore, D., Soccio, M., Laus, M.N., and Trono, D. (2013). The
uniqueness of the plant mitochondrial potassium channel. BMB Rep.
46:391–397.
Patron, M., Checchetto, V., Raffaello, A., Teardo, E., Vecellio Reane,
D., Mantoan, M., Granatiero, V., Szabo, I., De Stefani, D., and
Rizzuto, R. (2014). MICU1 and MICU2 finely tune the mitochondrial
Ca2+ uniporter by exerting opposite effects on MCU activity. Mol.
Cell 53:726–737.
Nazarenko, L.V., Andreev, I.M., Lyukevich, A.A., Pisareva, T.V., and
Los, D.A. (2003). Calcium release from Synechocystis cells induced
by depolarization of the plasma membrane: MscL as an outward
Ca2+ channel. Microbiology 149:1147–1153.
Paupe, V., Prudent, J., Dassa, E.P., Rendon, O.Z., and Shoubridge,
E.A. (2015). CCDC90A (MCUR1) is a cytochrome c oxidase
assembly factor and not a regulator of the mitochondrial calcium
uniporter. Cell Metab. 21:109–116.
Neuhaus, H.E., and Wagner, R. (2000). Solute pores, ion channels, and
metabolite transporters in the outer and inner envelope membranes
of higher plant plastids. Biochim. Biophys. Acta 1465:307–323.
Paynter, J.J., Andres-Enguix, I., Fowler, P.W., Tottey, S., Cheng, W.,
Enkvetchakul, D., Bavro, V.N., Kusakabe, Y., Sansom, M.S.,
Robinson, N.J., et al. (2010). Functional complementation and
genetic deletion studies of KirBac channels: activatory mutations
highlight gating-sensitive domains. J. Biol. Chem. 285:40754–40761.
Nieves-Cordones, M., Chavanieu, A., Jeanguenin, L., Alcon, C.,
Szponarski, W., Estaran, S., Cherel, I., Zimmermann, S.,
Sentenac, H., and Gaillard, I. (2014). Distinct amino acids in the
C-linker domain of the Arabidopsis K+ channel KAT2 determine its
subcellular localization and activity at the plasma membrane. Plant
Physiol. 164:1415–1429.
392
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Peltier, J.B., Ytterberg, A.J., Sun, Q., and van Wijk, K.J. (2004). New
functions of the thylakoid membrane proteome of Arabidopsis
thaliana revealed by a simple, fast, and versatile fractionation
strategy. J. Biol. Chem. 279:49367–49383.
Ion Channels in Plant Bioenergetic Organelles
Perocchi, F., Gohil, V.M., Girgis, H.S., Bao, X.R., McCombs, J.E.,
Palmer, A.E., and Mootha, V.K. (2010). MICU1 encodes a
mitochondrial EF hand protein required for Ca(2+) uptake. Nature
467:291–296.
Molecular Plant
have a dual localization in the cell but show a distinct role in
mitochondria. Plant Mol. Biol. 78:431–446.
Rocchetti, A., Sharma, T., Wulfetange, C., Scholz-Starke, J., Grippa,
A., Carpaneto, A., Dreyer, I., Vitale, A., Czempinski, K., and
Pedrazzini, E. (2012). The putative K(+) channel subunit AtKCO3
forms stable dimers in Arabidopsis. Front. Plant Sci. 3:251.
Petroutsos, D., Busch, A., Janssen, I., Trompelt, K., Bergner, S.V.,
Weinl, S., Holtkamp, M., Karst, U., Kudla, J., and Hippler, M.
(2011). The chloroplast calcium sensor CAS is required for
photoacclimation in Chlamydomonas reinhardtii. Plant Cell 23:2950–
2963.
Rocha, A.G., and Vothknecht, U.C. (2012). The role of calcium in
chloroplasts–an intriguing and unresolved puzzle. Protoplasma
249:957–966.
Petrussa, E., Casolo, V., Peresson, C., Braidot, E., Vianello, A., and
Macri, F. (2004). The K(ATP)+ channel is involved in a low-amplitude
permeability transition in plant mitochondria. Mitochondrion
3:297–307.
Rolland, N., Curien, G., Finazzi, G., Kuntz, M., Marechal, E., Matringe,
M., Ravanel, S., and Seigneurin-Berny, D. (2012). The biosynthetic
capacities of the plastids and integration between cytoplasmic and
chloroplast processes. Annu. Rev. Genet. 46:233–264.
Petrussa, E., Casolo, V., Peresson, C., Krajnakova, J., Macri, F., and
Vianello, A. (2008). Activity of a KATP+ channel in Arum spadix
mitochondria during thermogenesis. J. Plant Physiol. 165:1360–1369.
Safiarian, M.J., Pertl-Obermeyer, H., Lughofer, P., Hude, R., Bertl, A.,
and Obermeyer, G. (2015). Lost in traffic? the K(+) channel of lily
pollen, LilKT1, is detected at the endomembranes inside yeast cells,
tobacco leaves, and lily pollen. Front. Plant Sci. 6:47.
Pfeil, B., Schoefs, B., and Spetea, C. (2014). Function and evolution of
channels and transporters in photosynthetic membranes. Cell Mol.
Life Sci. 71:979–998.
Pisareva, T., Kwon, J., Oh, J., Kim, S., Ge, C., Wieslander, A., Choi,
J.S., and Norling, B. (2011). Model for membrane organization and
protein sorting in the cyanobacterium Synechocystis sp. PCC 6803
inferred from proteomics and multivariate sequence analyses. J.
Proteome Res. 10:3617–3631.
Pottosin, I.I. (1992). Single channel recording in the chloroplast envelope.
FEBS Lett. 308:87–90.
Pottosin, I.I. (1993). One of the chloroplast envelope ion channels is
probably related to the mitochondrial VDAC. FEBS Lett. 330:211–214.
Pottosin, I.I., and Schonknecht, G. (1995). Patch clamp study of the
voltage-dependent anion channel in the thylakoid membrane. J.
Membr. Biol. 148:143–156.
Pottosin, I.I., and Schonknecht, G. (1996). Ion channel permeable
for divalent and monovalent cations in native spinach thylakoid
membranes. J. Membr. Biol. 152:223–233.
Pottosin, I.I., Muñiz, J., and Shabala, S. (2005). Fast-activating channel
controls cation fluxes across the native chloroplast envelope. J.
Membr. Biol. 204:145–156.
Preston, G.M., Carroll, T.P., Guggino, W.B., and Agre, P. (1992).
Appearance of water channels in Xenopus oocytes expressing red
cell CHIP28 protein. Science 256:385–387.
Qin, G., Meng, X., Wang, Q., and Tian, S. (2009). Oxidative damage
of mitochondrial proteins contributes to fruit senescence: a redox
proteomics analysis. J. Proteome Res. 8:2449–2462.
Qiu, H., Price, D.C., Weber, A.P., Facchinelli, F., Yoon, H.S., and
Bhattacharya, D. (2013). Assessing the bacterial contribution to the
plastid proteome. Trends Plant Sci. 18:680–687.
Reintanz, B., Szyroki, A., Ivashikina, N., Ache, P., Godde, M., Becker,
D., Palme, K., and Hedrich, R. (2002). AtKC1, a silent Arabidopsis
potassium channel alpha -subunit modulates root hair K+ influx.
Proc. Natl. Acad. Sci. USA 99:4079–4084.
Rhee, H.W., Zou, P., Udeshi, N.D., Martell, J.D., Mootha, V.K., Carr,
S.A., and Ting, A.Y. (2013). Proteomic mapping of mitochondria
in living cells via spatially restricted enzymatic tagging. Science
339:1328–1331.
Rizzuto, R., De Stefani, D., Raffaello, A., and Mammucari, C. (2012).
Mitochondria as sensors and regulators of calcium signalling. Nat.
Rev. Mol. Cell Biol. 13:566–578.
Robert, N., d’Erfurth, I., Marmagne, A., Erhardt, M., Allot, M., Boivin,
K., Gissot, L., Monachello, D., Michaud, M., Duchene, A.M., et al.
(2012). Voltage-dependent-anion-channels (VDACs) in Arabidopsis
Sai, J., and Johnson, C.H. (2002). Dark-stimulated calcium ion fluxes in
the chloroplast stroma and cytosol. Plant Cell 14:1279–1291.
Salvi, D., Moyet, L., Seigneurin-Berny, D., Ferro, M., Joyard, J., and
Rolland, N. (2011). Preparation of envelope membrane fractions
from Arabidopsis chloroplasts for proteomic analysis and other
studies. Methods Mol. Biol. 775:189–206.
Sato, S., Shimoda, Y., Muraki, A., Kohara, M., Nakamura, Y., and
Tabata, S. (2007). A large-scale protein protein interaction analysis in
Synechocystis sp. PCC6803. DNA Res. 14:207–216.
Saviani, E.E., Orsi, C.H., Oliveira, J.F., Pinto-Maglio, C.A., and
Salgado, I. (2002). Participation of the mitochondrial permeability
transition pore in nitric oxide-induced plant cell death. FEBS Lett.
510:136–140.
Schleiff, E., and Becker, T. (2011). Common ground for protein
translocation: access control for mitochondria and chloroplasts. Nat.
Rev. Mol. Cell Biol. 12:48–59.
Schonknecht, G., Hedrich, R., Junge, W., and Raschke, K. (1988). A
voltage-dependent chloride channel in the photosynthetic membrane
of higher plant. Nature 336:589–592.
Schwacke, R., Schneider, A., van der Graaff, E., Fischer, K., Catoni, E.,
Desimone, M., Frommer, W.B., Flugge, U.I., and Kunze, R. (2003).
ARAMEMNON, a novel database for Arabidopsis integral membrane
proteins. Plant Physiol. 131:16–26.
Shanmughapriya, S., Rajan, S., Hoffman, N.E., Higgins, A.M., Tomar,
D., Nemani, N., Hines, K.J., Smith, D.J., Eguchi, A., Vallem, S., et al.
(2015). SPG7 is an essential and conserved component of the
mitochondrial permeability transition pore. Mol. Cell 60:47–62.
Shapiguzov, A., Lyukevich, A.A., Allakhverdiev, S.I., Sergeyenko, T.V.,
Suzuki, I., Murata, N., and Los, D.A. (2005). Osmotic shrinkage of
cells of Synechocystis sp. PCC 6803 by water efflux via aquaporins
regulates osmostress-inducible gene expression. Microbiology 151:
447–455.
Simm, S., Papasotiriou, D.G., Ibrahim, M., Leisegang, M.S., Muller, B.,
Schorge, T., Karas, M., Mirus, O., Sommer, M.S., and Schleiff, E.
(2013). Defining the core proteome of the chloroplast envelope
membranes. Front. Plant Sci. 4:11.
Singh, H., Lu, R., Bopassa, J.C., Meredith, A.L., Stefani, E., and Toro,
L. (2013). MitoBK(Ca) is encoded by the Kcnma1 gene, and a splicing
sequence defines its mitochondrial location. Proc. Natl. Acad. Sci. USA
110:10836–10841.
Slavikova, S., Vacula, R., Fang, Z., Ehara, T., Osafune, T., and
Schwartzbach, S.D. (2005). Homologous and heterologous
reconstitution of Golgi to chloroplast transport and protein import
into the complex chloroplasts of Euglena. J. Cell Sci. 118:1651–1661.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
393
Molecular Plant
Stael, S., Wurzinger, B., Mair, A., Mehlmer, N., Vothknecht, U.C., and
Teige, M. (2012). Plant organellar calcium signalling: an emerging field.
J. Exp. Bot. 63:1525–1542.
Stephan, A.B., and Schroeder, J.I. (2014). Plant salt stress status is
transmitted systemically via propagating calcium waves. Proc. Natl.
Acad. Sci. USA 111:6126–6127.
Strittmatter, P., Soll, J., and Bolter, B. (2010). The chloroplast protein
import machinery: a review. Methods Mol. Biol. 619:307–321.
Sweetlove, L.J., Taylor, N.L., and Leaver, C.J. (2007). Isolation of intact,
functional mitochondria from the model plant Arabidopsis thaliana.
Methods Mol. Biol. 372:125–136.
Szabo, I., and Zoratti, M. (2014). Mitochondrial channels: ion fluxes and
more. Physiol. Rev. 94:519–608.
Szewczyk, A., Skalska, J., Glab, M., Kulawiak, B., Malinska, D.,
Koszela-Piotrowska, I., and Kunz, W.S. (2006). Mitochondrial
potassium channels: from pharmacology to function. Biochim.
Biophys. Acta 1757:715–720.
Szewczyk, A., Jarmuszkiewicz, W., and Kunz, W.S. (2009).
Mitochondrial potassium channels. IUBMB Life 61:134–143.
Takahashi, Y., and Tateda, C. (2013). The functions of voltagedependent anion channels in plants. Apoptosis 18:917–924.
Takano, J., Wada, M., Ludewig, U., Schaaf, G., von Wiren, N., and
Fujiwara, T. (2006). The Arabidopsis major intrinsic protein NIP5;1 is
essential for efficient boron uptake and plant development under
boron limitation. Plant Cell 18:1498–1509.
Ion Channels in Plant Bioenergetic Organelles
Tunc-Ozdemir, M., Rato, C., Brown, E., Rogers, S., Mooneyham, A.,
Frietsch, S., Myers, C.T., Poulsen, L.R., Malho, R., and Harper,
J.F. (2013a). Cyclic nucleotide gated channels 7 and 8 are essential
for male reproductive fertility. PLoS One 8:e55277.
Tunc-Ozdemir, M., Tang, C., Ishka, M.R., Brown, E., Groves, N.R.,
Myers, C.T., Rato, C., Poulsen, L.R., McDowell, S., Miller, G.,
et al. (2013b). A cyclic nucleotide-gated channel (CNGC16) in pollen
is critical for stress tolerance in pollen reproductive development.
Plant Physiol. 161:1010–1020.
Uehlein, N., Otto, B., Hanson, D.T., Fischer, M., McDowell, N., and
Kaldenhoff, R. (2008). Function of Nicotiana tabacum aquaporins as
chloroplast gas pores challenges the concept of membrane CO2
permeability. Plant Cell 20:648–657.
van den Wijngaard, P.W., and Vredenberg, W.J. (1997). A 50picosiemens anion channel of the chloroplast envelope is involved in
chloroplast protein import. J. Biol. Chem. 272:29430–29433.
Veley, K.M., Maksaev, G., Frick, E.M., January, E., Kloepper, S.C., and
Haswell, E.S. (2014). Arabidopsis MSL10 has a regulated cell death
signaling activity that is separable from its mechanosensitive ion
channel activity. Plant Cell 26:3115–3131.
Velisetty, P., and Chakrapani, S. (2012). Desensitization mechanism
in prokaryotic ligand-gated ion channel. J. Biol. Chem. 287:18467–
18477.
Very, A.A., and Sentenac, H. (2002). Cation channels in the Arabidopsis
plasma membrane. Trends Plant Sci. 7:168–175.
Tampieri, E., Baraldi, E., Carnevali, F., Frascaroli, E., and De Santis, A.
(2011). The activity of plant inner membrane anion channel (PIMAC) can
be performed by a chloride channel (CLC) protein in mitochondria from
seedlings of maize populations divergently selected for cold tolerance.
J. Bioenerg. Biomembr. 43:611–621.
Vianello, A., Casolo, V., Petrussa, E., Peresson, C., Patui, S., Bertolini,
A., Passamonti, S., Braidot, E., and Zancani, M. (2012). The
mitochondrial permeability transition pore (PTP) - an example of
multiple molecular exaptation? Biochim. Biophys. Acta 1817:2072–
2086.
Teardo, E., Frare, E., Segalla, A., De Marco, V., Giacometti, G.M., and
Szabo, I. (2005). Localization of a putative ClC chloride channel in
spinach chloroplasts. FEBS Lett. 579:4991–4996.
Villarejo, A., Buren, S., Larsson, S., Dejardin, A., Monne, M., Rudhe, C.,
Karlsson, J., Jansson, S., Lerouge, P., Rolland, N., et al. (2005).
Evidence for a protein transported through the secretory pathway en
route to the higher plant chloroplast. Nat. Cell Biol. 7:1224–1231.
Teardo, E., Segalla, A., Formentin, E., Zanetti, M., Marin, O.,
Giacometti, G.M., Lo Schiavo, F., Zoratti, M., and Szabo, I. (2010).
Characterization of a plant glutamate receptor activity. Cell Physiol.
Biochem. 26:253–262.
Teardo, E., Formentin, E., Segalla, A., Giacometti, G.M., Marin, O.,
Zanetti, M., Lo Schiavo, F., Zoratti, M., and Szabo, I. (2011). Dual
localization of plant glutamate receptor AtGLR3.4 to plastids and
plasma membrane. Biochim. Biophys. Acta 1807:359–367.
Teardo, E., Carraretto, L., De Bortoli, S., Costa, A., Behera, S.,
Wagner, R., Lo Schiavo, F., and Formentin, E. (2015). Alternative
splicing-mediated targeting of the Arabidopsis GLUTAMATE
RECEPTOR3.5 to mitochondria affects organelle morphology. Plant
Physiol. 167:216–227.
Terashima, M., Petroutsos, D., Hudig, M., Tolstygina, I., Trompelt, K.,
Gabelein, P., Fufezan, C., Kudla, J., Weinl, S., Finazzi, G., et al.
(2012). Calcium-dependent regulation of cyclic photosynthetic
electron transfer by a CAS, ANR1, and PGRL1 complex. Proc. Natl.
Acad. Sci. USA 109:17717–17722.
Tester, M., and Blatt, M.R. (1989). Direct measurement of k channels in
thylakoid membranes by incorporation of vesicles into planar lipid
bilayers. Plant Physiol. 91:249–252.
Tomizioli, M., Lazar, C., Brugiere, S., Burger, T., Salvi, D., Gatto, L.,
Moyet, L., Breckels, L.M., Hesse, A.M., Lilley, K.S., et al. (2014).
Deciphering thylakoid sub-compartments using a mass spectrometrybased approach. Mol. Cell. Proteomics 13:2147–2167.
Trono, D., Laus, M.N., Soccio, M., and Pastore, D. (2014). Transport
pathways–proton motive force interrelationship in durum wheat
mitochondria. Int. J. Mol. Sci. 15:8186–8215.
394
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
Vincill, E.D., Bieck, A.M., and Spalding, E.P. (2012). Ca(2+) conduction
by an amino acid-gated ion channel related to glutamate receptors.
Plant Physiol. 159:40–46.
Voelker, C., Schmidt, D., Mueller-Roeber, B., and Czempinski, K.
(2006). Members of the Arabidopsis AtTPK/KCO family form
homomeric vacuolar channels in planta. Plant J. 48:296–306.
von Charpuis, C., Meckel, T., Moroni, A., and Thiel, G. (2015). The
sorting of a small potassium channel in mammalian cells can be
shifted between mitochondria and plasma membrane. Cell Calcium
58:114–121.
Waditee, R., Hossain, G.S., Tanaka, Y., Nakamura, T., Shikata, M.,
Takano, J., Takabe, T., and Takabe, T. (2004). Isolation and
functional characterization of Ca2+/H+ antiporters from cyanobacteria.
J. Biol. Chem. 279:4330–4338.
Wagner, S., Behera, S., De Bortoli, S., Logan, D.C., Fuchs, P.,
Carraretto, L., Teardo, E., Cendron, L., Nietzel, T., Fussl, M., et al.
(2015). The EF-Hand Ca2+ binding protein MICU choreographs
mitochondrial Ca2+ dynamics in Arabidopsis. Plant Cell 27:3190–3212.
Wang, D., Xu, Y., Li, Q., Hao, X., Cui, K., Sun, F., and Zhu, Y. (2003).
Transgenic expression of a putative calcium transporter affects the
time of Arabidopsis flowering. Plant J. 33:285–292.
Ward, J.M., Maser, P., and Schroeder, J.I. (2009). Plant ion channels:
gene families, physiology, and functional genomics analyses. Annu.
Rev. Physiol. 71:59–82.
Weber, A.P., Schwacke, R., and Flugge, U.I. (2005). Solute transporters
of the plastid envelope membrane. Annu. Rev. Plant Biol. 56:133–164.
Ion Channels in Plant Bioenergetic Organelles
Weis, B.L., Schleiff, E., and Zerges, W. (2013). Protein targeting to
subcellular organelles via MRNA localization. Biochim. Biophys. Acta
1833:260–273.
Wilson, M.E., Basu, M.R., Bhaskara, G.B., Verslues, P.E., and Haswell,
E.S. (2014). Plastid osmotic stress activates cellular stress responses
in Arabidopsis. Plant Physiol. 165:119–128.
Witt, H.T. (1979). Energy conversion in the functional membrane of
photosynthesis. Analysis by light pulse and electric pulse methods.
The central role of the electric field. Biochim. Biophys. Acta
505:355–427.
Xu, H., Martinoia, E., and Szabo, I. (2015). Organellar channels and
transporters. Cell Calcium 58:1–10.
Yin, L., Vener, A.V., and Spetea, C. (2015). The membrane proteome of
stroma thylakoids from Arabidopsis thaliana studied by successive insolution and in-gel digestion. Physiol. Plant. 154:433–446.
Yogev, O., and Pines, O. (2011). Dual targeting of mitochondrial proteins:
mechanism, regulation and function. Biochim. Biophys. Acta
1808:1012–1020.
Molecular Plant
Zanetti, M., Teardo, E., La Rocca, N., Zulkifli, L., Checchetto, V.,
Shijuku, T., Sato, Y., Giacometti, G.M., Uozumi, N., Bergantino,
E., et al. (2010). A novel potassium channel in photosynthetic
cyanobacteria. PLoS One 5:e10118.
Zifarelli, G., and Pusch, M. (2010). CLC transport proteins in plants.
FEBS Lett. 584:2122–2127.
Zoratti, M., and Szabo, I. (1995). The mitochondrial permeability
transition. Biochim. Biophys. Acta 1241:139–176.
Zottini, M., and Zannoni, D. (1993). The use of Fura-2 fluorescence to
monitor the movement of free calcium ions into the matrix of plant
mitochondria (Pisum sativum and Helianthus tuberosus). Plant
Physiol. 102:573–578.
Zottini, M., Barizza, E., Costa, A., Formentin, E., Ruberti, C., Carimi, F.,
and Lo Schiavo, F. (2008). Agroinfiltration of grapevine leaves for fast
transient assays of gene expression and for long-term production of
stable transformed cells. Plant Cell Rep. 27:845–853.
Molecular Plant 9, 371–395, March 2016 ª The Author 2016.
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