Plant Science 255 (2017) 1–11
Contents lists available at ScienceDirect
Plant Science
journal homepage: www.elsevier.com/locate/plantsci
Review article
Dithiol disulphide exchange in redox regulation of chloroplast
enzymes in response to evolutionary and structural constraints
Desirée D. Gütle a,b,c,d,∗,1 , Thomas Roret a,b,1 , Arnaud Hecker a,b , Ralf Reski c,d,e ,
Jean-Pierre Jacquot a,b,∗
a
Université de Lorraine, UMR 1136 Interactions Arbres Microorganismes, F-54500 Vandœuvre-lès-Nancy, France
INRA, UMR 1136 Interactions Arbres Microorganismes, F-54280 Champenoux, France
c
Plant Biotechnology, Faculty of Biology, University of Freiburg, Schänzlestr. 1, 79104 Freiburg, Germany
d
Spemann Graduate School of Biology and Medicine (SGBM), University of Freiburg, 79104 Freiburg, Germany
e
BIOSS − Centre for Biological Signalling Studies, University of Freiburg, Schänzlestr. 18, 79104 Freiburg, Germany
b
a r t i c l e
i n f o
Article history:
Received 13 September 2016
Received in revised form 4 November 2016
Accepted 5 November 2016
Available online 8 November 2016
Keywords:
Redox regulation
Disulphide
Dithiol
Ferredoxin
Thioredoxin
Calvin-Benson cycle
a b s t r a c t
Redox regulation of chloroplast enzymes via disulphide reduction is believed to control the rates of CO2
fixation. The study of the thioredoxin reduction pathways and of various target enzymes lead to the
following guidelines:
i) Thioredoxin gene content is greatly higher in photosynthetic eukaryotes compared to prokaryotes;
ii) Thioredoxin-reducing pathways have expanded in photosynthetic eukaryotes with four different
thioredoxin reductases and the possibility to reduce some thioredoxins via glutaredoxins;
iii) Some enzymes that were thought to be strictly linked to photosynthesis ferredoxin-thioredoxin
reductase, phosphoribulokinase, ribulose-1,5-bisphosphate carboxylase/oxygenase, sedoheptulose1,7-bisphosphatase are present in non-photosynthetic organisms;
iv) Photosynthetic eukaryotes contain a genetic patchwork of sequences borrowed from prokaryotes
including ␣–proteobacteria and archaea;
v) The introduction of redox regulatory sequences did not occur at the same place for all targets. Some
possess critical cysteines in cyanobacteria, for others the transition occurred rather at the green algae
level;
vi) Generally the regulatory sites of the target enzymes are distally located from the catalytic sites. The
cysteine residues are generally not involved in catalysis. Following reduction, molecular movements
open the active sites and make catalysis possible;
vii) The regulatory sequences are located on surface-accessible loops. At least one instance they can be cut
out and serve as signal peptides for inducing plant defence.
© 2016 Elsevier Ireland Ltd. All rights reserved.
Contents
1.
2.
Principles of redox regulation of chloroplast enzymes via dithiol-disulphide exchange, what are the actors, what are the targets? . . . . . . . . . . . . . . . . . 2
1.1.
Diversification of the electron donor systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
1.2.
The complex TRX gene family of plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
Why did redox regulation via dithiol reduction appear? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2
2.1.
Oxidative stress increases during evolution of higher plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3
2.2.
Functional modifications of target proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
∗ Corresponding authors at: Université de Lorraine, UMR 1136 Interactions Arbres Microorganismes, Vandœuvre-lès-Nancy, F-54500, France.
E-mail addresses: [email protected] (D.D. Gütle), [email protected] (J.-P. Jacquot).
1
Joint first authors.
http://dx.doi.org/10.1016/j.plantsci.2016.11.003
0168-9452/© 2016 Elsevier Ireland Ltd. All rights reserved.
2
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
3.
4.
5.
6.
When and how did redox regulation via dithiol reduction appear? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
3.1.
Occurrence of TRX reductases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
3.2.
Integration of redox sensitive cysteine residues in target protein sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4
In response to which structural constraints did redox regulation happen on molecular targets? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
4.1.
ATP synthase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6
4.2.
NADP-malate dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
4.3.
Glucose 6-phosphate dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
4.4.
Fructose-1,6-bisphosphatase and sedoheptulose-1,7-bisphosphatase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
4.5.
Phosporibulokinase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
4.6.
Glyceraldehyde 3-phosphate dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8
Do the regulatory sequences have additional roles in signalling? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Potential new avenues of research in the redox field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9
Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10
1. Principles of redox regulation of chloroplast enzymes via
dithiol-disulphide exchange, what are the actors, what are
the targets?
and the catalytic disulphide. Those four reduction pathways in land
plants are schematized in Fig. 1, providing also a list of chloroplast
target enzymes for which molecular structural details are available
and which will be discussed later in more detail in this review.
1.1. Diversification of the electron donor systems
Redox regulation by dithiol-disulphide exchange reactions
together with the thiol-dependent repair systems (peroxiredoxins (PRX) and methionine sulfoxide reductases (Msr)) is getting
ever more attention in all biological systems, but it is clearly in
plants and especially in land plants that these processes have
become amazingly diversified [1–5]. Ubiquitously present, thioredoxin proteins (TRX) are central to those reactions. They have
unmatched ability to reduce (and oxidize) disulphide bonds on target enzymes/proteins that perform reduction reactions. TRX are
then reduced at the expense of NADPH via an NADPH-dependent
thioredoxin reductase (NTR) in bacteria [6]. The situation is quite
similar in animal/mammalian cells with NADPH as the primary
electron provider and NTR as a signal transmitter. However, as animal cells evolved multiple compartments, the reducing systems
became more diversified with the existence of both cytosolic and
mitochondrial systems. In animal cell cytosolic and mitochondrial
NTR components are modified by a C-terminal extension possessing a selenocysteine. All animal NTRs have similar properties
forming a homodimer with FAD and a redox active selenosulfide together with a disulphide in each subunit. One complication
with mammalian thioredoxin reductase genes is their capacity to
undergo differential exon splicing leading to the generation of multiple forms of the enzyme in both compartments [7]. Conversely,
plants are even more complicated than animal or fungal cells, possessing an extra compartment, the chloroplast which performs
oxygenic photosynthesis. Accordingly, currently four different TRX
reduction pathways have been described in plant cells [8]. Two of
those are similar to those found in animal cells i.e. mitochondrial
and cytosolic NTR-based systems. There is though a major difference; NTRs in cytosolic and mitochondrial land plant are of the
prokaryotic type, containing the disulphide and FAD but lacking the
C-terminal extension and the selenocysteine residue. Selenocysteines are not present in photosynthetic eukaryotes after the green
algae lineage [9]. The two other TRX reduction systems are located
in the chloroplast; one of these is called NTRC and it is actually a
variant of the prokaryotic NTR with a built-in TRX domain in the
C-terminus [10–12]. The other is the ferredoxin-thioredoxin system consisting of a redox cascade involving the photosystems, and
the following stromal components: ferredoxin (FDX), ferredoxinthioredoxin reductase (FTR) and chloroplast-located TRXs [13,14].
FTR is a heterodimer with a so called “variable subunit”, with a likely
protective task [15], and a somewhat larger and more conserved
catalytic subunit. The catalytic subunit bears both a [4Fe-4S] centre
1.2. The complex TRX gene family of plants
The diversification of the TRX reductions modes is not the only
significant difference between plant cells and other organisms. One
additional indicator is the number of TRX genes. Strictly speaking,
a TRX should bear a WCGPC redox-active site although WCPPC signature can be quite frequent especially in plants, but other more
deviant motifs can also be found [16,17]. Anyway, when considering only the classical (and more numerous) WCGPC containing
TRXs, bacterial and animal species contain but only few of those
(1–3 sequences on average) [2,18], while land plants have a large
content of those genes (nearly 20 on average) with an unequal distribution between compartments with most in chloroplasts (f, m, x,
y, z isoforms) or cytosol (h isoforms), whilst mitochondria contain h
and o isoforms [3,5,19]. The expansion of the diversity of TRX genes
in plants is not unique to this antioxidant protein. It is even more
enhanced for its cousin protein glutaredoxin (GRX) with more than
40 genes of that category in plant genomes, some of them unique to
land plants [20]. The reasons for this remarkable enhanced diversity of proteins of the broad redoxin lineage will be discussed in
a later section. It is, however, of interest to note that there are
connections between the TRX and GRX systems and at least one
plant TRX is reduced by a GRX system. It can also be palmitoylated
and has the capacity to move from cell to cell [21,22]. Likewise,
there are proteins clearly more related phylogenetically to TRXs
but with non-canonical active sites which in reality are reduced by
glutathione (GSH) and behave as GRXs [17]. The complex interplay
between the TRX and GRX systems has been described in a number of recent reviews [3,4,23,24] and recent paper discuss how FTR
and NTRC systems can back up one another and, as a consequence,
the analysis of mutants of the regulatory cascades needs to be done
with circumspection [25–27].
2. Why did redox regulation via dithiol reduction appear?
As noted above, two indicators reflect the expansion of the
redox networks in land plants, i.e. the diversification of the reduction routes and the expansion and diversification of the TRX and
GRX genes. Actually, a third criterion can be added to that list: the
appearance of enzyme target sequences containing critical disulphides needed for ON/OFF regulation in chloroplasts. During the
dark phase, plastid enzymes are generally under an oxidized state
and turn out inactive whereas in the light phase, enzymes are
rather in reduced form and are active. In contrast, plastid glucose
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
3
Fig. 1. Schematic representation of the reduction pathways found in land plants localized in the three sub-compartments mitochondria, chloroplast and cytosol. NTR −
NADPH-thioredoxin reductase; TRX − thioredoxin; Fd − ferredoxin; FTR − ferredoxin-thioredoxin reductase; GR − glutathione reductase; GSH − reduced gluthathione; GRX
− glutaredoxin; PS − photosystem. Pink regions of NTRC indicate TRX domain.
6-phosphate dehydrogenase (G6PDH), behaves the opposite way.
With this observation in mind, two questions immediately arise:
why and when did redox regulation appear?
2.1. Oxidative stress increases during evolution of higher plants
In response to the why, essentially two hypotheses have been
put forward. The first one is based on the coincidence of the
advent of redox regulatory pathways with oxygenic photosynthesis. It is believed that oxygen began to accumulate on the earth
after the elaboration of oxygenic photosynthesis based on the
capacity of organisms able to cleave water molecules with the
release of protons, electrons and molecular oxygen. Oxygenic photosynthesis is believed to have been initiated in cyanobacteria
and further developed in algae and land plants. In this scenario,
as the oxygen content gradually increased on the still primitive
earth, various reactive oxygen species (ROS) have been and are
still being produced (singlet oxygen, hydroxyl radicals, superoxide ions, hydrogen peroxide) as well as reactive nitrogen species
(RNS). Several of ROS and/or RNS compounds are used in cellular
signal transduction pathways, but when their levels become uncontrolled they can damage macromolecules, including lipids, nucleic
acids and proteins. The proteins are mostly affected at the level
of methionine and cysteine, the two of the most reactive amino
acids. As a consequence, all living cells evolved detoxification and
repair systems. The detoxification systems include catalase in peroxisomes (destroys H2 O2 ), ascorbate and glutathione peroxidases
and superoxide dismutases in various sub-compartments (reduce
superoxide ions) but also some very abundant enzymes called PRX
or thiol-peroxidases. The PRX are present in nearly all cell compartments and they can reduce simple peroxides as H2 O2 but also
more complex ones as lipid hydroperoxides (compounds that catalase cannot reduce) [28]. The catalytic cysteine of PRXs becomes
oxidized to a sulfenic acid In the catalytic cycle that is regenerated
via either the TRX or the GRX systems [29].
Methionine sulfoxide reductases that can reduce methionine
sulfoxide back to methionine and thus repair damaged proteins are
also essential in the repair systems. As there are two stereoisomers
of methionine sulfoxide (R and S), there are accordingly two different types of methionine sulfoxide reductases (MsrA and MsrB).
MsrA and MsrB, as PRXs, generate a sulfenic acid (R-SOH) moiety in their catalytic cycle which is reduced back to a thiol via
either the TRX or the GRX system [30–32]. As explained before,
the TRX and GRX systems of higher plants have the greatest known
diversity in nature and it is certainly a reasonable hypothesis to
propose that TRX and GRX and their regeneration systems are very
much involved in overcoming oxidative stress in plants containing chloroplasts i.e. with an additional (and very intense) site of
ROS generation. Still it is puzzling to observe that cyanobacteria,
which are also able to perform oxygenic photosynthesis lack several
characteristics elucidated in algae and land plants and in particular they do not show this expansion of the TRX and GRX genes
although that if one considers cyanobacterial cells as the equivalent of a chloroplast, the difference is somewhat reduced [18]. This
leads to postulate that either cyanobacteria might have lower oxygen yields and thus generate reduced levels of ROS or that they
possess physical/biochemical characteristics that limit ROS diffusion and damage. There is no easy answer to this hypothesis, as
comparison of the quantum yield of cyanobacteria and green algae
varies depending on the methodology used, and it has been hypothesized that their O2 evolving capacities are not greatly different. But
there might be large differences in the rates of the O2 consuming
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D.D. Gütle et al. / Plant Science 255 (2017) 1–11
Mehler reaction (reduction of molecular oxygen (O2 ) to superoxide
anion (O2 − ) by donation of an electron in photosystem I) leading to
the generation of H2 O2 after conversion by superoxide dismutase
[33]. Cyanobacteria are much more sensitive to hydrogen peroxide than green algae. This physiological difference has been used to
clean lakes polluted by cyanobacterial blooms [34,35]. There is little
doubt that this is due at least partially to their reduced anti-oxidant
capacities. Also to be noted is that cyanobacteria differ notably from
eukaryotes in their PRX content [36].
2.2. Functional modifications of target proteins
A second hypothesis that has been put forward is limited to
a couple enzymes located in the chloroplast. This compartment
hosts both a fructose-1,6-bisphosphatase (FBPase) able to cleave
fructose-1,6-bisphosphate and to generate fructose-6-phosphate
and inorganic phosphate and also a phosphofructokinase (PFK)
which does the reverse reaction at the expense of ATP. It has been
reasoned that if both enzymes functioned simultaneously in this
compartment, it would lead to a futile cycle with useless consumption of ATP. So one hypothesis put forward was that FBPase is active
only in the light phase (through reduction through the FTR system)
while conversely PFK operates only in the dark phase in this compartment [37]. While this argument has some strength it ignores
the fact that there are also FBPase and PFK enzymes in the cytosol of
plants and they are apparently not subject to this type of regulation.
Alternatively, metabolic channelling and the formation of enzyme
super complexes might be another way that circumvent the problem of futile cycles [38]. Another example of a potential futile cycle
could be the sequence F6P->G6P->->->R5P->Ru5P) which is found
in the oxidative pentose pathway and in reverse in the Calvin Benson cycle (Ru5P->->->->F6P->G6P).
There are very large lists of possible molecular targets of the
TRX and GRX systems in various sub-cellular plant compartments
mostly from proteomic data [39–42]. In general, the proteins have
been isolated following a similar procedure involving the replacement of the second cysteine of TRX (or GRX) into a serine, trapping
the targets as disulphides, releasing them with a reductant and
then identifying them by mass spectrometry analysis of peptides. It
has become immediately noticeable that some, if not many, of the
potential targets do not contain two cysteines, sometimes a single
one is found and sometimes none, casting some doubt on the actual
physiological nature of the latter targets. Still reasonable explanations have been provided for single cysteine containing targets
(reviewed in [4]) that can be modified by glutathionylation, nitrosylation and a variety of other ROS-induced modifications. In addition
to the proteomics approach, the best biochemically and structurally
characterized redox-sensitive enzymes have been discovered in
earlier studies using more traditional biochemical approaches.
They essentially function in primary carbon metabolism. Still proteomics has uncovered that TRXs and GRXs extend their regulation
capacities to many other metabolic systems.
3. When and how did redox regulation via dithiol
reduction appear?
oxygenic photosynthetic organisms, being present in land plants
but also in some cyanobacteria [10,43]. NTRCs also occur in nonphotosynthetic tissues such as in plant root plastids, but a homolog
of this enzyme is also present in non-photosynthetic prokaryotes
such as Mycobacterium leprae [11,44]. The occurrence of fusions of
that type is actually quite common in pathogenic bacteria, as shown
for a PRX-GRX fusion in Neisseria meningitidis [45], or even in a red
alga (Gracilaria gracilis) for a GRX-GRX-Msr fusion [46]. Still concerning the reductases, FTR has long been thought to be absolutely
restricted to photosynthetic organisms, but genes with similar
sequences occur in genomes of bacteria and archaea. The protein is
also present in Arabidopsis roots as well as in non-photosynthetic
plant tissues as wheat endosperm [1,47–49]. Its function seems to
be related to methane metabolism [50] in methanogenic archaea.
Interestingly, in some prokaryotic organisms there are fusions
between GRX and the catalytic subunit of FTR, further suggesting
that the enzyme might possess additional unknown functions [8].
3.2. Integration of redox sensitive cysteine residues in target
protein sequences
A good deal of information concerning the ontogenesis of the
redox regulatory systems can be derived from the study of the
target regulatory enzymes. As pointed out earlier, many of the
target enzymes are involved in carbon metabolism and most
prominently in the Benson-Calvin cycle [51,52]. Initially, three
enzymes of the Benson-Calvin cycle (i.e. phosphoribulokinase
(PRK), ribulose 1,5-bisphosphate carboxylase/oxygenase (RubisCO)
and sedoheptulose-1,7-bisphophatase (SBPase)) were thought to
Arabidopsis thaliana
Oryza sativa
dicots
monocots
vascular system
Physcomitrella patens
mosses(loss of GADPH Gap B)
NADP-MDH N-term.
land colonization
Chlamydomonas reinhardtii
Cyanidioschyzon merolae
green algae NADP-MDH C-term.,
red algae
ATP synth. γ subunit,
GADPH Gap B
FBPase, SBPase,
G6PDH
plantae
Synechocystis sp. PCC 6803
cyanobacteria
PRK*, CP12
3.1. Occurrence of TRX reductases
As discussed above, a first event linked to the development
of redox regulation was the diversification of TRX and GRX
genes [18–20]. Obviously, this happened after the transition from
prokaryotes to eukaryotes and, as pointed out before, it may
have evolved to deal with an increase in ROS generation. While
the cytosolic and mitochondrial enzymes are of the prokaryotic
type (short NTR), the chloroplast NTRC are uniquely linked to
Fig. 2. Evolutionary appearance of the redox sensitive cysteine residues in redoxregulatory PRK (phosphoribulokinase), CP12, FBPase (fructose-1,6-bisphosphatase),
SBPase (sedoheptulose-1,7-bisphosphatase), G6PDH (glucose-6-phosphate dehydrogenase), NADP-MDH (NADP-malate dehydrogenase), ATP synthase ␥ subunit and
GADPH (glyceraldehyde 3-phosphate dehydrogenase) in the organisms Synechocystis sp PCC 6803 (cyanobacteria), Cyanidioschyzon merolae (red algae), Chlamydomonas
reinhardtii (green algae), Physcomitrella patens (moss), Oryza sativa and Arabidopsis
thaliana. The GADPH Gap B got lost in losses. *One cysteine residue got lost in red
algae.
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
5
Fig. 3. 3D structural models of redox targeted enzymes part 1. A. Model of the chloroplast ATP synthase from A. thaliana generated by the SWISS-MODEL server. Detail shows
Cys199 and Cys205 residues (in spheres) involved in the regulatory disulfide bridge in CF1 ␥-subunit. B. Crystal structure of the oxidized form of the chloroplast NADP-MDH
(NADP-malate dehydrogenase) from Sorghum bicolor at 2.4 Å resolution (PDB entry 7MDH). The N- and C-terminal extensions are coloured in red and grey, respectively.
The regulatory cysteines are labelled in both parts and the active site is highlighted together with its substrate oxaloacetate. The C- terminal extension blocks the active site by
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D.D. Gütle et al. / Plant Science 255 (2017) 1–11
be unique to this cycle. But genome sequencing has shown that
this was not the case, as representatives of those genes occur in
various microorganisms and especially in bacteria and archaea
[47,53]. In order to evaluate at which point in evolution redox regulatory cysteines were introduced into the target sequences we
have run sequence comparisons that included a cyanobacterium
(Synechocystis sp. PCC 6803), a red alga (Cyanidioschyzon merolae),
a green alga (Chlamydomonas reinhardtii), a moss (Physcomitrella
patens) and two angiosperms (Arabidopsis thaliana and Oryza sativa)
for the following enzymes: NADP-malate dehydrogenase (NADPMDH), PRK, FBPase, SBPase, the ␥-subunit of CF1 -ATPase, G6PDH,
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and CP12
(see Supplementary material Fig. S1 in the online version at DOI: 10.
1016/j.plantsci.2016.11.003). These analyses led to the following
conclusions:
i) Redox regulation has been achieved by single cysteine substitution in different routes (G6PDH, PRK, SBPase), or by insertion
of redox regulatory modules (FBPase, CF1 -ATPase) or even by additions of regulatory extensions either at the N- or C-termini or on
both sides of the sequence (NADP-MDH, GAPDH). Two versions of
the GAPDH sequence exist: an A4 enzyme, which recruits CP12
for its regulation, and an A2 B2 form, possessing a built-in C terminal extension that includes the regulatory cysteines in the B
subunit [54]. CP12 is actually a homolog of the extension with the
advantage as serving as a hub for recruiting super complexes of the
Calvin-Benson cycle. NADP-MDH is a very interesting example of
evolution in progress as the enzyme of C. rheinhardtii lacks the Nterminal regulatory sequence, but already possesses the C-terminal
one (Fig. S1 in the online version at DOI: 10.1016/j.plantsci.2016.
11.003).
ii) The appearance of redox properties on the regulatory
enzymes occurred at different times of evolution. The various
steps at which the regulatory sequences were integrated are
synthesized in Fig. 2. It is readily apparent that the integration
of redox properties did occur at different periods of evolution.
iii) The comparison of FBPase and SBPase is very revealing, showing that although the two sequences are clearly
related, they nevertheless originate from very different ancestors
− ␣-proteobacteria for FBPase and archaea for SBPase [53]. As
explained below, it also shows that there are several solutions possible to a given structural constraint.
iv) There are at least two examples which indicate that evolution relied on trial and error/success experiments. One of those is
RubisCO activase for which there is one sequence which includes
a regulatory disulphide but does not occur in all angiosperms (it is
apparently present in A. thaliana but absent in tobacco and maize)
[55]. A second example is the redox regulation of C. reinhardtii phosphoglycerate kinase which relies on two cysteines absent in land
plant homologs [56]. Analysis of future evolutionary events will
reveal whether these two examples are dead-ends of evolution or
whether they will be later more universally adopted. For GAPDH
B subunit the situation is actually quite complex as it is present in
green algae (Characeae and Ostreococcus) [57,58] but the sequence
has apparently been lost in mosses and somehow maintained in
angiosperms.
4. In response to which structural constraints did redox
regulation happen on molecular targets?
As a general rule, the generation of redox regulatory enzymes
from non-regulatory forms is driven by 3D structural constraints
specific to each enzyme. The goal is to shield the active site in the
inactive form, preventing access of at least one of the substrates.
Reduction of the regulatory disulphides is predicted to induce
structural rearrangements that result in an opening of the active
site, allowing catalysis. A very large panel of target enzymes has
been provided by proteomics studies and biochemistry. Genetics
has also opened the way to describing new functions in plants, such
as those linked to starch degradation or tetrapyrrole biosynthesis
[55,59,60]. Nevertheless, these studies have not yielded structural
models that could be useful to our study. We have thus limited
this section to those enzymes for which there are detailed structures available, or for which we can build credible 3D models using
related available structures (same six enzymes as those used for
phylogeny) (Figs. 3 and 4 ).
4.1. ATP synthase
The ATP synthase that produces ATP using the proton gradient generated by the photosynthetic electron flow in the light, is
a complex assembly of several subunits in which the CF1 extrinsic
catalytic domain is composed of three ␣- and ␤-subunits, the redox
regulatory ␥-subunit, one ␦- and ␧-subunits and b/b’-subunits
(coloured in red, green, blue, yellow, purple and orange, respectively). The CF0 membrane domain is composed of a ring of 14
c-subunits and a bundle of ␣-helices assigned to the ␣-subunit
(coloured in cyan and pale yellow, respectively, only 12 are shown
in our model based on mitochondrial F0 ) (Fig. 3A) [61]. ATP synthase
is mainly regulated by the FTR system [62,63], however, recent
studies postulate a regulation by NTRC especially under low light
conditions [64]. The regulatory site does not occur in cyanobacteria and red algae (Fig. S1 in the online version at DOI: 10.1016/j.
plantsci.2016.11.003), but it is highly conserved in higher plants.
CF0 -CF1 ATP synthase is a wonderful model of a rotating enzyme
converting kinetic energy into a high energy molecule and the rotor
part contains the ␥ subunit. Our model clearly shows that the redox
active site between Cys 199 and Cys 205 is readily accessible for
stromal components (e.g. TRX), being situated in a loop distant from
the adenine nucleotide binding sites situated in the CF1 part of the
molecule. How the reduction of the disulphide influences the catalytic activity of CF0 -CF1 ATP synthase is unknown. Still, one can
rather safely postulate that the formation of the disulphide might
be sufficient to slow down the movement of the rotating part and
thus the rate of catalysis. Conversely, its reduction could relax the
strain and improve catalysis. In connection with this possibility, a
nice artificial example of strain induced by disulphide bridge formation is roGFP where a disulphide engineered between two adjacent
␤ strands of the ␤ barrel constrains the protein and limits its fluorescence. Reducing the protein with GSH and GRX releases that
constraint and this probe can be used to monitor the intra-cellular
GSH status [65].
occupying the position of the substrate oxaloacetate (shown in spheres) from the X-ray crystal structure of the NADP-MDH from Aquaspirillum arcticum (PDB entry 1B8U).The
NADP molecule is coloured in purple. C. Model of chloroplast G6PDH (glucose 6-phosphate dehydrogenase) from A. thaliana obtained by homology modeling from the human
G6PDH (45% identity, PDB entry 1QKI). The G6PDH can be observed as a dimer or a dimer of dimers (separated by a dashed line) depending on specific conditions, the
regulatory cysteines are on position 149 and 157. Zoom of the active site with the G6P (glucose 6-phosphate) and NADP molecules obtained from the X-ray structure of the
G6PDH from Leuconostoc mesenteroides (PDB entry 1E7Y). D. Model of the reduced PRK (phosphoribulokinase) of A. thaliana. The dimer was obtained by homology modeling
of the PRK from Rhodobacter spheroides (24% identity, PDB entry 1A7J). The regulatory cysteine residues are highlighted and the active site of each monomer is shown as
blue surface. As indicated elsewhere, the low conservation of the primary sequence and the presence of the large insertion between the cysteines result in a model with
lower confidence. The overall structures are shown in cartoon representation with ␣-helices and ␤-strands in cylinders and arrows, respectively. The numbering of cysteine
residues was calculated excluding transit sequences.
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
7
Fig. 4. 3D structural models of redox targeted enzymes part 2. A.Crystal structure of tetrameric FBPase (fructose-1,6-bisphosphatase) from the moss P. patens at 3.0 Å
resolution (PDB entry 5IZ1). The redox active cysteine residues are highlighted and the active site is shown in detail with FBP (fructose bisphosphate) and Mg2+ ions. Both
binding sites were obtained from the X-ray structure of E. coli FBPase (PDB entry 2Q8 M). B. Crystal structure of dimeric SBPase (sedoheptulose-1,7-bisphosphatase) from P.
patens at 1.3 Å resolution (PDB entry 5IZ3). The two regulatory cysteines are labelled and the active site is shown in details together with the substrate SBP (sedoheptulose
bisphosphate) and magnesium binding sites which were obtained from the X-ray structure of E. coli FBPase (PDB entry 2Q8 M). SBP binding site is obtained by homology modeling followed by an energy minimization using the YAMBER force field. C. Model of the chloroplast A2 B2 -GAPDH (glyceraldehyde 3-phosphate dehydrogenase) from
8
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
4.2. NADP-malate dehydrogenase
The chloroplast NADP-MDH (Fig. 3B) is an unusual case concerning redox regulation, as it contains two or more redox cysteine
residue pairs per monomer in vascular plants. The NADP-MDH protein sequence harbours N- and C-terminal sequence extensions in
land plants. The enzyme is a head-to-tail-homodimer and the disulphides are located at the interface of the two subunits (N terminal
one) or at the outside corner of each subunit (C-terminal one). The
spacing between the cysteines involved in the disulphides includes
four N-terminal and eleven C-terminal residues. Both disulphides
must be reduced to obtain a fully activated enzyme, but the conformational changes occurring upon reduction are likely different.
The N-terminal part influences the catalytic rate. The C-terminus,
which mimics the substrate oxaloacetate, fills the active site in the
oxidized state, preventing access of oxaloacetate to this part of the
active site, and thus catalysis cannot occur. Both extensions occur
in algal protein sequences, but only the cysteines in the C-terminus
are present and are redox-sensitive and required for enzyme activation [66].
4.3. Glucose 6-phosphate dehydrogenase
We have modelled the structure of chloroplast G6PDH of closely
related sequences (Fig. 3C). This enzyme, which catalyzes the ratelimiting step of the oxidative pentose phosphate pathway, is a
homo tetramer and each subunit contains a regulatory disulphide,
having the cysteines separated by seven amino-acids. Here too,
the disulphides are surface-exposed at the outside corner of each
monomer and presumably easily accessible for reduction. As noted
before, the redox regulation of G6PDH is unique. The enzyme is
in the oxidized state in the dark and inactive in the reduced state
in the light [67–69]. As observed for the other enzymes, the regulatory disulphide is rather remote from the G6P binding site and
molecular reconfiguration is likely to be necessary for modifying
the catalytic properties of the enzyme [67].
4.4. Fructose-1,6-bisphosphatase and
sedoheptulose-1,7-bisphosphatase
FBPase and SBPase are two related enzymes although they
likely originate from different backgrounds: FBPase from ␣proteobacteria, SBPase from archaea [53]. At the subunit level there
is extensive homology between the two enzymes, but FBPase is
organized as a tetramer (Fig. 4A) and SBPase (Fig. 4B) as a dimer.
The tetrameric FBPase is actually composed of two pairs of dimers,
each pair quite similar to one SBPase molecule, so the homology
extends at least partially up to the quaternary structure. The regulatory disulphides of both enzymes are located on the surface of
the protein, away from the active site, and probably influence the
binding of the co-factor Mg2+ . Both enzymes share high similarities in their catalytic sites, their substrates being very similar (SBP
has one additional carbon compared to FBP). The biggest difference
concerning redox regulation is the position of the disulphides in the
monomers. The redox sensitive disulphide of FBPase is located on
the outer edges of the protein and the cysteines are in an insertion
separated by 16 to 19 amino acids (Fig. S1 in the online version
at DOI: 10.1016/j.plantsci.2016.11.003), whereas for SBPase it is
more hidden between the two monomers and less accessible for
the reductant TRX molecule (the spacing between the cysteines is
four residues). In contrast to earlier work carried out with proteins
isolated directly from plants, the availability of recombinant proteins has allowed to show that Trx m can also activate FBPase [70].
The more recent comparison of the redox properties of FBPase and
SBPase from P. patens has confirmed and extended this observation
to SBPase. Nevertheless, Trx f was still found to be the best reductant
and reduction was facilitated for FBPase with the more open redox
regulatory sites (those in SBPase are sandwiched between the two
subunits). This observation together with a tighter redox control of
the SBPase enzyme in vitro leads to the assumption that the location of redox-sensitive residues differentially impacts the quality
and extent of the redox control [53]. In terms of molecular movement following reduction, the comparison of a redox-insensitive
FBPase from pig kidney to the oxidized structure of the chloroplast pea enzyme suggests that the animal FBPase possesses a
pseudo-reduced conformation [71]. Comparing the oxidized and
pseudo-reduced structures has then lead us to propose that in the
pea enzyme, the reduction of the disulphide is likely to induce a
reversal of the orientation of two ␤-strands with the shifting of a
side chain glutamate that leads to better Mg2+ binding [69].
4.5. Phosporibulokinase
There is no current available structure of plant PRK, but we could
model this enzyme from the related structure of its Rhodobacter
sphaeroides homolog (Fig. 3D). As the sequence homology is low,
this model should be considered with caution. In our model, the
protein is a homodimer and the regulatory disulphide, the cysteines
of which are separated by 38 amino acids, is surface-exposed. Note
that these cysteines are also present in the cyanobacterial PRKs, but
the plant sequence has had an insertion of 17 amino acids during
evolution. The bacterial enzyme, on which we have based the model
does not contain the regulatory cysteines and is redox insensitive.
In the model, the sulfur atom of Cys 14 is close to the binding site of
ribulose-5-phosphate. Earlier biochemical studies have proposed
that both regulatory cysteines of PRK were also required for catalysis and close to the nucleotide binding site with a more marked role
for Cys 53 [72,73]. More recent papers have described the redox regulation of PRK in complex and the shape of the molecule via SAXS
[74,75].
4.6. Glyceraldehyde 3-phosphate dehydrogenase
The next enzyme of this series is GAPDH, which is present
in super complexes with PRK under oxidized conditions together
with the regulatory protein CP12, whereas the activation process
requires the TRX proteins. The photosynthetic GADPH occurs in two
different subunits, Gap A and Gap B, which share high sequence
similarity, but with an extension of the C-terminus at the Gap B
subunit that harbours the redox regulatory cysteine residues and
the homology with the CP 12 probably evolved from a fusion of
GADPH A subunit with CP12 protein. So far the Gap B subunit is
restricted to higher plants [54] (Fig. S1 in the online version at
DOI: 10.1016/j.plantsci.2016.11.003). The enzyme appears in two
conformations, either as a homotetramer A4 , without redox sensitivity, or as a heterotetramer with two A and two B subunits
A. thaliana. The tetrameric model was obtained by homology modeling from the photosynthetic GAPDH from Spinacia oleracea (85% identity, PDB entry 2PKQ). Only subunit B
contains the regulatory cysteine residues highlighted. The G3P (glyceraldehyde 3-phosphate) and NADP binding sites, shown in detail, were obtained from the GAPDH X-ray
structure from Geobacillus stearothermophilus (PDB entry 3CM3). D. Crystal structure of the binary complex of photosynthetic A4 -GAPDH with the intrinsically disordered
protein CP12-2 (PDB entry 3QV1). The subunits A of the GAPDH tetramer and CP12-2 are coloured in green and cyan, respectively. The first 58 residues in CP12-2 (79
residues) are missing in the X-ray structure showing a highly disordered N-terminal part. In the folded C-terminal part, Cys73 establishes a disulfide bridge with Cys64 and
probably mimics the regulatory disulfide of A2 B2 -GAPDH. The overall structures are shown in cartoon representation with ␣-helices and ␤-strands in cylinders and arrows,
respectively. The numbering of cysteine residues was calculated excluding transit sequences.
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
(A2 B2 ). The latter is very sensitive to redox regulation through TRX
with an auto-inhibitory process (similar to NADP-MDH) by occupying the active site with the C-terminus extension in the oxidized
state. The A4 GADPH is kept inactive via interaction with CP12 and
PRK and after the reductive activation of the two cysteine pairs of
CP12 the super complex dissociates and releases active A4 GADPH
[76,77]. Our structural model of A2 B2 GADPH (Fig. 4C) was constructed using the A. thaliana sequence and the crystal structure of
Spinacia oleracea as a basis. The two binding sites of the substrate
glyceraldehyde 3-phosphate and NADP were included accordingly
to the structure from Geobacillus stearothermophilus (PDB entry
3CM3). The crystal structure of the photosynthetic A4 homotetramer was solved in complex with two CP12 proteins (Fig. 4D)
[76].
5. Do the regulatory sequences have additional roles in
signalling?
The structural studies indicate that, as a general rule, the redoxsensitive disulphide bonds of the target enzymes are distal from
the active site and present in surface-exposed loops. There is no
sequence similarity whatsoever in these regulatory parts and it
looks as if each of the regulatory enzymes was modified during evolution in response to its own structural constraints. The
FBPase/SBPase story [53] indicates that there are several solutions
to solve a similar 3D constraint. It is likely, that the ubiquitous
surface-exposed position is strategic for making the regulatory
sites accessible to TRX. However, this positioning renders these
sequences potential targets of endoproteases. There are examples
of endoproteases that are able to cleave before or after 16 of the
20 amino acids. We could not find adequate endoproteases in the
literature for C, N, P and Q, with proline having a strongly negative effect when adjacent to one of the permissive amino acids for
peptide hydrolysis.
It is highly puzzling that in cowpea the peptide inceptin was isolated and identified as a fragment of the ␥ subunit of the CF0 -CF1
ATP synthase [78]. Inceptin is an eleven amino-acid disulphidebridged peptide that is released by the plant after attack by insect
herbivores such as Spodoptera frugiferda [79]. Inceptin is found in
the oral secretions of the pest and it induces ethylene production by the plant at concentrations as low as 1 fmol per leaf. In
turn, the plant produces ethylene which triggers the production of
the defence-related hormones jasmonic acid and salicylic acid. It
is thus believed that when the worm starts feeding on the plant,
the release of plant peptides acts as a defence signal. Amazingly
enough, inceptin has the sequence ICDINGVCVDA, which is nearly
exactly the redox regulatory insertion in the ␥ subunit of the chloroplast CF0 -CF1 ATP synthase (see Fig. S1 in the online version at
DOI: 10.1016/j.plantsci.2016.11.003). The surrounding sequence
from which inceptin is generated is GE/ICDVNGVCVDA/AE, and
thus one can imagine that a combination of a glutamyl endopeptidase (that cuts after E) and of an elastase-like peptidase (can cut
after A) is sufficient to generate this peptide. An interesting hypothesis concerning inceptin is that it is probably easily cut out by the
proteases because it is surface-exposed, but also possibly stabilized by its disulphide bond thus improving its potential as a signal
transmitter. Our structural study indicates that these properties
are certainly shared by all redox-regulated enzymes. In particular, the regulatory insertion of FBPase could be easily released by
the same combination of a glutamyl endopeptidase and elastase.
Other simple combinations of various endoproteases or even the
use of a single endoprotease could easily release the regulatory
sequences of SBPase, GAPDH, NADP-MDH, PRK, CP12, G6PDH and
RubisCO activase. Whether those potential fragments are indeed
released and could serve as signalling molecules in plant defence,
9
as demonstrated for inceptin, could be an exciting research area for
the future.
6. Potential new avenues of research in the redox field
A number of problems remain to be elucidated concerning the
regulatory enzymes and their interaction with TRXs, but also about
the multiplicity of reduction systems. We have pointed out that
the area of recognition on the target enzymes is enriched in acidic
residues with resulting negatively charged electrostatic surfaces
(Fig. S2A–H in the online version at DOI: 10.1016/j.plantsci.2016.
11.003).Interestingly, the most efficient TRXs (especially the f type)
have a compatible positively charged surface (Fig. S2I in the online
version at DOI: 10.1016/j.plantsci.2016.11.003). The interaction
would then be governed at least in part by electrostatic interactions.
The active site of TRX is still primarily hydrophobic and these interactions should also play an essential role [80]. A number of recent
papers have dealt with TRX specificity although in general they do
not address the problem at the molecular structural level [81,82]. To
better understand the target enzyme/TRX interaction it is desirable
to generate stable heterodimers by replacing backup cysteines by
serines and thus generating stable mixed disulphides. This technology has been used to generate ternary complexes of FDX-FTR-TRX,
the structure of which could then be determined by X ray crystallography [13]. Those protein-protein interactions are essential for
governing the reduction of enzymes soluble in the stromal phase of
the chloroplasts, a location where the biomolecules could be significantly distant from the reductant TRX. The problem is not as crucial
in membrane-bound electron transfer chains where the positions
of all components are well defined. It has already been demonstrated that FDX recognizes its protein partners (FTR, FNR) via a
combination of charge and hydrophobic recognition [13,83,84].
This could possibly serve as an earlier example of what the TRX
target enzyme interaction could be. Protein-protein interactions
can also be tested using fluorescence methods, isothermal titration
calorimetry and/or surface plasmon resonance to obtain binding
parameters. The reactivity between a given TRX and its target can
also be described/predicted in terms of redox potentials. In general
TRXs possess disulfides with lower redox potentials than those of
the targets making the reaction thermodynamically favourable. An
abundant literature dealing with those aspects has already been
reviewed in [1–6].
We have also pointed out to the need for molecular movements
in the redox-regulated enzymes for reaching their active conformation. At this point, the only report dealing with that aspect is
an NMR study showing modifications of resonances related to the
C-terminus of NADP-MDH after reduction [85]. Given the size of the
proteins studied, it remains to be seen whether this very exciting
approach could be applied to other enzymes or whether this was a
lucky draw (which has nevertheless required a lot of perseverance
in this case). Another possibility would be to try to solve both oxidized and reduced structures of the target enzymes, and possibly
using anaerobiosis for elucidating the reduced forms.
The origins of the regulatory sequences present as insertions
or additions are also of evolutionary interest. Do they result from
slow evolution processes or are these pieces of DNA borrowed
from other sequences? This is very difficult to answer because the
sequence additions/insertions are short and quite variable. Nevertheless, when BLASTing the regulatory sequence of the ␥ subunit of
CF0 -CF1 ATPase, one obtains a hit with 50% identity to a seemingly
unrelated chloroplast protein, the 30S ribosomal protein S13 from
Porphyridium purpureum (see below).
CF1 gamma ATPase 236 DGKCVDAADDEIFKL 250
Conserved residues
D KC D DDE+ K+
DTKCKDLQDDEVVKI 56
30 S rib protein
10
D.D. Gütle et al. / Plant Science 255 (2017) 1–11
This sequence is not totally coincident with the regulatory insertion, but it overlaps it partially containing in particular one of the
cysteines. Of course, given the short length of the sequence, it might
still be coincidental, but if other examples of that type arise, then it
may well be that the genes coding for the redox regulated targets
may also be a composite of sequences borrowed elsewhere in the
genome.
Another area that warrants interest is to understand why redundant TRX reduction systems are present in plant cells (five different
possibilities when including the GSH GRX-dependent systems).
In fact, several mutants of NTR A, B and C and FTR have already
been generated, sometimes in combination but so far no complete
knockout of FTR has been reported.
Acknowledgements
This work was supported by a grant from by the French National
Research Agency (ANR) as part of the “Investissements d’Avenir”
program ANR-11-LABX-0002-01, Lab of Excellence ARBRE, and by
the Excellence Initiative of the German Federal and State Governments (EXC 294).
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