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\begin{document}

\title{Redox regulation in chloroplast thylakoid lumen: The pmf changes
everything, again.}



\author[1]{Donghee Hoh}%
\author[1]{John E. Froehlich}%
\author[1]{David M. Kramer}%
\affil[1]{MSU-DOE Plant Research Laboratory}%


\vspace{-1em}



  
  \date{May 25, 2024}


\begingroup
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\selectlanguage{english}
\begin{abstract}
Photosynthesis is the foundation of all life on Earth, providing oxygen
and energy. However, if not well regulated, it can also generate toxic
reactive oxygen species (ROS), which can cause photodamage. Regulation
of photosynthesis is highly dynamic, responding to both environmental
and metabolic cues, and occurs at many levels, from light capture to
energy storage and metabolic processes. One general mechanism of
regulation involves the reversible oxidation and reduction of protein
thiol groups, which can affect the activity of enzymes and the stability
of proteins. Such redox regulation has been well studied in stromal
enzymes, but more recently evidence has emerged of redox control of
thylakoid lumenal enzymes. This review/hypothesis paper summarizes the
latest research and discusses several open questions and challenges to
achieving effective redox control in the lumen, focusing on the distinct
environments and regulatory components of the thylakoid lumen, including
the need to transport electrons across the thylakoid membrane, the
effects of pH changes in the stromal and lumenal compartments, and the
observed differences in redox states. These constraints suggest that
activated oxygen species are likely to be major regulatory contributors
to lumenal thiol redox regulation, with key components and processes yet
to be discovered.%
\end{abstract}%



\sloppy


PCE Special issue Redox regulation of plant stress and development

\textbf{Redox regulation in chloroplast thylakoid lumen: The
\emph{pmf}changes everything, again.}

Donghee Hoh\textsuperscript{1}, John E. Froehlich\textsuperscript{1,2},
David M. Kramer\textsuperscript{1,2}

\textsuperscript{1}MSU-DOE Plant Research
Laboratory,\textsuperscript{2}Department of Biochemistry and Molecular
Biology, Michigan State University, East Lansing, MI 48824 USA

\textbf{Abstract}

Photosynthesis is the foundation of all life on Earth, providing oxygen
and energy. However, if not well regulated, it can also generate toxic
reactive oxygen species (ROS), which can cause photodamage. Regulation
of photosynthesis is highly dynamic, responding to both environmental
and metabolic cues, and occurs at many levels, from light capture to
energy storage and metabolic processes. One general mechanism of
regulation involves the reversible oxidation and reduction of protein
thiol groups, which can affect the activity of enzymes and the stability
of proteins. Such redox regulation has been well studied in stromal
enzymes, but more recently evidence has emerged of redox control of
thylakoid lumenal enzymes. This review/hypothesis paper summarizes the
latest research and discusses several open questions and challenges to
achieving effective redox control in the lumen, focusing on the distinct
environments and regulatory components of the thylakoid lumen, including
the need to transport electrons across the thylakoid membrane, the
effects of pH changes in the stromal and lumenal compartments, and the
observed differences in redox states. These constraints suggest that
activated oxygen species are likely to be major regulatory contributors
to lumenal thiol redox regulation, with key components and processes yet
to be discovered.

Keywords: Photosynthesis, redox regulation, thiol-disulfide redox
regulation, lumen, photoprotection, non-photochemical quenching (NPQ)

\textbf{Introduction}

Photosynthesis is a biological process that converts light energy into
chemical energy, which is then used to fix CO\textsubscript{2} into
organic compounds. This process is essential for life on Earth, as it
provides the food and energy that all living things need. However, at
the same time, photosynthesis can be a dangerous process if it is not
finely tuned and regulated (Kanazawa et al. 2020; Kramer et al. 2004;
Davis et al. 2017; Foyer 2018; Gururani et al. 2015; Murata et al. 2007;
Aro et al. 1993; Raven 2011). For instance, the under stressful
conditions, photosynthetic process can generate reactive oxygen species
(ROS) which can ultimately damage numerous cellular components (Asada
1996; Nishiyama, Allakhverdiev \& Murata 2006; Nawrocki \emph{et
al.}2021b).

To avoid photodamage, photosynthetic organisms have evolved a
photoprotective mechanism, called non-photochemical quenching (NPQ)
(Demmig-Adams \& Adams 1996; M\selectlanguage{ngerman}üller, Li \& Niyogi 2001; Li \emph{et al.}
2004). NPQ itself is composed of several distinctive components. Each
component is defined by an underlying mechanism and rate of formation
and relaxation (Bellafiore, Barneche, Peltier \& Rochaix 2005). The most
rapid responding form of NPQ, is qE (energy-dependent of quenching)
which is activated by acidification of lumen (the \selectlanguage{greek}Δ\selectlanguage{english}pH component of
proton motive force (\emph{pmf} )) via protonation of photosystem II
subunit S (PsbS) (Li\emph{et al.} 2004; Niyogi, Li, Rosenberg \& Jung
2005) as well as activation of violaxanthin de-epoxidase (VDE), which
catalyzes the conversion of violaxanthin (Vx) to antheraxanthin and then
zeaxanthin (Zx) (Niyogi, Grossman \& Bj\selectlanguage{ngerman}örkman 1998). There are
additionally, more slowly responding forms of NPQ, including sustained
quenching (qH) (Malnoë\emph{et al.} 2018), photoinhibition (qI) which is
related to the photodamage of PSII centers and subsequent repair
(Andersson \& Aro 2004; Murata, Takahashi, Nishiyama \& Allakhverdiev
2007; Nawrocki, Liu, Raber, Hu \& De Vitry 2021a), zeaxanthin-dependent
quenching (qZ) which involves the accumulation of Zx but not the PsbS
(Demmig-Adams \& Adams 1996; Müller \emph{et al.} 2001; Li \emph{et al.}
2004; Nilkens\emph{et al.} 2010), and finally, qT, which involves
antenna state transitions (where LHCII migrates to PSI) (Quick \& Stitt
1989). qZ, qH, qI and qT are likely to be too slow to respond to rapid
fluctuations in light but likely act as backup processes when qE fails.

It is important to note that the processes listed above are involved in
balancing the tradeoffs between photoprotection and photosynthetic
efficiency (Kramer\emph{et al.} 2004; Zhu, Long \& Ort 2010; Kromdijk
\emph{et al.} 2016; Davis \emph{et al.} 2017; Kanazawa \emph{et al.}
2020). For instance, slow onset of photoprotection causes photodamage
when light intensity rapidly increases (Kramer\emph{et al.} 2004; Davis
\emph{et al.} 2017; Kanazawa \emph{et al.}2020), whereas slow recovery
leads to losses of photosynthetic efficiency when light intensity
suddenly decreases (Kramer\emph{et al.} 2004; Zhu \emph{et al.} 2010;
Kromdijk \emph{et al.} 2016; Davis \emph{et al.} 2017; Kanazawa \emph{et
al.} 2020). Understanding such ``fine-tuning'' mechanisms that are
involved in maintaining energy balance when plants are subjected to
constantly changing environmental conditions will be required if one
wants to achieve more robust and resilient photosynthesis, and thus,
improve overall crop productivity.

So how do plants adjust to their constantly changing environmental
conditions such as light availability and quality? One approach involves
the regulation and/or fine-tuning of protein function using a
thiol-disulfide redox mechanism (Buchanan \& Balmer 2005; Waszczak
\emph{et al.} 2015). In this approach, we can exploit the fact that,
under physiological conditions, pairs of the amino acid cysteine (Cys),
exist in two forms: when reduced, the Cys residues will be in their
(thiol, SH) forms, whereas when oxidized, may form a disulfide (S-S)
form, linked by a covalent bond (Cremers \& Jakob 2013). In the
chloroplast, thiol-disulfide reactions are involved in numerous
functions such as, in protein folding, in regulating the activity of
countless enzymes, and in ROS detoxification (Buchanan and Balmer 2005;
Kieselbach 2013; Balsera and Buchanan 2019; Yoshida and Hisabori 2016;
Montrichard et al. 2009). These processes are mediated by a complex
network of redox-sensing and redox-regulated enzymes. The reductive and
oxidative activities of this intricate system are essential for
achieving regulatory redox balance.

While many questions still remain, this general scheme for stromal redox
control appears to explain much of the known data on regulation of
stromal enzymes as illustrated in Figure 1. In the light, electron flow
from photochemistry reduces regulatory thiols through the
ferredoxin/thioredoxin and NADPH-dependent thioredoxin reductase C
(NTRC) systems in the stroma. This activation or deactivation of key
enzymes is mediated by the redox state of the thiol groups. Meanwhile,
H\textsubscript{2}O\textsubscript{2} reoxidizes the regulatory thiol
pools both in the light and dark (Cejudo, Ojeda, Delgado-Requerey,
González \& Pérez-Ruiz 2019).{]}

Recently, it has been recognized that reactions at the thylakoid
membrane and within the lumen are also redox-regulated. However, the
roles and mechanisms of these reactions are not well understood.

Several key proteins involved in photoprotection (e.g., violaxanthin
de-epoxidase (VDE)), partitioning of \emph{pmf} into \selectlanguage{greek}Δψ \selectlanguage{english}and \selectlanguage{greek}Δ\selectlanguage{english}pH (the
H\textsuperscript{+}/K\textsuperscript{+} antiporter KEA3), PSII
stability and repair (Deg1 and PsbO), state transition (STN7) have
redox-active thiol groups that are exposed to the thylakoid lumen
(Str\selectlanguage{ngerman}öher \& Dietz 2008; Hall \emph{et al.} 2010; Kieselbach 2013; Yu,
Lu, Du, Peng \& Wang 2014; Simionato \emph{et al.} 2015; Hallin, Guo \&
Åkerlund 2015; Wang \emph{et al.} 2017a; Wu \emph{et al.} 2021). These
thiol groups can undergo redox transitions, meaning that they can be
oxidized or reduced. Interestingly, while the reduction of regulatory
thiols in the stroma tends to activate enzymes involved in
photosynthesis, the opposite seems to be true for lumenal proteins
(e.g., VDE) (Yu\emph{et al.} 2014; Simionato \emph{et al.} 2015; Hallin
\emph{et al.}2015). This suggests that oxidative reactions may be
critical for adjusting the activities of lumenal proteins and preventing
the buildup of damaged PSII centers.

This review will discuss the possible important role(s) of redox
regulation in photoprotection in preventing plants from photodamage,
focusing on redox-regulated proteins in the thylakoid lumen, which is a
unique environment with different properties than the stroma. Also, we
will discuss the challenges of achieving redox regulation in the lumen
and point out that it involves a distinct mode of regulation that links
redox changes, reactive oxygen species generation, and stress responses.

\textbf{2. Redox regulation in the stroma}

In chloroplasts, thiol-disulfide redox regulation is essential for both
the stroma and lumen, but redox regulation in the stroma has been more
extensively investigated than in the lumen (Buchanan, Kalberer \& Arnon
1967; Wolosiuk \& Buchanan 1977; Meyer, Belin, Delorme-Hinoux, Reichheld
\& Riondet 2012; Buchanan, Holmgren, Jacquot \& Scheibe 2012; Balsera,
Uberegui, Schürmann \& Buchanan 2014; Buchanan 2016a). However, in order
to fully understand lumenal redox regulation, we need to have a good
understanding of the stroma side as well. This is because the stroma and
lumen are interconnected, and the redox status of the stroma can affect
the redox status of the lumen.

During oxygenic photosynthesis, electrons are transported from
photosystem I (PSI) to ferredoxin (Fd) in the stroma. Fd carries
electrons between donor and acceptor pairs. In the stroma, the electrons
from Fd along with ATP and NADPH derived from electron and proton
transport reactions respectively are utilized to fix carbon dioxide
through a series of enzymes that constitute the Calvin-Benson-Bassham
(CBB) cycle. Redox regulation in the stroma is also essential for
modulating the activity of various enzymes of the CBB as well as
controlling numerous other metabolic pathways. Hence, the redox
regulation system has been long considered an important means to
synchronize photosynthetic reactions to respond to both light
availability and quality (Pearcy, Krall \& Sassenrath-Cole 2004; Kaiser
\emph{et al.} 2015). Indeed, several enzymes are known to be regulated
by light-dependent reductive activation, for example, the gamma subunit
of chloroplast ATP synthase (CF1-\selectlanguage{greek}γ\selectlanguage{english}) and several key enzymes in the CBB
cycle such as fructose-1,6 bisphosphatase (FBPase), sedoheptulose -1,7
bisphosphatase (SBPase), Phosphoribulokinase (PRK),
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and rubisco activase
(Rca) (Michelet\emph{et al.} 2013).

The following subsections will briefly discuss two current stroma
reduction systems as well as a recently discovered new oxidation system.
For more detailed information on these systems, the reader is encouraged
to the following relevant papers (Nikkanen \& Rintam\selectlanguage{ngerman}äki 2014; Yoshida,
Hara \& Hisabori 2015; Cejudo \emph{et al.}2019; Cejudo, González \&
Pérez-Ruiz 2021; Gurrieri, Fermani, Zaffagnini, Sparla \& Trost 2021)

\textbf{2.1 The stroma has two distinct thiol reduction systems (FTR-Trx
and FNR-NTRC)}

The ferredoxin-thioredoxin reductase (FTR)-thioredoxin (Trx) system is
only active in light, as it requires electrons from photosynthesis. The
system works by transferring electrons from photosystem I (PSI) to
ferredoxin (Fd), which then transfers these electrons to
ferredoxin-thioredoxin reductase (FTR), which then transfers the
electrons to thioredoxin (Trx), and then finally transfers these
electrons to various target enzymes, such as FBPase and SBPase (Figure
1A) (Schürmann and Buchanan 2008; Michelet et al. 2013).

Trx contains cysteine residues that are redox-active and can reversibly
transfer the reducing potentials from light reactions to thiol-regulated
enzymes. Initially, two forms of Trx, -f and -m, were proposed to be
involved in this redox process (Buchanan 1980) but later, more than 20
additional isoforms of Trx were found by sequencing of the
\emph{Arabidopsis thaliana} genome. The newly identified chloroplast
Trxs were categorized into five classes, Trx-f, m, x, y and z
(Collin\emph{et al.} 2003; Yoshida, Matsuoka, Hara, Konno \& Hisabori
2014; Geigenberger, Thormählen, Daloso \& Fernie 2017). Essentially, all
Trxs have the conserved sequence motif (WCGPC) which allows them to
interact more specifically with subsets of target enzymes (Schürmann \&
Jacquot 2000; Collin \emph{et al.} 2003; Yoshida \emph{et al.} 2014;
Geigenberger \emph{et al.} 2017).

A second reduction system, NADPH-dependent thioredoxin reductase C
(NTRC), was discovered relatively recently as a chloroplast
thiol-regulatory system specifically in oxygenic organisms (Serrato,
Pérez-Ruiz, Spínola \& Cejudo 2004; Pulido \emph{et al.} 2010; Carrillo,
Froehlich, Cruz, Savage \& Kramer 2016) (Fig.1B). In contrast to the FTR
system, NTRC uses NADPH as a reducing power to deliver electrons to
target enzymes. NTRC appears to be critical under lower light
conditions, operating with even low LEF, or in the dark using NADPH
generated from the oxidative pentose phosphate pathway (OPPP) (Neuhaus
\& Emes 2000).

\textbf{2.2 The stroma has multiple sulfhydryl oxidation systems}

The majority of research on redox regulation in chloroplasts has focused
on light-dependent reduction of regulatory thiols in the light. More
recently, attention has shifted to the mechanism of ``thiol oxidation'',
the ``reversal'' of light-induced reduction important for determining
the steady-state redox poise of regulatory thiols.

The thiol-modulated enzymes in the stroma have redox potentials
relatively close to those of the major redox carrier, NADPH (see also
below), so that the reaction should be thermodynamically reversible
(Kramer et al. 1990). This means that a net reduction or oxidation of
NADPH, should result in concomitant changes in the thiol regulatory
status. For instance, relative changes in the rates of the light and
assimilatory reactions should alter redox balance, in turn affecting the
activation of a range of thiol-modulated enzymes (Kramer et al. 1990;
Ort et al. 1990; Stitt 2004). The extent to which such modulation
occurs, however, will depend on the relative redox potentials of the
target enzymes. Some of these (e.g. FBPase) have relatively negative
redox potentials and should be more rapidly reversed, whereas others,
notably the chloroplast ATP synthase, have less negative potentials, and
will only be reoxidized after extensive oxidation of NADPH.

This simple, redox quasi-equilibrium model has, in recent years, been
challenged by the identification of redox proteins that specifically
oxidize specific sets of regulatory thiols. For example, the newly
discovered 2-Cys peroxiredoxin (2CP) is involved in oxidizing
reductively activated proteins in the dark (Fig. 1B)~(Yoshida, Hara,
Sugiura, Fukaya \& Hisabori 2018; Yoshida, Yokochi \& Hisabori 2019;
Vaseghi \emph{et al.} 2018; Ojeda, Pérez-Ruiz \& Cejudo 2018; Yokochi,
Fukushi, Wakabayashi, Yoshida \& Hisabori 2021). The proposed mediators
for thiol oxidation of target proteins are Trx-f, Trx-like2 (TrxL2),
atypical Cys His-rich Trx (ACHT), which have a less negative redox
potential (more oxidizing) than the thiols on typical regulatory
proteins (Eliyahu, Rog, Inbal \& Danon 2015; Yoshida \emph{et al.} 2018,
2019; Vaseghi\emph{et al.} 2018; Ojeda \emph{et al.} 2018; Yokochi
\emph{et al.}2019, 2021). Those mediators transfer reducing power from
redox-regulated proteins to 2CP. Reduced 2CP then reduces
H\textsubscript{2}O\textsubscript{2} to H\textsubscript{2}O. The
protein-oxidizing activity of mediators such as TrxL2 is strongly
dependent on 2CP and
H\textsubscript{2}O\textsubscript{2}(Yoshida\emph{et al.} 2018, 2019;
Vaseghi \emph{et al.} 2018; Ojeda \emph{et al.} 2018; Yokochi \emph{et
al.} 2021).

Finally, other forms of peroxiredoxins, such as Prx IIE and Prx Q, did
not show a similar role to 2CP in the dark transition. However, there
are contradictory results for PrxQ (Yoshida\emph{et al.} 2018, 2019;
Vaseghi \emph{et al.} 2018; Ojeda \emph{et al.} 2018; Telman, Liebthal
\& Dietz 2020; Yokochi \emph{et al.} 2021). It was also found that
NTRC-modulated 2CP contributes to the control of chloroplast redox
homeostasis (Pérez-Ruiz, Naranjo, Ojeda, Guinea \& Cejudo 2017). Several
thiol-regulated enzymes were revealed from these studies to be oxidized
by 2CP, including CF1-\selectlanguage{greek}γ, \selectlanguage{english}FBPase, SBPase, Rca, and NADPH-malate
dehydrogenase (MDH) (Yoshida\emph{et al.} 2018, 2019; Vaseghi \emph{et
al.} 2018; Ojeda \emph{et al.} 2018; Yokochi \emph{et al.} 2021).

\textbf{3. Connecting thiol regulation across the thylakoid membrane}

Recent work has shown that the stromal redox pool communicates with
redox-regulated proteins in the lumen, and there is strong evidence that
several lumenal enzymes or proteins are modulated by thiol redox state
(Str\selectlanguage{ngerman}öher \& Dietz 2008; Hall \emph{et al.} 2010; Kieselbach 2013; Yu,
Lu, Du, Peng \& Wang 2014; Simionato \emph{et al.} 2015; Hallin, Guo \&
Åkerlund 2015; Wang \emph{et al.} 2017a; Wu \emph{et al.} 2021). The
special properties of the chloroplast compartments impose distinct
requirements for the stromal and lumenal compartments. As with the
stromal system, three functionalities are required to operate an
effective redox regulatory network: 1) controlled injection of reducing
equivalents (electrons, hydrogen); 2) balanced removal of electrons; and
3) adjustment of redox potentials and kinetics to match the specific
regulatory requirements.

Upon illumination, the electron and proton transport reaction deposits
protons into the lumen, generating a proton motive force (\emph{pmf} ),
which is composed of two components, the electric field (\selectlanguage{greek}Δψ\selectlanguage{english}) and the
proton gradient ([?]pH). Both \emph{pmf} components drive the synthesis of
ATP at the chloroplast ATP synthase (Kramer et al. 2003). The initial
form of \emph{pmf} is exclusively stored as the electric field component
due to the lower capacity of electric capacitance and higher buffering
capacity of [?]pH in the lumen (Kanazawa \& Kramer 2002; Cruz \emph{et
al.} 2005; Takizawa, Kanazawa \& Kramer 2008). As counterions move into
the lumen, the proton gradient builds up, resulting in acidification of
the lumen to values as low as about 5.5 (Kramer et al. 1999; Takizawa et
al. 2007). By contrast, stromal pH tends to increase in the light to
about 7.5-8.0 (Werdan and Heldt 1972; Heldt et al. 1973; Werdan et al.
1975; Reardon-Robinson 1985; Wu and Berkowitz 1992). In addition, lumen
acidification is known to have crucial photoprotective regulatory roles,
such as inducing PSII photoprotection mechanisms (section 4) and
``photosynthetic control'' of electron flow by slowing down the electron
transfer rate at the Cyt\emph{b\textsubscript{6}f} complex, thus
preventing PSI photodamage.

However, each of these functions is strongly impacted by the specific
properties of the lumen and its role in photosynthetic energy
transduction, requiring the lumenal system to be distinct in several
ways, as described in the following.

\emph{The need for transthylakoid redox exchange} . To fulfill its role
in chemiosmotic energy transduction, the thylakoid lumen must be an
electrochemically sealed compartment, to prevent the leakage of energy
stored in the \emph{pmf} . There must then be machinery to allow for
transmembrane thiol-disulfide exchange. While such systems have been
extensively studied in bacteria, how this occurs in chloroplasts is only
beginning to emerge. For example, in the well-characterized oxidative
protein folding disulfide bond isomerization pathway in Gram-negative
bacteria, the Dsb family (Ito \& Inaba 2008; Reardon-Robinson \&
Ton-That 2015) protein has been imported into the periplasm by the Sec
transport apparatus, is facilitated by protein oxidation by DsbA
(Grauschopf\emph{et al.} 1995). Subsequently, the reduced DsbA is
oxidized by DsbB in the inner membrane (IM), which transfers reducing
equivalents to quinones (Bader, Muse, Ballou, Gassner \& Bardwell 1999).
However, when a newly imported protein is subjected to oxidation, this
can lead to misfolding and inactivation. To prevent or repair these
situations, Trx in the cytoplasm transfers reducing power to DsbD in the
inner membrane which then transfers this reducing power to DsbC, which
maintains proteins in reduced (active or foldable) states (Krupp, Chan
\& Missiakas 2001; Herrmann, Kauff \& Neuhaus 2009) (Fig 2A).

An analogous system likely operates in thylakoid membranes, but may also
function in reversible thiol regulation of enzymes. The protein
components needed to establish a functional thiol-disulfide exchange
system across the thylakoid membrane, while not complete, are slowly
being identified, as discussed in the following sections. Unlike the
thiol-disulfide redox regulation in the stroma, stromal soluble electron
carriers, such as Trx or Trx-like, proteins have not yet been identified
in the lumen (Buchanan 2016b). However, several thiol-modulated enzymes
or proteins have been identified (Str\selectlanguage{ngerman}öher \& Dietz 2008; Hall \emph{et
al.} 2010; Kieselbach 2013; Yu, Lu, Du, Peng \& Wang 2014; Simionato
\emph{et al.} 2015; Hallin, Guo \& Åkerlund 2015; Wang \emph{et al.}
2017a; Wu \emph{et al.} 2021) and multiple H-carriers (redox
transporters) have been reported to transfer reducing equivalents from
the stroma across the thylakoid membrane to the thylakoid lumen
(Motohashi \& Hisabori 2006, 2010; Brooks, Jansson \& Niyogi 2014;
Motohashi \& Hisabori 2006, 2010; Karamoko, Cline, Redding, Ruiz \&
Hamel 2011; Brooks \emph{et al.} 2014).

\textbf{3.1 HCF164, CcdA and SOQ1 transfer reducing equivalent from
stroma to lumen}

HCF164 was first identified by Meurer, Meierhoff \& Westhoff as a gene
that when mutated in plants resulted in high chlorophyll fluorescence
phenotype (Meurer\emph{et al.} 1996; Meurer, Plücken, Kowallik \&
Westhoff 1998). This phenotype was later found to be caused by a defect
in the assembly of cytochrome
\emph{b\textsubscript{6}f}(Lennartz\emph{et al.} 2001). HCF164 is
anchored in the thylakoid membrane via a single transmembrane domain
(TMD) with the bulk of HCF164 orientated towards the thylakoid lumen
(See Fig. 2B). HCF164 possesses a Trx-like domain, localized to the
thylakoid lumen with disulfide reductase activity (Motohashi \& Hisabori
2006, 2010). Further, HCF164 can interact with potential target proteins
such as PSI-N, Cytf, and Rieske FeS (a subunit of the Cyt
\emph{b\textsubscript{6}f} ) through Trx affinity chromatography
experiments (Motohashi \& Hisabori 2006, 2010; Brooks \emph{et al.}
2014). Intriguingly, it has also shown that Trx-m type is an electron
donor for HCF164 (Motohashi \& Hisabori 2006, 2010; Brooks \emph{et al.}
2014). However, because HCF164 has its Cys residues on the lumenal side,
and not on the transmembrane side, it was proposed that chloroplasts
must possess a system for transferring redox equivalents across the
thylakoid membrane from the stroma to the lumen. Motohashi \& Hisabori
found evidence that CcdA serves precisely this function: a thylakoid
membrane protein that acts to transfer reducing equivalents from the
stroma to the thylakoid lumen. CcdA is a homolog of the prokaryotic
thiol-disulfide transporter, and it was previously reported to be
required for the assembly of the Cyt\emph{b\textsubscript{6}f}complex
(Page\emph{et al.} 2004). Motohashi and Hisabori (2010) further showed
that CcdA's redox state is modulated in the thylakoids by stromal m-type
thioredoxins.

The suppressor of quenching (SOQ1) was identified during the process of
a suppressor screening of non-photochemical quenching (NPQ) (Brooks,
Sylak-Glassman, Fleming \& Niyogi 2013). SOQ1 is anchored in the
thylakoid membrane via a single transmembrane domain (TMD) with the bulk
of SOQ1 localized within the thylakoid lumen (Fig. 2B). SOQ1 appears to
transfer reducing equivalents from the stroma to the plastid lipocalin
(LCNP), thereby suppressing the formation of a sustainable form of NPQ
called qH (Malnoë\emph{et al.} 2018) (see Section 4.3 for more details).
The lumenal thioredoxin-like domain and a \selectlanguage{greek}β-\selectlanguage{english}propeller NCL-1, HT2A, and
LIN-41 (NHL) domain of SOQ1 have been shown to be required for the
transfer of reducing equivalents from the stroma to the lumen (Slack \&
Ruvkun 1998; Brooks \emph{et al.} 2013). Finally, another domain of
SOQ1, the stromal-located halo-acid dehalogenase-like hydrolase (HAD)
domain with a transmembrane helix, has been shown to be not involved in
the suppression of qH (Brooks\emph{et al.} 2013). Interestingly, recent
work from Yu et al (Yu\emph{et al.} 2022) showed that the independent
\selectlanguage{greek}β-\selectlanguage{english}stranded C-terminal domain (CTD), which has structural homology to the
N-terminal domain of DsbD, is essential for the regulation of qH,
suggesting that it is involved in transferring redox equivalents from
the stroma side to the lumen (Yu\emph{et al.} 2022). The electron donor
to SOQ1 has not been specified yet, but it is possible that it is either
CcdA or another unknown mediator.

Taken together, these examples suggest a mechanism for redox regulation
of lumen proteins as illustrated in Fig. 2B in which disulfide-thiol
redox control across the thylakoid membrane, in which reducing
equivalents are transferred from the stroma (e.g., from Trx-m) to CcdA
(or other, yet to be identified) carriers, and then to HCF164, which
then delivers them to target proteins.

\textbf{3.2 The acidification of the lumen by \emph{pmf} alters the
redox potentials of regulatory thiols, altering the mechanism of redox
balancing}

At physiological pH (less than about 8.0), the two-electron reduction of
protein disulfide occurs with the uptake of 2 H\textsuperscript{+}, as
in

-S-S- + 2e\textsuperscript{-} + 2H\textsuperscript{+} =
(-SH)\textsubscript{2}

\textsubscript{}

The involvement of one proton per electron in the redox reaction implies
that the redox midpoint potentials for the regulatory thiols will shift
by -0.06 eV / pH unit, i.e., the lower the pH, the less reducing (more
oxidizing) the regulatory thiols will become. The redox potentials of
the stromal and lumenal thiol couples will change in opposite
directions. Interestingly, reducing the stromal NADP+/NADPH couple,
involves only one proton per two electrons, so its redox potential will
only shift by about -30 mV/pH unit.

Figure 4 illustrates what would happen if the lumen and stromal pH
values started at about 7.0 in the dark and changed to 6.0 and 8.0 in
the light. The midpoint potential of thiol couples in the stroma should
become more negative relative to dark (more difficult to reduce) and in
some cases (e.g., NADPH-MDH) will become more negative than the
NADP+/NADPH couple. In this case, reversal of the electron transfer
(oxidizing the thiols and reducing NADP+) should result in net oxidation
of the regulatory thiols, as discussed above, which can account for at
least some of the redox modulation (Kramer et al. 1990).

By contrast, the acidification of the lumen should shift the lumenal
thiol couples to less reducing (more oxidizing) redox potentials, making
the transfer of electrons from stromal carriers (like NADPH or Trx)
strongly favorable, and inhibiting oxidation by reversal processes.
Extrapolating from published values, we estimate that VDE and HCF164
will have redox midpoint potentials of -202 and -164 mV at pH = 6.0,
reducing in a strongly favorable transfer of electrons from stromal
NADPH or Trx, effectively making the transfer of electrons irreversible
and preventing stromal carriers from acting as oxidants for lumenal
thiol couples.

In the stroma, light-induced electron transport typically results in a
net reduction of thiol modulation enzymes that activate assimilation
(e.g., FBPase) and inactivate catabolism (e.g., glucose-6-phosphate
dehydrogenase). In contrast to stromal thiol-modulated enzymes, which
are activated by becoming more reduced, lumenal thiol-modulated enzymes
are often activated by becoming more oxidized (compare Figure 3A to 3C,
see section 4). For example, VDE is inactivated by the artificial
treatment of dithiothreitol, a strong reducing agent (Bilger and
Bj\selectlanguage{ngerman}örkman 1990). This occurs despite the fact that acidification of the
lumen should make thiol reduction more favorable, strongly implying that
the kinetics of oxidative processes, rather than thermodynamic
constraints, control the overall redox poise.

The question then is, how do electrons get out of the lumen in a
chloroplast system? The above arguments imply that alternative,
lumen-associated components are absolutely required for redox balancing
and that their redox potentials must be more oxidizing than the least
negative thiol component, i.e. midpoint potential at pH = 6 higher than
about -160 mV. Molecular oxygen was proposed as an oxidant in the lumen
(Buchanan and Luan 2005; Gopalan et al. 2004) (Figure 3B). However, this
is unlikely, as the oxygen concentration does not change significantly
in the lumen and O\textsubscript{2} is a relatively stable form. Obvious
candidates include the plastoquinone (PQ) pool and some form of
activated oxygen species, e.g. ROS species like
H\textsubscript{2}O\textsubscript{2}, as seen in 2CP-mediated oxidation
in the stroma.

In the Gram-positive bacterial system, thiol oxidation performed by DsbA
(Fig. 2A) and a homologous proteins, 2CP, was proposed to oxidant in the
stroma (Yoshida, Hara, Sugiura, Fukaya \& Hisabori 2018; Yoshida,
Yokochi \& Hisabori 2019; Vaseghi \emph{et al.} 2018; Ojeda, Pérez-Ruiz
\& Cejudo 2018; Yokochi, Fukushi, Wakabayashi, Yoshida \& Hisabori
2021). However, 2CP is unlikely to have direct access to lumenal thiols.

LTO1 was proposed as a lumen-localized thiol oxidase by Karamoko et al
(2011). LTO1 is an oxidoreductase that belongs to a distinct class of
disulfide bond-forming enzymes in bacteria. It has two domains: a
lumenal thioredoxin-like domain, which is functionally similar to DsbA,
and functions to oxidize (forms disulfide bonds) proteins and a
transmembrane domain, that has homology to the mammalian vitamin K
epoxide reductase (VKOR) catalytic domain, which is also similar to DsbB
(Karamoko\emph{et al.} 2011; Onda 2013). LTO1 has been shown to oxidize
lumenal proteins such as PsbO, STN7, and VDE (Kieselbach 2013; Lu
\emph{et al.} 2013, 2015; Yu \emph{et al.} 2014; Wu \emph{et al.} 2021)
(see Section 4 for details). However, the final acceptor of LTO1 is
still unknown (see Section 5).

Molecular oxygen was suggested as an oxidant in the lumen (Buchanan and
Luan 2005; Gopalan et al. 2004) (Figure 3B). However, the oxygen
concentration does not change significantly in the lumen and
O\textsubscript{2} is a relatively stable form.

\textbf{4. A model for the role of lumenal redox regulation in
photoprotection}

In the previous sections, we discussed redox regulation in the stroma,
the trans-membrane mediators and the unique redox environments in the
lumen, which are all affected by pH changes. In this section, we will
explore the implications of the thylakoid \emph{pmf} , not only on the
canonical NPQ mechanisms, but on interactions with lumenal redox
regulation, as summarized in Fig. 5.

There are several forms of NPQ, defined by their rates of activation and
deactivation (Pinnola \& Bassi 2018). These forms include the most rapid
form, energy-dependent quenching (qE) (Li\emph{et al.} 2004; Niyogi
\emph{et al.} 2005; Niyogi\emph{et al.} 1998, 2005; Li \emph{et al.}
2004), state-transition quenching (qT) (Quick \& Stitt 1989),
Z-dependent quenching (qZ) (Demmig-Adams \& Adams 1996; Müller \emph{et
al.} 2001; Li \emph{et al.} 2004; Nilkens\emph{et al.} 2010), sustained
quenching (qH) (Malnoë\emph{et al.} 2018), and photoinhibitory quenching
(qI) (Andersson \& Aro 2004; Murata \emph{et al.} 2007; Nawrocki
\emph{et al.} 2021a). In the following subsections, we will discuss the
essential role of redox regulation on each NPQ form.

\textbf{4.1 Energy-dependent quenching, qE}

Energy-dependent quenching (qE) is the most rapidly-responding form of
NPQ. It is triggered by the acidification of the lumen (proton gradient
(\selectlanguage{english}[?]pH) component of proton motive force (\emph{pmf} ) ) which results in
the protonation of PsbS as well as the activation of violaxanthin
de-epoxidase (VDE), which catalyzes the conversion of violaxanthin (Vx)
to zeaxanthin (Zx). Zx subsequently dissipates excessive light energy
and induces qE along with PsbS (Niyogi\emph{et al.} 1998).

Recent studies have shown that both the proton gradient (\selectlanguage{english}[?]pH) and also
the redox status of the thiol-disulfides within in VDE play a role in
controlling its function (Arnoux, Morosinotto, Saga, Bassi \& Pignol
2009; Yu \emph{et al.} 2014; Hallin\emph{et al.} 2015; Arnoux\emph{et
al.} 2009; Simionato \emph{et al.} 2015). Specifically, the proton
gradient (\selectlanguage{english}[?]pH) facilitates a structural change in VDE causing it to
change from a monomer to a homo-dimer complex upon lumen acidification
(Arnoux\emph{et al.} 2009). In addition, to VDE undergoing a structural
conformational change, VDE activity is also regulated in a
thiol-disulfide-dependent manner (Hall\emph{et al.} 2010; Kieselbach
2013; Yu \emph{et al.} 2014; Simionato\emph{et al.} 2015; Hallin
\emph{et al.} 2015). This elaborate regulation is achieved due to the
unique structure of VDE. The VDE protein consists of three domains: a
Cys-rich N-terminal domain, a lipocalin-like domain (predicted to bind
violaxanthin) (Saga\emph{et al.} 2010) and a glutamate-rich C-terminal
domain. VDE has a total of 13 conserved Cys residues, 12 of which are at
the N-terminal and these Cys residues play a major role in protein
function. Consequently, VDE is active only when fully oxidized, with six
disulfide bonds, is VDE active (Simionato\emph{et al.} 2015), and this
corresponds to a more compact, rigid, thermostable form of the protein
(Hallin\emph{et al.} 2015).

It has been proposed that the electron donor for VDE is Trx-m through
CcdA and HCF164 (Motohashi \& Hisabori 2006, 2010) and the oxidase is
LTO1(Lu\emph{et al.} 2013, 2015; Yu \emph{et al.} 2014; Wu \emph{et al.}
2021) (Fig. 4). Coincidently, an \emph{lto1-2} mutant line showed a
lower de-epoxidation state of xanthophyll cycle pigment de-epoxidase
index (DEI), suggesting that de-epoxidation of Vx to Zx was suppressed
in the\emph{lto1-2} mutant line\emph{(Yu et al. 2014; Lu et al. 2015)}.

Another component that can affect the kinetics of qE is
K\textsuperscript{+} efflux antiporter3 (KEA3), a potassium/proton
antiporter. KEA3 exchanges protons (out of the lumen) and potassium ions
(into the lumen) (Armbruster\emph{et al.} 2014). Consequently, when
light transitions from high to low, the KEA3 antiporter function, helps
relax qE. Furthermore, it has been recently reported that KEA3 has five
Cys residues, but only one Cys residue is located at the N-terminal
which extends into the lumen (Wang\emph{et al.} 2017a). The location of
this Cys residue suggests that KEA3 may be regulated by a redox
mechanism, acting as adimer. However, the possible electron donors and
oxidases involved in regulating KEA3 need further investigation.

\textbf{4.2 Redistribution of excitation energy, qT}

In response to light quality and intensity, the balance of excitation
energy between the two photosystems is dynamically adjusted to avoid
photodamage. This is achieved through serine/threonine-protein kinase
(STN7) (Bellafiore\emph{et al.} 2005). STN7 is a transmembrane protein
having its catalytic domain extending into the stroma. STN7 also has two
conserved cysteines located on the lumen side of the thylakoid
(Lemeille\emph{et al.} 2009; Bergner \emph{et al.} 2015).

The catalytic domain of STN7 is responsible for the phosphorylation of
light-harvesting complex II (LHCII) (Puthiyaveetil 2011).
Phosphorylation of LHCII changes its structure, which causes it to
detach from PSII and migrate to PSI. This process is important for
balancing the amount of light energy that is captured by each
photosystem under dynamic environmental changes (Bellafiore et al.
2005).

The proposed model for STN7 functions is that plastoquinol
(PQH\textsubscript{2}) binds to the Qo site of
Cyt\emph{b\textsubscript{6}f} and activates STN7 by forming
intermolecular disulfide bridges between the conserved cysteines in the
lumenal domain of STN7. However, the dimer formed by these disulfide
bridges is very transitory and can be easily converted back to a monomer
(Lemeille\emph{et al.} 2009; Wunder \emph{et al.} 2013; Bergner \emph{et
al.}2015; Shapiguzov \emph{et al.} 2016). Wu\emph{et al.} (2021) showed
that the lumenal cysteines of STN7 are maintained in the oxidized state
by the Trx-like domain of LTO1, which then transfers reducing power to
the VKOR domain (Wu\emph{et al.} 2021). Under conditions where the PQ
pool is oxidized, e.g. when PSI is preferentially excited by light, STN7
becomes inactivated, LTO1 no longer oxidizes STN7, and the antenna adopt
the state 1 configuration (Fig. 5). However, it is noteworthy that state
transitions are strongly decreased in the \emph{lto1} mutant, but they
are not completely abolished, unlike in the \emph{stn7} mutant
(Wu\emph{et al.} 2021). This result suggests that other factors, in
addition to LTO1, are likely involved in keeping STN7 oxidized and
active in the acidic lumenal environments. In addition, it has shown
that STN7 is inactivated by Trx-m, in coordination with HCF164 and CcdA
(Ancin\emph{et al.} 2019) (Fig. 5). In this scenario, Trx-m breaks the
intermolecular disulfide bridges in STN7, leading to the formation of an
intramolecular disulfide bond, which inactivates STN7 by monomerizing
it. So once again, the function of STN7 is heavily impacted by its redox
regulation However, the exact oxidative mechanism by which STN7
functions in the acidified lumen, where the redox midpoint potentials of
the regulatory thiols increase, still needs further investigation.

\textbf{4.3 Sustained quenching, qH}

qH is a form of sustained and slowly reversible quenching that occurs in
the peripheral antenna of PSII prior to photoinhibitory quenching (qI)
(Malnoe\emph{et al.} 2018). Bru\emph{et al.} showed that the qH
quenching site is a major trimeric antenna complex that consist of
Lhcb1, Lhcb2, Lhcb3, which can form homotrimeric and heterotrimeric
complexes (Bellottari et al 2012, Crepin et al 2018). Interestingly, Bru
et al. also found specific isoforms of the major trimeric antenna
complex that are not required for qH induction (Bru\emph{et al.} 2021).

qH is known to be redox-regulated but independent of protonation of
PsbS, accumulation of Zx, and phosphorylation by STN7 (Brooks\emph{et
al.} 2013; Malnoe \emph{et al.} 2018; Bru \emph{et al.} 2021). It has
been shown that qH is induced by oxidative stress conditions such as
cold and high light (Levesque-Tremblay, Havaux \& Ouellet 2009; Brooks
\emph{et al.} 2013; Malnoe \emph{et al.}2018; Bru \emph{et al.} 2021).

The molecular players of qH are SOQ1 (Brooks\emph{et al.} 2013; Malnoe
\emph{et al.} 2018; Malnoe 2018; Bru \emph{et al.} 2021), plastid
lipocalin (LCNP) (Brooks\emph{et al.} 2013; Malnoe \emph{et al.} 2018;
Bru \emph{et al.} 2021), and relaxation of qH1 (ROQH1) (Amstutz\emph{et
al.} 2020) (Fig. 5). LCNP is a major component that induces qH. SOQ1 is
a trans-membrane protein (see section 3.1) that inhibits LCNP, while
ROQH1 relaxes qH. LCNP is a soluble lumenal protein that induces qH by
binding to LHCII and thus induces a conformational change within LHCII
causing it to undergo a transformation from a light-harvesting state to
a dissipated state (Yu\emph{et al.} 2022). In addition, the redox status
of the six conserved cysteines within LCNP play a critical role in
regulating LCNP's activity (Malnoe\emph{et al.} 2018).

Consequently, the reduced form of LCNP is inactive, while the oxidized
form is active. Mechanistically, LCNP can be proposed to work as
follows: SOQ1, transfers reducing equivalents to LCNP, which inhibits
LCNP under non-stress conditions. However, under stress conditions
(oxidative condition), SOQ1 no longer transfers reducing equivalents to
LCNP, which keeps LCNP in an oxidized state and thus allows it to induce
qH (Malnoe 2018). As a result, the function of LCNP and qH is being
modulated by the redox state of the chloroplast (See Fig. 5).

\textbf{4.4 PSII stability and repair of PSII, qI}

Under stress conditions, such as high light intensity or extreme
temperature, PSII is highly susceptible to photodamage of its core
proteins, specifically the D1 protein. This photodamage can lead to the
irreversible damage of PSII, which requires repair (Andersson \& Aro
2004; Murata \emph{et al.} 2007; Nawrocki \emph{et al.} 2021a). However,
PSII function can be maintained through an intricate mechanism involving
the turnover and replacement of damaged D1 protein using an elaborate
PSII repair cycle (Tikkanen, Mekala \& Aro 2014). Accordingly, when the
rate of damage of D1 exceeds the rate of repair of D1, this unbalance
can ultimately lead to photoinhibition.

Briefly, the PSII repair cycle begins with the monomerization and
detachment of the light-harvesting complex (LHCII) from the PSII core by
phosphorylation through STN8 (STN7) kinase. The monomerized PSII core
then migrates to the unstacked region of the thylakoid membrane, where
it is dephosphorylated by the phosphatase, PBCP. The damaged D1 subunit
is then degraded by the proteases Deg and FtsH. Once the damaged D1
subunit has been degraded, a new D1 subunit is synthesized and inserted
into the PSII core (Kirchhoff 2014). Coincidently, the key proteases,
Deg and FtsH, that are involved in D1 protein turnover are reported to
be redox-regulated during this process.

By dynamic thiol-disulfide redox proteomics, Deg1 and Deg2 were found to
be redox regulated (Stroher \& Dietz 2008). Deg1 is peripherally
attached to the thylakoid lumenal side (Chassin, Kapri-Pardes, Sinvany,
Arad \& Adam 2002) and is subsequently regulated by both the proton
gradient (\selectlanguage{english}[?]pH) and redox state of the thylakoid membrane (Stroher \&
Dietz 2008; Knopf \& Adam 2018). The formation of Deg1 homo-hexamers is
dependent on \selectlanguage{english}[?]pH (Knopf \& Adam 2018). Further, Deg1 showed maximal
activity under reducing conditions and less activity under mild
oxidative stress conditions (Stroher \& Dietz 2008). Although Deg1 has
only one Cys residue in its mature form (Table 1), the fact that it
forms homo-hexamers and changes its activity depending on
oxidizing/reducing condition suggests that is regulated by redox
regulation upon formation of homo-hexamers, however this possible unique
regulation needs to be studied further. Conversely, Deg2 is located in
the stroma and has an opposite redox-dependent mechanism, showing higher
proteolytic activity under oxidizing conditions (Haussuhl, Andersson \&
Adamska 2001). Other interacting factors of Deg protease, such as
electron donors, and oxidizers have not yet been determined.

FtsH is an ATP-dependent zinc metalloprotease that exists as two types
of subunits: A (FtsH1 and FtsH5) and B (FtsH2 and FtsH8) (Zaltsman, Ori
\& Adam 2005; Kato \& Sakamoto 2018). Both types have a similar
structure, with a C-terminal extension orientated towards the stroma and
an N-terminal transmembrane domain (Lindahl\emph{et al.} 1996). The
C-terminal domain has ATP hydrolysis activity and conserved cysteines
(Sauer \& Baker 2011; Wang \emph{et al.} 2017b). FtsH cooperates with
Deg proteases to degrade the D1 protein in the PSII repair cycle. It has
been suggested that FtsH degrades D1 after it has been cleaved by
Deg-mediated proteolysis (Nishimura, Kato \& Sakamoto 2016; Kato \&
Sakamoto 2018). FtsH is active under reducing conditions, and it has
been proposed that the possible electron donors for FtsH are members of
the stromal Trx family (Wang\emph{et al.} 2017b). However, the oxidant
for FtsH remains unknown.

Given that photoinhibition decreases the efficiency of photosynthesis
and redox regulation directly affects protease activity and rate of
repair, further research will be needed to fully understand the role
that redox regulation has on these proteases.

Finally, besides proteases, PsbO, a subunit of the oxygen-evolving
complex (OEC), is also associated with PSII stability and repair. The
redox state of two cysteine residues in PsbO is the key determinant of
its stability (Hall\emph{et al.} 2010; Kieselbach 2013). The oxidized
form of PsbO is stable, while the reduced form is unstable and becomes a
target for proteolysis, leading to increased accessibility of PSII core
proteins for degradation. For example, when D1 in PSII is damaged, PsbO
in its stable oxidized form is less accessible to degradation, which
further inhibits the repair cycle. Therefore, the redox state of PsbO
must be tightly regulated. It has been shown that LTO1 oxidizes PsbO
(Karamoko\emph{et al.} 2011), but the reducing pathway for PsbO remains
to be determined. PsbO was found to be a potential target of Trx
(Lee\emph{et al.} 2004; Marchand, Le Marechal, Meyer \& Decottignies
2006; Hall \emph{et al.} 2010), but given the location of Trx and PsbO
in the stroma and lumen respectively, a mediator such as CcdA and HCF164
or another unknown factor may be involved in the transfer of electrons
to PsbO (Fig. 4).

\textbf{5. \emph{What are the missing pieces in the lumen
thiol-disulfide regulation mechanism?}}

So far, we have discussed the redox regulation from the stroma to the
transmembrane region to the lumen and its photoprotective roles. The
molecular players in the redox regulation and relevant photoprotective
forms are summarized in Figure 5. In brief, reducing equivalents from
the stroma are transferred through CcdA and HCF164 to VDE and STN7.
These proteins are inactive in their reduced forms, as indicated by the
arrows pointing in the opposite direction (-). Another reducing
mediator, SOQ1, transfers reducing power to LCNP, which is inactivated
in its reduced form. The electron donors for KEA3-1, PsbO, and Deg1 are
still unknown, as indicated by the dashed arrows. The oxidizing system
appears to be mediated by LTO1. LTO pulls electrons from VDE, STN7, and
PsbO. The oxidizing mediators for KEA3-1, Deg1, and LCNP are not known.
All of the redox-regulated proteins in the lumen, including VDE (qE,
qZ), KEA3 (\emph{pmf} partitioning into \selectlanguage{english}[?]\selectlanguage{greek}ψ \selectlanguage{english}and [?]pH and effects on qE),
STN7 (qT), LNCP (qH), Deg1, and PsbO (qI), orchestrate photoprotective
mechanisms.

However, as seen in Figure 5, many of the pathways in lumenal redox
regulation are still hypothetical (so many dashed lines!). Certainly, we
have just begun to scratch the surface of this complex lumenal redox
regulation network. While there are many unanswered questions regarding
this complex redox network, there are two significant yet unresolved
questions we would like to focus on:

How do electrons get out of the lumen? Recall that LTO1 may be involved
in numerous oxidizing events but the final acceptor of LTO1 has not been
identified (See Figure 5).

What keeps fine-tuning redox potentials and kinetics to achieve the
appropriate regulatory levels in the ``acidic lumen?'' As we discussed
in section 3.2, due to the [?]pH component of \emph{pmf} , the lumen has
quite a unique redox environment, where the redox midpoint potentials of
the regulatory thiols substantially increase upon illumination (See
Figure 4).

One hypothetical model that could answer both of these questions is that
reactive oxygen species (ROS) act as a strong oxidant. It has been shown
that in the stroma, the 2CP-mediated oxidation system pulls electrons
from target enzymes and transfers them to the final acceptor hydrogen
peroxide (H\textsubscript{2}O\textsubscript{2}), which is then reduced
to water. In the hypothetical model for lumenal redox regulation, the
Trx-domain of LTO1 would pull electrons from target enzymes, and these
electrons would then be transferred to the VKOR domain of LTO1 in the
thylakoid membrane. From there, the electrons could be transferred to an
unknown oxidizing mediator (marked as ''?'' in the figure), which could
further reduce H\textsubscript{2}O\textsubscript{2}. Another possibility
is that there is another oxidoreductase in the thylakoid membrane that
uses H\textsubscript{2}O\textsubscript{2} as a final electron acceptor.
This hypothesis is supported by the results of state transition
experiments in the \emph{lto1} mutant, which showed that state
transitions were not completely abolished in this mutant (Wu et al
2021). This result strongly suggests that other factors are also
involved in keeping STN7 oxidized and active. The other possible strong
oxidants could be plastoquinone (PQ) pool. More research will be needed
to determine the exact mechanisms involved.

\textbf{Concluding remarks}

Redox regulation is a key mechanism that enables plants to adapt to
unpredictable changes in their environment, such as variations in light
intensity or temperature. We have shown that numerous lumenal proteins
involved in photoprotection are subject to regulation by their redox
state, although there are still several unanswered questions regarding
redox regulatory mechanisms in the lumen. It is critically important to
keep in mind that \textbf{the \emph{pmf} changes everything} . For
instance, an acidified lumen makes it more challenging to maintain
oxidizing environments in the lumen, as the redox midpoint potentials of
regulatory thiols change and increase.

In future studies, it will be important to focus on the following
aspects of redox regulation in the chloroplast thylakoid lumen:

\begin{itemize}
\tightlist
\item
  Study the effects of lower pH in the lumen that impacts the redox
  midpoint potentials of proteins.
\item
  Identify the strong thiol-oxidizing system that extracts electrons
  from the thylakoid lumen in response to changes in ROS or redox
  status.
\item
  Determine the final acceptor of the oxidation pathway that leads to
  the oxidation of lumenal proteins.
\item
  Consider the possibility that there are more trans-thylakoid membrane
  thiol mediators that can reduce and oxidize lumenal proteins.
\item
  Investigate the roles and interactions of target enzymes in the redox
  regulation network.
\end{itemize}

By considering these recommended focal points for future investigations,
we should attain a better understanding of the complex mechanisms of
redox regulation and photoprotection in the chloroplast. This new
knowledge will contribute to our understanding of how plants survive and
adapt to ever changing environments and it may also provide us with new
insights as to how we can potentially improve photosynthesis.

\textbf{Acknowledgment}

The authors thank Dr Atsuko Kanazawa, Dr Liana Acevedo-Siaca, Shannon,
Donnelly and Dr Roderick Van Daniel for their insightful discussions and
critical feedback on the manuscript. This work was supported by the
Chemical Sciences, Geoscience and Biosciences Division, Office of Basic
Energy Sciences, Office of Science, U.S. Department of Energy (award
number DE-FG02-91ER20021 and DE-SC0007101).

\textbf{Conflict of interest statement}

The authors declare no conflict of interest.

\textbf{Data availability statement}

Data sharing is not applicable to this article because no datasets were
generated during the current study.

\textbf{Figures}\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image1/image1}

\end{center}
\end{figure}

\textbf{Figure 1. Scheme of chloroplast redox regulation in the stroma.}

A schematic representation of two reduction pathways (A), FNR/NTRC and
FTR/Trx under different light availability, and (B) oxidation pathway
under dark is shown. (A) Under high light intensity, the FTR regulatory
system dominantly reduces thiol-regulated target enzymes. Under limited
light availability, the NTRC system mainly functions. (B) Under dark,
2CP mediates oxidizing the target enzymes. More details can be found in
Section 2. The arrows indicate the flow of reducing equivalents, with
blue representing reduction and pink representing oxidation.

Abbreviations: ETC, Electron transport chain; FNR, ferredoxin-NADP+
reductase; FTR, ferredoxin-thioredoxin reductase; NTRC, NADPH-dependent
thioredoxin reductase C;

2CP, 2 Cys-peroxiredoxin\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image2/image2}

\end{center}
\end{figure}

\textbf{Figure 2. Comparison of the models of reducing equivalents
transport pathway across (A) inner membrane (IM) of Gram-negative
bacteria and (B) the thylakoid membranes chloroplast.} (A) Hypothetical
model of transferring reducing equivalents across the IM that involves
protein folding in the periplasm. This oxidative protein folding
disulfide bond isomerization pathway in Gram-negative bacteria through
the Dsb family. The protein is transported to the periplasm by SecYEG,
which is oxidized by DsbA. Reduced DsbA then transfers reducing
equivalents to DsbB in the IM. Reduced DsbB then transfer reducing
equivalents to quinone. When the protein is mis-oxidized, Trx in the
cytoplasm transfers reducing power to DsbD in the cytoplasmic membrane,
which then transfers it to DsbC for proper folding. (B) Hypothetical
model of reducing equivalents across the thylakoid membrane that may be
involved in possible protein folding and enzyme activity within the
lumen. Reducing equivalents from the stroma are transferred through CcdA
and HCF 164, followed by reduced redox-regulated proteins in the lumen.
LTO1 oxidizes redox-regulated proteins. The arrows indicate the flow of
reducing equivalents, blue for reduction and pink for oxidation. (Note:
Figure A was adapted from From Melissa E. Reardon-Robinson, and Hung
Ton-That J. Bacteriol. (2016) 198:746-754; while Figure B was adapted
from Frontiers in Plant Sci (2017) doi: 10.3389/fpls.2017.01313)\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image3/image3}

\end{center}
\end{figure}

\textbf{Figure 3. Distinct redox active statuses in the stroma and the
lumen.}

Light-dependent thiol/disulfide-regulated (A) stromal proteins are
considered to be in their active form when they are reduced. Conversely,
(B,C) lumenal proteins are considered to be in their active form when
they become oxidized. The proposed oxidation models are: (B) oxygen
(O\textsubscript{2}) and (C) LTO1.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image4/image4}

\end{center}
\end{figure}

\textbf{Figure 4. Scheme of hypothetical changes in redox potential of
selected thiol/disulfide redox-regulation proteins upon the energized
system by light, which induces increased and decreased pH in the stroma
(blue box) and lumen (pink box) respectively} . Redox-regulated proteins
in the stroma are shown in the orange box. Proteins that have the Cys
residues on the lumenal side are shown in the blue box. Note that HCF164
and LTO1 is transmembrane protein (section 3), but the shown redox
potentials here are for the soluble domain of those. The asterisk
indicates indicates changes in redox potential, two asterisks (**) for a
changes of -60mV/pH and one asterisk (*) for changes of -30mV/pH.
Lumenal thiols tend to easily gain reducing equivalents upon pH changes
(increased redox potential) while stromal thiols are the opposite.\selectlanguage{english}
\begin{figure}[H]
\begin{center}
\includegraphics[width=0.70\columnwidth]{figures/image5/image5}

\end{center}
\end{figure}

\textbf{Figure 5. Hypothetical model for redox regulation in the stroma
and the lumen.}

The redox-regulated stromal enzymes are shown in the upper panel (for
details, see section 2) and the redox-regulated thylakoid membrane and
lumenal proteins (section 4) are at the bottom of the panel. The arrows
indicate the flow of reducing equivalents. Hypothetical electron flow is
shown as dashed lines. The reductive pathway is indicated by the blue
and cyan arrows for stromal, and lumenal respectively, and oxidative
pathways are shown in red and pink arrows for stromal, and lumenal
respectively. To clarify, the use of a plus (+) sign in reductive arrows
denotes that the reduced state of the protein in the direction of the
arrow is in an active or stable form. On the other hand, the presence of
a minus (-) sign in the reductive arrow indicates that the reduced state
of the protein in the direction of the arrow is in an inactive or
unstable form. In summary, reducing equivalents from the stroma are
transferred to the lumen via CcdA and HCF164 to VDE and STN7. These
proteins are inactive when in their reduced forms, as indicated by the
minus (-) sign in the arrows. Additionally, SOQ1 also serves as a
reducing mediator, transferring reducing power to LCNP, which is
deactivated when reduced. The oxidizing system is mediated by LTO1,
which pulls electrons from VDE, STN7, and PsbO. However, the oxidizing
mediators for KEA3, Deg1, and LCNP are still not known. LTO1 mediates
the oxidizing system, extracting electrons from VDE, STN7, and PsbO.
Nevertheless, the oxidizing mediators for KEA3, Deg1, and LCNP remain
unknown. All of these redox-regulated proteins in the lumen, including
VDE (qE, qZ), KEA3 (qE), STN7 (qT), LNCP (qH), Deg1, and PsbO (qI), are
cooperating to induce photoprotective mechanisms. The hypothetical model
for removing electrons from the lumen is shown that reactive oxygen
species (ROS) act as a strong oxidant via unknown strong oxidant. The
Trx-domain of LTO1 would pull electrons from target enzymes, and these
electrons would then be transferred to the VKOR domain of LTO1 in the
thylakoid membrane. From there, the electrons could be transferred to an
unknown oxidizing mediator (marked as ''?'' in the figure), which could
further reduce H\textsubscript{2}O\textsubscript{2.} See more details in
Table 1 and section 3 and 4.

\textbf{Table 1.}

Summary of redox-regulated proteins that contribute to photoprotection
in the thylakoid lumen and transmembrane\selectlanguage{english}
\begin{longtable}[]{@{}llllllllll@{}}
\toprule
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Categorized role\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Protein\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Accession\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Redox potential\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Localization\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Known function\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Roles in redox pathway or Redox status of Active form / Stable
form\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Interactions ( Trx, HCF164, LTO1)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
number of cysteines in mature proteins\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
references\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trans- thylakoid thiol mediators\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
CCDA\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT5G54290\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Assembly of Cyt\emph{b\textsubscript{6}f}\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
transfer redox equivalents\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trx, HCF164\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
4\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Page et al. 2004, Motohashi \& Hisabori 2010\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
HCF164\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT5G23120\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
-224mV (pH 7.0)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Assembly of Cyt\emph{b\textsubscript{6}f}\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
transfer redox equivalents\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trx, CCDA\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
3\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Lennartz et al. 2001, Motohashi \& Hisabori 2006\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
SOQ1\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
AT1G56500\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Photoprotection\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
transfer redox equivalents\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
LCNP\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
2\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Brooks et al 2013\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
LTO1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT4G35760\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
-180mV (pH 7.0)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Assembly of PSII\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
oxidant\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
VDE, STN7, PsbO\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
8\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Karamoko, Cline, Redding, Ruiz \& Hamel 2011, Lu et al. 2013; Yu, Lu,
Du, Peng \& Wang 2014\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Photoprotection (qE)\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
KEA3-1\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
AT4G04850\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
\emph{pmf} partitioning\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
5\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Wang et al. 2017\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
VDE\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT1G08550\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
-317.3mV (pH 7.9)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Lumen\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
catalyzes the conversion of violaxanthin (Vx) to zeaxanthin (Zx)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
oxidized state is active\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
LTO1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
13\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Arnoux et al. 2009, Yu 2014, Hallin 2015, Simionato 2015\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Sustained quenching (qH)\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
LCNP\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
AT3G47860\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Lumen\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
photoprotection\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
oxidized state is active\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
SOQ1\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
6\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Malno\selectlanguage{ngerman}ë et al. 2018\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
State transition (qT)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
STN7\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT1G68830\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
thylakoid membrane\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
state transition\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
oxidized state is active\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trx, LTO1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
2\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Rintam\selectlanguage{ngerman}äki, Martinsuo, Pursiheimo \& Aro 2000, Pesaresi et al. 2009,
Lemeille et al. 2009, Wu et al. 2021\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
PSII stability and repair (qI)\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
PsbO1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT5G66570\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Lumen\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
PSII subunit, OEC\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
oxidized state is stable\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trx, LTO1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
2\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Hall et al. 2010, Karamoko et al. 2011\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
PsbO2\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
AT3G50820\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Lumen\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
PSII subunit, OEC\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
oxidized state is stable\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Trx, LTO1\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
2\strut
\end{minipage} & \begin{minipage}[t]{0.07\columnwidth}\raggedright\strut
Lundin, Hansson, Schoefs, Vener \& Spetea 2007, Karamoko et al.
2011\strut
\end{minipage}\tabularnewline
\begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
DEG1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
AT3G27925\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Unknown\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Lumen\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
protease, degradation of D1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
reduced state is stable\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Trx\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
1\strut
\end{minipage} & \begin{minipage}[t]{0.10\columnwidth}\raggedright\strut
Chassin et al. 2002, Str\selectlanguage{ngerman}öher \& Dietz 2008, Sun et al. 2010\strut
\end{minipage}\tabularnewline
\bottomrule
\end{longtable}\selectlanguage{ngerman}

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