Redox regulation in the thylakoid lumen

Accepted Manuscript Title: Redox regulation in the thylakoid lumen Author: Zhen-Hui Kang Gui-Xue Wang PII: DOI: Reference:

S0176-1617(16)00009-2 http://dx.doi.org/doi:10.1016/j.jplph.2015.12.012 JPLPH 52285

To appear in: Received date: Revised date: Accepted date:

1-11-2015 4-12-2015 4-12-2015

Please cite this article as: Kang Zhen-Hui, Wang Gui-Xue.Redox regulation in the thylakoid lumen.Journal of Plant Physiology http://dx.doi.org/10.1016/j.jplph.2015.12.012 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Redox regulation in the thylakoid lumen Zhen-Hui Kang, Gui-Xue Wang** [email protected] Key Laboratory of Biorheological Science and Technology (Chongqing University), Ministry of Education, Bioengineering College of Chongqing University, Chongqing 400030, China **

 

Corresponding author. Tel: +8623 65112672; fax: +8623 65112672.

1

Highlights



Redox regulation is crucial for photosystem biogenesis/function in varying light.



Control of disulfide bond formation by thioredoxins (Trx) is vital to this process.



Compared to chloroplast stroma, little is known about this process in the lumen.



The thylakoid lumen contains over 30 proteins with disulfide bonds.



The discovery of lumenal Trx highlights the role of redox regulation in the lumen.

 

2

Abstract Higher plants need to balance the efficiency of light energy absorption and dissipative photo-protection when exposed to fluctuations in light quantity and quality. This aim is partially realized through redox regulation within the chloroplast, which occurs in all chloroplast compartments except the envelope intermembrane space. In contrast to the chloroplast stroma, less attention has been paid to the thylakoid lumen, an inner, continuous space enclosed by the thylakoid membrane in which redox regulation is also essential for photosystem biogenesis and function. This sub-organelle compartment contains at least 80 lumenal proteins, more than 30 of which are known to contain disulfide bonds. Thioredoxins (Trx) in the chloroplast stroma are photo-reduced in the light, transferring reducing power to the proteins in the thylakoid membrane and ultimately the lumen through a trans-thylakoid membrane-reduced, equivalent pathway. The discovery of lumenal thiol oxidoreductase highlights the importance of the redox regulation network in the lumen for controlling disulfide bond formation, which is responsible for protein activity and folding and even plays a role in photo-protection. In addition, many lumenal members involved in photosystem assembly and non-photochemical quenching are likely required for reduction and/or oxidation to maintain their proper efficiency upon changes in light intensity. In light of recent findings, this review summarizes the multiple redox processes that occur in the thylakoid lumen in great detail, highlighting the essential auxiliary roles of lumenal proteins under fluctuating light conditions.

 

3

Keywords: Disulfide; photosynthetic efficiency; redox; thiol; thylakoid lumen.

 

4

Introduction During the past decade, much progress has been made in understanding the role of the thylakoid lumen in photosynthesis (Spetea and Lundin, 2012; Jarvi et al., 2013; Karamoko et al., 2013). The thylakoid lumen was originally considered to be a vacant space used only for photosynthetic electron transport and proton gradient formation. The discovery of the lumenal peripheral and soluble proteins in a few higher plants, such as pea, spinach and Arabidopsis by mass spectrometry (Kieselbach et al., 1998; Peltier et al., 2000; Peltier et al., 2002; Schubert et al., 2002; Kieselbach and Schroder, 2003; Friso et al., 2004) has led to the gradual realization that the thylakoid lumen is far more complicated and has its own proteome. The thylakoid lumen in Arabidopsis contains at least 80 proteins, with a protein density of approximately 10 mg mL-1 (Kieselbach, 2013). Although seemingly narrow and limited, the space of thylakoid lumen is highly flexible (Kana et al., 2009). High light induces the dimensional extension of the lumen, making it favorable for protein diffusion and pH maintenance to control the activities of lumenal proteins (Granlund et al., 2009; Kirchhoff et al., 2011). Under dark conditions, the pH of the chloroplast stroma is neutral (approximately 7.0). In the light, its pH increases to 7.5-8.0. By contrast, the pH of the thylakoid lumen decreases to 5.4-6.0 in the light from around 7.0 in dark or very low light (Kramer et al., 1999; Cruz et al., 2001; Tikhonov, 2013). All lumenal proteins identified to date are encoded by the nuclear genome and are targeted to the chloroplast by their transit peptides, across the chloroplast envelope membrane via the TOC/TIC channel (Soll and Schleiff, 2004) and into the lumen through the

 

5

trans-thylakoid membrane Sec/Tat translocation pathway (Aldridge et al., 2009). During their transport, many lumenal proteins are post-translationally modified through reversible phosphorylation and thiol/disulfide modulation (Gopalan et al., 2004; Spetea et al., 2004). The exchange of the oxidation and reduction states of sulfhydryl groups (disulfide bridges and -SH) is a common phenomenon. In addition to important structural and catalytic functions, in many cases, breakage and reforming of disulfide bonds are tightly regulated. The thylakoid lumen of higher plants evolved from bacterial periplasm; thiol oxidation in both structures shares common principles but unrelated machineries (Herrmann et al., 2009). In bacteria periplasm, the key components of the thiol oxidation pathway consist of soluble protein DsbA and membrane protein DsbB. In the disulfide-reducing pathway, thiol-disulfide transporter DsbD transmits reducing equivalents to target proteins from the cytosol to the periplasm, which require reduced thiols for activation (Heras et al., 2009; Kadokura and Beckwith, 2010; Depuydt et al., 2011). The discovery of the control of disulfide bond formation and reduction by trans-thylakoid redox pathways in Chlamydomonas and Arabidopsis has opened a new frontier in the field of photosynthesis (Karamoko et al., 2013). Each compartment of the chloroplast (except for the inner space of the envelope) contains targets of Thioredoxin (Trx), including the thylakoid lumen. More than 40% of lumenal proteins can form disulfide bonds, which are potentially regulated by redox signaling. Therefore, redox regulation in the thylakoid lumen in multiple life processes in higher plants, such as protein translocation and folding, as well as protein

 

6

activity, cannot be ignored (Hall et al., 2010). Like the bacterial plasma membrane, our understanding of the thiol metabolism pathway also sheds light on the thylakoid membrane connecting the chloroplast stroma and lumen (Karamoko et al., 2013). Redox regulation controls the activity and function of many lumenal proteins, making it indispensable for the assembly and photoprotection of the photosystem and photosynthetic electron transport. In this review, we present a detailed discussion of the redox regulation of these lumenal proteins.

1. Trx-m in the chloroplast stroma function as electron donors for thylakoid lumenal proteins Light-dependent regulation of disulfide/thiol exchange in Trx has long been thought to be the basic mechanism underlying chloroplast metabolism. Trx-mediated redox control is a common characteristic of these important pathways, such as the Calvin cycle, nitrogen metabolism and oxidative stress (Lindahl and Kieselbach, 2009). The Arabidopsis genome encodes 19 isoforms of Trx, which are divided into six groups, with f-, m-, x- and y-type Trx distributed in the chloroplast. Trx contains a conserved WC(G/P)PC motif that is reduced by chloroplast ferredoxin-thioredoxin reductase (FTR) or NADPH-dependent ferredoxin reductase (FNR), which in turn function as electron donors (hydrogen) by reducing other target proteins (Serrato et al., 2013). During photosynthesis, because FTR receives electrons from photoreduced ferredoxin, Trx acts as an “eye” to sense light. Trx also links enzyme activity with the levels of oxidants generated under stress conditions via a mechanism designated

 

7

“oxidative regulation” (Balmer et al., 2003). The target proteins of Trx are widely distributed in the suborganelle compartments of chloroplasts, from the envelope to the thylakoid lumen. The chloroplast stroma contains 79 known targets of Trx, while in the thylakoid lumen, 30 known proteins contain one or more Cys residues (mature form, Table 1), and 19 proteins have been identified as targets of Trx (Hall et al., 2010). The functions of these lumenal proteins are clearly Trx dependent. The occurrence of Trx target proteins in the thylakoid lumen suggests that regulation by Trx is involved in photosystem complex assembly and protease/protein activity, as well as non-photochemical quenching (NPQ). Identification of Trx target proteins is mainly achieved through the detection of disulfide bonds via two different methods. First, as WC(G/P)PC is the redox-active sequence in all classical Trx, the Cys residue at the N-terminus of this motif can form disulfide intermediates with target proteins, which are subsequently broken by nucleophilic attack of the second Cys residue, leading to the production of an oxidized Trx and a reduced target protein (Berndt et al., 2008). Exposed thiol groups in the newly formed target protein can be labeled by certain thiol reagents, such as the fluorescent dye monobromobimane and radioactive iodoacetamide (Marchand et al., 2006), and the resulting complexes can be visualized by CCD cameras (Yano et al., 2001). Second, Trx affinity chromatography and mass spectrometry, which take advantage of the characteristics of Trx reduction of special disulfides, can be used for Trx target protein detection. Before the disulfide bond in the target protein is completely reduced, transient iso-disulfides form between Trx and its interacting

 

8

protein. If only one of the two Cys is mutated, the transient iso-disulfides will be stabilized. The target protein can then be cleaved by DTT, which covalently combines with Trx (Motohashi et al., 2001). Target proteins of Trx in the thylakoid lumen are listed in Table 1. The thylakoid lumen contains no soluble Trx, and the reducing power is transferred from reduced Trx-m into the lumen via the thylakoid membrane protein

CCDA,

a

member

of

the

recently

determined

trans-thylakoid

thiol-metabolizing pathway (Karamoko et al., 2013)(see section 2).

2. Trans-thylakoid thiol-metabolizing pathways control disulfide bond formation and reduction of lumenal proteins Many enzymes that catalyze disulfide bond formation in proteins are members of the Trx superfamily. Unlike the chloroplast stroma, no soluble Trx has been identified in the thylakoid lumen. Increasing evidence strongly indicates that the reducing equivalents are transferred from the chloroplast stroma into the thylakoid lumen (Page et al., 2004; Motohashi and Hisabori, 2010). The reduction of disulfides to free thiols (-S-S- to 2SH) and the reverse reaction, thiol oxidation to disulfides (2SH to -S-S-), is an integral part of biological processes in the bacterial periplasm, the mitochondrial intermembrane space (IMS) and the thylakoid lumen; these processes are involved in the regulation of protein folding and function. Although the IMS and thylakoid lumen are evolutionary products of bacterial periplasm, the oxidoreductases in the three compartments are not the same. The process of protein secretion into the periplasmic space in bacteria is mediated by a protein transposon channel (Dsb family proteins)

 

9

(Inaba et al., 2006). In brief, either during or after their translocation across the inner membrane, the sulfhydryl groups in the transferred proteins are initially oxidized by soluble protein DsbA (Grauschopf et al., 1995). The reduced DsbA is then reoxidized and recovered by the membrane protein DsbB, with the electrons transferred to quinone (Bader et al., 1999). DsbC and DsbG counteract protein oxidation of DsbA until a protein or protein complex adopts its native structure, which is essential for isomerization and folding of periplasmic proteins with nonconsecutive disulfide bonds (Hiniker and Bardwell, 2004). DsbD supplies DsbC and DsbG with electrons for their reduction again (Krupp et al., 2001). DsbD is located in the cytoplasmic membrane and functions as a thiol-disulfide transporter, representing a link between DsbC and DsbG and the cytoplasmic Trx systems (Herrmann et al., 2009). Erv1 and Mia40 transport Cys-rich proteins into the IMS, relying on the oxidative folding mechanism (Hell, 2008). For the reduction of disulfides in the lumenal proteins, the reducing equivalent transfer system functions across the thylakoid membrane from the chloroplast stroma to the thylakoid lumen. This system comprises the transmembrane transporter protein CCDA, as well as thylakoid membrane-anchored lumenal proteins HCF164 and SOQ1 (Karamoko et al., 2013). Stromal FTR and/or FNR-dependent Trx are usually used as hydrogen donors in this trans-membrane Dsb-like system. 2.1 CCDA is a transporter across the thylakoid membrane with reducing power Plant CCDA is a prokaryotic homolog of thiol disulfide transporter (Page et al., 2004). CCDA in Arabidopsis is a polytopic thylakoid membrane protein containing six transmembrane regions, which can be reduced by Trx-m and functions as a mediator

 

10

in transferring reducing equivalents from the stoma to the thylakoid lumen (Motohashi and Hisabori, 2010). HCF164, a downstream factor of CCDA, is also reduced by Trx-m; the specificity is determined by CCDA reacting with Trx-m before HCF164 (section 2.2). CCDA is an indispensable factor for Cyt b6f complex assembly (Page et al., 2004). Mutants of ccda show similar phenotypes to hcf164 mutants, such as their inability to synthesize Cyt b6f complex (Lennartz et al., 2001). Although chloroplasts contain four types of Trx, including m-, f-, x- and y-type, it is currently unclear why CCDA only uses m-type Trx as an electron donor. Perhaps this occurs because f-type has a eukaryotic origin, while m-type has a prokaryotic origin. However, neither x- nor y-type Trx, which have prokaryotic origins, participate in this system; this issue requires further study. A direct interaction between HCF164 and CCDA has not yet been detected. CCDA contains only one disulfide bond, with two Cys residues embedded within the membrane. Therefore, other factors involved in the transfer of reducing equivalents should exist in the thylakoid membrane, such as a thylakoid membrane-anchored protein with a Trx domain oriented to the stroma. In addition, the active cysteines in the two proteins are not close enough for disulfide bond formation. Thus, biochemical analyses are needed to elucidate how these proteins carry out thiol-disulfide exchange. CCDA and HCF164 were identified through the study of mutants with abnormal assembly of Cyt b6f complex. HCF153, a nuclear-encoded factor, is also necessary during a post-translational step in biogenesis of the Cyt b6f complex (Lennartz et al., 2006). It will be interesting to elucidate the relationship between CCDA and HCF153,

 

11

as the biochemical parameters of the latter are still unclear. Arabidopsis HCF153 shares limited similarity with CCS4, which encodes the algal ortholog in Chlamydomonas. CCS4 is a thylakoid membrane-anchored protein required for the assembly of the Cyt b6f complex, which interacts with CCDA by stabilizing the protein and/or controlling its activity, possibly via its C-terminal domain (Gabilly et al., 2011). Although CCS4 does not contain sequence motifs with suggestive redox or heme-binding function, biochemical and genetic complementation experiments suggest that it plays a role in the disulfide-reducing pathway required for heme attachment to apoforms of Cyt f and c6 (Gabilly et al., 2011). Therefore, it is important to investigate whether HCF153 is the third factor of the CCDA/HCF164 pathway. 2.2 HCF164 and SOQ1 serve as disulfide reducing components in the lumen Plant plastid c-type Cyt biogenesis is necessary for the assembly of the Cyt b6f complex, which occurs in the thylakoid lumen. The reduction state of sulfhydryls in its CxxCH motif of apocytochrome c must be maintained before the covalent attachment of heme. The hcf164 mutant exhibits electron transfer defects, in which subunits of the Cyt b6f complex cannot accumulate, suggesting that HCF164 is involved in the assembly and maturation of the Cyt b6f complex (Lennartz et al., 2001). HCF164 is a thylakoid membrane Trx-like protein. Recombinant HCF164 protein, whose hydrophilic C-terminus contains thiol groups facing the thylakoid lumen, displays in vitro redox activity (Cain et al., 2009). Topological analysis revealed that HCF164 functions as a component of the transmembrane disulfide reducing pathway and is similar to bacterial CcmG/ResA/CcsX, reducing disulfide in

 

12

the CxxCH heme-binding site of apocytochrome c prior to heme ligation to cysteines (Bonnard et al., 2010; Sanders et al., 2010). As the electron donor, chloroplast stromal Trx-m can reduce HCF164 in vitro, facilitating the transmission of reducing power to the catalytic domain of HCF164 residing at the lumenal side (Motohashi and Hisabori, 2006). PSI-N, a subunit of PSI, as well as cytochrome f and Rieske FeS proteins of the Cyt b6f complex, were identified as targets of HCF164 by Trx affinity chromatography (Fig. 1). Moreover, in vitro and in organello experiments showed that reduction of the disulfide bonds in PSI-N is dependent on HCF164. It is still unclear whether HCF164 reduces its lumenal targets and Trx-m reduces CCDA in the dark or in the light. Hall et al. speculated that the transfer of reducing equivalents to lumenal target proteins might be restricted to dark periods and that the reduction of the transmembrane system might alternatively be catalyzed in the dark by NTRC (Hall et al., 2010). This assumption appears to be reasonable, because the acidic conditions in the lumen in the light are certainly not favorable for the reduction of HCF164. NTRC is a light-independent NADPH-disulfide reductase in the chloroplast stroma (Michalska et al., 2009; Bernal-Bayard et al., 2012). Thus, a reversible regulatory mechanism for lumenal proteins might involve reduction in the dark and oxidation in the light. This process may be helpful for enzymes such as violaxanthin de-epoxidase (VDE), a typical lumenal enzymes that converts violaxanthin to zeaxanthin, which is reductively inactivated in the dark and oxidatively reactivated in the light (Spinola et al., 2008). However, the reduction of Trx-m by NTRC has not yet been reported. Further experiments are needed to confirm

 

13

these assumptions. Both HCF164 and VDE were recently shown to interact with Lumen Thiol Oxidoreductase1 (LTO1) in vitro (Lu et al., 2014), suggesting that HCF164 is inactive in its oxidized form and that its reducing power does not function in the light, whereas it reopens in the dark for the reduction of lumenal targets. Future studies should investigate the conditions under which HCF164 interacts with LTO1 and reduces its lumenal targets. In addition to HCF164, another Trx-like protein, suppressor of quenching 1 (SOQ1), was recently identified in the thylakoid lumen (Brooks et al., 2013). SOQ1 is thought to preserve the efficiency of light harvesting under high-light conditions as well as maintain target proteins in the reduced state in the oxidized environment of the thylakoid lumen (Fig. 1). The soq1 mutants have high NPQ, even in the absence of PSII subunit S (PsbS, npq4), and are light density-dependent, with slow recovery dynamic kinetics. The role of SOQ1 is to prevent the occurrence of a slowly reversible form of antenna quenching, directly or indirectly preventing qI-type quenching of excitation energy of light-harvesting antenna proteins distinct from known NPQ components (qE, qZ, qT and qI). In the absence of SOQ1, one or more quenching site(s) forms in the proximal and/or peripheral PSII antenna, which reduces the chlorophyll excited-state lifetime. High light stimulation induces PSII reorganization in soq1, and the absence of SOQ1 affects the organization of disordered PSII supercomplexes and weakens the interactions among antenna complexes (Onoa et al., 2014). SOQ1 contains a thioredoxin-like/β-propeller domain localized in the lumen, which is critical to its function (Brooks et al., 2013). However,

 

14

whether SOQ1 receives reducing power from CCDA or directly from stromal Trx has not yet been determined. 2.3 LTO1 is a disulfide bond-forming catalyst in the lumen LTO1 has thiol-oxidizing catalyst activity in the thylakoid lumen (Karamoko et al., 2011). LTO1 is a thylakoid membrane protein containing a VKOR domain fused to a Trx-like domain. It was previously unclear whether the disulfide bond formation of thylakoid lumen proteins was catalyzed by oxidase or whether it was a non-enzymatic reaction process, namely, random oxidation of oxygen produced by oxygen evolving complex (OEC). The high concentration of oxygen produced by the light reaction at the thylakoid lumenal side suggests that free oxygen must be the oxidant that oxidizes lumenal proteins. It was indeed suggested that sufficient levels of free oxygen are present to induce the formation of disulfide bonds in the newly inserted proteins in the thylakoid lumen. However, many thylakoid lumenal proteins contain one or more disulfide group, and disulfide bonds can form in more than one motif, indicating that random oxidation does not lead to disulfide bond formation in these lumenal proteins. Both Cyt c6A and plastocyanin were also suggested to be oxidants for catalyzing disulfide bond formation in lumenal proteins. However, they cannot facilitate reversible regulation of enzyme activity because it would occur by a one-way process. Therefore, an unknown oxidoreductase is probably present in the lumen. This speculation was finally confirmed with the recent discovery of LTO1 (Karamoko et al., 2011; Karamoko et al., 2013). The Arabidopsis lto1 mutant has a photoautotrophic defect and inhibited

 

15

photosynthetic electron transfer from PSII. LTO1 interacts with the PSII subunit PsbO, and disulfide bonds of PsbO are introduced by the Trx-like domain of LTO1 in vitro. The redox state of thiols in PsbO plays a decisive role in its stability and in PSII accumulation. When transferred to the thylakoid membrane, PsbO can easily be degraded, as it is in the transition state. Disulfide bonds in newly transported PsbO are clearly reduced (Kieselbach, 2013). Disulfide bond formation and rapid folding of PsbO are required to ensure stable folding and binding to PSII. However, it is still unknown whether LTO1 assists in the translocation of PsbO into the thylakoid lumen. There are two models of the membrane topology of LTO1 based on Pho and Laz receptors. In both models, the Trx-like domain in its C-terminus is exposed to the thylakoid lumen, but this protein may have either four or five transmembrane helices due to the topology of the VKOR-like domain. The Trx-like domain of LTO1 is functionally equivalent to bacterial DsbA, while the VKOR-like domain is similar to DsbB (Singh et al., 2008; Dutton et al., 2010; Wang et al., 2011). The VKOR-like domain oxidizes the Trx-like domain in vitro and receive electrons from the latter. The final electron acceptor of LTO1 has not yet been identified. It is generally believed that phylloquinone may be the electron receptor of the LTO1-dependent disulfide formation pathway (Furt et al., 2010), as VKOR reduces Vitamin K using phylloquinone as a cofactor (Tie and Stafford, 2008). LTO1 was recently shown to display disulfide bond isomerase activity in vitro, which is critical for proteins containing more than two Cys. LTO1 introduces disulfide bonds in AtFKBP13 in vitro, indicating that it is essential for maintaining the activity

 

16

of thylakoid lumen proteins in oxidation (Lu et al. 2013). In addition to PsbO and AtFKBP13, at least eight targets of LTO1 have been identified in the lumen, including HCF164 and VDE (Lu et al., 2014). Since 22 lumenal proteins have two or more Cys, the lumen may contain other oxidoreductases besides LTO1. 2.4 Other potential oxidants that function as electron acceptors in the lumen Cyt c6A is a unique dithio-cytochrome in plants that shares a similar core structure to that of its bacterial and algal counterpart Cyt c6. However, Cyt c6A is unable to transfer electrons from Cyt f to PSI due to its low redox potential (+140 mV) (Marcaida et al., 2006). Cyt c6A represents an evolutionary “bricolage”. A key feature of Cyt c6A is that the midpoint potential of its heme group is lower than that of Cyt c6, mainly because Gln in the heme pockets was replaced by Val during the evolution from Cyt c6 (Worrall et al., 2007; Worrall et al., 2008). Cyt c6A in Arabidopsis is responsible for disulfide bond formation in response to changes in intracellular redox state, subsequently transferring the reducing equivalents to plastocyanin. Unlike Cyt c6, Cyt c6A possesses a conserved insertion of 12 amino acids, including two absolutely conserved Cys residues (Chida et al., 2006; Howe et al., 2006; Schlarb-Ridley et al., 2006).

3. Activities of several lumenal proteins are redox regulated during the photoprotection and assembly of photosystems When light intensity exceeds a certain limit, photosynthesis reaches a saturation point. The resulting excess energy, if not dissipated promptly, is transmitted to the

 

17

surrounding oxygen, producing reactive oxygen species (ROS). ROS induce photodamage in the photosynthetic membrane and reversely inhibit photosynthesis (Goss and Lepetit, 2015). During the long process of evolution, the photosystems developed a series of protective light quenching mechanisms to minimize this damage (Standfuss et al., 2005). 3.1. Non-photochemical quenching (NPQ) Thermal dissipation depending on the xanthophyll cycle, also referred to as NPQ, is considered to be the main mechanism for protecting the photosynthetic apparatus from photodamage due to excess light (Muller et al., 2001). NPQ comprises four components, qE, qZ, qT and qI, which were designated according to the sense of their known dependent factors as well as their induction and relaxation kinetics (Nilkens et al., 2010). Reduction of disulfide bonds inhibits the activity of lumenal VDE, which is required to activate qE (the pH-dependent regulation of photosynthetic light harvesting) to dissipate excess absorbed light energy as heat (Pfundel and Dilley, 1993). VDE catalyzes the de-epoxidation of violaxanthin to antheraxanthin and finally to zeaxanthin (Rockholm and Yamamoto, 1996). The generation of zeaxanthin is related to the activity of VDE, which is strictly under the control of pH in the thylakoid lumen (Li et al., 2009; Fufezan et al., 2012). When the pH in the thylakoid lumen is less than 6.2, and with ascorbic acid (ASC) as a cofactor and electron acceptor, VDE is activated (Hieber et al., 2000; Muller-Moule et al., 2002). VDE contains a lipocalin domain, as well as Cys- and Glu-rich domains, which are required

 

18

for its function. VDE comprises a single eight-stranded antiparallel β-barrel, which adopts an open conformation that facilitates ligand access to the active site in the dimeric state (Arnoux et al., 2009). Reduction of the disulfide bonds in VDE results in the loss of its rigid structure (Hallin et al., 2015). VDE contains 13 Cys, at least 12 of which are conserved. The activity of VED depends on its oxidation state: it is active only when six disulfide bonds form, which is inhibited by DTT, a VDE-specific inhibitor (Simionato et al., 2015). At pH 7, VDE is a soluble lumenal protein that exists as a monomer. However, under acidic conditions, it adopts a stable dimeric conformation and attaches to the thylakoid membrane (Arnoux et al., 2009). Through redox regulation, plants can dynamically and rapidly control VDE levels, which is vital for adaptation of the thylakoid lumen to fluctuating light conditions. Reductively inactivated VDE may be degraded by protease or regenerated by forming new disulfide bonds in the thylakoid lumen. To optimize growth in the light, plants must maintain the correct redox environment in chloroplasts to balance the functioning of PSII and PSI (Pesaresi et al., 2011). STN7 kinase is involved in the reversible phosphorylation of LCHII, an important state transition mechanism controlling energy distribution between PSII and PSI under low-light conditions (Bellafiore et al., 2005). High light induces the migration of LHCII trimer from PSII to PSI to prevent excessive activation of PSII (Bonardi et al., 2005). However, the phosphorylation of LHCII is inhibited by high light, and the dynamic distribution of light energy between photosystems is

 

19

unnecessary (Martinsuo et al., 2003). The homodimer of STN7 is active upon binding of reduced plastoquinone (PQ) to the Qo site of the Cyt b6f complex (Betterle et al., 2015). The deactivation of STN7 achieves the light intensity-dependent inhibition of LHCII phosphorylation under high-light conditions (Wunder et al., 2013). STN7 is a thylakoid membrane protein with catalytic sites at the stromal side, but its redox-active Cys is located in the lumen, which is essential for its kinase activity (Pesaresi et al., 2009). The reduction of intra/inter disulfide bonds of STN7 may induce monomerization of this protein, thereby breaking its association with Cyt b6f. In their review, (Ruban and Belgio, 2014) suggested that HCF164 and LTO1 may be responsible for the reduction and oxidation of STN7. However, we speculate that STN7 is probably reduced/deactivated through a light density-dependent reducing power-transferring pathway and that it receives electrons from Trx-m reduced by FTR. It appears that SOQ1 can reduce STN7 but not HCF164 (Hall et al., 2010; Brooks et al., 2013). Due to the low pH in the thylakoid lumen, SOQ1 may retain its reducing potential with the help of its NHL domain, which also faces the lumen, as NHL has diverse functions including catalysis, ligand binding, and protein-protein interactions (Loedige et al., 2014). Further experiments are needed to investigate whether the recombinant Trx+NHL domain alone has high activity at low pH in vitro. A recent study showed that LTO1 does not interact with STN7 (Lu et al., 2014), implying that it is not the oxidant of STN7. S-sulfocysteine may be involved in the oxidation of STN7 due to its special properties under high-light conditions (see section 4).

 

20

3.2 Photosystem assembly Light is associated with changes in the redox state of chloroplast enzymes and post-translational regulation. By linking a hydrogen donor to Cys-containing proteins at their transition state, redox reactions control the activity of target proteins through disulfide exchanges. The thylakoid lumen proteome contains a large proportion of immunophilins with peptidyl-prolyl isomerase (PPIase) activity, which are generally correlated with the biogenesis and maintenance of supramolecular complexes (Ratajczak et al., 2003; Lima et al., 2006; Edvardsson et al., 2007). The elucidation of the relationship between the immunophilin FKBP13 and Trx-dependent redox regulation in Arabidopsis has opened up a new frontier in redox regulation in the thylakoid lumen (Gopalan et al., 2004; Buchanan and Luan, 2005). The redox regulation of the PPIase activity of a few immunophilins by the redox signal in the thylakoid lumen plays an indispensable role in photosystem assembly. In the chloroplast stroma, enzymes participating in biosynthesis are functionally activated via reduction by the Trx system, whereas FKBP13 acquires its active form by oxidizing sulfhydryl groups in the light (Romano et al., 2005). FKBP13 is transmitted into the chloroplast as a protein precursor and is processed into its mature form while being targeted to the thylakoid lumen, which relies on the ΔpH Tat pathway (Gupta et al., 2002b). In Arabidopsis, AtFKBP13 is essential for the formation of the Cyt b6f complex, and its precursor (but not the mature form) interacts with Rieske and FeS protein (Gupta et al., 2002b; Vasudevan et al., 2014). Another potentail target of AtFKBP13 is Cyt c6A (Gupta et al., 2002a; Schlarb-Ridley et al.,

 

21

2006), which is believed to be derived from Cyt c6 that serves as a substitute for the copper-containing protein plastocyanin during electron transfer in cyanobacteria and eukaryotic algae when they are short of Cu2+ (Haddadian and Gross, 2006; Bell et al., 2009). Two conserved disulfides are located at its C- and N-terminus of AtFKBP13 respectively; both can be reduced by Trx derived from the chloroplast (m-type) or E. coli (Gopalan et al., 2004). AtFKBP13 loses its PPIase activity when the two disulfide bridges are reduced or mutated, indicating that these two disulfide bonds are required for the PPIase activity of this protein (Edvardsson et al., 2007; Ingelsson et al., 2009). Rearrangement of the redox active sites in its reduced form and the redox-linked variation of secondary structure at residues 50-53 in its PPIase domain are important reasons for the inactivation of AtFKBP13 (Gopalan et al., 2006; Shapiguzov et al., 2006). Another lumenal immunophilin, FKBP20-2, which is involved in PSII complex assembly, is also reactive with Trx. This protein contains two conserved Cys residues at its C-terminus. Recombinant FKBP20-2 can accept electrons from E. coli Trx, but the reduction of FKBP20-2 does not affect its PPIase activity (which is relatively low) (Lima et al., 2006). AtCYP38 is another lumenal immunophilin target of Trx (Hall et al., 2010), which plays an important role in PSII biogenesis and assembly, especially under high-light conditions. The mature sequence of AtCYP38 contains only a single Cys residue, C256, which is conserved in plant homologs and is located at the interface between the acidic domain and the PPIase-domain of the protein (Shapiguzov et al., 2006). However, the role played by Trx-mediated signaling in

 

22

regulating AtCYP38 remains unknown. LTO1 was recently found to catalyze disulfide bond formation in AtFKBP13, as demonstrated in vitro (Lu et al., 2013). However, reducing signals in the lumen for the conversion of disulfide to thiols in AtFKBP13 have not yet been experimentally verified, although both HCF164 and SOQ1 can potentially transfer reducing power from the chloroplast stroma to the thylakoid lumen (Lennartz et al., 2006; Motohashi and Hisabori, 2006, 2010; Brooks et al., 2013). Meanwhile, additional candidate enzymes involved in the reduction and oxidation of FKBP20-2 and AtCYP38 remain to be identified. FKBP16-2, another PPIase in Arabidopsis (He et al., 2004), is present in the lumenal multi-protein subcomplex NAD(P)H dehydrogenase-like complex (NDH), which facilitates cyclic electron flow (CEF) around PSI (Peng et al., 2009). Deficiency of FKBP16-2 disrupts the assembly of functional NDH and affects the formation of the NDH-PSI supercomplex. FKBP16-2 is a homolog of FKBP13 in higher plants; both possess cysteine pairs at the identical positions (Gollan and Bhave, 2010). Therefore, the activity of FKBP16-2 is probably regulated by lumenal redox signals, suggesting that it links redox signaling with the assembly and/or activity of NDH, which is associated with CEF. However, FKBP16-2 was not identified as a target of Trx and does not interact with LTO1, indicating that it is regulated by another thiol oxidizing pathway.

4. Redox potentials of lumenal proteins in the oxidative stress response The oxidative stress response is a complicated process in photosynthetic organisms

 

23

that is differentially regulated in different types of cells and subcellular organelles (Foyer and Noctor, 2009; Lindahl and Kieselbach, 2009). The thylakoid lumen contains the S-sulfocysteine synthase CS26, whose activity is essential for long-day light-dependent redox regulation within the chloroplast (Bermudez et al., 2010). Chloroplasts contain two O-acetylserine(thiol)lyase homologs, which are encoded by OAS-B and CS26 in Arabidopsis (Romero et al., 2014). Recombinant CS26 protein possesses S-sulfocysteine synthase activity and lacks O-acetylserine(thiol)lyase activity. Although it is present in the thylakoid lumen at relatively low levels, CS26 is highly important for chloroplast function (Gotor et al., 2010). Cs26-deficient mutants exhibit strong photoinhibition and ROS stress, which may interfere with the reduction of sulfite or may oxidize sulfide to form thiosulfate (Bermudez et al., 2012). CS26 is quite sensitive to thiosulfate accumulation and may function as a ROS-sensing protein in the thylakoid lumen. In addition, CS26 uses thiosulfate as a substrate to produce S-sulfocysteine, which triggers protective mechanisms of the photosynthetic apparatus and plays a key role in monitoring the redox environment in the thylakoid lumen, especially under excess light conditions (Gotor and Romero, 2013). Perhaps S-sulfocysteine acts as a mild oxidant to activate regulatory proteins such as STN7 in the lumen. Lipocalins are ligand binding proteins with simple ternary structures that can combine with small hydrophobic molecules (Charron et al., 2005). Seven lipocalins have been identified in plants. These proteins are categorized as temperature-induced lipocalins and chloroplast lipocalins (CHLs), such as VDE and zeaxanthin epoxidase

 

24

(ZEP), which are responsible for the xanthophyll cycle (Jahns et al., 2009). Only AtCHL, a CHL in Arabidopsis, is present in the lumen (Levesque-Tremblay et al., 2009). AtCHL is involved in photoprotection of thylakoid membrane lipids. AtCHL accumulates in the thylakoid lumen, protecting thylakoid membranes from peroxidation, especially when singlet oxygen is produced under high-light conditions (Boca et al., 2013). Peroxiredoxins (Prx) are thiol-dependent peroxidases that usually use Trx as electron donors (Moon et al., 2006). Prxs are present in many organisms ranging from eubacteria to mammals; their known biological functions include both oxidant defense and signal transduction (Wakita et al., 2007). Chloroplast Prx Q is located in the thylakoid lumen, but this protein does not appear function in ROS detoxification; its function requires further study.

5. Concluding remarks Redox signaling in the thylakoid lumen appears to regulate the turnover and functioning of lumenal proteins, which participate in photosynthesis and adaptation to high-light intensity. Several lumenal proteins in Arabidopsis are known to be disulfide bonded, and the discovery of trans-thylakoid redox pathways controlling disulfide bond formation and reduction highlights the impact of thiol-mediated signal transduction in the thylakoid lumen. Future studies should focus on expanding our knowledge of the following: (1) The roles of lumenal oxidants and electron acceptors. Both Cyt c6A and S-Cys are potential oxidants in the lumen. Cyt c6A has high midpoint potential, and S-Cys may act as mild oxidant, inducing redox changes. Glutathione

 

25

(GSH) is undetectable in the lumen, although it functions as an electron acceptor. The roles of ASC and phylloquinone in receiving electrons remain to be determined. ASC is an alternative electron acceptor when plants are short of plastocyanin. Phylloquinone is thought to be the final electron acceptor of the VKOR-like domain of LTO1. (2) Further elucidating the redox regulation network in the lumen. At least 30 proteins have lumenal thiols, but not all are targets of LTO1 and HCF164. Therefore, other Trx-like domain-containing proteins may exist in the lumen. The Trx-like domain is necessary for SOQ1 function, which implies that its redox potential is required to reduce these lumenal disulfide-bonded proteins. Thus, the next question to address is how SOQ1 prevents the occurrence of qI-type quenching of excitation energy in LHCII.

Acknowledgements This work was supported by the National Genetically Modified Organisms Breeding Major Projects of China (2009ZX08009-109B), the National Natural Science Foundation of China (30900882), the National Key Technology R&D Program of China (2011BAD35B 02-05), as well as the Public Experiment Center of State Bioindustrial Base (Chongqing), China.

 

26

References Aldridge, C., Cain, P., Robinson, C., 2009. Protein transport in organelles: Protein transport into and across the thylakoid membrane. FEBS J. 276, 1177-1186. Arnoux, P., Morosinotto, T., Saga, G., Bassi, R., Pignol, D., 2009. A structural basis for the pH-dependent xanthophyll cycle in Arabidopsis thaliana. Plant Cell 21, 2036-2044. Bader, M., Muse, W., Ballou, D.P., Gassner, C., Bardwell, J.C., 1999. Oxidative protein folding is driven by the electron transport system. Cell 98, 217-227. Balmer, Y., Koller, A., del Val, G., Manieri, W., Schurmann, P., Buchanan, B.B., 2003. Proteomics gives insight into the regulatory function of chloroplast thioredoxins. Proc. Natl. Acad. Sci. U. S. A. 100, 370-375. Bell, P.D., Xin, Y., Blankenship, R.E., 2009. Purification and characterization of cytochrome c(6) from Acaryochloris marina. Photosynth. Res. 102, 43-51. Bellafiore, S., Barneche, F., Peltier, G., Rochaix, J.D., 2005. State transitions and light adaptation require chloroplast thylakoid protein kinase STN7. Nature 433, 892-895. Bermudez, M.A., Galmes, J., Moreno, I., Mullineaux, P.M., Gotor, C., Romero, L.C., 2012. Photosynthetic adaptation to length of day is dependent on S-sulfocysteine synthase activity in the thylakoid lumen. Plant Physiol. 160, 274-288. Bermudez, M.A., Paez-Ochoa, M.A., Gotor, C., Romero, L.C., 2010. Arabidopsis S-sulfocysteine synthase activity is essential for chloroplast function and long-day light-dependent redox control. Plant Cell 22, 403-416. Bernal-Bayard, P., Hervas, M., Cejudo, F.J., Navarro, J.A., 2012. Electron transfer pathways and dynamics of chloroplast NADPH-dependent thioredoxin reductase C (NTRC). J. Biol. Chem. 287, 33865-33872. Berndt, C., Lillig, C.H., Holmgren, A., 2008. Thioredoxins and glutaredoxins as facilitators of protein folding. Biochim. Biophys. Acta 1783, 641-650. Betterle, N., Ballottari, M., Baginsky, S., Bassi, R., 2015. High light-dependent phosphorylation of photosystem II inner antenna CP29 in monocots is STN7  

27

independent and enhances nonphotochemical quenching. Plant Physiol. 167, 457-471. Boca, S., Koestler, F., Ksas, B., Chevalier, A., Leymarie, J., Fekete, A., et al., 2013. Arabidopsis lipocalins AtCHL and AtTIL have distinct but overlapping functions essential for lipid protection and seed longevity. Plant Cell Environ. 37, 368-381. Bonardi, V., Pesaresi, P., Becker, T., Schleiff, E., Wagner, R., Pfannschmidt, T., et al., 2005. Photosystem II core phosphorylation and photosynthetic acclimation require two different protein kinases. Nature 437, 1179-1182. Bonnard, G., Corvest, V., Meyer, E.H., Hamel, P.P., 2010. Redox processes controlling the biogenesis of c-type cytochromes. Antioxid. Redox Signal. 13, 1385-1401. Brooks, M.D., Sylak-Glassman, E.J., Fleming, G.R., Niyogi, K.K., 2013. A thioredoxin-like/beta-propeller protein maintains the efficiency of light harvesting in Arabidopsis. Proc. Natl. Acad. Sci. U. S. A. 110, E2733-2740. Buchanan, B.B., Luan, S., 2005. Redox regulation in the chloroplast thylakoid lumen: a new frontier in photosynthesis research. J. Exp. Bot. 56, 1439-1447. Cain, P., Hall, M., Schroder, W.P., Kieselbach, T., Robinson, C., 2009. A novel extended family of stromal thioredoxins. Plant Mol. Biol. 70, 273-281. Charron, J.B., Ouellet, F., Pelletier, M., Danyluk, J., Chauve, C., Sarhan, F., 2005. Identification, expression, and evolutionary analyses of plant lipocalins. Plant Physiol. 139, 2017-2028. Chida, H., Yokoyama, T., Kawai, F., Nakazawa, A., Akazaki, H., Takayama, Y., et al., 2006. Crystal structure of oxidized cytochrome c(6A) from Arabidopsis thaliana. FEBS Lett. 580, 3763-3768. Cruz, J.A., Sacksteder, C.A., Kanazawa, A., Kramer, D.M., 2001. Contribution of electric field (Delta psi) to steady-state transthylakoid proton motive force (pmf) in vitro and in vivo. Control of pmf parsing into Delta psi and Delta pH by ionic strength. Biochemistry 40, 1226-1237. Depuydt, M., Messens, J., Collet, J.F., 2011. How proteins form disulfide bonds. Antioxid. Redox Signal. 15, 49-66. Dutton, R.J., Wayman, A., Wei, J.R., Rubin, E.J., Beckwith, J., Boyd, D., 2010.  

28

Inhibition of bacterial disulfide bond formation by the anticoagulant warfarin. Proc. Natl. Acad. Sci. U. S. A. 107, 297-301. Edvardsson, A., Shapiguzov, A., Petersson, U.A., Schroder, W.P., Vener, A.V., 2007. Immunophilin AtFKBP13 sustains all peptidyl-prolyl isomerase activity in the thylakoid

lumen

from

Arabidopsis

thaliana

deficient

in

AtCYP20-2.

Biochemistry 46, 9432-9442. Foyer, C.H., Noctor, G., 2009. Redox regulation in photosynthetic organisms: signaling, acclimation, and practical implications. Antioxid. Redox Signal. 11, 861-905. Friso, G., Giacomelli, L., Ytterberg, A.J., Peltier, J.B., Rudella, A., Sun, Q., et al., 2004. In-depth analysis of the thylakoid membrane proteome of Arabidopsis thaliana chloroplasts: new proteins, new functions, and a plastid proteome database. Plant Cell 16, 478-499. Fufezan, C., Simionato, D., Morosinotto, T., 2012. Identification of key residues for pH dependent activation of violaxanthin de-epoxidase from Arabidopsis thaliana. PLoS One 7, e35669. Furt, F., Oostende, C., Widhalm, J.R., Dale, M.A., Wertz, J., Basset, G.J., 2010. A bimodular oxidoreductase mediates the specific reduction of phylloquinone (vitamin K(1)) in chloroplasts. Plant J. 64, 38-46. Gabilly, S.T., Kropat, J., Karamoko, M., Page, M.D., Nakamoto, S.S., Merchant, S.S., et al., 2011. A novel component of the disulfide-reducing pathway required for cytochrome c assembly in plastids. Genetics 187, 793-802. Gollan, P.J., Bhave, M., 2010. Genome-wide analysis of genes encoding FK506-binding proteins in rice. Plant Mol. Biol. 72, 1-16. Gopalan, G., He, Z., Balmer, Y., Romano, P., Gupta, R., Heroux, A., et al., 2004. Structural analysis uncovers a role for redox in regulating FKBP13, an immunophilin of the chloroplast thylakoid lumen. Proc. Natl. Acad. Sci. U. S. A. 101, 13945-13950. Gopalan, G., He, Z., Battaile, K.P., Luan, S., Swaminathan, K., 2006. Structural comparison of oxidized and reduced FKBP13 from Arabidopsis thaliana.  

29

Proteins 65, 789-795. Goss, R., Lepetit, B., 2015. Biodiversity of NPQ. J. Plant Physiol. 172, 13-32. Gotor, C., Alvarez, C., Bermudez, M.A., Moreno, I., Garcia, I., Romero, L.C., 2010. Low abundance does not mean less importance in cysteine metabolism. Plant Signal. Behav. 5, 1028-1030. Gotor, C., Romero, L.C., 2013. S-sulfocysteine synthase function in sensing chloroplast redox status. Plant Signal. Behav. 8, e23313. Granlund, I., Hall, M., Kieselbach, T., Schroder, W.P., 2009. Light induced changes in protein expression and uniform regulation of transcription in the thylakoid lumen of Arabidopsis thaliana. PLoS One 4, e5649. Grauschopf, U., Winther, J.R., Korber, P., Zander, T., Dallinger, P., Bardwell, J.C., 1995. Why is DsbA such an oxidizing disulfide catalyst? Cell 83, 947-955. Gupta, R., He, Z., Luan, S., 2002a. Functional relationship of cytochrome c(6) and plastocyanin in Arabidopsis. Nature 417, 567-571. Gupta, R., Mould, R.M., He, Z., Luan, S., 2002b. A chloroplast FKBP interacts with and affects the accumulation of Rieske subunit of cytochrome bf complex. Proc. Natl. Acad. Sci. U. S. A. 99, 15806-15811. Haddadian, E.J., Gross, E.L., 2006. A Brownian dynamics study of the effects of cytochrome f structure and deletion of its small domain in interactions with cytochrome c6 and plastocyanin in Chlamydomonas reinhardtii. Biophys. J. 90, 566-577. Hall, M., Mata-Cabana, A., Akerlund, H.E., Florencio, F.J., Schroder, W.P., Lindahl, M., et al., 2010. Thioredoxin targets of the plant chloroplast lumen and their implications for plastid function. Proteomics 10, 987-1001. Hallin, E.I., Guo, K., Akerlund, H.E., 2015. Violaxanthin de-epoxidase disulphides and their role in activity and thermal stability. Photosynth. Res. 589, 919-923. He, Z., Li, L., Luan, S., 2004. Immunophilins and parvulins. Superfamily of peptidyl prolyl isomerases in Arabidopsis. Plant Physiol. 134, 1248-1267. Hell, K., 2008. The Erv1-Mia40 disulfide relay system in the intermembrane space of mitochondria. Biochim. Biophys. Acta 1783, 601-609.  

30

Heras, B., Shouldice, S.R., Totsika, M., Scanlon, M.J., Schembri, M.A., Martin, J.L., 2009. DSB proteins and bacterial pathogenicity. Nat. Rev. Microbiol. 7, 215-225. Herrmann, J.M., Kauff, F., Neuhaus, H.E., 2009. Thiol oxidation in bacteria, mitochondria and chloroplasts: common principles but three unrelated machineries? Biochim. Biophys. Acta 1793, 71-77. Hieber, A.D., Bugos, R.C., Yamamoto, H.Y., 2000. Plant lipocalins: violaxanthin de-epoxidase and zeaxanthin epoxidase. Biochim. Biophys. Acta 1482, 84-91. Hiniker, A., Bardwell, J.C., 2004. In vivo substrate specificity of periplasmic disulfide oxidoreductases. J. Biol. Chem. 279, 12967-12973. Howe, C.J., Schlarb-Ridley, B.G., Wastl, J., Purton, S., Bendall, D.S., 2006. The novel cytochrome c6 of chloroplasts: a case of evolutionary bricolage? J. Exp. Bot. 57, 13-22. Inaba, K., Murakami, S., Suzuki, M., Nakagawa, A., Yamashita, E., Okada, K., et al., 2006. Crystal structure of the DsbB-DsbA complex reveals a mechanism of disulfide bond generation. Cell 127, 789-801. Ingelsson, B., Shapiguzov, A., Kieselbach, T., Vener, A.V., 2009. Peptidyl-prolyl isomerase activity in chloroplast thylakoid lumen is a dispensable function of immunophilins in Arabidopsis thaliana. Plant Cell Physiol. 50, 1801-1814. Jahns, P., Latowski, D., Strzalka, K., 2009. Mechanism and regulation of the violaxanthin cycle: the role of antenna proteins and membrane lipids. Biochim. Biophys. Acta 1787, 3-14. Jarvi, S., Gollan, P.J., Aro, E.M., 2013. Understanding the roles of the thylakoid lumen in photosynthesis regulation. Front. Plant Sci. 4, 434. Kadokura, H., Beckwith, J., 2010. Mechanisms of oxidative protein folding in the bacterial cell envelope. Antioxid. Redox Signal. 13, 1231-1246. Kana, R., Prasil, O., Mullineaux, C.W., 2009. Immobility of phycobilins in the thylakoid lumen of a cryptophyte suggests that protein diffusion in the lumen is very restricted. FEBS Lett. 583, 670-674. Karamoko, M., Cline, S., Redding, K., Ruiz, N., Hamel, P.P., 2011. Lumen Thiol Oxidoreductase1, a disulfide bond-forming catalyst, is required for the assembly  

31

of photosystem II in Arabidopsis. Plant Cell 23, 4462-4475. Karamoko, M., Gabilly, S.T., Hamel, P.P., 2013. Operation of trans-thylakoid thiol-metabolizing pathways in photosynthesis. Front. Plant Sci. 4, 476. Kieselbach, T., 2013. Oxidative folding in chloroplasts. Antioxid. Redox Signal. 19, 72-82. Kieselbach, T., Hagman, Andersson, B., Schroder, W.P., 1998. The thylakoid lumen of chloroplasts. Isolation and characterization. J. Biol. Chem. 273, 6710-6716. Kieselbach, T., Schroder, W.P., 2003. The proteome of the chloroplast lumen of higher plants. Photosynth. Res. 78, 249-264. Kirchhoff, H., Hall, C., Wood, M., Herbstova, M., Tsabari, O., Nevo, R., et al., 2011. Dynamic control of protein diffusion within the granal thylakoid lumen. Proc. Natl. Acad. Sci. U. S. A. 108, 20248-20253. Kramer, D.M., Sacksteder, C.A., Cruz, J.A., 1999. How acidic is the lumen? Photosynth. Res. 60, 151-163. Krupp, R., Chan, C., Missiakas, D., 2001. DsbD-catalyzed transport of electrons across the membrane of Escherichia coli. J. Biol. Chem. 276, 3696-3701. Lennartz, K., Bossmann, S., Westhoff, P., Bechtold, N., Meierhoff, K., 2006. HCF153, a novel nuclear-encoded factor necessary during a post-translational step in biogenesis of the cytochrome bf complex. Plant J. 45, 101-112. Lennartz, K., Plucken, H., Seidler, A., Westhoff, P., Bechtold, N., Meierhoff, K., 2001. HCF164 encodes a thioredoxin-like protein involved in the biogenesis of the cytochrome b(6)f complex in Arabidopsis. Plant Cell 13, 2539-2551. Levesque-Tremblay, G., Havaux, M., Ouellet, F., 2009. The chloroplastic lipocalin AtCHL prevents lipid peroxidation and protects Arabidopsis against oxidative stress. Plant J. 60, 691-702. Li, Z., Ahn, T.K., Avenson, T.J., Ballottari, M., Cruz, J.A., Kramer, D.M., et al., 2009. Lutein accumulation in the absence of zeaxanthin restores nonphotochemical quenching in the Arabidopsis thaliana npq1 mutant. Plant Cell 21, 1798-1812. Lima, A., Lima, S., Wong, J.H., Phillips, R.S., Buchanan, B.B., Luan, S., 2006. A redox-active FKBP-type immunophilin functions in accumulation of the  

32

photosystem II supercomplex in Arabidopsis thaliana. Proc. Natl. Acad. Sci. U. S. A. 103, 12631-12636. Lindahl, M., Kieselbach, T., 2009. Disulphide proteomes and interactions with thioredoxin on the track towards understanding redox regulation in chloroplasts and cyanobacteria. J. Proteomics 72, 416-438. Loedige, I., Stotz, M., Qamar, S., Kramer, K., Hennig, J., Schubert, T., et al., 2014. The NHL domain of BRAT is an RNA-binding domain that directly contacts the hunchback mRNA for regulation. Genes Dev. 28, 749-764. Lu, Y., Du, J.J., Yu, Z.B., Peng, J.J., Xu, J.N., Wang, X.Y., 2014. Identification of Potential Targets for Thylakoid Oxidoreductase AtVKOR/LTO1 in Chloroplasts. Protein Pept. Lett. 22, 219-225. Lu, Y., Wang, H.R., Li, H., Cui, H.R., Feng, Y.G., Wang, X.Y., 2013. A chloroplast membrane protein LTO1/AtVKOR involving in redox regulation and ROS homeostasis. Plant Cell Rep. 32, 1427-1440. Marcaida, M.J., Schlarb-Ridley, B.G., Worrall, J.A., Wastl, J., Evans, T.J., Bendall, D.S., et al., 2006. Structure of cytochrome c6A, a novel dithio-cytochrome of Arabidopsis thaliana, and its reactivity with plastocyanin: implications for function. J. Mol. Biol. 360, 968-977. Marchand, C., Le Marechal, P., Meyer, Y., Decottignies, P., 2006. Comparative proteomic approaches for the isolation of proteins interacting with thioredoxin. Proteomics 6, 6528-6537. Martinsuo, P., Pursiheimo, S., Aro, E.M., Rintamaki, E., 2003. Dithiol oxidant and disulfide reductant dynamically regulate the phosphorylation of light-harvesting complex II proteins in thylakoid membranes. Plant Physiol. 133, 37-46. Michalska, J., Zauber, H., Buchanan, B.B., Cejudo, F.J., Geigenberger, P., 2009. NTRC links built-in thioredoxin to light and sucrose in regulating starch synthesis in chloroplasts and amyloplasts. Proc. Natl. Acad. Sci. U. S. A. 106, 9908-9913. Moon, J.C., Jang, H.H., Chae, H.B., Lee, J.R., Lee, S.Y., Jung, Y.J., et al., 2006. The C-type Arabidopsis thioredoxin reductase ANTR-C acts as an electron donor to  

33

2-Cys peroxiredoxins in chloroplasts. Biochem. Biophys. Res. Commun. 348, 478-484. Motohashi, K., Hisabori, T., 2006. HCF164 receives reducing equivalents from stromal thioredoxin across the thylakoid membrane and mediates reduction of target proteins in the thylakoid lumen. J. Biol. Chem. 281, 35039-35047. Motohashi, K., Hisabori, T., 2010. CcdA is a thylakoid membrane protein required for the transfer of reducing equivalents from stroma to thylakoid lumen in the higher plant chloroplast. Antioxid. Redox Signal. 13, 1169-1176. Motohashi, K., Kondoh, A., Stumpp, M.T., Hisabori, T., 2001. Comprehensive survey of proteins targeted by chloroplast thioredoxin. Proc. Natl. Acad. Sci. U. S. A. 98, 11224-11229. Muller-Moule, P., Conklin, P.L., Niyogi, K.K., 2002. Ascorbate deficiency can limit violaxanthin de-epoxidase activity in vivo. Plant Physiol. 128, 970-977. Muller, P., Li, X.P., Niyogi, K.K., 2001. Non-photochemical quenching. A response to excess light energy. Plant Physiol. 125, 1558-1566. Nilkens, M., Kress, E., Lambrev, P., Miloslavina, Y., Muller, M., Holzwarth, A.R., et al., 2010. Identification of a slowly inducible zeaxanthin-dependent component of non-photochemical quenching of chlorophyll fluorescence generated under steady-state conditions in Arabidopsis. Biochim. Biophys. Acta 1797, 466-475. Onoa, B., Schneider, A.R., Brooks, M.D., Grob, P., Nogales, E., Geissler, P.L., et al., 2014. Atomic force microscopy of photosystem II and its unit cell clustering quantitatively delineate the mesoscale variability in Arabidopsis thylakoids. PLoS One 9, e101470. Page, M.L., Hamel, P.P., Gabilly, S.T., Zegzouti, H., Perea, J.V., Alonso, J.M., et al., 2004. A homolog of prokaryotic thiol disulfide transporter CcdA is required for the assembly of the cytochrome b6f complex in Arabidopsis chloroplasts. J. Biol. Chem. 279, 32474-32482. Peltier, J.B., Emanuelsson, O., Kalume, D.E., Ytterberg, J., Friso, G., Rudella, A., et al., 2002. Central functions of the lumenal and peripheral thylakoid proteome of Arabidopsis determined by experimentation and genome-wide prediction. Plant  

34

Cell 14, 211-236. Peltier, J.B., Friso, G., Kalume, D.E., Roepstorff, P., Nilsson, F., Adamska, I., et al., 2000. Proteomics of the chloroplast: systematic identification and targeting analysis of lumenal and peripheral thylakoid proteins. Plant Cell 12, 319-341. Peng, L., Fukao, Y., Fujiwara, M., Takami, T., Shikanai, T., 2009. Efficient operation of NAD(P)H dehydrogenase requires supercomplex formation with photosystem I via minor LHCI in Arabidopsis. Plant Cell 21, 3623-3640. Pesaresi, P., Hertle, A., Pribil, M., Kleine, T., Wagner, R., Strissel, H., et al., 2009. Arabidopsis STN7 kinase provides a link between short- and long-term photosynthetic acclimation. Plant Cell 21, 2402-2423. Pesaresi, P., Pribil, M., Wunder, T., Leister, D., 2011. Dynamics of reversible protein phosphorylation in thylakoids of flowering plants: the roles of STN7, STN8 and TAP38. Biochim. Biophys. Acta 1807, 887-896. Pfundel, E.E., Dilley, R.A., 1993. The pH Dependence of Violaxanthin Deepoxidation in Isolated Pea Chloroplasts. Plant Physiol. 101, 65-71. Ratajczak, T., Ward, B.K., Minchin, R.F., 2003. Immunophilin chaperones in steroid receptor signalling. Curr. Top. Med. Chem. 3, 1348-1357. Rockholm, D.C., Yamamoto, H.Y., 1996. Violaxanthin de-epoxidase. Plant Physiol. 110, 697-703. Romano, P., Gray, J., Horton, P., Luan, S., 2005. Plant immunophilins: functional versatility beyond protein maturation. New Phytol. 166, 753-769. Romero, L.C., Aroca, M.A., Laureano-Marin, A.M., Moreno, I., Garcia, I., Gotor, C., 2014. Cysteine and cysteine-related signaling pathways in Arabidopsis thaliana. Mol. Plant 7, 264-276. Ruban, A.V., Belgio, E., 2014. The relationship between maximum tolerated light intensity and photoprotective energy dissipation in the photosynthetic antenna: chloroplast gains and losses. Philos. Trans. R. Soc. Lond. B Biol. Sci. 369, 20130222. Sanders, C., Turkarslan, S., Lee, D.W., Daldal, F., 2010. Cytochrome c biogenesis: the Ccm system. Trends Microbiol. 18, 266-274.  

35

Schlarb-Ridley, B.G., Nimmo, R.H., Purton, S., Howe, C.J., Bendall, D.S., 2006. Cytochrome c(6A) is a funnel for thiol oxidation in the thylakoid lumen. FEBS Lett. 580, 2166-2169. Schubert, M., Petersson, U.A., Haas, B.J., Funk, C., Schroder, W.P., Kieselbach, T., 2002. Proteome map of the chloroplast lumen of Arabidopsis thaliana. J. Biol. Chem. 277, 8354-8365. Serrato, A.J., Fernandez-Trijueque, J., Barajas-Lopez, J.D., Chueca, A., Sahrawy, M., 2013. Plastid thioredoxins: a "one-for-all" redox-signaling system in plants. Front. Plant Sci. 4, 463. Shapiguzov, A., Edvardsson, A., Vener, A.V., 2006. Profound redox sensitivity of peptidyl-prolyl isomerase activity in Arabidopsis thylakoid lumen. FEBS Lett. 580, 3671-3676. Simionato, D., Basso, S., Zaffagnini, M., Lana, T., Marzotto, F., Trost, P., et al., 2015. Protein redox regulation in the thylakoid lumen: The importance of disulfide bonds for violaxanthin de-epoxidase. FEBS Lett. 589, 919-923. Singh, A.K., Bhattacharyya-Pakrasi, M., Pakrasi, H.B., 2008. Identification of an atypical membrane protein involved in the formation of protein disulfide bonds in oxygenic photosynthetic organisms. J. Biol. Chem. 283, 15762-15770. Soll, J., Schleiff, E., 2004. Protein import into chloroplasts. Nat. Rev. Mol. Cell Biol. 5, 198-208. Spetea, C., Hundal, T., Lundin, B., Heddad, M., Adamska, I., Andersson, B., 2004. Multiple evidence for nucleotide metabolism in the chloroplast thylakoid lumen. Proc. Natl. Acad. Sci. U. S. A. 101, 1409-1414. Spetea, C., Lundin, B., 2012. Evidence for nucleotide-dependent processes in the thylakoid lumen of plant chloroplasts--an update. FEBS Lett. 586, 2946-2954. Spinola, M.C., Perez-Ruiz, J.M., Pulido, P., Kirchsteiger, K., Guinea, M., Gonzalez, M., et al., 2008. NTRC new ways of using NADPH in the chloroplast. Physiol. Plant 133, 516-524. Standfuss, J., Terwisscha van Scheltinga, A.C., Lamborghini, M., Kuhlbrandt, W., 2005. Mechanisms of photoprotection and nonphotochemical quenching in pea  

36

light-harvesting complex at 2.5 A resolution. EMBO J. 24, 919-928. Tie, J.K., Stafford, D.W., 2008. Structure and function of vitamin K epoxide reductase. Vitam. Horm. 78, 103-130. Tikhonov, A.N., 2013. pH-dependent regulation of electron transport and ATP synthesis in chloroplasts. Photosynth. Res. 116, 511-534. Vasudevan, D., Gopalan, G., Kumar, A., Garcia, V.J., Luan, S., Swaminathan, K., 2014. Plant immunophilins: a review of their structure-function relationship. Biochim. Biophys. Acta 1850, 2145-2158. Wakita, M., Masuda, S., Motohashi, K., Hisabori, T., Ohta, H., Takamiya, K., 2007. The significance of type II and PrxQ peroxiredoxins for antioxidative stress response in the purple bacterium Rhodobacter sphaeroides. J. Biol. Chem. 282, 27792-27801. Wang, X., Dutton, R.J., Beckwith, J., Boyd, D., 2011. Membrane topology and mutational analysis of Mycobacterium tuberculosis VKOR, a protein involved in disulfide bond formation and a homologue of human vitamin K epoxide reductase. Antioxid. Redox Signal. 14, 1413-1420. Worrall, J.A., Luisi, B.F., Schlarb-Ridley, B.G., Bendall, D.S., Howe, C.J., 2008. Cytochrome c6A: discovery, structure and properties responsible for its low haem redox potential. Biochem. Soc. Trans. 36, 1175-1179. Worrall, J.A., Schlarb-Ridley, B.G., Reda, T., Marcaida, M.J., Moorlen, R.J., Wastl, J., et al., 2007. Modulation of heme redox potential in the cytochrome c6 family. J. Am. Chem. Soc. 129, 9468-9475. Wunder, T., Liu, Q., Aseeva, E., Bonardi, V., Leister, D., Pribil, M., 2013. Control of STN7 transcript abundance and transient STN7 dimerisation are involved in the regulation of STN7 activity. Planta 237, 541-558. Yano, H., Wong, J.H., Lee, Y.M., Cho, M.J., Buchanan, B.B., 2001. A strategy for the identification of proteins targeted by thioredoxin. Proc. Natl. Acad. Sci. U. S. A. 98, 4794-4799.

 

37

Figure Captions

 

 

38

Fig. 1. Redox regulation in the thylakoid lumen plays important roles in the optimization of photosynthesis. The activities of several lumenal proteins are regulated by redox signals in the lumen. Chloroplast Trx-m functions as an electron donor for thylakoid lumen proteins. Trans-thylakoid thiol-metabolizing pathways control disulfide bond formation and reduction of lumenal proteins. Redox reactions also regulate thylakoid lumen proteins, functioning in the response to oxidative stress to protect the photosystem apparatus. Reducing equivalents are transferred to SOQ1, either directly or through CCDA, while its targets remain unclear. Proteins with lumenal disulfide bonds that are involved in biogenesis of the photosystem complex are indicated with an oval. The oxidants of underlined proteins have not yet been determined. Solid lines indicate that redox reactions between components have been experimentally determined, while dashed lines represent candidate substrates. S-Cys, S-sulfocysteine.

 

39

Tables Table 1 Components in the thylakoid lumen with disulfide bonds Cysteines Protein a

Accession

Trx

HCF164

targets b

targets c

LTO1 Phosphorylation in mature

Function targets d

References

site e proteins f

Assembly of Cyp38

At3g01480

Yes

─

─

─

1

(Sirpio et al. 2008)

─

─

─

─

2

(Kim et al. 2011)

Yes

─

Yes

─

4

(Gupta et al. 2002)

─

─

─

─

4

(Peng et al. 2009)

Yes

─

Yes

─

2

(Lima et al. 2006)

Yes

─

─

─

2

PSII Subunit of Cyp20-2

At5g13120 NDH Accumulation

FKBP13

At5g45680 of Rieske Subunit of

FKBP16-2

At4g39710 NDH Assembly of

FKBP20-2

At3g60370

PSII supercomplex Antioxidant?

PrxQ

At3g26060

(Petersson et al.

Signaling?

2006) (Lennartz et al.

Assembly of HCF164

Yes

At5g23120

─

Yes

─

3

2001; Motohashi and

Cyt b6f Hisabori 2006) (Page et al. 2004; Assembly of CCDA

AT5G54290

Yes

─

─

─

4

Motohashi and

Cyt b6f Hisabori 2010) SOQ1

At1g56500

LTO1

At4g35760

Photoprotection

Yes

─

─

─

2

─

─

─

─

8

Assembly of

(Karamoko et al.

PSII

2011)

Electron Cyt c6a

At5g45040

 

(Marcaida et al. ─

carrier?

(Brooks et al. 2013)

─

Yes

─

2 2006)

40

VDE

At1g08550

Photoprotection

Yes

─

Yes

─

13

(Arnoux et al. 2009) (Bermudez et al.

ROS CS26

At3g03630

─

─

─

─

2

2010; Bermudez et

detoxiation al. 2012) ROS CHL

At3g47860

(Levesque-Tremblay ─

─

─

─

12

detoxiation

et al. 2009)

Degradation of Deg1

At3g27925

Yes

─

─

─

1

(Sun et al. 2010)

Yes

─

─

─

2

(Sun et al. 2007)

Yes

─

─

─

2

(Hall et al. 2010)

Yes

─

─

─

2

(Lundin et al. 2007)

D1 Degradation of Deg5

At4g18370 D1 Oxygen

PsbO1

At5g66570 evolution Oxygen

PsbO2

At3g50820 evolution

TL15

At2g44920

Unknow

Yes

─

─

─

2

(Ni et al. 2011)

TL17

AT5G53490

Unknow

Yes

─

Yes

─

4

(Lu et al. 2014b)

TL19

AT3G63535

Unknow

Yes

─

─

1

(Friso et al. 2004)

TL20.3

AT1G12250

Unknow

Yes

─

─

4

(Schubert et al. Yes

2002) (Granlund et al. TL29

AT4G09010

Unknow

Yes

─

─

─

3

2009b; Lundberg et al. 2011)

Regulation of PsaN

AT5G64040

Yes

Yes

─

Yes

4

─

Yes

─

─

2

(Amunts et al. 2007)

PSI activity? (Gupta et al. 2002;

Electron Rieske

AT4G03280 transfer

 

Gollan et al. 2011)

STN7

AT1G68830

State transition

Yes

─

─

Yes

2

(Pesaresi et al. 2009)

PPD6

AT3G56650

Unknow

─

─

─

─

2

(Hall et al. 2012)

PsbP1

AT1G06680

Oxygen

Yes

─

─

Yes

1

(Allahverdiyeva et

41

evolution

al. 2013)

20 kDa PsbP

(Kleffmann et al. AT3G56650

Unknow

Yes

─

─

─

2

domain

2004)

protein (Pagnussat et al. EDA3

At2g34860

Unknow

─

─

Yes

─

11 2005)

CtD1

AT4G17740

Proteolysis

─

─

─

─

5

CtD1-like

AT5G46390

Unknow

─

─

─

─

4

(Che et al. 2013) (Schubert et al. 2002)

Starch APE1

At5g38660

─

─

─

─

4

─

─

─

─

2

(Ferro et al. 2010)

biosynthetic Thylakoid Thylakoid (Schubert et al.

lumenal At5g52970

membrane

2002)

15.0 kDa organization protein Maintenance of VIPP1

At1g65260

─

─

─

─

6

─

─

─

─

2

(Kroll et al. 2001)

thylakoids (Rutschow et al. MYM9

At3g23700

RNA binding

2008) a

See references (Hall et al. 2010; Jarvi et al. 2013; Karamoko et al. 2013);

b c

See reference (Motohashi and Hisabori 2006);

d e f

See reference (Hall et al. 2010);

See references (Karamoko et al. 2011; Lu et al. 2014a);

See reference (Spetea and Lundin 2012);

See references (Hall et al. 2010; Spetea and Lundin 2012; Jarvi et al. 2013; Karamoko et al. 2013;

Lu et al. 2014a).

 

42