Functional redox links between lumen thiol oxidoreductase1 and serine/threonine-protein kinase STN7.

In response to changing light quantity and quality, photosynthetic organisms perform state transitions, a process which optimizes photosynthetic yield and mitigates photo-damage. The serine/threonine-protein kinase STN7 phosphorylates the light-harvesting complex of photosystem II (PSII; light-harvesting complex II), which then migrates from PSII to photosystem I (PSI), thereby rebalancing the light excitation energy between the photosystems and restoring the redox poise of the photosynthetic electron transport chain. Two conserved cysteines forming intra- or intermolecular disulfide bonds in the lumenal domain (LD) of STN7 are essential for the kinase activity although it is still unknown how activation of the kinase is regulated. In this study, we show lumen thiol oxidoreductase 1 (LTO1) is co-expressed with STN7 in Arabidopsis (Arabidopsis thaliana) and interacts with the LD of STN7 in vitro and in vivo. LTO1 contains thioredoxin (TRX)-like and vitamin K epoxide reductase domains which are related to the disulfide-bond formation system in bacteria. We further show that the TRX-like domain of LTO1 is able to oxidize the conserved lumenal cysteines of STN7 in vitro. In addition, loss of LTO1 affects the kinase activity of STN7 in Arabidopsis. Based on these results, we propose that LTO1 helps to maintain STN7 in an oxidized active state in state 2 through redox interactions between the lumenal cysteines of STN7 and LTO1.

Introduction

Solar radiation is the energy source of all photosynthetic organisms, including algae, green plants, and photosynthetic bacteria on earth (Chmeliov et al., 2016). These organisms have evolved sophisticated mechanisms for maintaining optimal photosynthesis and for minimizing photo-damage in a constantly changing light environment (Rochaix, 2013, 2014). Energy-dependent nonphotochemical quenching (qE-NPQ) is triggered by light-induced acidification of the thylakoid lumen and promotes the dissipation of excess excitation energy as heat (Belgio et al., 2014; Goss and Lepetit, 2015). State transitions (STs) adjust the antenna cross-sections of the photosystems through the redistribution of mobile light-harvesting complex II (LHCII). This process rebalances the light excitation energy between photosystem I (PSI) and photosystem II PSII and restores the redox poise of the electron transport chain (Fork and Satoh, 1986; Bellafiore et al., 2005; Rochaix, 2014; Cutolo et al., 2019).

In recent years, several molecular components and regulatory mechanisms of STs have been uncovered in Arabidopsis thaliana. One of the key components is the serine/threonine kinase STN7 which phosphorylates the mobile LHCII and is activated when plastoquinone (PQH2) binds to the Qo site of Cyt b6f complex following PSII over-excitation relative to PSI (Depege et al, 2003; Bellafiore et al., 2005). Under these conditions a portion of phosphorylated LHCII migrates from PSII to PSI, increasing the antenna size of PSI and rebalancing the excitation energy between PSII and PSI (state 2; Shapiguzov et al., 2016). Under far-red (FR) light the PQ pool is oxidized, STN7 is inactivated, and LHCII is dephosphorylated by the phosphatase PPH1/TAP38 (protein phosphatase 1/thylakoid-associated phosphatase 38) and moves back to PSII (state 1; Pribil et al., 2010; Shapiguzov et al., 2010; Rantala et al., 2016). While the PPH1/TAP38 phosphatase appears to be constitutively active, the activity of STN7 is redox-regulated through a complex process which is still largely unknown (Rochaix, 2007, 2013). STN7 has a transmembrane helix linking the kinase catalytic domain on the stromal side with its amino terminus containing two conserved cysteine residues in the lumenal domain (LD) of thylakoids (Lemeille et al., 2009; Bergner et al., 2015). Changes of these lumenal cysteines by site-directed mutagenesis demonstrated that they are essential for the catalytic activity and STs in Chlamydomonas (Chlamydomonas reinhardtii) and Arabidopsis (Lemeille et al., 2009; Wunder et al., 2013). It was also shown that the lumenal cysteines of STN7 form an intra-molecular disulfide bond in state 1 as well as in state 2. Moreover, it was proposed that during activation of STN7 the intra-molecular disulfide bond is converted transiently into two intermolecular disulfide bonds giving rise to STN7 dimers (Wunder et al., 2013; Shapiguzov et al., 2016). Thus a key question in the redox regulation of STN7 is to identify the factor(s) which maintain the oxidized state of the lumenal cysteines and induce the switch between intra- and intermolecular form of the disulfide bonds. In this work, we report that the known lumen thioredoxin (TRX) lumen thiol oxidoreductase1 (LTO1) interacts with STN7 through their LDs (Karamoko et al., 2011; Lu et al., 2013; Yu et al., 2014; Lu, 2015). Based on genetic and biochemical studies, we propose that the conserved cysteines in the LDs of STN7 and LTO1 are important for maintaining the oxidized state of STN7 required for kinase activity during STs.

Results

LTO1 has a similar expression pattern as STN7

The lumenal cysteines of Stt7/STN7 are essential for the catalytic activity and STs in Chlamydomonas and Arabidopsis (Lemeille et al., 2009; Wunder et al., 2013; Shapiguzov et al., 2016). We set out to identify the chloroplast redox component(s) involved in the redox regulation of the STN7 kinase. As a first step, we analyzed gene co-expression profiles of STN7 from the ATTED-II database (http://atted.jp/), searched the chloroplast TRXs and ranked them according to the co-expression index, Logit-score (LS; Obayashi et al., 2018). Because the regulatory domain of STN7 is on the lumenal side, six proteins with predicted lumenal Trx domains (HCF164 (high chlorophyll fluorescence 164), SOQ1 (suppressor of quenching 1), LTO1, AT2G37240, AT4G10000, and AT1G21350) were selected (Supplemental Table S1; Shapiguzov et al., 2016). Among these candidates, the first two proteins may act as negative regulators of STN7 under high light conditions, whereas for the last three no clear subcellular localization and function are known (Motohashi and Hisabori, 2006; Brooks et al., 2013; Kang and Wang, 2016). We compared the expression of LTO1, a known TRX protein, with that of genes related to STs, including STN7, LHCII phosphatase PPH1/TAP38, PSII core proteins kinase STN8 (state transition 8), and phosphatase PBCP (photosystem II core phosphatase). From the RNAseq database, we found that there is a high co-expression between LTO1 and the above genes with Pearson correlation coefficients ranging between 0.60 and 0.77 (Figure 1). The LS and correlation of RNAseq (Cor-r) of STN7 and other genes related to STs are higher than for PsbO1/2, which are the known targets of LTO1 (Table 1; Karamoko et al., 2011). A similar expression correlation pattern was found with the transmembrane proteins CcdA and HCF164 which belong to the trans-thylakoid thiol-reducing pathway and have been proposed to be involved in the inactivation of STN7 under high light conditions (Figure 1 and Table 1; Rochaix, 2013; Kang and Wang, 2016). Recently, a quantitative atlas of the transcriptomes and proteomes of 30 tissues of the model plant A. thaliana was established. It provides a powerful bioinformatics tool to explore the network of Arabidopsis proteins (Samaras et al., 2019; Mergner et al., 2020). Compared to PsbO1/2, protein co-expression of LTO1 and STN7 in a heat-map of the ProteomicsDB database is higher, and expression of LTO1 is also similar to that of PPH1/TAP38, STN8, and PBCP (Supplemental Figure S1). These observations raise the possibility that LTO1 participates in the redox-dependent regulation of STN7 activity during STs.

Figure 1

Co-expression analysis based on the RNAseq database of LTO1. Co-expression data with RNAseq of LTO1 were obtained from the latest version (10.1) of the ATTED-II website (http://atted.jp/). STN7, LHCII kinase. STN8, PSII core protein kinase. TAP38, LHCII phosphatase. PBCP, PSII core phosphatase. CcdA and HCF164 operate in the trans-thylakoid thiol-reducing pathway. PsbO1/2, subunits of PSII complex. The PCC value of RNAseq database (Cor-r) is indicated at the top of each panel. LTO1 with itself has an obvious linear correlation (Cor-r = 1.00).

Table 1

Co-expression indexes of LS and Pearson correlation coefficient in co-expression analysis

LTO1	LTO1	STN7	STN8	TAP38	PBCP	HCF164	CcdA	PsbO1	PsbO2
LS	14.2	3.7	3.9	5.6	6.0	5.0	5.9	3.1	2.3
Cor-r	1.00	0.687	0.720	0.755	0.719	0.768	0.602	0.694	0.584

Co-expression indexes of lto1 and indicated genes originate from the ATTED-II database and the pattern are shown in Figure 1 and Supplemental Figure S1.

LS; higher LS indicates higher co-expression, LS = 0 indicates no co-expression. Correlation (Cor) is the Pearson Correlation Coefficient (PCC), calculated from the RNAseq database A PCC value near 1 indicates high co-expression, 0 indicates no co-expression, and −1 indicates opposite expression.

LTO1 directly interacts with STN7 both in vitro and in vivo

To investigate the potential function of LTO1 in regulating STN7 activity, we first examined whether these two proteins interact with each other. STN7 is known to have one transmembrane helix that links its small N-terminal lumen domain with the catalytic domain on the stromal side of thylakoids (Supplemental Figure S2; Rochaix, 2013; Shapiguzov et al., 2016). In the LTO1 protein, four or five predicted transmembrane helices embrace a vitamin K epoxide reductase (VKOR) domain and the C-terminal TRX-like domain in the lumen (Supplemental Figure S3; Karamoko et al., 2011; Kieselbach, 2013; Onda, 2013). To test for interactions between STN7 and LTO1, an in vitro pull-down assay was performed with the recombinant LDs of STN7 and LTO1. After incubation of bait (LTO1-LD-glutathione S-transferase [GST] and empty GST agarose) with prey (STN7-LD-human myelin basic protein [MBP]-His), the bound proteins were eluted. Compared to the negative control (empty GST), the His-tag was detected only in the elution of LTO1-LD-GST (Figure 2A), indicating that STN7 interacts with LTO1 on the lumenal side of thylakoids.

Figure 2

LTO1 directly interacts with STN7 in vitro and in vivo. A, Pull-down assay with LTO1 and STN7. The lumen domain of STN7 fused to MBP-His was incubated with immobilized empty GST and GST-tagged LTO1 lumenal-domain. After washing three times, bound proteins were eluted and fractionated by Tricine–PAGE for staining (CBB) or SDS–PAGE for immunoblot analysis with His antibody. B, Split ubiquitin-based yeast two-hybrid analysis. The full-length LTO1 was fused to the N and C termini of Cub-LexA-VP16 fragment in the pNCW and pCCW vectors as the prey, respectively. The same strategy was used for STN7 as the bait, fused with the NubG fragment of the pDSLNx and pDL2xN plasmids at the N and C termini. NubG, negative control; NubI, positive control. Plasmid construction and yeast two-hybrid analysis were performed as described in “Materials and methods” section. Left, yeast colonies on synthetic dropout medium without Trp-Leu-His (SD-Trp-Leu-His); Right, staining with X-Gal of the left plate. C, Protein overlay analysis of LTO1 protein interactions. Overlay of the reconstructed LTO1-LD with His tag and STN7-HA. Blotting of stn7 mutant or STN7-HAtransgenic Arabidopsis thylakoid proteins (20 μg per lane) was followed by immunodetection with anti-His and anti-HA antibody. Input, thylakoid proteins of stn7 mutant and STN7-HA transgenic Arabidopsis; Control, blotting membrane incubated with empty buffer. LTO1-His, blotting membrane incubated with LTO1-His-fused proteins. D, Co-immunoprecipitation analysis. Thylakoid proteins of STN7-HA transgenic Arabidopsis were immunoprecipitated with nonimmune serum (control) or antibodies against LTO1, then analyzed by immunoblotting. Input indicates that 5 mg (50%) and 10 mg (100%) protein was loaded on the gel. lto1, negative control for LTO1 antibody; cross-linked-IP, immunoprecipitation with the cross-linker DSP; Wash, third washing buffer; Elution, the elution buffer with bound proteins.

Since the LD of STN7 (forty amino acids) is too short to be used in the common yeast two-hybrid system, we inserted the full-length cytidinediphosphate diacylglycerol synthase (CDS) of STN7 and LTO1 into the split ubiquitin yeast two-hybrid plasmids. The results obtained with this system indicate that the intact STN7 and LTO1 proteins interact with each other (Figure 2B). These results were further confirmed by performing a protein overlay assay. As shown in Figure 2C, recombinant LTO1-LD-His protein bound to STN7 from extracts of transgenic Arabidopsis plants expressing STN7-HA. However, no signal was detected in a parallel overlay experiment using extracts of the stn7 mutant, indicating that the LD of LTO1 can interact with full-length STN7. Co-immunoprecipitation (IP) analysis further supported an interaction between STN7 and LTO1 in vivo (Figure 2D). Cross-linking is an effective method for stabilizing protein interactions. The interaction was indeed stronger after incubation with dithiobis(succinimidyl propionate) (DSP), a suitable cross-linker reagent for membrane proteins (Figure 2D; cross-linked-IP; Trakselis et al., 2005). We conclude from these results that STN7 interacts with LTO1 on the lumenal side of thylakoids both in vitro and in vivo.

The TRX-like domain of LTO1 oxidizes the conserved lumenal cysteines of STN7

The co-expression data and our results on the interactions between STN7 and LTO1 raise the possibility that STN7 may be one of the redox targets of LTO1. To test this possibility, recombinant proteins consisting of the LD of LTO1 and STN7 carrying different tags were purified and used in the redox assays in vitro. LTO1-LD was first oxidized with oxygen and subsequently reduced with increasing amounts of dithiothreitol (DTT) in the presence of the reagent 4‐acetoamide‐4′‐maleimidylstilbene‐2,2′‐disulfonic acid (AMS) which alkylates free cysteine sulfhydryl groups and thereby increases the molecular mass of the target protein. After incubation for 90 min, the protein samples were analyzed by polyacrylamide gel electrophoresis (PAGE; Figure 3A). Prior to DTT addition the bands corresponding to the fully oxidized (ox1), partly oxidized (ox2), and partly reduced (red1) protein as well as dimer and oligomer could be detected. Upon incubation with increasing concentrations of DTT, the bands of ox1/ox2 and oligomer disappeared and the amount of the red1 band increased and an additional band, red2 (fully reduced) appeared. The appearance of four shifted bands is due to the fact that the recombinant LTO1-LD contains four conserved cysteines which can form inter- and/or intra-molecular disulfide bonds (Supplemental Figure S3). Because the LD of STN7 is too short for the redox experiment, we fused it to the MBP protein (45 kDa) that does not contain any cysteines, and added a tobacco etch virus (TEV) protease cleavage site as a flexible linker between the two domains. Compared to air-oxidized protein (lane 3), more dimer and oligomer appear in oxygen-oxidized STN7-LD-TEV-MBP protein (lane 4; Figure 3B). When this protein was incubated with increasing concentrations of DTT (lanes 5–7), the level of dimer and oligomer both diminished gradually. The shifted STN7 monomer band cannot be detected because the molecular mass of AMS (0.54 kDa) is very small compared to the protein mass (50 kDa). We, therefore, used the STN7 dimer and/or oligomer as an indicator of the oxidized state of STN7-LD.

Figure 3

The thioredoxin-like domain of LTO1 catalyzes disulfide bond formation in the lumen domain of STN7. Recombinant proteins (lumen domain of STN7 and LTO1) were treated with AMS, separated by nonreducing SDS–PAGE and stained with CBB. The different states of LTO1 and STN7 are indicated on the right of the figure. AMS is an alkylating reagent and AMS treatment of exposed thiols in reduced LTO1 and STN7 results in an increased molecular mass of the alkylated molecules. The conserved cysteines of STN7 and LTO1 in the lumen domains can form dimers and/or oligomers under oxidizing conditions. A, DTT-dependent reduction of the disulfides in LTO1. Lane 1, recombinant LTO1-lumen domain in reducing buffer as the control. Lane 2, marker. Lanes 3–7, oxygen-oxidized LTO1 was reduced with increasing concentrations of DTT for 90 min then loaded with nonreducing buffer. LTO1 monomer is present in several shifted bands corresponding to fully oxidized (ox1), partially oxidized (ox2), partially reduced (red1), fully reduced (red2) forms under different incubation conditions. The dimer and oligomer can also be detected. B, DTT-dependent reduction of the disulfide in STN7. Lane 1, recombinant STN7-lumen domain with MBP tag in reducing buffer as the control. Lanes 2 and 8, marker. Lanes 3–7, O2-oxidized STN7 was reduced with increasing concentrations of DTT for 90 min, then loaded with nonreducing buffer. The shifted and unshifted forms of the STN7 monomer cannot be resolved because the molecular weight of AMS (0.54 kDa) is very small compared to the fused protein (50 kDa). The dimer and oligomer of the fusion proteins can be detected under oxidizing conditions and gradually decrease with increasing concentration of DTT. C, LTO1-dependent oxidation of STN7 sulfhydryls. Lanes 1 and 10, the control in reducing buffer. Lanes 2 and 9, markers. Lanes 3 and 8, oxygen-oxidized LTO1 and STN7, respectively. Lanes 4 and 7, DTT-reduced LTO1 and STN7, respectively. Lanes 5 and 6, co-incubation of oxidized LTO1 and reduced STN7. The concentration (mg/mL) of loaded samples is shown above the figures.

To test the possibility that oxidized LTO1-LD catalyzes disulfide bond formation in STN7-LD, we co-incubated oxygen-oxidized LTO1-LD and reduced STN7-LD-TEV-MBP with two different concentrations (Figure 3C, lanes 5 and 6). Under these conditions, the dimer/oligomer of STN7-LD-TEV-MBP and the reduced forms of LTO1 (red1 and red2) distinctly increased, and the dimer of LTO1-LD disappeared (Figure 3C). These results indicate that the two conserved cysteines in the LD of STN7 can form disulfide bonds catalyzed by the TRX-like domain of LTO1.

Comparison of the sequences of STN7 and LTO1 from different photosynthetic organisms including algae (Ostreococcus lucimarinus and C. reinhardtii), moss (Physcomitrium patens), monocotyledons (Hordeum vulgare, Oryza sativa, Triticum aestivum, and Zea mays), and dicotyledons (A. thaliana and Nicotiana attenuata) reveals that amongst the six cysteines of AtSTN7, the two in the LD are the most conserved in all species. Moreover, the other two cysteines are conserved except for C. reinhardtii (Supplemental Figure S2). In the transmembrane regions of LTO1, four partially conserved cysteines belong to the VKOR domain (Supplemental Figure S3; Du et al., 2015). In the LD of LTO1, another four cysteines forming two CxxC motifs (Supplemental Figure S4) are also conserved although the last two cysteines are 15 amino acids apart (Supplemental Figure S3). To test the function of the cysteines in the LD in the redox assay, the lumenal conserved cysteines of LTO1 and STN7 were changed to alanine giving rise to the variants (4cm-LTO1 and 2cm-STN7). No shifted band of the monomers/dimer of 2cmSTN7 or 4cm-LTO1 was detected upon treatment with oxygen or DTT (Supplemental Figure S5, A and B). When 4cm-LTO1 was incubated with wild-type (WT) STN7-LD, no redox change of STN7 was observed (Supplemental Figure S5C; lanes 5 and 6) although this change occurred when STN7 was oxidized with O2 (Supplemental Figure S5C; lanes 7 and 8). Similarly, when reduced 2cmSTN7 was incubated with WT LTO1, no STN7 dimer was formed (Supplemental Figure S5D). In conclusion, these results suggest that the conserved cysteines in the LD of STN7 and LTO1 interact with each other and that they may be involved in the redox regulation of STN7.

Loss of LTO1 affects STs and LHCII phosphorylation in Arabidopsis

Since the above results showed that LTO1 can oxidize the lumenal Cys of STN7, we tested whether LTO1 plays a role in STs using the WT, stn7 and lto1 mutant (Supplemental Figure S6). First, we checked STs as depicted in Supplemental Figure S7 by monitoring fluorescence changes. Seedlings were dark-adapted for 30 min and illuminated with blue light and after 12 min with FR light. This treatment allows one to estimate (Fs1–Fs2)/Fs1 where Fs1 and Fs2 are the fluorescence values at the end of the blue light illumination and after switching on FR light, respectively (Shapiguzov et al., 2010; Shapiguzov et al., 2016). Although this measurement does not provide a direct estimation of ST, it corresponds well with the extent of STs and can be used as an indicator (Shapiguzov et al., 2016). Brightfield images, chlorophyll fluorescence, and STs values given on a false-color scale of WT, stn7 and lto1 are shown in Supplemental Figure S7, A–C. When grown on Murashige–Skoog (MS)-medium with sugar, 1- and 2-week-old lto1 mutant seedlings display a reduced signal compared to WT with this assay for ST. The quantum yield of PSII (Fv/Fm fluorescence) is lower than that in WT and stn7 (Supplemental Figure S7, A and B), which is consistent with a previous report (Karamoko et al., 2011). When grown in the soil under photo-autotrophic conditions, 4-week-old plants show a similar phenotype (Supplemental Figure S7C). Fluorescence traces of STs for WT (Supplemental Figure S7D), stn7 (Figure S7E), and lto1 (Figure S7F) illustrate these differences further. To eliminate the possibility that the decreased STs in lto1 is an indirect consequence of the lower Fv/Fm value in this mutant, we checked several mutants with decreased PSII activity, including ffc (lacking LHC transport function; Ouyang et al., 2011), and ohp2-R (affected in PSII assembly; Li et al., 2019). Consistent with previous research, these three mutants have a lower Fv/Fm ratio, but ST is similar to that of WT as measured with this assay with false-color (Supplemental Figure S8A) and fluorescence traces (Supplemental Figure S8B).

A characteristic feature of STs is the occurrence of LHCII phosphorylation in state 2, a process that is significantly decreased in stn7 (Bellafiore et al., 2005; Wunder et al., 2013). To test this, the seedlings of WT, stn7 and lto1 were treated with FR light for 90 min (state 1) and subsequently with white light for 90 min (state 2). Thylakoid membrane proteins from the seedlings were extracted and immunoblotted with phospho-threonine antibodies. Compared to the WT, the phosphorylation level of LHCII was significantly decreased in lto1 (Figure 4A). In these seedlings, the protein levels of LHCII, phosphorylated D1/D2 and phosphorylated CP43 were similar, indicating that LTO1 and STN7 are not involved in the phosphorylation of D1/D2 and CP43. To obtain more quantitative data, the relative levels of the phosphorylation bands and LHCII bands in each lane of Figure 4A were scanned and normalized to the amount of WT in state 2. The LHCII phosphorylation level of lto1 in state 2 is between that of WT and stn7 (Figure 4B). These results show that LTO1 plays some role in STs though to a smaller extent than the STN7 kinase.

Figure 4

Phosphorylation patterns in lto1 and stn7 mutants during STs. A, Thylakoid membrane proteins extracted from WT, lto1 and stn7 mutants in state 1 (ST1, far-red light) and state 2 (ST2, white-light) were separated by SDS–PAGE and immunoblotted with anti-phosphothreonine antiserum (top) and LHCII antibodies (medium). Equal protein loading was verified with CBB staining. B, Relative level of phosphorylated proteins in figure A. The amount of protein of the relevant bands was estimated by Image J software and analyzed with three biological repeats. The relative levels of the phosphorylated bands and LHCII bands in every lane were normalized with the amount of WT under ST2 condition. Bars indicate standard deviations. Asterisks indicate significant differences (**P <0.01 by Student’s t test) compared to WT.

The cysteines of STN7 on the lumenal side are essential for the kinase activity and the corresponding Cys variants have a defective STs phenotype (Lemeille et al., 2009; Shapiguzov et al., 2016). To test the role of the Cys of LTO1 in STs, the four cysteines of two CxxC motifs (named as C12 and C34 in the following text) in the LTO1 LD were changed to alanine and STs were examined by measuring the maximal fluorescence (Fm) in state 1 and state 2 (Figure 5, A and B). As shown in Figure 5C, STs of C12 and C34 are slightly lower than in WT but higher than in the lto1 mutant and C1234 variant. The latter two mutants have a similar phenotype indicating defective STs (Figure 5, C and D). In addition, the pigment content of the leaves and Fv/Fm fluorescence are similar in lto1 and the cysteine variants of LTO1 (Figure 5, E and F). These results were further confirmed with a STs fluorescence assay using method 1. State transitions of the LTO1 cysteine variants whether estimated by false-color images or by fluorescence traces are deficient compared to WT (Supplemental Figure S9). These results indicate that the four cysteines in the LD of LTO1 are at least partially needed for STs.

Figure 5

Cysteines in the lumen domain of LTO1 affect STs. The four cysteines in the lumen domain of LTO1 were changed to alanine. These Cys variants which include double sites (named as C12 and C34) and four sites (named as C1234) mutants were introduced into the lto1 mutant. The phenotype of WT, stn7, lto1, LTO1 complemented line (LTO1/lto1) and Cys variants were examined by fluorescence imaging. A, B, STs measured with Method-2 in WT (A) and stn7 (B). D, dark-adaptation. B, blue-light. B+FR, blue and far-red light. Fo, minimum fluorescence after dark-adaptation; Fm, maximum fluorescence after dark-adaptation; Fm1, maximum fluorescence in state 1. Fm2, maximum fluorescence in state 2. a.u., arbitrary units. C, False color of ST fluorescence using method-2. D, State transition (qST) was estimated by (Fm1-Fm2)/Fm1. This value is positive in WT and close to zero or negative in stn7. The mean values obtained from three independent measurements of qST are shown. Bars indicate standard deviations. ns, not significant. Asterisks indicate significant differences (**P <0.01 by Student’s t test) compared to WT. E, Brightfield image. F, Fv/Fm was obtained from (Fm-Fo)/Fm. Three independent measurements were performed. Bars indicate standard deviations. ns, not significant. Asterisks indicate significant differences (**P <0.01 by Student’s t test) compared to WT.

Discussion

During acclimation of photosynthetic organisms to changes in light conditions, STN7 plays an important role both in STs and in the long-term response (Pesaresi et al., 2009; Rochaix, 2014). State transitions require the kinase STN7 which upon activation, phosphorylates the mobile LHCII antenna and redistributes the excitation energy between the two photosystems in a fluctuating light environment so as to achieve a balanced redox state of the photosynthetic electron transport chain (Depege et al, 2003; Bellafiore et al., 2005). STN7 can be activated by the reduced plastoquinone pool and the two conserved cysteines in its LD are essential for its activity, indicating that the chloroplast redox power is an important factor in the redox regulation of this protein kinase (Lemeille et al., 2009; Pesaresi et al., 2009; Rochaix, 2013). However, the redox component(s) for the redox regulation of STN7 remain unknown to a large extent. In this work, we report that the TRX protein LTO1 functions in maintaining the oxidized active state of STN7 in STs through redox interactions between their LDs.

Here, we have provided evidence that supports the functional redox link between LTO1 and STN7 in STs. First, we showed that STN7 and LTO1 interact with each other based on co-IPs, yeast two-hybrid, pull-down, and protein overlay assays. More specifically, our results revealed that the TRX-like domain of LTO1 interacts with the N-terminal domain of STN7. Importantly, this domain is the regulatory site for kinase activity and, as the TRX-like domain of LTO1, it is located on the lumen side of the thylakoid membrane (Lemeille et al., 2009; Karamoko et al., 2011; Shapiguzov et al., 2016). Second, we showed that STN7 is one of the targets of LTO1, a protein known to catalyze disulfide bond formation with active CxxC motifs in the thylakoid lumen (Karamoko et al., 2011). In particular, we performed redox assays in vitro to show that the reduced LD of STN7 is oxidized by the oxygen-treated TRX-like domain of LTO1, which implies electron transfer from STN7 to LTO1. Previous work demonstrated that the cysteines in the LD of both STN7 and LTO1 are essential for disulfide bond formation and for their physiological function (Lemeille et al., 2009; Karamoko et al., 2011; Wunder et al., 2013). These cysteines, two for STN7 and four for LTO1, are highly conserved in several photosynthetic organisms (Supplemental Figure S3). The four conserved cysteines of LTO1 form two CxxC motifs in the predicted spatial structure although the last two cysteines are 15 amino acids apart. We verified that the redox interactions between STN7 and LTO1 no longer occur when either the cysteines of STN7 or LTO1 were changed (Supplemental Figure S5). These findings imply that LTO1 interacts with STN7 and promotes the oxidation of the two conserved cysteines in the LD of STN7. Third, analysis of chlorophyll fluorescence and LHCII phosphorylation patterns in vivo both revealed that STs are partially deficient in the lto1 mutant (Figures 4 and 5), and a similar deficiency was observed with the cysteine variants of LTO1 (Figure 5). Based on these results, we propose that the highly conserved cysteines of LTO1 participate in the regulation of STN7 activity via redox reaction on the lumenal side of thylakoids.

We noticed that the Fv/Fm ratio is lower in lto1 than in WT (Figure 5; Supplemental Figures S7 and S9), which is consistent with a previous report (Karamoko et al., 2011). This raises the question whether the impairment of STs observed in lto1 and the cysteine variants of LTO1 is due to the compromised redox reactions between LTO1 and STN7 or whether it results indirectly from the impairment of PSII activity, which could impact the redox state of the plastoquinone pool and thereby affect STs. To test this possibility, we measured STs of other mutants similarly affected in PSII activity with lower Fv/Fm such as ffc (lacking cpSRP54) and ohp2-R (affected in PSII assembly) and found that STs in these mutants are similar as in WT (Supplemental Figure S8). A previous report also showed that the phosphorylation level of LHCII, low-temperature fluorescence, and PSI–LHCI–LHCII super-complex are normal in the ffc mutant in contrast to stn7 (Wang and Grimm, 2016). These observations indicate that the defect in STs observed in lto1 does not result from its decreased PSII activity. In conclusion, these findings suggest that LTO1 interacts with STN7 and promotes the oxidation of two conserved cysteines in the lumen domain of STN7.

In the bacterial periplasm, disulfide bonds of nascent peptides are formed and broken through the disulfide-bond formation (Dsb) system which includes thiol oxidizing (DsbA–DsbB) and reducing (DsbC–DsbD) pathways (McCarthy et al., 2000; Kadokura and Beckwith, 2002; Ito and Inaba, 2008; Herrmann et al., 2009). Chloroplasts have evolved from cyanobacteria through endosymbiosis and have inherited a similar pathway for shuttling redox power across membranes (Koehler and Tienson, 2009; Sideris and Tokatlidis, 2010; Kieselbach, 2013; Onda, 2013). In the chloroplast stroma, proteins are maintained in a reduced state by the ferredoxin (Fd)/TRX system which functions as a light-dependent reductase (Oelze et al., 2008). This reducing power is transferred to the thylakoid lumen via membrane-anchored CcdA/HCF164 of the trans-thylakoid thiol-reducing pathway which has been proposed to inactivate STN7 in high light (Lennartz et al., 2001; Page et al., 2004; Gabilly et al., 2011). It is possible that CcdA/HCF164-mediated thiol-reducing equivalents break the intermolecular dimer of STN7 or disturb the structural conformation of the STN7/Cyt b6f complex (Motohashi and Hisabori, 2006; Motohashi and Hisabori, 2010). A recent study confirms that the overexpression of Trx-m which feeds electrons to the TRX system inactivates STN7 in Nicotiana tabacum under high light conditions (Ancin et al., 2019).

In the thylakoid lumen of the chloroplast, LTO1 belongs to the thiol-oxidizing pathway which catalyzes Dsb (Kang and Wang, 2016). Based on the results obtained in this study, we propose the following model for the redox regulation of STN7 though the trans-thylakoid thiol pathway including LTO1 and CcdA/HCF164 (Figure S10). The trans-membrane region of STN7 separates the LD with the redox-responsive Cys from the catalytic domain on the stromal side of the thylakoids (Willig et al., 2011; Shapiguzov et al., 2016). Stt7, the orthologous protein of STN7 in C. reinhardtii (Depege et al., 2003; Bellafiore et al., 2005) is known to interact with subunit IV (Dumas et al., 2017) and the Rieske iron–sulfur protein (ISP; Lemeille et al., 2009) of the Cyt b6f complex which forms a dimer with contacts at the interface between Cyt b6 and ISP in the recent cryo-EM structure of spinach (Malone et al., 2019). Two conserved cysteines in the LD of STN7 can form inter- and/or intra-molecular disulfide bridges that appear to be essential for the kinase function of STN7 (Lemeille et al., 2009; Wunder et al., 2013; Shapiguzov et al., 2016). Upon over-excitation of PSII relative to PSI, the reduced PQH2 binds to the Qo site of Cyt b6f and activates STN7 through a still unknown mechanism. Previous studies proposed that the conformation of Cyt b6f complex changes upon binding of PQH2 to Qo when ISP moves from the proximal to the distal position during electron transfer in a similar way as in the mitochondrial bc1 complex (Rochaix, 2007; Shapiguzov et al., 2016). It is also likely that the conformational change of the chlorophyll phytyl tail near the Qo pocket (PQ oxidation site) at the interface between cyt b6 and subunit IV participates in the regulation of the activity of STN7 (Zito et al., 2002; Saif Hasan et al., 2013; Malone et al., 2019). Site-directed mutagenesis of conserved cysteines suggested that activation of STN7 requires dimer formation with two intermolecular disulfide bridges when the Qo site is occupied by PQH2. However, these intermolecular disulfide bonds are transient and are quickly converted to intra-molecular bonds in each monomer (Wunder et al., 2013; Shapiguzov et al., 2016). The question arises how the lumenal Cys of STN7 are oxidized to form inter- and intra-molecular disulfide bonds upon binding of PQH2 to the Qo site. Our results indicate that LTO1 is a potential candidate for this model. LTO1 contains two conserved domains, the TRX-like domain with a similar activity as DsbA and VKOR that is functionally equivalent to DsbB and transfers electrons from TRX to the final acceptor (vitamin K epoxide) in PSI (Karamoko et al., 2011; Onda, 2013). It is likely that when the Qo site is occupied by PQH2 under state 2 condition, the TRX-like domain of LTO1 interacts with the N-terminal domain of STN7 and maintains the conserved cysteines of STN7 in an oxidized state. The oxidized cysteines could form transient intermolecular disulfide bridges that are converted to the more stable intra-molecular disulfide of monomeric STN7 (Shapiguzov et al., 2016). Together with the conformational change of the Cyt b6f complex, these processes may promote a suitable conformation of STN7 for its activation (Rochaix, 2013; Shapiguzov et al., 2016). Electrons liberated by the formation of the disulfide bond of the LD of STN7 sequentially transfer to the TRX-like domain, then to the VKOR domain. When PSI is reduced relative to PSII (state 1), STN7 is inactivated and LTO1 does not participate in electron shuttling from STN7. However, it is noticeable that the redox interactions between LTO1 and STN7 are not fully needed for STs. Although this process is strongly decreased, it is not completely abolished in the lto1 mutant in contrast to the stn7 mutant. We, therefore, postulate that LTO1 acts as a helper to maintain the STN7 LD in its oxidized state for optimizing and maintaining kinase activity and that most likely other factors participate in this task.

Materials and methods

Plant materials

The Arabidopsis (A. thaliana)Col-0 ecotype and the transfer DNA (T-DNA) insertion lines of stn7 (SALK073254; Bellafiore et al., 2005) were used. The transgenic lines ohp2-R was obtained from Prof. Lianwei Peng (Li et al., 2019). The lto1 (SALK_151963C) and ffc (CS850421) mutants were obtained from the European Arabidopsis Stock Centre. The precise T-DNA insertion sites of the SALK073254 and SALK_151963C mutants were confirmed by PCR with primers lto1-LP/RP and LBb1.3 for LTO1 (all primer sequences are listed in Supplemental Table S2), primers stn7-LP/RP and LBb1.3 for STN7 (Supplemental Figure S6). After ripening, seeds were sterilized with 10% (v/v) sodium hypochlorite solution and sown on MS media supplemented with 3% (w/v) sucrose. Seedlings and plants were grown under controlled growth-chamber conditions as described previously (Ouyang et al., 2011).

Molecular cloning and plant transformation

For the cysteine substitution experiments, the two cysteines of STN7 and four cysteines of LTO1 in the thylakoid lumen domain were mutated to alanines with the Fast Mutagenesis System Kit (Trans, FM111), respectively. For the complementation experiments, the full-length coding sequences (CDS) of STN7 and LTO1 were amplified by PCR using the primers STN7-F/R and LTO1-F/R. The amplified products of the cysteine variants and CDS were digested with KpnI and SalI and subcloned into the pSN1300-3x Flag tag vector under the control of the cauliflower mosaic virus 35S promoter, and STN7-CDS products were transferred into the pCF399-HA tag vector (Bellafiore et al., 2005). Recombinant plasmids were transferred into Agrobacterium tumefaciens GV3101 by electroporation and then transformed into Arabidopsis plants using the floral-dipping method. The T1 generation of the transgenic lines were selected with hygromycin B (pSN1300) or Basta (pCF399) and grown in a greenhouse to produce seeds for PCR analysis. Successful complementation was confirmed by immunoblot analysis.

Antiserum production

The CDS sequences of mature STN7 and LTO1 were amplified using the specific primers mentioned above. The amplified products were subcloned into the pET28a vector and transformed into Escherichia coli strain DE3 (BL21). The recombinant proteins containing an N-terminal His tag were expressed and extracted under urea-denaturation conditions. The antigens were purified with nickel–nitrilotriacetic acid agarose (Novagen) according to the manufacturer’s instructions. Polyclonal antibodies were raised in rabbits with purified antigen (Mao et al., 2015).

Protein preparation and immunoblot analysis

Thylakoid membrane proteins extraction and immunoblot analysis were performed as previously described (Peng et al., 2006). Briefly, thylakoid membrane proteins were extracted from homogenizing Arabidopsis leaves in an ice-cold extraction buffer containing 400 mM sucrose, 10 mM NaCl, 2 mM MgCl2 and 50 mM HEPES-KOH, pH 7.8. For immunoblot analysis, equal amounts of chlorophyll were separated by sodium dodecyl sulphate (SDS)–PAGE and transferred to nitrocellulose membranes. The membranes were incubated with specific primary antibodies (Phospho-Threonine/Tyrosine Antibody, 9381S, Cell Signaling Technology; LHCII, made by our laboratory, Peng et al., 2006). The secondary antibody (goat anti-rabbit IgG, Origene, ZB-2301) conjugated to horseradish peroxidase (HRP) was used at a dilution of 1/10,000 for detection using the Pro-Light HRP Chemiluminescent Kit (Tiangen Biotech; Ouyang et al., 2020).

In vitro redox assays

The nucleotide sequences encoding the lumen domain of STN7 and LTO1 were amplified with the following primers: STN7-redox-F/R and LTO1-redox-F/R, respectively. For the cysteine substitution experiments, two G nucleotides of STN7 and four Gs of LTO1 in the lumen domain were replaced with Cs using the Fast Mutagenesis System (Trans, FM111). The PCR fragments were subcloned into the pETMALc-H and pET28a vectors and transformed into E. coli strain DE3 (BL21). To induce the recombinant proteins, E. coli strains were grown for 8 h at 37°C and further grown at 17°C overnight after isopropyl β-d-1-thiogalactopyranoside was added to the cultures. Cells were then harvested by centrifugation at 8,000g for 10 min at 4°C, and the pellets of recombinant LTO1 were purified using nickel–nitrilotriacetic acid resin (Novagen) with resuspension buffer (20 mM Tris–HCl, 200 mM NaCl, pH 7.0). Recombinant STN7 with maltose-binding protein (MBP) tag was purified using MBP sepharose (NEB). Reduced states of LTO1 and STN7 were obtained by incubation with 200 mM DTT for 1 h on ice and DTT was subsequently displaced with resuspension buffer using Amicon Centriprep tubes (Ultracel-10 membrane; Millipore). Proteins were oxidized using the following methods. For weak oxidation, air or oxygen gas was bubbled into the protein solution. For strong and irreversible oxidation, the proteins were incubated with 50 mM CuSO4 for 1 h on ice. In order to prevent oxidation by air, all tubes and solutions were treated with nitrogen gas. Proteins were precipitated with isopycnic 20% trichloroacetic acid at −20°C overnight, washed with ice-cold 80% acetone and then dissolved in a buffer containing 50 mM Tris–HCl, pH 7.8, 2% SDS, 10 mM AMS. After a 90-min incubation, the proteins were fractionated by 15% SDS–PAGE with nonreducing loading buffer and visualized by Coomassie-Brilliant Blue (CBB) staining (Motohashi et al., 2001; Karamoko et al., 2011).

Overlay analysis

Thylakoid proteins were extracted as described above and separated by SDS–PAGE. Proteins were then transferred to nitrocellulose membranes, blocked with Tween tris-buffered saline (TTBS) buffer (20 mM Tris–HCl, pH 7.6, 0.137 M NaCl, and 0.1% Tween-20) containing 5% skimmed milk and incubated with the fused LTO1 protein carrying a poly-histidine tag at a concentration of 0.1 mg ml−1 in TTBS buffer with 1% skimmed milk. The membrane was washed five times with TTBS buffer, probed with an anti-His-tag antibody (TA150088, Origene) and anti-HA-tag antibody (26D11, Abmart). The secondary antibody (goat anti-mouse IgG, Origene, ZB-2305) conjugated to HRP was used at a dilution of 1/10,000 for detection using the Pro-Light HRP Chemiluminescent Kit (Tiangen Biotech; Ouyang et al., 2011).

Pull-down assays

The lumen domain of LTO1 and STN7 were fused with GST and MBP-His tag, respectively. Then the recombinant proteins were purified using glutathione beads (Novagen) and amylose resin (New England Biolabs), according to the manufacturer’s instructions. Fifty microliter aliquots of 50% glutathione-agarose beads were incubated with four micrograms of LTO1-LD-GST and empty GST. After washing three times with buffer (20 mM Tris–HCl, pH 7.0, 200 mM NaCl), 6 µg of STN7-LD-MBP-His were added and incubated for 2 h at 4°C. Unbound proteins were removed by washing three times. The bound proteins eluted from the beads by boiling in 50 μL of 2× sampling buffer were loaded onto a 15% SDS–PAGE and analyzed by immunoblot analysis with an anti-His-tag antibody or CBB staining (Feng et al., 2016).

Co-IP assay

Thylakoid proteins of the lto1 mutant and STN7-HA transgenic Arabidopsis were extracted with phosphate-buffered saline (PBS) buffer (added final concentration 1% (wt/vol) dodecyl-β-d-maltopyranoside). 200 μL protein A/G agarose (Abcam) was incubated with non-immune serum (control) or antibodies against LTO1. Then the agarose was cross-linked to the antibodies using the cross-linker dimethylpimelidate dihydrechloride. For redox IP experiments, the thylakoid protein solution was treated with 50 μM glutathione disulfide or glutathione. Then the protein A/G agaroses with LTO1 antibodies was incubated with thylakoid proteins of STN7-HA overnight at 4°C. For cross-linking IP, the reaction mixture was treated with the cross-linker DSP (dithiobis (succinimidyl propionate) for 2 h at 4°C. After incubation, the protein A/G agarose was washed with PBS buffer containing 0.5% Nonidet P-40 six times and the bound proteins were eluted with 0.1 M glycine (pH 3.0), separated by SDS–PAGE and analyzed by immunoblotting with LTO1 and STN7 antibodies.

Split ubiquitin-based yeast two-hybrid assays

The yeast two-hybrid assay for membrane proteins was performed using the yeast (Saccharomyces cerevisiae) strain NMY32. The pCCW vector encoding the Cub-LexA-VP16 fragment was used to construct the bait plasmid. The prey plasmids were constructed from the vector pDSLNx, which encodes NubG (Dualsystems Biotech). Plasmids constructed for the yeast assays are displayed in Figure 2B annotated as mentioned above. Interactions were determined by growing diploid yeast colonies on SD-Trp-Leu-His plates and by β-galactosidase activity using an X-gal (5-bromo-4-chloro-3-indolyl-b-d-galactopyranoside) filter assay. NMY32 containing bait protein-Cub-LexAVP16 was transformed with the NubI-Alg5 expression plasmid as a positive control, and NMY32 containing x-Cub-LexA-VP16 was transformed with the plasmid expressing NubG-Alg5 as a negative control (Ouyang et al., 2011).

Chlorophyll fluorescence analysis

Room temperature chlorophyll fluorescence was monitored with Chlorophyll Fluorescence Imager (Technologica) and FluorCam800MF kinetic imaging fluorimeter (PhotoSystems Instruments). Seedlings grown on MS agar medium or soil were dark-adapted for 30 min before fluorescence measurements. For the measurement of STs, two different methods were used. In Method 1, a dark-adapted seedling was exposed to blue light to induce state 2 and a first checkpoint (Fs1) was set at 12 min. To promote a transition to state 1, FR light was turned on at 12 min and a second checkpoint (Fs2) was set after 3 min. The value of (Fs1-Fs2)/Fs1 is inversely related to the extent of state transitions. These values were shown with false-color. In the legend of the false-color scale, red and black/dark-blue represent the deficient and normal STs, respectively. In Method-2, the minimum fluorescence yield (Fo) was measured at the beginning and Fm yield (Fm) was measured during exposure to a saturating flash (0.8 s, 6,000 μmol m−2 s−1) after dark-adaptation. Then the seedling was illuminated with 80 μmol m−2s−1 blue light (PSII light) for 15 min. Subsequently, a FR light (PSI light) with a peak wavelength at 735 nm was switched on, and the maximal Fo in state 1 (Fm1) was checked after 15 min. The FR light was switched off and the maximum fluorescence yield (Fm2) was measured after 15 min of blue light excitation. Fv/Fm, representing the maximal quantum yield of PSII, and STs (qST) were calculated as (Fm-Fo)/Fm and (Fm1-Fm2)/Fm1, respectively (Bellafiore et al., 2005).

Statistical analysis

The statistical significance between three means of measurements was determined using Student’s t test, with a P <0.01 or <0.05.

Accession numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers STN7: AT1G68830; LTO1: AT4G35760.

Supplemental data

The following materials are available in the online version of this article.

. Ranking list of genes co-expressed with STN7.

. Primers used in this study.

. Heat-map of transcriptomics and proteomics of LTO1, STN7, and PsbO1/2.

. Sequence alignment of Stt7/STN7 in various species.

. Sequence alignment of LTO1 in various species.

. Structural fitting of the thioredoxin-like domain of AtLTO1 and SyVKOR.

. Cysteines in the lumen domain of LTO1 and STN7 are essential for its redox state.

. Identification of the stn7 and lto1 mutant.

. Phenotype characterization of lto1 and stn7 mutants.

. Independence between high fluorescence phenotype and the defect of state transitions.

. Phenotype characterization of cysteines variants of LTO1.

. Model for functional redox interactions between STN7 and LTO1.

Click here for additional data file.