The Variegated Mutants Lacking Chloroplastic FtsHs Are Defective in D1 Degradation and Accumulate Reactive Oxygen Species^,,

Abstract

In the photosynthetic apparatus, a major target of photodamage is the D1 reaction center protein of photosystem II (PSII). Photosynthetic organisms have developed a PSII repair cycle in which photodamaged D1 is selectively degraded. A thylakoid membrane-bound metalloprotease, FtsH, was shown to play a critical role in this process. Here, the effect of FtsHs in D1 degradation was investigated in Arabidopsis (Arabidopsis thaliana) mutants lacking FtsH2 (yellow variegated2 [var2]) or FtsH5 (var1). Because these mutants are characterized by variegated leaves that sometimes complicate biochemical studies, we employed another mutation, fu-gaeri1 (fug1), that suppresses leaf variegation in var1 and var2 to examine D1 degradation. Two-dimensional blue native PAGE showed that var2 has less PSII supercomplex and more PSII intermediate lacking CP43, termed RC47, than the wild type under normal growth light. Moreover, our histochemical and quantitative analyses revealed that chloroplasts in var2 accumulate significant levels of reactive oxygen species, such as superoxide radical and hydrogen peroxide. These results indicate that the lack of FtsH2 leads to impaired D1 degradation at the step of RC47 formation in PSII repair and to photooxidative stress even under nonphotoinhibitory conditions. Our in vivo D1 degradation assays, carried out by nonvariegated var2 fug1 and var1 fug1 leaves, demonstrated that D1 degradation was impaired in different light conditions. Taken together, our results suggest the important role of chloroplastic FtsHs, which was not precisely examined in vivo. Attenuated D1 degradation in the nonvariegated mutants also suggests that leaf variegation seems to be independent of the PSII repair.

Excessive light often limits the growth of photosynthetic organisms by irreversibly inactivating the photosynthetic apparatus, a process called photoinhibition (for review, see Barber and Andersson, 1992; Aro et al., 1993). A major target of photodamage is PSII (for review, see Barber and Andersson, 1992; Aro et al., 1993; Murata et al., 2007), a large pigment-protein complex in the thylakoid membrane. In particular, the reaction center D1 protein, which binds cooperatively to D2 and carries cofactors required for electron flow from the manganese cluster of the water-oxidizing complex to the plastoquinone pool (Zouni et al., 2001; Loll et al., 2005), is the primary target of light-induced irreversible oxidative damage (Mattoo et al., 1981; Ohad et al., 1990). Because D1 can be damaged by even low light intensities, photosynthetic organisms cannot avoid photodamage (Tyystjärvi and Aro, 1996; for review, see Barber and Andersson, 1992). To overcome this, photosynthetic organisms have evolved an efficient PSII repair cycle, which involves disassembling PSII, degrading photodamaged D1, and replacing newly synthesized D1 (for review, see Baena-Gonzalez and Aro, 2002). The rate of photodamage is proportional to light energy. When the light intensity exceeds the repair capacity, damaged D1 accumulates, resulting in photoinhibition.

In the PSII repair, recent studies in Synechocystis species PCC 6803 and Arabidopsis (Arabidopsis thaliana) suggest important roles of prokaryotic proteases (Lindahl et al., 1996, 2000; Bailey et al., 2002; Sakamoto et al., 2003; Silva et al., 2003; Komenda et al., 2006; Sun et al., 2007; Kapri-Pardes et al., 2007). Among them, FtsH appears to be a major protease. It is a membrane-anchored ATP-dependent zinc metalloprotease that belongs to the ATPases associated with a variety of cellular activities (AAA)+ protein family (for review, see Patel and Latterich, 1998; Ogura and Wilkinson, 2001). The ATPase and protease domain of FtsH was shown to form a hexameric-ring structure (Suno et al., 2006). In Synechocystis species PCC 6803, deletion of one of the thylakoidal FtsHs, slr0228, results in light-sensitive growth, impairment of the PSII repair cycle, and slower D1 degradation under high-light conditions (Silva et al., 2003). In Arabidopsis, 12 FtsH homologues were identified, nine of which are targeted to chloroplasts (Sakamoto et al., 2003). FtsH2 and FtsH5 are the most abundant among all chloroplastic FtsHs and are located in thylakoid membranes (Sakamoto et al., 2003; Yu et al., 2004, 2005). Chloroplastic FtsHs predominantly exist as a heterocomplex consisting of at least two types of isomers, A and B, represented by FtsH1/5 and FtsH2/8, respectively. These two types are functionally distinguishable from each other (Sakamoto et al., 2003; Yu et al., 2004, 2005; Zaltsman et al., 2005b).

We have extensively studied Arabidopsis mutants lacking chloroplast FtsHs. A mutant lacking FtsH5 (called yellow variegated1 [var1]) or lacking FtsH2 (var2) is highly vulnerable to PSII photodamage under high light (Chen et al., 2000; Lindahl et al., 2000; Takechi et al., 2000; Sakamoto et al., 2002, 2004). One notable feature in these mutants, in addition to the defective PSII repair, is the leaf-variegated phenotype that displays two sectors in the same leaf (green sectors containing normal chloroplasts and white sectors containing abnormal plastids lacking thylakoid membranes; Supplemental Fig. S1). White sectors are made by living cells and appear to be comparable with green sectors, except for lacking photosynthetic proteins (Kato et al., 2007). These results demonstrated that white sectors in var2 are arrested in chloroplast development. It is thus proposed that FtsH is not only involved in PSII repair but also in the formation of thylakoid membranes. Moreover, a series of genetic studies enabled us to identify trans-acting mutations that suppressed leaf variegation in var2 (Park and Rodermel, 2004; Miura et al., 2007; Yu et al., 2008). Many suppressors appeared to be associated with chloroplast translation, suggesting that the formation of variegated sectors is not simply governed by a specific factor but rather by factors related to chloroplast development (Miura et al., 2007; Yu et al., 2008). Since the variegated phenotype complicates our biochemical study in Arabidopsis, unlike cyanobacteria, a combined usage of var and its suppressors is necessary to further investigate the role of FtsH in the PSII repair cycle.

In this study, we show that chloroplasts in green sectors accumulate less PSII supercomplex and more PSII partial complexes than wild-type chloroplasts, likely due to the compromised PSII repair. Interestingly, we found that chloroplasts in green sectors accumulated significantly high levels of reactive oxygen species (ROS), suggesting that var2 indeed suffers from photooxidative stress. Given the essential role of FtsH, we evaluated impaired D1 degradation by the lack of FtsH2 and FtsH5 under different light conditions. Although a defect in degrading PSII reaction center proteins in var2 has been reported to occur under a photoinhibitory light condition (Bailey et al., 2002), D1 degradation examined in this study was very limited. This is because in vivo degradation of D1 is very difficult to measure in variegated leaf tissues (e.g. the presence of green and white leaf sectors interferes with protein normalization). To overcome the difficulty of handling variegated sectors, nonvariegated suppressor lines were subjected to these experiments. Our D1 degradation assays demonstrated that the lack of FtsH2 or FtsH5 significantly impairs D1 degradation. Collectively, our data corroborate important roles of FtsH2 and FtsH5 in avoiding photooxidative stress in chloroplasts.

RESULTS

PSII Protein Complexes in Thylakoid Membranes of the Wild Type and var2

The functional PSII core complex of higher plants is a dimer and is associated with a light-harvesting antenna chlorophyll complex (LHCII). In the repair cycle of the PSII complex, assembly and disassembly of PSII are stepwise processes that involve the formation of several intermediates, such as a PSII core complex lacking CP43 (RC47) and a monomeric core complex with a reaction center core (PSII monomer; Baena-Gonzalez and Aro, 2002). Thus, monitoring intermediates of PSII assembly/disassembly and analyzing them in the wild type and var2 should allow us to assess the repair state of PSII and the involvement of FtsH. To examine the repair intermediates, we purified chloroplasts using a Percoll gradient, extracted thylakoid membranes, solubilized the fraction with 0.4% n-dodecyl-β-d-maltoside, and finally separated PSII and other complexes using two-dimensional blue native (BN)/SDS-PAGE. We simultaneously performed an immunoblot analysis to also detect D1 in these preparations. For BN/SDS-PAGE, proteins from the wild type and var2-1 were equally loaded based on total chlorophyll content. To monitor PSII assembly under a minimized photooxidative condition, we first isolated thylakoid protein complexes from leaves adapted at 20 μmol m⁻² s⁻¹ (referred to as low light condition) for 1 d prior to the extraction experiment, and BN/SDS-PAGE was performed as shown in Figure 1. Silver staining of the gels revealed that the PSII-LHCII supercomplexes were detectable in var2-1 as well as in the wild type under low light, indicating that the assembly of the supercomplex proceeds normally in var2-1.

Figure 1.

BN/SDS-PAGE analysis of thylakoid protein complexes from mature leaves of wild-type Col and var2-1. A, Thylakoid membranes (0.5 mg mL⁻¹ chlorophyll) were purified from intact and broken chloroplasts of Col and var2-1 leaves (approximately 8-week-old plants). Thylakoid protein complexes were solubilized with 0.4% n-dodecyl-β-d-maltoside and separated on 4% to 12% BN/PAGE gels (10 μg chlorophyll per lane). Thylakoid membrane proteins were further separated by 14% SDS-PAGE and stained with silver. Thylakoid membranes were isolated from low light-adapted leaves (pretreated for 1 d at 20 μmol m⁻² s⁻¹; left two panels) and from leaves illuminated constantly at 150 μmol m⁻² s⁻¹ followed by low light adaptation (right two panels). B, Two-dimensional gels were also electroblotted onto polyvinylidene difluoride membranes and subjected to immunodetection of D1 protein. The positions corresponding to PSII supercomplexes, PSII dimer, PSII monomer, and RC47 are indicated at the bottom. To confirm D1, CP47, and CP43 in each complex, spots on the gel were subjected to matrix-assisted laser-desorption ionization time of flight mass spectrometric analysis (data not shown).

We subsequently isolated thylakoid proteins from leaves illuminated constantly at 150 μmol m⁻² s⁻¹ (referred to as normal light condition). Under this normal light condition, the PSII-LHCII supercomplexes were hardly detectable in var2-1, whereas these complexes were still detectable in Columbia ecotype (Col), indicating that increased light intensity affected the assembly of PSII supercomplexes in var2-1. Further immunoblot analysis of the PSII complex using D1 antibodies confirmed the decreased level of a PSII supercomplex and the PSII dimer in var2-1 constant illumination under normal light. More importantly, the level of RC47 was higher in var2-1. RC47 is a known intermediate that is formed during a disassembly step of PSII. Studies in cyanobacteria have demonstrated that detachment of CP43 from damaged PSII complexes is a prerequisite for FtsH to access and degrade photodamaged D1 (Komenda et al., 2006). Thus, the preferential accumulation of RC47 in var2-1 after light irradiation strongly suggests that the selective D1 turnover was impaired at the step of the formation of RC47 in var2. Normal accumulation of RC47 in var2 adapted under low light also suggests the possibility that the residual level of FtsHs is sufficient or that other proteases considerably participate in the PSII repair at low light intensity.

High ROS Accumulated in var2

As a consequence of defects in the PSII repair, var2 leaves are very likely to receive photooxidative stress. Given the partial PSII complexes accumulated in var2, we assumed that var2 leaves are defective in transmitting excitation energy into water oxidation reaction even under normal light and accumulate substantial ROS as a result of the oxidative stress. To test this possibility, we histochemically detected superoxide radical (O₂⁻) and hydrogen peroxide (H₂O₂) by nitroblue tetrazolium (NBT) staining and 3,3′-deaminobenzidine (DAB) staining, respectively. NBT forms blue formazan in the presence of O₂⁻, and polymerization of DAB by peroxidase is visible as a brown precipitate in the presence of H₂O₂.

Our NBT staining showed that O₂⁻ preferentially accumulated in var2 (Fig. 2A). Detailed in situ observation in Col and var2 leaves revealed that Col had limited amounts of NBT stains, whereas var2 leaves contained the blue precipitates that appeared to overlap green sectors (Fig. 2B). Observation of NBT stains under light microscopy demonstrated that the NBT stains in green sectors were detected within organelles whose morphologies are typical of chloroplasts (Fig. 2C). Likewise, we performed DAB staining with different light intensities, and the results showed that H₂O₂ was detectable in green sectors of var2 leaves (Fig. 2D). Although DAB did not reveal the difference between Col and var2 leaves as clearly as NBT precipitates, we observed that generation of H₂O₂ appeared to be light dependent: dark-grown leaves had little DAB stains, whereas a photoinhibitory light condition (800 μmol m⁻² s⁻¹) deepened DAB stains both in Col and var2. Our results thus supported our assumption that the defective PSII repair capacity in var2 leads to high ROS production in chloroplasts.

Figure 2.

ROS accumulation in var2 green sectors. A, In situ detection of O₂⁻ by staining with NBT (blue, bottom panels) in 4-week-old wild-type Col and var2. B, In situ detection of O₂⁻ by staining with NBT (blue, bottom panels) in 4-week-old wild-type Col and var2 leaves. C, Higher magnification of var2 leaves from A. D, In situ detection of H₂O₂ by DAB staining (dark brown, bottom panels) in 3-week-old wild-type Col and var2 grown on Murashige and Skoog plates under different light conditions (growth light, 100 μmol m⁻² s⁻¹; high light, 800 μmol m⁻² s⁻¹). Plants that were dark adapted for 2 d are also shown (dark).

To further characterize ROS accumulation in var2 quantitatively, we measured levels of three ROS from Col and var2 leaves as described in “Materials and Methods.” Because ROS were preferentially observed in chloroplasts, ROS amounts were normalized based on total chlorophyll content (Fig. 3). First, we assessed the levels of thiobarbituric acid-reactive substances (TBARS), including malondialdehyde, as an index for lipid peroxidation. TBARS was 3.5-fold higher in var2 than in Col (Fig. 3A). Second, H₂O₂ and O₂⁻ were quantitatively measured to confirm our histochemical experiments. To detect both ROS, we performed this measurement using not only 3-week-old seedlings grown on Murashige and Skoog medium but also 10-week-old mature leaves (Fig. 3, B and C). This was because our initial experiments showed that ROS levels varied in different developmental stages (data not shown). H₂O₂ levels increased dramatically in both Col and var2-1 mature leaves, but its levels were constantly higher in var2 than in Col (4.4-fold in young seedlings and 5.1-fold in mature leaves; Fig. 3B). In addition, O₂⁻ levels were also shown to be constantly higher in var2 than in Col (10.4-fold in young seedlings and 5.5-fold in mature leaves; Fig. 3C). No significant difference in O₂⁻ was detected between dark-treated var2 and Col seedlings (covered with aluminum foil for 2 d), which was consistent with our DAB stains and suggests that O₂⁻ generation is light dependent. We also calibrated each ROS level based on fresh weight, and higher O₂⁻ and H₂O₂ levels in var2 mature leaves were evident (Supplemental Fig. S2). Taken together, these results demonstrated that var2 leaves indeed suffer from photooxidative stress and generate substantial ROS in chloroplasts in a light-dependent manner.

Figure 3.

Quantification of ROS in var2 compared with wild-type Col. A, The level of lipid peroxidation in 35-d-old plants without roots (young seedling) was measured by TBARS assay. B, H₂O₂ accumulation in 30-d-old plants without roots (young seedling) and 9-week-old leaves (mature plant) was quantified using potassium iodide methods. C, The level of blue formazan as O₂⁻ in 4-week-old leaves (young seedling), 9-week-old leaves (mature plant), or 4-week-old leaves after dark adaptation for 2 d (dark). NBT formazan precipitates were completely eluted, and the amount was determined at 560 nm. Values of each graph were normalized to chlorophyll (Chl) content. (mean ± sd, n = 3). Asterisks indicate significant difference from the wild type using Student's t test (** P < 0.01).

In Vivo Assessment of Impaired D1 Degradation in var2 Using a Variegation Suppressor

Because photooxidative stress accessed by ROS production appeared to take place in var2 leaves under normal light, we attempted to design an experiment by which defective PSII repair cycle, as represented by D1 turnover, can be monitored in different light conditions. To perform this experiment, we encountered difficulty in utilizing variegated var2 leaves, because D1 levels that can be detected by immunoblots were shown to fluctuate among leaf discs (Supplemental Fig. S3). Moreover, variegated sectors were very difficult to normalize protein contents (Kato et al., 2007). In vivo measurement of D1 turnover in var2 thus gave limited results, as reported previously by Bailey et al. (2002). To minimize such an experimental problem, a nonvariegated var2 mutant (var2 fug1) was used in this study. The FU-GAERI1 (FUG1) locus encodes a chloroplastic translation initiation factor 2 (cpIF2). A complete lack of cpIF2 in fug1-2 results in an embryo-lethal phenotype, whereas two leaky mutations in fug1-1 and fug1-3 are viable and recover leaf variegation when combined with var2 (Supplemental Figs. S1 and S3). In this study, D1 turnover in the absence of FtsHs was assessed in var2 fug1, and we used fug1 as a control.

Prior to D1 measurement by immunoblots, leaf discs from Col and other mutant lines were infiltrated with lincomycin to inhibit chloroplast protein synthesis. In the presence of lincomycin, wild-type leaves showed a rapid decrease in maximum photochemical efficiency of PSII in the dark-adapted state (F_v/F_m) under high light conditions of 1,200 μmol m⁻² s⁻¹ (approximately 20% after 210 min; Fig. 4A). The rapid decrease of F_v/F_m in var2-1, fug1-3, and var2-1 fug1-3 was comparable to that in the wild type. We also measured F_v/F_m values of var2-1, fug1-3, and var2-1 fug1-3 under growth and low light conditions (100 and 20 μmol m⁻² s⁻¹ for 8 h, respectively; Fig. 4, B and C). In the absence of lincomycin, a decrease in PSII activity was not detected in any of the genotypes, suggesting that PSII photodamage is within the capacity of the PSII repair cycle under these conditions. On the other hand, addition of lincomycin decreased F_v/F_m in all genotypes, and as expected, photoinhibitory effects appeared to be light dependent. These results suggest that infiltration of lincomycin into leaf discs efficiently inhibits D1 synthesis and that PSII photodamage was similar in the wild type and the mutants.

Figure 4.

Maximal photochemical efficiency of PSII in the presence or absence of lincomycin. F_v/F_m was measured in leaf discs from Col (circles), var2-1 (diamonds), fug1-3 (triangles), and var2-1 fug1-3 (squares). Leaf discs were incubated for 0, 60, 120, and 210 min under high light condition (1,200 μmol photons m⁻² s⁻¹ [A]) or for 0, 120, 240, 360, and 480 min under growth and low light conditions (100 and 20 μmol photons m⁻² s⁻¹, respectively [B and C]) in the presence (gray symbols) or absence (white symbols) of lincomycin. At each time point, leaf discs were dark adapted for 10 min prior to measurement. Values are means ± sd (n = 3).

In Vivo D1 Degradation Assay in var2 fug1

Following lincomycin infiltration, leaf discs were first incubated for 0, 60, 120, and 210 min under high light (1,200 μmol m⁻² s⁻¹). To detect D1 levels during these periods, thylakoid membranes were isolated and subjected to immunoblot analysis (Fig. 5A). Rates of D1 degradation in fug1-3 and var2-1 fug1-3 were estimated based on the ratio of immunoreacted D1 to Coomassie Brilliant Blue-stained LHCII, as shown in Figure 5. Neither obvious decrease nor significant change of LHC levels was detected between the wild type and mutants in the conditions we employed (Supplemental Fig. S4). After 210 min of incubation, D1 levels in fug1-3 rapidly decreased to about 20% of the initial value. In contrast, D1 levels in var2-1 fug1-3 remained at 40%. We also performed a D1 degradation assay in the wild type, which had a D1 degradation rate that was similar to fug1-3 (Supplemental Fig. S5). These results are consistent with the previous study, which showed that the D1 degradation in var2 was significantly slower than that in the wild type when plants were exposed to high light (Bailey et al., 2002).

Figure 5.

Immunoblot analysis of D1 protein in fug1-3 and var2-1 fug1-3 mutants under three different light conditions. Leaf discs harvested from fully expanded mature leaves of fug1-3 and var2-1 fug1-3 (approximately 8-week-old plants grown under normal conditions) were infiltrated with 5 mm lincomycin. Leaf discs were incubated for 0, 60, 120, and 210 min under high light condition (1,200 μmol m⁻² s⁻¹[A]) or for 0, 120, 240, 360, and 480 min under growth and low light conditions (100 and 20 μmol m⁻² s⁻¹, respectively [B and C]). Thylakoid membrane proteins were separated by 14% SDS-PAGE and stained with Coomassie Brilliant Blue. A representative immunoblot using anti-D1 antibodies and the band corresponding to Coomassie Brilliant Blue-stained LHCII are shown. Signals of immunoblots from three biological repeats were quantified using the ImageJ program and normalized to the amount of Coomassie Brilliant Blue-stained LHCII (sd with bars, n = 3). Gray and white circles indicate fug1-3 and var2-1 fug1-3, respectively. To compare D1 levels, ratios at 0 min were adjusted to 1.

Next, we examined D1 degradation in fug1-3 and var2-1 fug1-3 during nonphotoinhibitory growth and low light irradiance (100 and 20 μmol m⁻² s⁻¹, respectively; Fig. 5, B and C). Under growth light, D1 levels in fug1-3 decreased to 40% of the initial amount, whereas var2-1 fug1-3 remained at 70% of the initial D1 amount after 8 h of irradiation. Likewise, under low light, D1 levels in fug1-3 decreased to 70% of the initial amount, whereas no or little D1 degradation was detected in var2-1 fug1-3. These results suggest that the rate of D1 degradation was significantly delayed and slowed in var2-1 fug1-3 when compared with the control. We also performed a D1 degradation assay in the dark and found that D1 protein levels remained unchanged even after 480 min of incubation (Supplemental Fig. S6). These data suggest that var2-1 fug1-3 leaves are compromised in light-dependent D1 degradation under nonphotoinhibitory light conditions.

To further characterize D1 turnover in vivo and exclude a possible secondary effect of lincomycin, we estimated D1 degradation by pulse labeling of chloroplast proteins (Fig. 6). De novo synthesized protein in leaf discs was labeled with [³⁵S]Met for 60 min and subsequently chased in unlabeled medium for 60, 120, and 180 min as reported previously (Bonardi et al., 2005). Our previous data showed that after 60 min of labeling, D1 proteins detectable in fug1-3 were comparable with those in Col (Miura et al., 2007). The bands corresponding to D1 (confirmed by immunoblot; data not shown) were shown to turn over very rapidly in fug1-3 during the chase period, whereas we consistently observed that D1 turnover was impaired in var2-1 fug1-3. Collectively, our in vivo D1 degradation assays reinforced the important role of FtsHs in the PSII repair cycle.

Figure 6.

Pulse-chase analysis of thylakoid membrane proteins in fug1-3 and var2-1 fug1-3 mutants. Leaf discs were labeled for 60 min at 100 μmol m⁻² s⁻¹ and subsequently chased for 60, 120, 180, and 240 min at a light intensity of 600 μmol m⁻² s⁻¹. Thylakoid membrane proteins were separated by 14% SDS-PAGE, and labeled proteins were detected. A representative labeled D1 protein signal is indicated by the arrowhead. The bands corresponding to LHCII on the Coomassie Brilliant Blue-stained gel are also shown at the bottom. Similar results were obtained in four additional independent experiments.

In Vivo D1 Degradation Assay in var1 fug1

Although FtsH2 was shown to play a role in D1 degradation, the involvement of other FtsH isomers such as FtsH5 is also possible. FtsH2 and FtsH5 are major FtsHs that were suggested to constitute a heterocomplex (Sakamoto et al., 2003; Yu et al., 2004; Zaltsman et al., 2005b). To test whether FtsH5 is indeed involved in D1 protein degradation, var1-1 fug1-1 was subjected to the aforementioned D1 degradation assay (photosynthetic phenotypes of fug1-1 and fug1-3 were essentially identical). Immunoblot analysis of the D1 protein levels was performed at three different light conditions (Fig. 7). Similar to var2, D1 levels in var1-1 fug1-1 remained high during high light irradiation, whereas the control plant fug1-1 showed a rapid decrease of D1 (Fig. 7A). Relative amounts of D1 estimated by immunoblot signals showed that the rate of D1 degradation in var1-1 fug1-1 was slower than that in fug1-1 (Fig. 7). The defect in D1 protein degradation in var1-1 fug1-1 was also observed when plants were grown at growth and low light intensities (Fig. 7, B and C). We conclude that FtsH5 as well as FtsH2 are functioning in the PSII repair cycles.

Figure 7.

Immunoblot analysis of D1 protein in fug1-1 and var1-1 fug1-1 mutants under three different light conditions. Leaf discs harvested from fully expanded, mature leaves of fug1-1 and var1-1 fug1-1 (approximately 8-week-old plants grown under normal conditions) were infiltrated with 5 mm lincomycin. The leaf discs were incubated for 0, 60, 120, and 210 min under high light condition (1,200 μmol m⁻² s⁻¹ [A]) or for 0, 120, 240, 360, and 480 min under growth and low light conditions (100 and 20 μmol m⁻² s⁻¹, respectively [B and C]). Thylakoid membrane proteins were separated by 14% SDS-PAGE and stained with Coomassie Brilliant Blue. A representative immunoblot using anti-D1 antibodies and the band corresponding to Coomassie Brilliant Blue-stained LHCII are depicted. Signals of immunoblots from three biological repeats were quantified as indicated in Figure 5.

PSII Activities of fug1-3 and var2-1 fug1-3 upon High Light Exposure

To assess the photoinhibitory status of var2 along with fug1, we measured the F_v/F_m in leaf discs prepared from fully expanded rosette leaves (8 weeks old, grown in soil). The results shown in Figure 4 indicated that F_v/F_m values were similar between the wild type, fug1-3, and var2-1 fug1-3, suggesting that the coexistence of fug1 and var2 not only recovers leaf variegation but also mitigates PSII photoinhibition. Because the effect of fug1 under long-term light irradiation complicates the evaluation of photoinhibitory status of PSII activity, we calibrated another photosynthetic parameter, PSII electron transport rate (ETR), in this study. Light-dependent changes of ETRs were measured in leaves from the wild type, var2-1, fug1-3, and var2-1 fug1-3 (Fig. 8). No significant difference in ETRs was observed between the wild type and the mutants when light intensity was below 100 μmol m⁻² s⁻¹. A slower increase of ETRs in var2-1 and var2-1 fug1-3 was detectable at relatively low light intensity (over 200 μmol m⁻² s⁻¹), whereas ETRs in the wild type and fug1-3 elevated well until 400 μmol m⁻² s⁻¹. At 1,200 μmol m⁻² s⁻¹, ETRs in var2-1 and var2-1 fug1-3 declined and reached only 70% of ETR in the wild type. We also found that fug1-3 itself does not affect ETR.

Figure 8.

PSII capacities during high light exposure. ETR was measured in leaf discs from wild-type Col (white circles), var2-1 (black circles), fug1-3 (triangles), and var2-1 fug1-3 (squares) in response to increasing light intensities. PAR, Photosynthetically active radiation. Means ± sd (n = 3) are shown.

In addition to ETR, we also measured capability of recovery from photoinhibition in the wild type and the mutants. To analyze this, leaf discs from Col, var2-1, fug1-3, and var2-1 fug1-3 were illuminated until the initial F_v/F_m was reduced to approximately 50%, and subsequent recoveries of F_v/F_m were monitored under a low light condition (20 μmol m⁻² s⁻¹). As shown in Supplemental Figure S7, recovery of F_v/F_m was always faster in Col than in other mutant lines and reached above 85% of the initial value after 3 h. Similar to Col, fug1-3 exhibited a fast recovery rate, whereas recoveries in var2-1 and var2-1 fug1-3 were apparently slower than in Col and reached only 70% after 3 h. Taken together, these results appeared to reflect the fact that the lack of FtsH2 has a significant effect on the PSII repair cycle.

DISCUSSION

An essential role of FtsHs in chloroplasts has been well documented previously by us and other groups (Chen et al., 2000; Takechi et al., 2000; Sakamoto et al., 2002, 2003). Involvement of FtsHs in protecting photosystems has also been implicated in cyanobacteria (Mann et al., 2000; Silva et al., 2003). In particular, characterization of Arabidopsis var2 mutants demonstrated that the lack of FtsH2 leads to an impaired capacity in the PSII repair cycle and a defective D1 turnover (Bailey et al., 2002; Sakamoto et al., 2004). However, measurement of PSII activity was mostly conducted under photoinhibitory light conditions, where we cannot rule out secondary effects of excess light energy on damaging photosystems. Here, we show that the impairment of the PSII repair cycle occurs under nonphotoinhibitory light conditions. First, comparison of PSII supercomplex and other PSII complexes in var2 and Col leaves revealed that var2 results in accumulating less functional PSII supercomplex and more RC47 intermediates than Col (Fig. 1). Second, we found that var2 leaves accumulate more ROS in chloroplasts than Col, suggesting that photooxidative stress is induced due to the impaired PSII cycle under growth conditions (Figs. 2 and 3). Finally, D1 degradation monitored in nonvariegated var2 fug1 mutants was compromised under both growth and weak light conditions (Fig. 5). Collectively, our results demonstrated the critical role of FtsHs in the PSII repair cycle not only under photoinhibitory but also nonphotoinhibitory light conditions, which was not precisely investigated in the previous studies (Bailey et al., 2002).

Our histochemical and quantitative analyses clearly indicated that chloroplasts in green sectors produced high levels of O₂⁻, which has a short lifetime and is converted rapidly to H₂O₂ by superoxide dismutase. Because O₂⁻ cannot permeate across membranes, it is probably converted to H₂O₂ by chloroplastic superoxide dismutase (for review, see Asada, 2006). H₂O₂ can then be exported to cytosol or to other organelles to ultimately convert it to water. As a consequence, high ROS in chloroplasts may cause damage in photosynthetic proteins. In contrast, ROS in var2 leaves may affect the response to environmental stresses, since H₂O₂ not only causes cytotoxicity but also acts as a signaling molecule for various cellular activities (for review, see Apel and Hirt, 2004). Further studies may allow us to investigate a physiological role of chloroplast ROS. In photosynthetic electron transport, it is known that ROS can be mainly produced by Mehler reaction around PSI (Asada, 2006). However, considering that FtsH is involved in the PSII repair cycle, O₂⁻ and H₂O₂ accumulated in var2 were likely generated around PSII. Other ROS, such as singlet oxygen and hydroxy radicals, may also be generated around PSII. To date, in vitro investigation of electron transport processes of PSII suggested ROS generation on the electron accepter and donor sides of PSII under the reducing and oxidizing conditions, respectively. However, these observations have yet to be confirmed in vivo (for review, see Pospíšil, 2009). Interestingly, high ROS generation in var2 is correlated with our finding that the less functional partial PSII complexes accumulate. It is thus possible that partial PSII complexes contribute to ROS production. An effective PSII repair can be required not only to provide functional PSII but also to avoid cytotoxicity caused by these partial PSII complexes.

Proteases involved in the PSII repair cycle have been extensively studied in Synechocystis species PCC6803, in which FtsH (slr0228) was shown to play major roles in degrading photodamaged D1 (Silva et al., 2003; Komenda et al., 2006). Phylogenetic analysis indicated that slr0228 is most closely related to Arabidopsis FtsH2 (Chen et al., 2000; Sakamoto et al., 2003). The participation of other cyanobacterial FtsH isomers in thylakoidal proteolysis is currently unknown. Loss of slr0228 was first suggested to affect PSI (Mann et al., 2000). However, later it was demonstrated that loss of slr0228 function results in high light-sensitive growth, which was suggested to be due to defects in the PSII repair cycle (Silva et al., 2003; Komenda et al., 2006). Similar to the results obtained with var2 in this study, a FtsH (slr0228) strain attenuated D1 degradation and accumulated RC47 (Silva et al., 2003; Komenda et al., 2006). These observations further supported important roles of FtsH in light-dependent D1 degradation in photosynthetic organisms. In addition, deletion of 20 amino acids from the N-terminal end of D1 in Synechocystis was shown to significantly affect D1 degradation, suggesting that D1 turnover is processively carried out by FtsH from its N-terminal end (Komenda et al., 2007). Damaged D1 probably causes a conformational change of the PSII dimer, leading to partial disassembly of the PSII monomer, thus allowing access of FtsH to RC47 and degradation of D1 from the N-terminal end.

Our D1 degradation assays showed that D1 turnover still occurred even in the absence of FtsH2 or FtsH5 (Figs. 5 and 7), suggesting the presence of a yet unidentified D1 degradation pathway in addition to the FtsH2/FtsH5 protease pathway in the repair cycle. One possibility is the contribution of other FtsH subunits, such as FtsH8 and FtsH1, because loss of FtsH2 or FtsH5 in var2 or var1 is partially compensated by an elevated level of FtsH8 or FtsH1, respectively (Yu et al., 2004, 2005). Transcriptional analysis of several FtsH genes showed a remarkable increase of FtsH8 and FtsH1 mRNA levels during high light exposure (Sinvany-Villalobo et al., 2004). An alternative possibility is the contribution of other proteases such as Deg, which encodes an ATP-independent Ser-type endopeptidase and which is peripherally attached to thylakoid membranes (Itzhaki et al., 1998). Deg1, Deg5, and Deg8, located in the thylakoid lumen, were suggested to act on PSII repair under high light conditions (Kapri-Pardes et al., 2007; Sun et al., 2007). Deg was proposed to act as an endopeptidase to aid in processive degradation that is preceded by other proteases. Therefore, the putative cooperation between FtsH and Deg in D1 degradation is feasible. Under our experimental conditions, however, we could not detect remarkable accumulation of D1 degradation fragments, which were previously reported in the wild type but not in the deg mutants (data not shown). In contrast to the reports in Arabidopsis, physiological experiments using a triple deg mutant in Synechocystis species PCC 6803 showed that Deg is not essential for in vivo D1 degradation (Barker et al., 2006). Different roles of Deg in Arabidopsis and Synechocystis may indirectly suggest that Deg acts supplementary to the FtsH-dependent D1 degradation pathway. Finally, we cannot rule out the possibility that unknown proteases contribute to D1 degradation under certain conditions.

Previous studies in var mutants have demonstrated that leaf variegation is not a consequence of the defect in the PSII repair cycle. This hypothesis is supported by the facts that (1) the degree of variegation is independent of light intensities and is rather dependent on plant development (Zaltsman et al., 2005a) and (2) white leaf sectors in var2 are composed of living cells with undifferentiated plastids (Kato et al., 2007). In addition to these observations, our D1 degradation assays showed that PSII repair is attenuated in the nonvariegated var2 fug1 and var1 fug1 mutants, further supporting the fact that variegation is independent of PSII repair. Thus, it is likely that FtsH plays an additive and distinct role in thylakoid development at an early stage of chloroplast differentiation, which is unrelated to PSII photodamage but is related to the variegation phenotype. This is supported by the facts that fug1 shows a phenotypic effect on chloroplast development in early developmental stages but the mutant does not exhibit an altered phenotype at later developmental stages (Miura et al., 2007). Whether the protease activity of FtsH is required for this additive function remains unclear and requires further investigation.

It is interesting that photosynthetic organisms apparently possess multiple genes for proteases in thylakoid membranes, as exemplified by FtsH (nine in Arabidopsis and four in Synechocystis) and Deg (four in Arabidopsis and three in Synechocystis; Sokolenko et al., 2002; Sakamoto et al., 2003). Although this raises the question of functional divergence between isomers, no experimental data on differential roles such as tissue specificity have been reported so far. One exception is FtsH6, which was suggested to play a specific role in the degradation of LHCII during senescence, but the presence of an FtsH6 homocomplex has not yet been confirmed (Zelisko et al., 2005). Rather, our study demonstrates a redundant role of FtsH2 and FtsH5 in the PSII repair cycle. Given the fact that two types of FtsH isomers, FtsH5 in type A and FtsH2 in type B, were shown to be required for heteromeric complex formation, it is reasonable to assume that FtsH2 and FtsH5 equally contribute to PSII repair. This is supported by the fact that the degree of photoinhibition as assessed by a decline in F_v/F_m was similar in var1 and var2 (Sakamoto et al., 2004). Therefore, our data suggest that in Arabidopsis, the type A/B FtsH heterocomplex plays a major role in degrading D1 and possibly other thylakoid proteins. Other isomers such as FtsH1 and FtsH8, the loss of which does not show any detectable phenotypes, may act supplementary to the major FtsH2/H5 isomers. Thus, the presence of multiple FtsH isomers may be required to ensure proper quality control of thylakoid proteins that are highly vulnerable to photooxidative damage.

MATERIALS AND METHODS

Plant Materials and Growth Conditions

Arabidopsis (Arabidopsis thaliana) Col was used as the wild type. The mutant lines used in this study, fug1-1, fug1-3, var1-1 fug1-1, var2-1, and var2-1 fug1-3, were described previously. All mutants are in the Col background (Miura et al., 2007). Plants were germinated and grown on 0.7% (w/v) agar plates containing Murashige and Skoog medium supplemented with Gamborg's vitamins (Sigma-Aldrich), 2 mm MES, pH 5.8, and 1.5% (w/v) Suc. Plants were maintained under 12 h of light (approximately 60 μmol m⁻² s⁻¹) at a constant temperature of 22°C. When plants were 4 weeks old, they were transferred onto soil and maintained under 12 h of light (100 μmol m⁻² s⁻¹) at a constant temperature of 22°C.

Detection and Measurements of ROS

In situ detection of O₂⁻ was performed by treating leaves with NBT as described by Kawai-Yamada et al. (2004). Leaves were detached from seedlings, vacuum infiltrated with 10 mm NaN₃ in 10 mm potassium phosphate buffer (pH 7.8) for 1 min using a syringe, and incubated in 0.1% (w/v) NBT in 10 mm potassium phosphate buffer (pH 7.8) for 120 min at room temperature under light (100 μmol m⁻² s⁻¹) or dark (covered with aluminum foil). Stained leaves were cleared by boiling in acetic acid:glycerol:ethanol (1:1:3, v/v/v) solution before photographs were taken. To quantify formazan generation, washed leaves were boiled in dimethyl sulfoxide until formazan precipitates were eluted completely (Yaeno et al., 2004). The amount of formazan was determined spectrophotometrically by measuring A₅₆₀. In situ detection of H₂O₂ was performed by treating leaves with 5 mm DAB at pH 3.8, as described previously (Orozco-Cardenas and Ryan, 1999). Stained leaves were completely cleared by boiling in ethanol. H₂O₂ levels were determined according to Velikova et al. (2000). Leaves (>200 mg) were homogenized in liquid nitrogen with 500 μL of 0.1% (w/v) TCA. The homogenate was centrifuged at 15,000g for 15 min at 4°C, and 200 μL of the supernatant was added to 20 μL of 100 mm potassium phosphate buffer (pH 7.0) and 160 μL of 5 m potassium iodide. Absorbance of the supernatant was read at 390 nm. The content of H₂O₂ was determined using a standard curve. Malondialdehyde was measured according to the TBARS method (Yaronskaya et al., 2003). Malondialdehyde equivalents were calculated as described (Yaronskaya et al., 2003).

Lincomycin Treatment, Protein Extraction, and SDS-PAGE

Three milligrams of leaf discs were harvested from approximately 8-week-old plants using a 5-mm-diameter biopsy punch (Kai Medical). The leaf discs were placed in a glass vial containing 5 mm lincomycin in 0.2% (v/v) Tween 20, and the vial was sealed with a rubber cap. A syringe needle was pierced through the rubber cap, and lincomycin was infiltrated into the leaf discs by repeated pressuring and depressuring of the syringe for 60 s. Leaf discs treated with lincomycin were immediately placed on wet filter papers and irradiated with three different light intensities (20, 100, and 1,200 μmol m⁻² s⁻¹). To isolate thylakoid proteins, three leaf discs were collected at each time point. SDS-PAGE sample preparation of thylakoid membrane proteins and SDS-PAGE were carried out as described previously (Kato et al., 2007). Sample loadings were normalized based on fresh weight.

In Vivo Labeling of Chloroplast Proteins

The in vivo labeling was carried out as described previously (Bonardi et al., 2005) with slight modifications. To label the chloroplast proteins, leaf discs of approximately 8-week-old plants were vacuum infiltrated by syringe and incubated with 0.1 mCi mL⁻¹ [³⁵S]Met in 0.2% Tween 20 at a light intensity of 100 μmol m⁻² s⁻¹ for 60 min. After labeling, leaf discs treated with 0.1 mCi mL⁻¹ [³⁵S]Met were immediately washed twice with 10 mm unlabeled l-Met and 0.2% Tween 20 and further incubated in the presence of unlabeled l-Met at a light intensity of 600 μmol m⁻² s⁻¹ for 60, 120, 180, and 240 min after labeling. Two leaf discs were collected to isolate thylakoid proteins at each time point. SDS-PAGE sample preparation and SDS-PAGE were carried out as described previously (Kato et al., 2007). To detect labeled proteins, gels were stained by Coomassie Brilliant Blue, dried, and exposed to imaging plate (Fuji Photo Film). Radiolabels were detected by the BAS 1000 image analyzer (Fuji Photo Film).

Immunoblot Analysis

For immunoblot analysis, proteins were electroblotted onto a polyvinylidene difluoride membrane (ATTO) after SDS-PAGE. Prior to immunoreaction, the membrane was blocked with 1% (w/v) bovine serum albumin in 50 mm sodium phosphate buffer, pH 7.5, containing 155 mm NaCl and 0.05% (v/v) Tween 20 (PBST buffer) for 1 h. After two washes with PBST buffer, the membrane was incubated with anti-D1 (dilution 1:5,000). After two washes with PBST buffer, the membrane was incubated with secondary antibodies. Immunodetection was performed as described previously using the enhanced chemiluminescence system (GE Healthcare). Relative amounts of signals from immunoblots were quantified by ImageJ (developed at the U.S. National Institutes of Health and available at http://rsb.info.nih.gov/) and normalized to the amount of LHCII detected by Coomassie Brilliant Blue.

Preparation of Thylakoid Membranes and BN/SDS-PAGE

For BN/PAGE, samples were prepared according to the protocol described previously with the following modifications (Suorsa et al., 2004). The plants were low light adapted (20 μmol m⁻² s⁻¹, 10 h of light/14 h of dark) for 1 d prior to the start of light treatment. Thylakoid membranes were isolated from low light-adapted leaves and from leaves illuminated for 2 h at 150 μmol m⁻² s⁻¹. To isolate chloroplasts, 2 g of wild-type and var2-1 leaves from approximately 8-week-old plants was harvested. Chloroplasts were isolated according to the protocol described previously (Miura et al., 2007). The homogenate containing intact and broken chloroplasts was diluted 10 times with homogenation buffer and centrifuged at 60g for 2 min. The pellet was washed with 50 mm HEPES, pH 7.5, containing 5 mm mannitol and centrifuged at 2,500g for 4 min. The pellet was then resuspended in 50 mm HEPES, pH 7.5, containing 100 mm mannitol and 10 mm MgCl₂. After centrifugation at 2,500g for 4 min, the pellet was resuspended in the same buffer, and total chlorophyll concentration was measured. Thylakoid membrane suspensions containing 100 μg of chlorophyll were centrifuged at 2,500g for 4 min, and the pellet was washed with wash buffer (50 mm bis-Tris, 330 mm mannitol, pH 7.5). After centrifugation, thylakoid membranes were resuspended in buffer (25 mm bis-Tris, 20% [w/v] glycerol, pH 7.5) to a final concentration of 0.5 mg mL⁻¹ chlorophyll. To solubilize thylakoid membranes, n-dodecyl-β-d-maltoside was added to a final concentration of 0.4% (w/v). After centrifugation at 14,000g for 20 min, the supernatant was supplemented with one-tenth volume of loading buffer (50 mm bis-Tris, 500 mm ε-amino-n-capronic acid, 5% [w/v] Coomassie Brilliant Blue G-250, pH 7.5) and loaded onto a 4% to 12% gradient native gel. Electrophoresis was performed at 4°C overnight at 50 V. After electrophoresis, each lane was excised from the gel and incubated in equilibration buffer (50 mm Tris-Cl, 6 m urea, 30% [w/v] glycerol, 2% [w/v] SDS, 0.05% [w/v] BPB, 10 mm dithiothreitol, pH 8.8) for 30 min at room temperature. Then, proteins were separated by SDS-PAGE on a 10% polyacrylamide gel. After electrophoresis, the resolved gels were used for immunoblot analysis or stained using the PlusOne Silver Staining Kit (GE Healthcare) according to the manufacturer's instructions. Chlorophyll content was determined using 80% (v/v) acetone extracts of thylakoid membranes. The chlorophyll concentration was determined as described previously (Porra et al., 1989).

Fluorescence Measurements

Photosynthetic ETR was calculated as Φ II × PAR × 0.5 × 0.84, where Φ II represents the overall photochemical quantum yield (F_v/F_m′), PAR is actinic irradiance in μmol m⁻² s⁻¹, 0.5 is the assumed proportion of photons absorbed by pigments associated with PSII, and the 0.84 standard factor is the incident quanta absorbed by the leaf (for review, see Baker, 2008). Chlorophyll fluorescence for ETR using fully expanded leaves was determined using the chlorophyll fluorometer PAM-2000 (Walz Effertrich). Changes in F_v/F_m were measured by FluorCam700MF (Photon Systems Instruments). Prior to the measurement, leaf discs were kept in the dark for 10 min to fully oxidize the plastoquinone pool. The results shown are averages of three biological repeat experiments.

Recovery Treatment

The recovery treatment was carried out as described previously (Ishihara et al., 2007). Saturating light (1,200 μmol m⁻² s⁻¹) was illuminated on leaf discs until F_v/F_m reached about 50% of the initial F_v/F_m value in Col, fug1-3, var2-1, and var2-1 fug1-3. The discs were transferred under low light condition (20 μmol m⁻² s⁻¹), and the change of F_v/F_m values was measured at 30, 60, 90, 120, 180, and 360 min. Prior to the measurement, leaf discs were kept in the dark for 10 min. The chlorophyll fluorescence was measured by FluorCam700MF (Photon Systems Instruments).

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers VAR2 (NM179825), VAR1 (NM123592), and FUG1 (NM101583).

Supplemental Data

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

• Supplemental Figure S1. Plant materials used in this study.

• Supplemental Figure S2. Quantification of ROS in var2 compared with the wild type (Col).

• Supplemental Figure S3. Accumulation of D1 protein in leaf discs prepared from var2 and var2 fug1 leaves.

• Supplemental Figure S4. LHCII levels in the wild type and mutants after light irradiation.

• Supplemental Figure S5. Immunoblot analysis of D1 protein in wild-type Col.

• Supplemental Figure S6. D1 degradation in darkness.

• Supplemental Figure S7. Recovery curves of the photochemical efficiency after photoinhibition.