Proteomic evidence for genetic epistasis: ClpR4 mutations switch leaf variegation to virescence in Arabidopsis


  • Wenjuan Wu,

    1. National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China
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    • These authors contributed equally to this work.
  • Ying Zhu,

    1. National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China
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    • These authors contributed equally to this work.
  • Zhaoxue Ma,

    1. National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China
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  • Yi Sun,

    1. National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China
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  • Quan Quan,

    1. Department of Chemistry, The University of Hong Kong, Hong Kong, China
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  • Peng Li,

    1. Center for Bioinformatics and Computational Biology, Shanghai Key Laboratory of Regulatory Biology, The Institute of Biomedical Sciences and School of Life Sciences, , East China Normal University, Shanghai 200241, China
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  • Pengzhan Hu,

    1. Center for Bioinformatics and Computational Biology, Shanghai Key Laboratory of Regulatory Biology, The Institute of Biomedical Sciences and School of Life Sciences, , East China Normal University, Shanghai 200241, China
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  • Tieliu Shi,

    1. Center for Bioinformatics and Computational Biology, Shanghai Key Laboratory of Regulatory Biology, The Institute of Biomedical Sciences and School of Life Sciences, , East China Normal University, Shanghai 200241, China
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  • Clive Lo,

    1. School of Biological Sciences, The University of Hong Kong, Hong Kong, China
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  • Ivan K. Chu,

    1. Department of Chemistry, The University of Hong Kong, Hong Kong, China
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  • Jirong Huang

    Corresponding author
    1. National Key Laboratory of Plant Molecular Genetics, Institute of Plant Physiology and Ecology, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences, Shanghai 200032, China
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Chloroplast development in plants is regulated by a series of coordinated biological processes. In this work, a genetic suppressor screen for the leaf variegation phenotype of the thylakoid formation 1 (thf1) mutant combined with a proteomic assay was employed to elucidate this complicated network. We identified a mutation in ClpR4, named clpR4-3, which leads to leaf virescence and also rescues the var2 variegation. Proteomic analysis showed that the chloroplast proteome of clpR4-3 thf1 is dominantly controlled by clpR4-3, providing molecular mechanisms that cause genetic epistasis of clpR4-3 to thf1. Classification of the proteins significantly mis-regulated in the mutants revealed that those functioning in the expression of plastid genes are oppositely regulated while proteins functioning in antioxidative stress, protein folding, and starch metabolism are changed in the same direction between thf1 and clpR4-3. The levels of FtsHs including FtsH2/VAR2, FtsH8, and FtsH5/VAR1 are greatly reduced in thf1 compared with those in the wild type, but are higher in clpR4-3 thf1 than in thf1. Quantitative PCR analysis revealed that FtsH expression in clpR4-3 thf1 is regulated post-transcriptionally. In addition, a number of ribosomal proteins are less expressed in the clpR4-3 proteome, which is in line with the reduced levels of rRNAs in clpR4-3. Furthermore, knocking out PRPL11, one of the most downregulated proteins in the clpR4-3 thf1 proteome, rescues the leaf variegation phenotype of the thf1 and var2 mutants. These results provide insights into molecular mechanisms by which the virescent clpR4-3 mutation suppresses leaf variegation of thf1 and var2.


Chloroplast development is coordinately regulated by a large number of genes, which could be involved in signaling transduction, gene expression, plastome stability, photosynthesis, and metabolism (Lopez-Juez and Pyke, 2005; Pogson and Albrecht, 2011). To date, a large number of mutants defective in chloroplast development have been identified and clustered into four classes based on the severity of phenotypes in plant growth and development: embryo lethal, seedling lethal or albino, seedling lethal but conditional survival, and autotrophy with pale-green, virescent, or variegated leaves (Kim et al., 2009). Among these mutants, leaf variegation has been used as a model system to elucidate molecular mechanisms of chloroplast development (Sakamoto, 2003; Aluru et al., 2006; Yu et al., 2007). In Arabidopsis, dozens of leaf-variegated mutants including var1/ftsh5, var2/ftsh2, im, and thf1 have been reported; of these var2/ftsh2 is best characterized in terms of physiological and biochemical functions (Putarjunan et al., 2013).

FtsH2 is an ATP-dependent metalloprotease which forms a hexameric complex with other three subunits – FtsH1, VAR1/FtsH5, and FtsH8 (Yu et al., 2004, 2005). FtsH protease is directly involved in turnover of the photosystem II (PSII) reaction center D1 protein and other processes required for chloroplast development (Lindahl et al., 2000; Zaltsman et al., 2005; Kato et al., 2009). The four FtsH subunits are divided into two groups: type A (FtsH1 and FtsH5) and type B (FtsH2 and FtsH8). Members in the same type are functionally redundant, and loss of function of either type results in embryonic or seedling lethality. Based on these findings, a threshold model has been proposed to explain the mechanism of var2 variegation; namely, above the threshold of FtsH activity normal chloroplasts are produced while below the threshold white plastids occur (Chen et al., 2000). The FtsH threshold is dynamic from plastid to plastid due to their intrinsic differences in a cell (Liu et al., 2010a). This model has gained genetic support from the study of the var2 suppressors. Either enhancing FtsH expression (Yu et al., 2004, 2005; Zhang et al., 2009) or reducing the FtsH threshold can rescue the variegation phenotype in the absence of FtsH2 (Yu et al., 2008; Liu et al., 2010b). In addition, Miura et al. (2007) proposed another model in which the balance between protein biosynthesis and degradation is an important factor determining chloroplast development in var2, based on the finding that mutations reducing protein biosynthesis in plastids suppress leaf variegation of var2. In essence, this hypothesis is in agreement with the threshold model, because reduced biosynthesis of plastid protein may lower the threshold for chloroplast development. However, molecular evidence is lacking to support the idea that suppressor genes could reduce the threshold level of FtsH protease.

Among reported var2 suppressors, three suppressors (clpC1, clpC2, and clpR1) were found to be subunits of the ATP-dependent caseinolytic protease (Clp) (Park and Rodermel, 2004). In Arabidopsis, the Clp system is composed of the Clp core complex, which is formed with the heptameric P-ring (Clp3–6 subunits) and R-ring (ClpP1, ClpR1–4 subunits), two plant-specific subunits (ClpT1 and -T2) required for the assembly of the Clp core complex, three Clp AAA+ chaperones (ClpC1, -C2, and -D) to recruit the unfolded substrates into the narrow proteolytic chamber of ClpP, and a potential substrate affinity regulator ClpS (Adam et al., 2001; Peltier et al., 2004; Clarke et al., 2005; Sjogren and Clarke, 2011). In addition, an unfoldase (ClpB3) is present in Arabidopsis, and has been demonstrated to play an important role in the removal of aggregated proteins coordinately with Clp (Zybailov et al., 2009). Recent reports have demonstrated that knockout of Clp leads to embryo or seedling lethality (Kuroda and Maliga, 2003; Kim et al., 2009), while knockdown leads to leaf virescence (Rudella et al., 2006; Sjogren et al., 2006; Zheng et al., 2006; Koussevitzky et al., 2007). Thus, like FtsH, Clp protease is essential for chloroplast development as well as plant development and growth.

THF1, also called Psb29, is encoded by a single copy gene conserved in oxygenic photosynthetic organisms ranging from cyanobacteria to flowering plants, and has been proposed to participate in the biogenesis of PSII (Keren et al., 2005; Huang et al., 2013). Loss of function of THF1 leads to leaf variegation in Arabidopsis (Wang et al., 2004). We previously reported that FtsH protease is a key factor controlling the leaf variegation phenotype of the thf1 mutant, and that overexpressing the constitutively active form of the G-protein alpha subunit (GPA1) rescues leaf variegation of both thf1 and var2 (Zhang et al., 2009). To elucidate the mechanism of thf1-mediated leaf variegation, we performed genetic screening for the second-site suppressors of thf1. In this study, we found that clpR4 suppresses the leaf variegation phenotype of both thf1 and var2 at least in part via an increase in FtsH accumulation.


ClpR4 mutation suppresses thf1 leaf variegation

To elucidate the molecular mechanisms underlying thf1-mediated leaf variegation, we created an ethyl methanesulfonate (EMS) mutagenized population in the thf1 genetic background. Eight thf1 suppressor lines without leaf variegation were isolated from 8000 independent M2 lines. 47-2, one of the suppressor lines, was selected for further analysis. 47-2 displays pleiotropic phenotypes, such as virescent and serrated leaves and a smaller plant body than the wild-type (WT) (Figure 1a,b). Genetic analysis showed that leaf variegation is suppressed by a single recessive mutation, designated suppressor of thf1 7 (sot7), in 47-2. The sot7 single mutant isolated from the F2 progenies of the cross between 47-2 and WT exhibits the same phenotype as 47-2 (Figure 1a). Analysis of the chlorophyll content demonstrated that sot7 and 47-2 mutants contained about 43% of the WT chlorophyll content in young leaves, reaching about 65% of WT in mature leaves (Figure 1c). This was consistent with the ultrastructural change in the chloroplasts observed during 47-2 and sot7 leaf greening. Yellow leaves from 47-2 and sot7 had smaller chloroplasts and less thylakoid membrane than those from WT (Figure 1d). The ultrastructural difference between WT and 47-2 or sot7 disappeared when the leaf turned green (Figure 1d). In contrast, the plastids in the yellow sector of thf1 leaves had many membranous vesicles. These data indicate that sot7 is epistatic to thf1 in the regulation of chloroplast development.

Figure 1.

Identification of a thf1 suppressor line 47-2. (a) Phenotypes of 30-day-old wild type (WT), thf1, sot7, and 47-2 plants grown under short-day conditions (8-h light/16-h dark). Bars = 1 cm. (b) Phenotypes of 10-day-old WT, thf1, sot7, and 47-2 seedlings. (c) Chlorophyll contents in green and yellow leaves of 30-day-old plants. FW, fresh weight. Error bars are SD; n = 4. (d) Chloroplast ultrastructure observed in WT, thf1 (yellow), sot7 (green and yellow), and 47-2 (green and yellow) leaves of 30-day-old plants. Bars = 1 μm. Red arrows indicate membranous vesicles.

To clone sot7, we established an F2 mapping population by crossing 47-2 with Landsberg erecta (Ler). Map-based cloning results showed that the mutation site was located within the 360-kb region between simple sequence length polymorphism (SSLP) markers FCA6-37 and FCA8 on the lower arm of chromosome 4 (Figure 2a). Since sot7 displays a very similar phenotype to the reported mutants defective in Clp subunits (Rudella et al., 2006; Koussevitzky et al., 2007) and the ClpR4 gene is located in this region, we sequenced the genomic fragment of ClpR4 (At4 g17040) amplified from 47-2. A single nucleotide mutation from G to A was detected at the start site of the second intron in ClpR4 (Figure 2a). Reverse transcriptase-PCR analysis showed that the clpR4 locus produced three different transcripts in sot7 (Figure 2b), while DNA sequencing revealed that the longest transcript contained intron 2 (band A) while other two transcripts were mis-spliced products (band B and C) (Figure 2c). Interestingly, the shortest transcript (band C) can be translated into a protein that is five amino acids longer than the endogenous ClpR4. The other two transcripts contain a premature stop codon. Given that knock-out mutants of ClpR4, e.g. clpR4-1 and clpR4-2, are seedling lethal (Kim et al., 2009), we infer that the clpR4 protein is partially functional in the sot7 mutant. Thus, we renamed sot7 as clpR4-3.

Figure 2.

SOT7 encodes a caseinolytic protease (Clp) subunit ClpR4. (a) Map-based cloning identifies a mutation of G to A at the second intron of ClpR4. (b) Three mis-spliced mRNAs (A, B, C) amplified in clpR4-3. gDNA, genomic DNA. (c) The mis-spliced sequences in (b). (d) The clpR4-3 leaf virescence is complemented by ClpR4. Bars = 1 cm. (e) 47-2 is recaptured by ClpR4. The transgenic plant was screened from the cross between 47-2 and a clpR4-3 complementation line. Bars = 1 cm. Thirty-day-old plants grown under 100 μmol photons m−2 sec−1 at 22°C were photographed.

To validate the map-based cloning result, the full-length ClpR4 cDNA under the control of the cauliflower mosaic virus 35S promoter was introduced into clpR4-3 and 47-2. Ectopic expression of ClpR4 rescued clpR4-3 and 47-2 to the WT and thf1 phenotypes, respectively (Figure 2d,e). Furthermore, overexpression of the full-length ClpR4 cDNA fused to green fluorescent protein (p35S:ClpR4-GFP) also rescued the mutant phenotype of clpR4-3 plants. The GFP signal was overlapped with the chlorophyll autofluorescence, indicating that ClpR4 is localized in chloroplasts (Figure S1 in Supporting Information). Taken together, our data suggest that thf1 leaf variegation in the suppressor line 47-2 is suppressed by clpR4-3.

clpR4 partially rescues thf1 hypersensitivity to high light

To know whether clpR4 could rescue the hypersensitivity of thf1 to high light (Keren et al., 2005), we measured the ratio of variable fluorescence to maximum fluorescence (Fv/Fm) in the green part of mutant and WT leaves. The Fv/Fm ratio reflects the maximal photochemical efficiency of PSII photochemistry. As shown in Figure 3, PSII activity was significantly lower in thf1 (0.09 ± 0.03) than clpR4-3 (0.28 ± 0.05) or WT (0.45 ± 0.02) after exposure to high light for 6 h. Fv/Fm values of 47-2 were between those of the single thf1 and clpR4-3 mutants. Likewise, clpR4-3 and 47-2 showed faster recovery of Fv/Fm than thf1 in the dark, but did not reach the WT value (Figure 3). These results indicate that clpR4 partially rescues the hypersensitivity of thf1 to high light.

Figure 3.

Photosystem II (PSII) activity of 40-day-old wild type (WT), thf1, clpR4-3, and 47-2 plants treated with high light. The ratio of variable to maximum fluorescence (Fv/Fm) was measured in detached leaves at indicated times. Leaves were exposed to high light (800 μmol photons m−2 sec−1) for 6 h, and then recovered (re) in the dark for 2–14 h. (Bars indicate SD; n = 4).

Reduced levels of the Clp complex suppress leaf variegation

To investigate whether clpR4-3 was able to suppress var2 leaf variegation, we generated the clpR4-3 var2 double mutant by crossing clpR4-3 with var2. Our results showed that clpR4-3 var2 displayed virescence as clpR4-3, but grew more slowly than the var2 and clpR4-3 single mutants (Figure 4a), suggesting that the two proteases function independently. We found that heterozygosity at the ClpR4 locus could also rescue var2 leaf variegation to a great degree (Figure 4a), indicating that suppression of leaf variegation by clpR4-3 is dose dependent. In turn, we addressed whether thf1 leaf variegation is suppressed by clpC1 and clpC2, which have been shown to suppress var2 leaf variegation (Park and Rodermel, 2004; Yu et al., 2008). Interestingly, neither clpC1 thf1 nor clpC2 thf1 produced variegated leaves (Figure 4b). Thus, reducing the levels of the Clp complex can suppress leaf variegation of both var2 and thf1.

Figure 4.

Reduced levels of the Clp complex suppress thf1 and var2 leaf variegation. A representative 40-day-old plant grown under long-day (16 h light/8 h dark) conditions is shown for each genotype. Red arrows indicate a white/yellow sector. Bars = 1 cm. (a) Phenotypes of var2, var2 ClpR4+/−, var2 clpR4-3, and clpR4-3 plants. (b) Phenotypes of thf1, clpC1, clpC1 thf1, clpC2, and clpC2 thf1 plants.

Levels of mature plastid rRNAs are reduced in clpR4-3

To investigate the effect of the clpR4 mutation on plastid rRNA processing, we examined levels of rRNAs (23S, 16S, 5S, and 4.5S) using RNA blotting with gene-specific probes. The plastid rrn operon shown in Figure 5(a) is transcribed as a large polycistronic precursor RNA, which is processed subsequently by a series of endo- and exo-ribonucleases to yield mature rRNAs and tRNAs (Harris et al., 1994a). Except for mature 5S and 16S rRNAs, 4.5S and three mature 23S (1.2-, 1.0- and 0.5-kb) rRNAs were clearly reduced in clpR4-3 and 47-2 mutants, compared with WT (Figure 5b). However, the mutants accumulated higher levels of several rRNA precursors, e.g. 4.5S plus 23S (23S 3.2-kb fragment) and 16S precursors, than WT (Figure 5b). In contrast, thf1 produced the same levels of rRNAs as WT (Figure S2). Thus, ClpR4 mutations lead to a slowdown of plastid rRNA processing.

Figure 5.

Effect of ClpR4 mutations on the levels of mature chloroplast rRNAs. (a) A diagram of the chloroplast rrn operon. The probes used for RNA gel blot analysis in (b) are indicated by thick black lines under the individual rRNA genes. (b) Northern blot analysis of chloroplast rRNAs in wild type (WT), clpR4-3 and 47-2 plants. One microgram of total RNA extracted from 40-day-old plants was loaded in each lane. The level of 25S rRNA stained with ethidium bromide is shown as a loading control.

The 47-2 chloroplast proteome is dominantly controlled by clpR4-3

To investigate the molecular mechanisms underlying genetic epistasis of clpR4 to thf1, we performed quantitative proteomic analysis of chloroplasts isolated from 2-week-old WT, thf1, clpR4-3, and 47-2 seedlings using isobaric tags for relative and absolute quantification (iTRAQ) (Lau et al., 2011). In triplicate analyses, we identified a total of 1629 proteins [false discovery rate (FDR) ≤ 1%] from isolated chloroplasts using the UniProt database. Among these proteins, 1071 are common to all three triplicates (Data S1). A search of The Arabidopsis Information Resource (TAIR) database showed that 105 of the 1629 proteins are produced from 52 genes, and 31 could not be linked to the currently known gene loci. Collectively, we identified 1545 genes which encode 1598 proteins (Data S1). Functional annotations obtained from the TAIR and Gene Ontologies databases revealed that, among 1545 gene products, 1313 (84.98%) were annotated in chloroplasts. The other proteins (232) could be from contamination by other organelles or true plastid-localized proteins that need to be confirmed experimentally. Proteomic data for each mutant were expressed as the ratios of the mutant versus WT protein accumulation. The Spearman's rank correlation coefficient (Sr) was used to evaluate the systematic relationship between the mutants. The results showed that the Sr between 47-2/WT and clpR4-3/WT for the 1066 proteins was the highest (0.818) (Figure 6a) while between 47-2/WT and thf1/WT it was 0.539 (Figure 6b), suggesting that the 47-2 chloroplast proteome is dominated by clpR4-3. The correlation coefficient between clpR4-3/WT and thf1/WT was 0.481 (Figure 6c), indicating that ClpR4 and THF1 function in a distinct manner.

Figure 6.

Spearman correlation coefficients between proteomes of the mutants. The scatter plots show the correlation of 47-2/WT versus clpR4-3/WT (a), 47-2/WT versus thf1/WT (b) and clpR4-3/WT versus thf1/WT (c), respectively, based on the means of the protein levels detected in two biological duplicates. WT, wild type.

To further analyze the proteomic data, we selected the roteins with statistically significant changes (P-value ≤ 0.05; threshold values smaller than 1.4-fold for underexpression and larger than 1.4-fold for overexpression) between the WT and mutants in two biological repeats, and clustered them according to metabolic pathways and protein complexes (Table 1). We found that 18, 57, and 55 proteins were significantly mis-regulated in thf1, clpR4-3, and 47-2, respectively. As expected, ClpR4 and THF1 were downregulated to the greatest extent in the proteomes of clpR4-3 and thf1, respectively. To evaluate the weight of thf1 and clpR4 on remodeling in the double mutant proteome, we classified the 55 mis-regulated proteins in 47-2 into four groups, one for common in all mutants, two for common between 47-2 and thf1 or clpR4-3, and one for unique in 47-2 (Table 1). In all mutants, six proteins (THF1, FtsH2, FtsH8, PRX Q, CPN21, and CP12-1) were commonly mis-regulated at the same direction. Of the remaining 49 proteins, the THF1 mutation contributed only one protein (2.0%) while the ClpR4 mutation contributed 26 proteins (53.1%), suggesting that the proteome of 47-2 is dominated by clpR4-3. Twenty-two proteins (44.9%) were unique for the double mutant (Table 1), indicating an additive interaction between THF1 and ClpR4. Taken together, our data provide proteomic evidence supporting the conclusion that clpR4-3 is epistatic to thf1.

Table 1. Proteins mis-regulated significantly in thf1, clpR4-3 and 47-2 chloroplast proteomes
Gene locusNamethf1/WTclpR4-3/WT47-2/WTMis-regulated in
  1. Mis-regulated proteins that are common in all or two mutants or unique for single mutants are grouped. The data are expressed by means of three repeats in two biological duplicates (B1 and B2).

  2. NA, not available (not significantly detected or not detected).

AT2G20890THF1−45.78−50.35−1.73−1.70−57.81−58.99All mutants
AT3G26060PRX Q−2.47−2.41−13.94−22.23−11.62−12.16
AT5G16710DHAR3−1.85−4.11−2.35−2.09NA−2.84thf1 and clpR4-3
AT3G43540DUF 1350−2.49−2.45−2.66−2.39NA−2.82
AT5G45680FKBP13−2.19−6.92NA−7.22−3.49−6.67thf1 and 47-2
AT5G42270FTSH5−8.13−8.33NANA−5.26NA thf1
AT5G47860DUF 1350−2.56−2.64NANA−1.85NA
AT4G17040CLPR4NANA−24.56−32.87−24.84−37.29clpR4-3 and 47-2
AT4G26900HIS HFNANA−2.20−2.46−2.25−2.43
AT1G12410CLPR2NANA−3.62−2.99−3.18NA clpR4-3
AT4G02530Lumenal proteinNA−7.53−2.37−17.02NA−9.52
AT5 g24490Ribosomal proteinNA1.80−9.75−3.00−2.81NA
AT4 g34090UnknownNANA−4.14−5.31−3.27NA
AT1G49970CLPR1NANANA−2.31−2.46−2.18 47-2
AT1G56070EF-2 likeNANANANA3.905.15
AT1G21440PEPC familyNANANANA−3.65−7.02
AT1 g56050UnknownNANANA−2.70−5.30−3.82

Accumulation of FtsH is dramatically reduced in thf1

In thf1, there were six upregulated and 12 downregulated proteins (Table 1). The levels of FtsH2/VAR2, FtsH8 and FtsH5/VAR1 were dramatically reduced in thf1. This result is consistent with our previously report that thf1 leaf variegation can be attributed to the reduced level of FtsH protease (Zhang et al., 2009). Other downregulated proteins are associated with oxidative defense (DHAR3 and PRX Q), plastid gene transcription (PTAC16), protein folding in the thylakoid lumen (FKBP13), amino acid biosynthesis (ASP5), carotenoid metabolism (CCD4), and two unknown proteins containing the DUF 1350 domain. In contrast, six upregulated proteins are involved in PSI biogenesis (PsaC), plastid protein translation (PSRP2), oxidative stress (PRXII E and ENH1), and two chaperonins, CPN21/GroES and CP12-1, that are required for the complex assembly of Rubisco and glyceraldehyde-3-phosphate dehydrogenase (GAPDH), respectively. Taken together, our proteomic results strongly support that the FtsH protease is a key mediator for THF1-regulated chloroplast development.

Mis-regulated proteins in the clpR4-3 proteome

In clpR4-3, we detected 57 mis-regulated (23 downregulated and 34 upregulated) proteins in two biological duplicates (Table 1). These proteins are mainly enriched in the processes of protein folding and degradation, PSII biogenesis, plastid gene expression, oxidative defense, and carbohydrate metabolism. Beside ClpR4, another Clp subunit, ClpR2, was also down-regulated by about three-fold. This result is consistent with the previous finding that ClpP1 and ClpR1–4 form a heptameric R-ring and are regulated at a post-translational level (Sjogren and Clarke, 2011). We found that some factors related to protein folding and unfolding factors (Cpn60B2, HSP90.5, and ClpB3) and peptidase (At5G42390) were upregulated in clpR4-3. These results imply that retrograde signaling from plastids to the nucleus may exist to alleviate the stress of mis-folded proteins accumulated in clpR4 chloroplasts. In addition, as many as 10 mis-regulated proteins in clpR4-3 are involved in the process of plastid gene expression. Of these, three (RPL1, RPL12C and AT5G24490) ribosomal subunits are downregulated. This result is consistent with the decreased levels of mature chloroplast rRNAs (Figure 5). Furthermore, translational activity in clpR4-3 was assessed by the ribosomal association of plastidic mRNAs, rbcL and psbA following sucrose gradient fractionation. The proportion of polysomes associated with unassociated transcripts allows estimation of the efficiency of translation initiation and elongation (Barkan, 1993). Analysis of polysome profiles showed that the peaks of rbcL and psbA transcripts were obviously shifted to lighter fractions in the mutants (Figure S4), suggesting that plastid translation is indeed impaired in the clpR4-3 mutants. In contrast, three RNA-binding proteins (CP33, RBP31, and RBP29), three translation elongation factors (SCO/EF-G, EF-Tu, and EF-Ts), and one exoribonuclease (PNPase) that mediates global RNA decay were upregulated. Taking these data together, we suggest that ClpR4 mutations lead to multiple effects on plastid gene expression.

Several important proteins related to PSII biogenesis/function were downregulated in the clpR4-3 proteome. Besides FtsH protease (FtsH2 and FtsH8) and THF1, the level of PsbS that plays a critical role in non-photochemical quenching in high light was also reduced. In addition, VIPP1 required for the assembly of the photosynthetic apparatus in collaboration with Alb3 and four luminal proteins (OE33/PsbO1 and PsbQ in the PSII oxygen-evolving complex, PetA in the cytochrome b6f complex and one unknown protein) were downregulated. On the other hand, five proteins (DHAR3, Fe-SOD, PRX Q, PRX M1, and GSTF8) involved in antioxidative defense were downregulated in clpR4-3, supporting our observation that clpR4-3 is hypersensitive to high light. In terms of plant metabolism, the clpR4-3 mutation led to a decline in proteins functioning in the Calvin–Benson cycle (RBCL, RPE, and SBPase), de novo fatty acid biosynthesis (CAC3), nucleotide metabolism (AMK2) and amino acid biosynthesis (GS2 and His-HF), implying that primary metabolism is compromised with slow chloroplast development. Interestingly, three chaperonins were included among the upregulated proteins, two (CP12-1 and CP12-2) of which have been reported to play an important role in the assembly of supramolecular complexes between GAPDH and phosphoribulokinase (PRK) (Marri et al., 2008) and another (CPN21) which may be involved in Rubisco assembly. In nitrogen assimilation, leaf-specific GS2 was downregulated, whereas root-specific NADH-GOGAT was upregulated. This phenomenon may be attributed to a high level of nitrogen nutrition in the growth media. Other upregulated proteins are related to starch metabolism (AMY3 and SBE2.2), chlorophyll biosynthesis (HEMB), the methylerythritol phosphate (MEP) pathway (ISPG), ABA-mediated photoprotection (PGL35), and jasmonic acid biosynthesis (LOX2). In summary, we propose that the low level of the Clp complex causes protein-folding stress, slows down plastid gene expression and PSII biogenesis, and ultimately leads to a leaf virescent phenotype.

Mis-regulated biological processes in 47-2 are similar to those in clpR4-3

As we described above, 22 unique mis-regulated proteins were identified in 47-2. However, most of these proteins function in the same processes as described for clpR4. For example, ClpR1 is a subunit of the heptameric R-ring in the ClpPR protease; RPL11, RPL9, FUG, EF-2 like, and NAI are involved in plastid gene expression; PPOX and CHLM are enzymes in chlorophyll biosynthesis; APS1, APL1, GWD1, phosphoglucomutase, PPDK, and PEPC in starch or sugar metabolism; and ASS and EAD9 in amino acid biosynthesis. Other proteins such as PER1, COR15B, and AOR have been reported to function in the resistance of plants to stress. Thus, we conclude that the biological processes in 47-2 are primarily controlled by clpR4-3.

Because of the important role of FtsH activity in controlling thf1 leaf variegation, FtsH levels were inspected in the mutants. We found that in the double mutant 47-2 type B accumulated at a slightly higher level in one biological repeat but at about 60% of that in thf1 in another repeat, whereas no significant difference in the type A level was observed (Table 1), suggesting that FtsH expression is improved in the suppressor line 47-2. Immunoblotting analysis showed that VAR2 accumulation was apparently increased in 47-2 compared with thf1 (Figure S3). Quantitative PCR analysis showed that transcript levels of FtsHs were not altered among the mutants (Figure S5), indicating that FtsH accumulation in chloroplasts is post-transcriptionally regulated. Thus, we infer that FtsH expression is enhanced and should not be a rate-limiting factor for chloroplast development in 47-2.

Leaf variegation is suppressed by mutations in ribosomal proteins

80-1 is another thf1 suppressor line that we identified from the EMS-mutagenized seeds, and displays phenotypes of leaf virescence and slow growth (Figure 7a). We cloned the mutated gene, named sot6, from 80-1 using a map-based cloning strategy. The suppressor gene, sot6, turned out to be prpl11 (Figure 7b), which encodes the plastid ribosomal large subunit 11. A single mutation from G to A was detected at the last nucleotide of the third intron in the PRPL11 gene. A RT-PCR analysis showed that the mutation leads to the generation of two differential transcripts (Figure 7c). DNA sequencing demonstrated that the whole of intron 3 was retained in the longer transcript (band A) while in the shorter one (band B) the 3′ splicing site of the third intron was 34-bp nucleotides ahead of the WT site (Figure 7d). Both transcripts are predicted to have a premature stop codon, suggesting that sot6 is a null mutant. We named sot6 as prpl11-2 following another allele mutant of prpl11-1 (Pesaresi et al., 2001). Transforming the genomic DNA of PRPL11 under the control of the 35S promoter into 80-1 led to the leaf variegation phenotype of the transgenic plants (Figure 7a), indicating that the map-based cloning result was correct. Like the other suppressor genes discussed above, prpl11-2 can suppress the var2 leaf variegation phenotype too (Figure 7e). These data are consistent with our proteomic results showing that PRPL11 accumulation was significantly reduced in 47-2. Thus, it is likely that quantitative proteomics can be utilized as an alternative tool to identify new suppressor genes for leaf variegation.

Figure 7.

Leaf variegation is suppressed by mutations in PRPL11. (a) Phenotypes of 25-day-old wild type (WT), thf1, sot6, 80-1, and 80-1 complementation plants. 80-1, a suppressor line of thf1; sot6, the prpl11-2 single mutant isolated from the F2 generation between 80-1 and WT; com, an 80-1 complementation line. (b) Map-based cloning identifies a mutation of G to A at the end of the third intron in the PRPL11 gene. (c) Two mis-spliced mRNAs (A, B) amplified from prpl11-2. gDNA, genomic DNA. (d) The mis-spliced sequences in (c). B and A contains intron three with a premature stop codon, and band B is a mis-spliced product with a premature stop codon. (e) PRPL11 mutations also suppress leaf variegation of var2.


We previously reported genetic evidence that the leaf variegation of thf1 is attributed to the low activity of FtsH protease (Zhang et al., 2009). In this study, we provide several lines of evidence to support this conclusion. First, clpR4-3, identified as the first suppressor for the leaf variegation phenotype of thf1, can also rescue the phenotype of var2; in turn two var2 suppressors (clpC1 and clpC2) also suppress thf1 leaf variegation. Second, chloroplast proteomic analysis revealed that in thf1 type B subunits (FtsH2 and FtsH8) of the FtsH complex were the two most reduced proteins except for THF1 itself, and one (FtsH5/VAR1) of the type A subunits also dramatically decreased. Third, another suppressor gene (prpl11-2) of thf1 also rescues var2 leaf variegation. How THF1 regulates FtsH accumulation is an interesting question that needs to be investigated in the future.

Leaf virescence of clpR4-3 is associated with slow assemblies of plastid ribosome and PSII

Since complete knockout of ClpR4 results in seedling lethality (Kim et al., 2009), we infer that the leaf virescent phenotype of clpR4-3 is caused by the reduced level of Clp. Accordingly, the virescent phenotype has been observed in other clp mutants, such as clpR1, clpR2, clpC1, and antisense clpP (clpP4 and -6) mutants (Sjogren et al., 2006; Zheng et al., 2006; Koussevitzky et al., 2007). It remains unclear how decrease in the level of Clp protease causes leaf virescence. A previous study showed that chloroplasts contain lower levels of mature rRNAs but higher levels of rRNA precursors in clpR1 and clpC1 than in WT (Yu et al., 2008), indicating that rRNA processing was disturbed. In this study, we observed the same result in clpR4-3 (Figure 5). Consistently, our proteomic analysis showed that three plastid ribosomal proteins were significantly downregulated in clpR4-3. In contrast, a number of chloroplast rRNA processing factors, such as ribonuclease (PNPase/RIF10) and RNA-binding proteins (CP33, RBP31, and RBP29) (Barkan et al., 2007; Germain et al., 2011; Asakura et al., 2012), were upregulated in clpR4-3. These data imply that the limited ribosomal proteins restrict plastid ribosome assembly, and ultimately slow down the rate of plastid rRNA processing by a feedback regulatory mechanism. However, downregulation of the ribosomal proteins was not detected in clpR4-1 and clpR2-1 proteomes (Kim et al., 2009; Zybailov et al., 2009). The reason could be due to different materials or clpR4 mutations used for chloroplast proteomics analysis. In the current study, we used total proteins from isolated chloroplasts for proteomics assay, which could increase analytical resolution. On the other hand, several key proteins (SCO1, EF-Tu, and EF-Ts) functioning in plastid translation are strongly upregulated, implying that the protein translation rate may increase to compensate for the reduced amount of ribosome in clpR4. Nevertheless, polysome analysis showed that ClpR4 mutations impair plastid translation (Figure S4), which could lead to a slowdown of chloroplast development.

Concurrently, many nuclear-encoded chloroplast proteins that are components of photosynthetic complexes and involved in the complex assembly showed lower accumulation in the clpR4-3 proteome. PSII biogenesis is a crucial process for chloroplast development. It is well documented that the PSII supercomplex is composed of three components: the PSII reaction center, an oxygen evolving complex (OEC), and a light harvesting complex (Hankamer et al., 1997). Compared with WT, the clpR4-3 proteome accumulated significantly lower levels of two OEC subunits, PsbO and PsbQ, which have been recognized to be the Mn- and Ca2+/Cl-stabilizing proteins, respectively (Semin et al., 2012). This result supports the idea that assembly of PSII supercomplexes may be delayed as a result of altered subcomplex stoichiometry. In addition, two other proteins (AT4G02530 and PetA) located in the lumen were also downregulated in clpR4-3. Recently, Lo and Theg (2012) reported that VIPP1 enhances protein transport across the thylakoid membrane via the cpTat pathway. We found that VIPP1 is significantly downregulated in clpR4-3 and may be related to the reduced accumulation of luminal proteins. In general, our data demonstrate that the coordinated assembly of PSII is disturbed in clpR4-3. This result is also in agreement with the previous report that the Clp system is closely involved in the biogenesis of the photosynthetic apparatus (Kim et al., 2009).

Both ClpR4-3 and THF1 mutations affect plastid gene expression but in a distinct manner

Suppressor screens for the leaf variegation phenotype have provided us with insights into the linkage of leaf variegation with various biological processes, such as rRNA processing, translation, protein folding and complex assembly (Liu et al., 2010a). To date, however, systematic analysis of various biological processes has not been conducted in variegation mutants and their suppressors. Our proteomic analysis showed that many more proteins were significantly mis-regulated in clpR4-3 than thf1, which is in support of the ClpR4 mutation having a greater and broader effect on chloroplast development and function than does the thf1 mutation. For example, clpR4-3 also displays defects in leaf development. Importantly, we found that the chloroplast proteome of 47-2 is dominantly controlled by clpR4-3. Proteomics data revealed that nine unique mis-regulated proteins identified in thf1 actually function in the same biological processes, such as plastid gene expression, metabolism and antioxidative defense, as those mis-regulated in clpR4 and 47-2. It is interesting that mis-regulated proteins involved in plastid gene expression were oppositely regulated in thf1 and clpR4-3. For example, some ribosomal proteins upregulated in thf1 are downregulated in clpR4-3 and 47-2. These data imply that THF1 and Clp protease function in the same biological processes in chloroplasts, but in an opposite manner. It needs to be further investigated whether the oppositely regulated plastid gene expression in thf1 and clpR4 is related to the patterns of chloroplast development.

Based on the genetic results in this study and several previously reported suppressors of var2 (Liu et al., 2010a), virescent clp mutants suppress leaf variegation of var2 and thf1. The suppression is dependent on the expression level of ClpR4-3 since heterozygosity at the ClpR4 locus also rescues var2 leaf variegation to a great degree (Figure 4a). Furthermore, we found that levels of FtsH in the clpR4-3 thf1 double mutant were increased compared with those in thf1 and were regulated at the post-transcriptional level. This is also consistent with our previous reported data that overexpression of FtsH subunits can rescue thf1 leaf variegation (Zhang et al., 2009). According to the threshold model, the lack of FtsH activity in var2 can be compensated for either by an increase in FtsH per se or by lowering the threshold demand for chloroplast development (Yu et al., 2007; Liu et al., 2010a). Our data favor the hypothesis that leaf variegation is suppressed, at least in part, by an increase in FtsH levels. Liu et al. (2010b) demonstrated that svr7 is epistatic to var2 at 22°C while var2 is epistatic to svr7 at 8°C. It will be interesting to know whether the level of FtsH expression is significantly reduced at the low temperature. If an increase in FtsH expression levels is common in suppressor lines, it is critical to elucidate molecular mechanisms by which FtsH levels are regulated at a post-transcriptional level, such as pre-protein import into plastids, protein folding and complex assembly.

Lastly, we simply summarize possible mechanisms underlying the suppression of leaf variegation based on available evidence (Figure 8). In thf1 and var2, the level of FtsH protease is under the threshold required for chloroplast development. Most of the suppressor lines display a leaf virescent phenotype, whereas a small number of them have no defect in chloroplast development. It is worth noting that leaf virescence and variegation are two distinctive patterns controlling chloroplast development. Chloroplasts ultimately form in mature leaves of virescent mutants while plastids stop at the very early development stage in the white/yellow sector of leaf-variegated mutants (Wang et al., 2004; Sjogren et al., 2006). We divide suppressor genes of thf1 and var2 into two classes. One class functions to promote expression of FtsH genes and/or FtsH import into plastids. An example of this class is GPA1 (Zhang et al., 2009). Activation of GPA1 stimulates expression of FtsH genes and probably also import of FtsH apoprotein into chloroplasts via unknown mechanisms. Another class plays a role in removing the FtsH threshold by enhancing FtsH activity (FtsH folding and complex assembly) and/or by reducing the FtsH threshold by impairing gene expression in plastids. These suppressor genes include svr1, svr2, svr3, svr4, svr7, svr8, clpc1, clpc2, pdf1b, fug1, and sco1 (Putarjunan et al., 2013). Many of these suppressors directly or indirectly inhibit the process of plastid gene expression. We found that the clpR4-3 mutation results in an enhanced level of FtsH as well as impaired plastid gene expression in the suppressor line. More evidence is required to distinguish whether leaf variegation is rescued by an increase in FtsH accumulation or a decrease in the FtsH threshold. To this end, quantitative proteomics will be a powerful tool to elucidate molecular mechanisms underlying the suppression of leaf variegation in the future.

Figure 8.

Summary of possible mechanisms underlying suppression of thf1 and var2 leaf variegation. Based on the threshold model, a certain level of FtsH protease is required for chloroplast development, and leaf variegation can be suppressed by removing the threshold either via an increase in FtsH per se or via a decrease in the FtsH threshold. N, nucleus; C, chloroplast.

Experimental Procedures

Plant materials, map-based cloning, plasmid construction, and transformation

All Arabidopsis plants used in this study were of Columbia ecotype background, except for the Ler for map-based cloning. Seeds were surface-sterilized and stratified for 3 days, and then germinated on half-strength Murashige and Skoog (MS) medium with 1% sucrose under approximately 100 μmol photons m−2 sec−1 (photo-period of 16-h light/8-h dark) at 22°C. Seven-day-old seedlings were transferred to soil and grown in greenhouse under long-day (16-h light/8-h dark) or short-day (8-h light/16-h dark) conditions. Two suppressor lines, 47-2 and 80-1, for thf1 leaf variegation were crossed to Ler, respectively, to generate the mapping populations. The clpR4-3 (sot7) and prpl11-2 (sot6) single mutants were isolated from F2 generations of the crosses of 47-2 × Col and 80-1 × Col, respectively. Genomic DNA was isolated from F2 seedlings showing the suppressor-like phenotype. Linkage analysis was performed by SSLP markers ( Plasmid con-struction and plant transformation were performed as described by Zhou et al. (2008). All primers used in this study are listed in Table S1. The GFP fluorescence was observed with a confocal laser scanning microscope (FITC488, Zeiss LSM500,

Analyses of chlorophyll content, PSII activity, and chloroplast ultrastructure

Chlorophyll was extracted with 80% acetone at 4°C for 24 h in the dark. Chlorophyll content was determined by measuring the absorbance at 652 nm using a spectrophotometer (UV-2102PCS, UNIC, Chlorophyll fluorescence emission was measured by an Imagining PAM 101 (Walz, according to the manufacturer's instructions. Forty-day-old leaves were cut off and put on a piece of wet paper. After dark adaption for 15 min, the ratio of variable to maximum fluorescence (Fv/Fm) was measured. Then, the detached leaves were exposed to high light (800 μmol photons m−2 sec−1) for 6 h and recovered in the dark. Fv/Fm was measured at the indicated times during the process. To examine chloroplast ultrastructure, leaves from 40-day-old WT and mutant plants grown in short-day conditions were cut into small pieces and fixed in a solution of 4% glutaraldehyde, and then processed, embedded, and viewed under transmission electron microscopy as described by Harris et al. (1994b).

RT-PCR, quantitative PCR, polysome experiments, RNA blotting, and immunoblotting analyses

Total RNA was isolated using the RNAgents kit (Promega, according to the manufacturer's protocol. Semi-quantitative RT-PCR was carried out using a reverse transcription system (Promega). Quantitative PCR was performed using SYBR® Premix Ex Taq™ (Takala, with the primers in Table S1. Polysomes were isolated from leaf tissue of 4-week-old plants as described (Barkan, 1998). Northern blotting was conducted as described by Wu et al. (2011). The gene-specific primers used to label the probes are listed in Table S1. Total protein was extracted and quantified using Bio-Rad Protein Assay Dye Reagent ( Following SDS-PAGE, proteins were transferred to Hybond-ECL nitrocellulose membrane and probed with an antibody raised against VAR2. Signals were detected using a chemiluminescence detection system (Amersham Biosciences, according to the manufacturer's instructions.

Intact chloroplast isolation and quantitative proteomic analysis

Sixteen-day-old seedlings grown on 1/2 MS media were homogenized with SHE buffer (330 mm sorbitol, 50 mm HEPES-KOH, 2 mm EDTA-2Na, 5 mm ascorbic acid, pH 7.8) and filtered through Miracloth (Calbiochem, Chloroplasts were precipitated by low-speed centrifugation at 1000 g for 5 min. Then they were resuspended in SHE buffer and purified on a 40/70% discontinuous Percoll gradient. After centrifugation at 4500 g for 5 min, intact chloroplasts were taken from the interface of 70 and 40% Percoll, and washed twice with SHE buffer. All steps were performed at 4°C. Two independent biological replicates of intact chloroplasts for each genotype were isolated using 16-day-old seedlings grown under the same conditions.

For proteomic analysis, proteins were extracted from chloroplasts by 0.7% SDS at 95°C for 5 min. For each biological replicate, 100 μg of proteins from each genotype were concentrated to 50 μl, followed by the addition of SDS, TCEP [tris(2-carboxyethyl)phosphine] and methyl methanethiosulfonate (MMTS) for cysteine blocking according to the instructions of the iTRAQ Reagents kit. Samples were proteolyzed with 1:33 sequencing-grade trypsin at 37°C overnight. Each trypsinized sample was labeled with a corresponding iTRAQ label following the manufacturer's instructions. For iTRAQ, three replicates were performed in each biological replicate as described by Lau et al. (2011).

All mass spectrometry (MS) data were generated using an AB Sciex QSTAR XL or Pulsar quantitative time-of-flight mass spectrometer (AB Sciex,, controlled using Analyst QS 1.1 software. The acquired MS/MS data were analyzed directly using the Paragon algorithm in the ProteinPilot 4.0 software from AB Sciex. Briefly the MS/MS spectra were matched against theoretical spectra generated from sequences in the UniProt15.8 Arabidopsis thaliana database of the UniProtKB database ( (Boutet et al., 2007). Trypsin was set as the enzyme; MMTS was set as the cysteine alkylation agent. Thorough identification with biological modification settings, which contained a number of built-in biological and artifact peptide modifications, was performed; for all quantification experiments, iTRAQ 8-plex peptide-labeled quantification was selected. Precursor mass accuracy and product ion mass accuracy were predetermined by the software according to the instrument settings chosen. The identified peptides from the Paragon algorithm were grouped into minimal non-redundant protein sets by the ProGroup algorithm of the software. For protein identifications to be considered, a minimal unused ProtScore of 2.0 was required. The FDR of protein identification was analyzed using the Proteomics System Performance Evaluation Pipeline add-on of ProteinPilot and a decoy database of reverse sequences. A 5% cutoff of the local (or instantaneous) FDR was used for protein identification; this value is more accurate than the usually calculated global FDR, and thus improves the confidence level of protein identifications. The mass spectrometry proteomics data including all peptides identified in this study and the peptide quantification have been deposited in the ProteomeXchange Consortium ( via the PRIDE partner repository (Vizcaino et al., 2013) with the dataset identifier PXD000166.


This study was supported by grants from the National Science Fund for Distinguished Young Scholars (30925005), the National Natural Scientific Foundation of China (31070214 and 31171264), National Special Grant for Transgenic Crops (2011ZX08009-003-005) and the postdoctoral fund of SIBS (2012KIP508). The VAR2 antibody was kindly provided by Professor Sakamoto (Okayama University).