Repression of Essential Chloroplast Genes Reveals New Signaling Pathways and Regulatory Feedback Loops in Chlamydomonas

Silvia Ramundo; Michèle Rahire; Olivier Schaad; Jean-David Rochaix

doi:10.1105/tpc.112.103051

. 2013 Jan 4;25(1):167–186. doi: 10.1105/tpc.112.103051

Repression of Essential Chloroplast Genes Reveals New Signaling Pathways and Regulatory Feedback Loops in Chlamydomonas^{^[W]}

Silvia Ramundo ^a, Michèle Rahire ^a, Olivier Schaad ^b, Jean-David Rochaix ^a,¹

PMCID: PMC3584532 PMID: 23292734

Conditional repression of two essential chloroplast genes involved in plastid transcription and translation in the alga Chlamydomonas reinhardtii leads to cell growth arrest and overaccumulation of many plastid transcripts. It also identifies multiple negative regulatory chloroplast feedback loops and alters expression of nuclear genes implicated in chloroplast biogenesis, protein turnover, and stress.

Abstract

Although reverse genetics has been used to elucidate the function of numerous chloroplast proteins, the characterization of essential plastid genes and their role in chloroplast biogenesis and cell survival has not yet been achieved. Therefore, we developed a robust repressible chloroplast gene expression system in the unicellular alga Chlamydomonas reinhardtii based mainly on a vitamin-repressible riboswitch, and we used this system to study the role of two essential chloroplast genes: ribosomal protein S12 (rps12), encoding a plastid ribosomal protein, and rpoA, encoding the α-subunit of chloroplast bacterial-like RNA polymerase. Repression of either of these two genes leads to the arrest of cell growth, and it induces a response that involves changes in expression of nuclear genes implicated in chloroplast biogenesis, protein turnover, and stress. This response also leads to the overaccumulation of several plastid transcripts and reveals the existence of multiple negative regulatory feedback loops in the chloroplast gene circuitry.

INTRODUCTION

An important hallmark of eukaryotic photosynthetic organisms is the presence of a genetic system consisting of a genome and an autonomous protein synthesizing system within their plastids that cooperates closely with its nucleocytosolic counterpart in the biogenesis and functioning of chloroplasts. It is now well accepted that the origin of chloroplasts can be traced to an endosymbiotic event in which a cyanobacterium invaded an ancient host cell and became established in the cytoplasm followed by a massive transfer of genetic information to the host nucleus. However, it is not yet clear why this gene transfer was not fully completed and why a genome of modest size has been preserved in chloroplasts during evolution.

Chloroplast genes can be categorized into three major groups. The first includes genes coding for several subunits of the photosynthetic complexes photosystem II (PSII), photosystem I (PSI), cytochrome b₆f complex, ATP synthase, and ribulose-1,5-bis-phosphate carboxylase/oxygenase (Rubisco) as well as for assembly factors of some of these complexes. The second group includes genes involved in chloroplast gene expression, in particular those coding for subunits of the plastid DNA-dependent RNA polymerase, for rRNAs and tRNAs, for numerous plastid ribosomal proteins, and in some cases for the elongation factor EF-Tu. The third group consists of genes with various functions related to proteolysis, heme attachment, and general metabolism. In addition, several chloroplast genes have been identified whose function is still unknown.

Because homologous recombination occurs in the chloroplast, it has been possible to inactivate specific chloroplast genes with the aadA selectable marker conferring antibiotic resistance through chloroplast transformation in Chlamydomonas reinhardtii (Goldschmidt-Clermont, 1991; Boynton and Gillham, 1993) and tobacco (Nicotiana tabacum; Svab and Maliga, 1993). In this way, chloroplast reverse genetics has been used for elucidating the function of numerous chloroplast genes. Chloroplast genomes are present in multiple copies, and it is therefore necessary in most cases to disrupt all of the gene copies to study the phenotype resulting from a particular plastid gene disruption. Such a homoplasmic state can readily be achieved if the function of the disrupted gene is not essential, as is the case for genes involved in photosynthesis when the transformed strain is grown in the presence of a source of reduced carbon, which renders photosynthetic activity dispensable in C. reinhardtii. However, attempts to disrupt essential plastid genes resulted in a heteroplasmic state with a mixture of wild-type copies necessary for supporting growth and disrupted copies that confer resistance to the antibiotic used to select the transformants (Goldschmidt-Clermont, 1991). In the case of C. reinhardtii, it is likely that any gene involved in chloroplast gene expression is essential because among the numerous mutants isolated and characterized, none is completely deficient in chloroplast gene expression.

This hypothesis was further strengthened during attempts to disrupt the genes of the chloroplast RNA polymerase, which resulted in a heteroplasmic state (Goldschmidt-Clermont, 1991; Rochaix, 1995). However, one problem associated with these studies is that the selection of the transformants relies on the expression of aadA used as a selectable marker. It is therefore possible that the inability of obtaining homoplasmic mutants is due to the fact that such a homoplasmic state would prevent expression of aadA. Based on the criterion of heteroplasmy induced upon disruption of a chloroplast gene by transformation, other plastid genes of C. reinhardtii appear to be essential. They include clpP1 (Majeran et al., 2000), encoding a catalytic subunit of the ATP-dependent Clp protease, and ORF1995, which encodes a protein of unknown function (Boudreau et al., 1997).

A repressible/inducible chloroplast gene expression system should allow one to conditionally inactivate these essential chloroplast genes and study their function. Our previous studies (Surzycki et al., 2007) have already shown that artificial regulation of the chloroplast psbD mRNA of the PSII D2 reaction center protein can be achieved by placing the nuclear Nac2 gene under the control of the copper-repressible nuclear Cyc6 promoter (Merchant and Bogorad, 1987). In this system, conditional repression of chloroplast gene expression is based on the fact that the major target site of Nac2 is within the 5′-untranslated region (5′UTR) of the psbD mRNA and that this region is sufficient to confer Nac2-dependent expression for any chloroplast gene when it is driven by the psbD 5′UTR (Nickelsen et al., 1994). Although this system can be used for controlling expression of a heterologous gene (Surzycki et al., 2007), we have not succeeded in using it for the analysis of essential chloroplast genes.

Here, we established a new system based on the thiamine pyrophosphate (TPP)–responsive riboswitch from C. reinhardtii in which gene expression is regulated by exogenous thiamine (vitamin B1) (Croft et al., 2007). In this alga, the addition of thiamine to the culture prevents expression of Thi4 and ThiC as a result of alternative splicing of their transcripts (Croft et al., 2007). Using the Nac2 gene and the Thi4 5′UTR, which contains the TPP riboswitch, we engineered a strain in which we are able to control psbD expression with thiamine in a reversible way and to achieve conditional downregulation of essential plastid genes.

This system has allowed us to investigate the role of plastid translation and the function of the chloroplast RNA polymerase in C. reinhardtii. In particular, we used it for repressing rps12, a chloroplast gene that encodes the ribosomal protein Rps12 and rpoA, which encodes the α-subunit of the chloroplast DNA-dependent RNA polymerase. Our results demonstrate that both of these genes are essential for photo- and heterotrophic cell growth and survival of C. reinhardtii. This is in contrast with the situations in tobacco in which transplastomic plants with a homoplasmic disruption of the rpoB gene are still viable (Allison et al., 1996) and in barley (Hordeum vulgare) mutants in which plastid translation is fully deficient (Hess et al., 1994a). Similarly, in maize (Zea mays) or rapeseed (Brassica napus), embryo development is not arrested in the absence of chloroplast translation (Bryant et al., 2011; Lloyd and Meinke, 2012), whereas loss of plastid translation results in embryo lethality in Arabidopsis thaliana (Lloyd and Meinke, 2012). We show that, similar to the case of land plants, abrogation of either chloroplast protein synthesis or transcription in C. reinhardtii activates plastid-to-nucleus signaling pathways, causing the induction of several nuclear-encoded stress-responsive plastid proteins prior to cell death. We also identified a chloroplast feedback control system that compensates for the decreased level of translation or transcription of some important plastid mRNAs by increasing their stability or by altering their processing. Finally, understanding the function of essential chloroplast genes will shed new light on the question of why chloroplast genomes have been conserved throughout evolution since their endosymbiotic origin.

RESULTS

Experimental Design of a Repressible Chloroplast Gene Expression System

The experimental design is shown in Figure 1A. The nuclear Nac2 gene was fused to the Thi4 5′UTR, which contains a TPP riboswitch (Croft et al., 2007) (see Supplemental Figure 1 online). As a promoter, we included a region upstream of the MetE gene. Transcripts for MetE had previously been shown to be repressed by the presence of exogenous vitamin B12 (Croft et al., 2005), so this was another possible regulatory element. The construct (detailed in Supplemental Data Set 1 online) was introduced into the nac2-26 mutant strain of C. reinhardtii, which is deficient in Nac2 expression and therefore unable to perform photosynthesis (Kuchka et al., 1989) (Figure 1A). The Nac2 protein is targeted to the chloroplast where it is specifically required for the expression of the chloroplast psbD gene encoding the PSII reaction center protein D2, which in turn is needed for photoautotrophic growth. The transformants were then tested for restoration of growth on minimal medium in the absence of the vitamins B12 and thiamine, but not in their presence. In this way, we selected one transformant called Rep112 (Figure 1B). In order to be able to use this repressible/inducible system for any chloroplast gene without affecting phototrophic growth, we replaced the promoter and 5′UTR of psbD with those of psaA, thus making psbD expression no longer dependent on Nac2 (Figure 1A). As expected, all the homoplasmic transformants obtained with the psaA-psbD construct were able to grow on minimum medium in the presence or absence of vitamins and one of them, called A31, was chosen for further studies (Figure 1B; see Supplemental Table 1 online).

In photosynthetic organisms, F_v/F_m, the ratio between variable and maximum fluorescence provides a measure of the maximum quantum efficiency of PSII. Since the Nac2 protein is required for PSII activity in the Rep112 strain, the decline of F_v/F_m represents the repression of Nac2 expression. In Rep112, F_v/F_m decreased to nearly undetectable levels after 24 h of vitamin treatment but remained constant in the wild type and A31 (Figure 1C). Growth of Rep112 was retarded in comparison to the wild type (Figure 1D). In a reciprocal experiment, cells of Rep112 were first grown in the presence of vitamins. Upon replacing the medium with vitamin-free medium, F_v/F_m and growth were measured at different time points. PSII activity was recovered between 36 and 48 h and reached its maximum value after 72 h (Figures 1E and 1F).

The amount of Nac2 protein in Rep112 and A31 in the absence of vitamins was determined using immunoblotting. Mature Nac2 protein with a mass of 130 kD accumulated between 10 and 15% compared with the wild type in Rep112 but was undetectable in A31 (see Supplemental Figure 2 online). In spite of the greatly diminished amount of Nac2 in A31 compared with Rep112, both of these strains produced nearly equal amounts of psbD mRNA, close to 50% of wild-type levels in the absence of vitamins (Figure 2A). In the presence of vitamins, the psbD RNA was undetectable in Rep112 but accumulated normally in A31 (Figure 2A). Immunoblot analysis revealed that in Rep112, D2 protein accumulated to nearly 50% of wild-type levels in the absence of vitamins and was undetectable in the presence of vitamins (Figure 2B). By contrast, in A31, D2 was produced in the absence or presence of vitamins but in slightly higher amounts compared with Rep112, suggesting that the psaA-psbD chimeric RNA is translated more efficiently than the authentic psbD mRNA (Figure 2B). As expected, in these two strains, other chloroplast proteins, such as ClpP1 and PsaD, accumulated as in the wild type in the presence or absence of vitamins (Figure 2B).

Figure 2. — Vitamin-Mediated Repression and Induction of *psbD* Expression.

**(A)** RNA gel blot analysis of *psbD* in the indicated strains in the absence and presence of vitamins (vit). The *psaB* mRNA was used as control. A dilution series of wild-type (WT) extracts was used for estimating the amount of RNA.

**(B)** Immunoblot analysis of the chloroplast proteins D2, PRK, PsaD, and ClpP1 in the indicated strains.

**(C)** Immunoblot analysis of Nac2, ThiC, and Hsp90 in Rep112. Protein samples were examined at the indicated times after addition of vitamins.

**(D)** RNA gel blot analysis of *psbD* RNA in Rep112. The *atpB* mRNA was used as a loading control. RNA samples were examined at the indicated times (hours) after addition of vitamins.

**(E)** Immunoblot analysis of D2, D1, PRK, and ClpP1 in Rep112. Protein samples were examined at the indicated times (hours) after addition of vitamins.

**(F)** Time course of *psbD* mRNA recovery after removal of vitamins. RNA gel blot analysis was performed at different times after removal of vitamins with the indicated probes.

**(G)** Time course of D2 recovery after removal of vitamins. Immunoblot analysis was performed at different times with antibodies against D2 and the indicated proteins.

**(H)** Accumulation of D2 in the absence (−) or presence of thiamine (T) and vitamin B12 (B12) in the nac2 and Rep112 strains.

Mature Nac2 protein declined gradually upon addition of vitamins with a similar kinetics as ThiC, whose expression is controlled by an analogous TPP riboswitch (Figure 2C) (Croft et al., 2007). RNA gel blot analysis revealed that a decrease in psbD RNA was already detectable after 6 h, and the level of this RNA reached its minimal value at 14 h of vitamin treatment (Figure 2D). The decline of D2 protein was delayed and reached its minimum at 24 h due to the long half-life of the protein (Figure 2E). The amount of D1 decreased in a similar way as D2 because this protein is known to be unstable in the absence of D2 (Erickson et al., 1986). Moreover, in the absence of D2, D1 synthesis is reduced through a negative feedback exerted by unassembled D1 (Minai et al., 2006). By contrast, the levels of other chloroplast proteins encoded by either the chloroplast (ClpP1) or nuclear genome (phosphoribulose kinase [PRK]) were not affected by the addition of vitamins (Figure 2E).

When the cells were first grown in the presence of vitamins and then transferred to vitamin-free medium, the amount of psbD mRNA rose significantly between 36 and 48 h after the shift at which time it had reached its maximum (Figure 2F). The kinetics of D2 protein accumulation was similar to that of its mRNA (Figure 2G) and its partner D1 followed a similar pattern. ThiC was already detected between 24 and 36 h (Figure 2G).

To determine the range of vitamin concentrations at which the MetE promoter and the TPP riboswitch are responsive, the kinetics of inactivation of PSII in Rep112 was examined separately with different concentrations of B12 (see Supplemental Figure 3A online) and thiamine (see Supplemental Figure 3B online). These experiments showed that, at the concentrations of vitamin B12 used, the repression of PSII activity is not complete, in contrast with the addition of thiamine, where at 100 nM thiamine, the F_v/F_m is close to zero after 24 h. This demonstrates that the major contributor for the repression of Nac2 and psbD mRNA is the TPP-controlled riboswitch. The results were confirmed by immunoblotting, which showed a more significant decrease in the accumulation of D2 upon addition of thiamine versus vitamin B12 (Figure 2H), and they are in agreement with the studies of Croft et al. (2007) that showed that the Thi4 riboswitch is both necessary and sufficient for full repression of a reporter gene by thiamine. The reverse experiment after shift of the cells from vitamin-replete to vitamin-depleted medium was also performed (see Supplemental Figure 3C online).

Conditional Inactivation of Chloroplast Transcription and Translation

After having established that the vitamin-repressible riboswitch Nac2 system functions properly for controlling the expression of the chloroplast psbD gene, we tested whether it can also be used for chloroplast genes with essential functions. In this respect, genes involved in chloroplast transcription and translation are particularly interesting because it is not clear whether there is a unique chloroplast RNA polymerase in C. reinhardtii and whether chloroplast translation is essential in this alga. We therefore fused the psbD promoter and 5′UTR to rps12 encoding the ribosomal protein Rps12 and to the rpoA gene encoding the α-subunit of the chloroplast RNA polymerase. The aadA cassette was inserted upstream of the psbD5′-rpoA or psbD5′-rps12 gene for selection of the transformants on spectinomycin-containing plates. Both constructs were separately introduced into the A31 strain through biolistic chloroplast transformation, giving rise to numerous transformants in each case. After four to six recloning steps of the transformants on media with selective pressure (spectinomycin) but lacking vitamins, several of them were found to be homoplasmic (see Supplemental Figure 4 online). The DPF transformants (transcription inhibited) contain the psbD5′-rpoA chimeric gene and the RR transformants (translation inhibited) contain the psbD5′-rps12 chimeric gene. Spot tests revealed that growth of these transformants was indistinguishable from that of Rep112 and A31 in Tris-acetate phosphate (TAP) or high-salt minimal (HSM) medium but was blocked in TAP medium supplemented with vitamins (Figure 3A). Notably, when the RR5 and DPF1 transformants were plated on TAP and HSM plates supplemented with vitamins, individual colonies appeared at a frequency of 10⁻⁴ and 10⁻³, respectively, after several days (Figure 3A) (see Methods for further characterization of these suppressors).

Figure 3. — Growth Patterns upon Repression of Chloroplast Transcription in DPF1 and Translation in RR5.

**(A)** Growth patterns of DPF1, RR5, A31, and Rep112. Cells were spotted on TAP and HSM plates in the absence or presence of vitamins B12 (20 μg/L) and thiamine (20 μM) under an illumination of 60 μmol m⁻² s⁻¹.

**(B)** Growth curves and F_v/F_m measurements in the presence of vitamins. Cell concentration and F_v/F_m of A31, RR5, and DPF1 were measured at different times after addition of vitamins. To maintain cells in exponential growth during the time course, they were diluted to 0.5 × 10⁶ cells/mL when they reached a concentration between 2 and 4 × 10⁶ cells/mL and growth was continued. The experiment was repeated three times with similar results.

**(C)** Light- and dark-grown cell cultures of A31, DPF1, and RR5 in the presence (+) or absence (−) of vitamins.

**(D)** Light microscopy of cells from A31, DPF1, and RR5 in the presence (+) or absence (−) of vitamins. Cells were stained with Trypan blue after 6 d in the light (60 μmol m⁻² s⁻¹) or 12 d in the dark.

Of the transformants obtained, two of each type were used for further studies: DPF1 and DPF9 (containing psbD5′-rpoA) and RR5 and RR20 (containing psbD5′-rps12). The properties of DPF1 versus DPF9 and those of RR5 versus RR20 were indistinguishable, so only the results obtained with DPF1 and RR5 are shown. Growth of these strains was examined in liquid culture during several days (Figure 3B) and at first was very similar to that of A31, but decreased significantly after 25 h of vitamin treatment (Figure 3B). The effect on PSII activity was most pronounced for RR5 in which no PSII activity was detectable after 120 h.

To test whether photooxidative damage has an effect on cell survival upon repression of rps12 and rpoA, growth of DPF1 and RR5 was compared with growth of A31 both under an irradiance of 60 μmol m⁻² s⁻¹ white light and in the dark (Figure 3C). Cell bleaching was more pronounced after 6 d in the light compared with 12 d in the dark, suggesting that photooxidative damage had occurred and accelerated cell death. This was further confirmed by staining the cells with the vital stain Trypan blue (Figure 3D), which showed that many of these cells were dying.

Because the Rps12 protein plays an important role in the decoding center of chloroplast ribosomes, its absence is therefore expected to block chloroplast translation. This was confirmed by the analysis of chloroplast polysomes isolated from RR5 cells. To increase the proportion of non-membrane-bound ribosomes, cells were grown in the dark (Chua et al., 1973) in the absence or in the presence of vitamins for 48 h, at which time small amounts of Rps12 are still detected (Figures 4A and 4B). To stabilize the polysomes, cells were treated with chloramphenicol 10 min prior to cell harvesting to prevent ribosome run-off during polysome extraction. Ribosomes and polysomes from these cells were fractionated by Suc density gradient centrifugation (Figure 4A). The polysome profile of chloroplast mRNA was examined by hybridizing the RNA from each fraction with a psaB probe. Comparison of the RNA distribution between vitamin-treated and untreated cells revealed a significant shift of RNA from the polysome to the monosome region, confirming that translation is decreased under these conditions. A similar shift was also observed for the chloroplast psbD and tufA mRNA, although in the latter case, the shift was less pronounced. By contrast, no change was observed for the cytoplasmic ORM mRNA, confirming that the effect is specific to the chloroplast compartment. As expected, the polysomes could be destabilized by EDTA treatment (Figure 4A).

Upon vitamin addition, a decrease of Rps12 was detected after 24 h and the protein was undetectable after 72 h (Figure 4B). Because the loss of Rps12 leads to the inactivation of chloroplast ribosomes, the levels of other chloroplast-encoded proteins, such as ClpP1, α- and β-subunits of CF1, D2, Cytf, and Rubisco large subunit also declined but with different kinetics due to different half-lives of the proteins (Figure 4C). A similar decline was observed for the nucleus-encoded PsaD and Rieske iron proteins most likely as the result of the decrease of their chloroplast-encoded partner proteins in PSI and in the Cytb₆f complex, respectively. However, such a decrease was also observed for PRK, which functions in the Calvin-Benson cycle. The chloroplast-encoded TufA, RpoA, and RbcL proteins were remarkably stable and started to decrease only after 72 h (Figure 4B). The decrease of the nucleus-encoded chloroplast ribosomal proteins Rps20 was more pronounced than that of Rps21, which can be correlated with the fact that these two proteins are inserted early and late, respectively, during ribosome assembly (Held et al., 1974). The decrease of chloroplast ribosomes did not have any impact on the accumulation of the cytoplasmic Rpl37 protein and 25S rRNA, indicating that the cytosolic ribosome level is not affected by the loss of chloroplast translation (Figures 4B and 5A).

By contrast, the level of Vipp1 (Kroll et al., 2001; Aseeva et al., 2007) and Alb3.2 (Göhre et al., 2006), two proteins that play an important role in thylakoid membrane biogenesis, and the level of the chaperones Hsp70B and Hsp90C and of the cochaperone CDJ1 (DnaJ) increased significantly after 48 h (Figure 4D). The first two proteins are known to interact with each other and Hsp70B is involved in the refolding of stress-denatured proteins, especially under heat shock or high light where it plays a role in protection and photorepair of PSII. In C. reinhardtii, Vipp1 is associated with low density membranes (Zerges and Rochaix, 1998) and thylakoids (Drzymalla et al., 1996), and it is able to form ring-like structures that can assemble into rod-shaped complexes proposed to act as tracks for the transport of lipids or proteins during thylakoid biogenesis (Liu et al., 2005; Schroda and Vallon, 2009). Together with its cochaperone CDJ2, Hsp70B interacts with Vipp1 and may assist in the assembly and disassembly of the Vipp1 ring structures. It is possible that the upregulation of these proteins is required for membrane reorganization and/or maintenance when chloroplast translation is compromised.

The level of rps12 transcript in RR5 decreased gradually over a period of 72 h of vitamin treatment (Figure 5A). It is noteworthy that rps12 precursor transcripts detected in A31 were completely absent in the RR5 strain. This is due to the fact that for replacing the authentic rps12 gene with the psbD5′-rps12 construct, the aadA cassette used for selecting the transformants was inserted in opposite orientation between rps12 and the upstream psaJ gene (Liu et al., 1989). It is thus likely that in this strain the maturation of other precursor transcripts of rps12 containing upstream sequences is altered. Moreover, after 96 h, the level of rps12 RNA, but not of the protein, increased and a new larger transcript was detectable (Figures 4B and 5A), indicating changes in RNA processing and stability.

In RR5, the level of psbD mRNA remained stable as expected because its accumulation is no longer dependent on Nac2 (Figure 5A). By contrast, the levels of chloroplast rRNAs decreased slightly during the 120-h vitamin treatment. Strikingly, the amount of tufA and clpP1 RNAs increased substantially after 48 h (Figure 5A). In the case of ClpP1, an inverse correlation is apparent between the decrease of Clp1 protein and the increase of its mRNA (Figures 4B and 5A), suggesting that ClpP1 may be involved in this negative feedback loop. This hypothesis predicts that a specific decrease in ClpP1 translation would lead to an increase of its mRNA. This was indeed found to occur in the clpP-AUU mutant in which Clp1 accumulation is reduced to 25 to 40% of wild-type levels as a result of a change of its initiation codon (Figure 5B) (Majeran et al., 2000). No increase of tufA mRNA was detected in the clpP-AUU strain, indicating that the tufA mRNA level is not controlled by ClpP1. Finally, we note that while the mRNA level of the cytosolic Rpl37 protein and of cytosolic 25S rRNA remained constant in RR5 (Figure 5A), the levels of the cytosolic mRNAs of ubiquitin and Vipp1 increased after 72 h of vitamin treatment.

To obtain a more global view on the chloroplast transcriptome, we performed quantitative hybridizations with an array containing 68 chloroplast and 21 nuclear gene probes using Nanostring technology (see Supplemental Data Set 1 online) (Geiss et al., 2008). This method allows for a direct and very sensitive quantification of RNA abundance because the number of RNA molecules is estimated directly without the need of amplifying the transcripts through RT-PCR. It is thus very well suited for analysis of intronless transcripts for which removal of rRNAs and genomic DNA can be difficult. These probes were hybridized with RNA isolated from RR5 cells at 0, 12, 48, and 146 h after vitamin treatment grown in dim light. The results confirmed the upregulation of tufA, clpP1, and rps12 after 146 h of vitamin treatment and revealed that the expression of many chloroplast genes follows the same pattern (Figures 5 and 6; see Supplemental Data Set 1 online). They include the genes of the subunits of the plastid RNA polymerase, most of the ribosomal protein genes, introns, large open reading frames of unknown function, ycf3 and ycf4 encoding assembly factors for PSI, genes involved in light-independent chlorophyll synthesis, and, quite surprisingly, the short open reading frame encoded by the Wendy transposon-like element (Fan et al., 1995). Also included are a few subunits of ATP synthase, but expression of the large majority of the genes of subunits of the photosynthetic complexes is not significantly altered.

Figure 6. — Hierarchical Cluster Analysis of the Relative RNA Levels of RR5 and DPF1 upon Arrest of Chloroplast Translation and Transcription.

RNA was isolated from A31, RR5, and DPF1 at 0, 12, 48, and 146 h after addition of vitamins. RNA levels were quantified with a NanoString nCounter. The genes were first grouped by class and then within each class by Euclidian distance using the statistical function in Matlab 2012b (MathWorks). Genes are listed on the right with a different color for each class. The scale is in log₂ relative to A31 at each time point.

The analysis of the DPF1 strain revealed that the amount of rpoA RNA decreased gradually 48 h after addition of vitamins (Figure 5C). Surprisingly, after 72 h, the level of rpoA RNA increased, and after 144 h, it was significantly higher than in cells grown without vitamins or in the control strain A31. The tufA and rpoB RNAs markedly increased during the first 12 to 36 h and then declined followed by a smaller increase of tufA RNA after 96 h (Figure 5C). However, the rpoB RNA accumulated at a higher level than in cells grown without vitamins. For other transcripts, including atpB, rps12, and 16S RNA, the RNA level decreased gradually with time as expected (Figures 5C and 5D; see Supplemental Figure 5A online). By contrast, cytosolic 25S RNA and the mRNA of Rpl37 remained stable. The levels of the Vipp1 and ubiquitin mRNA significantly increased after 72 h (Figure 5D) as in the case of Rps12 depletion (Figure 5A).

These results were extended with Nanostring hybridizations using the same array of gene probes as for RR5 and RNA isolated at different times of vitamin treatment for DPF1 cells. As observed for tufA and rpoB, several genes were already upregulated after 12 h of vitamin treatment, including those of the chloroplast RNA polymerase, of several ribosomal proteins, of the enzymes involved in light-independent chlorophyll synthesis, and of the PSI assembly factors Ycf3 and Ycf4. Not surprisingly, this set of genes overlaps to a large extent with those upregulated when chloroplast translation was repressed. However, there were also marked differences in the transcriptional response between RR5 and DPF1. While some ribosomal protein genes (e.g., rpl14, rpl16, and rps18) had similar patterns in these two strains, others (e.g., rpl23, rpl36, rps4, rps7, and rps14) displayed opposite responses to vitamin treatment (Figure 6; see Supplemental Figure 6 online). Moreover, most genes coding for components of the photosynthetic complexes were downregulated upon addition of vitamins in DPF1, but they were upregulated or their transcript levels did not change significantly in RR5 (Figure 6; see Supplemental Figure 6 and Supplemental Data Set 1 online). These differences between RR5 and DPF1 may be attributed in part to the long half-lives of some chloroplast mRNAs for which any effect on translation is delayed in DPF1. As control, the nanostring hybridizations were also performed with the A31 strain (see Supplemental Figure 6 online). After 12 h of vitamin treatment, several of the same genes that were induced in RR5 and DPF1 were also upregulated, indicating that vitamin treatment could be partly responsible for this response. However, the levels of these RNAs returned to the initial level prior to vitamin treatment after prolonged vitamin treatment. In a few cases, we also noticed differences in gene expression between A31, RR5, and DPF1 prior to the addition of vitamins. The causes for these differences are not clear.

To directly assess transcriptional activity, run-on experiments were performed with DPF1 grown for 48 h in the absence and presence of vitamins (Figure 5E). Cells were permeabilized by a freeze-thaw cycle and labeled with ³²P-UTP for 15 min (Gagné and Guertin, 1992). RNA was extracted and hybridized to a filter containing several chloroplast gene probes. These experiments clearly indicated a significant decrease of transcriptional activity in the presence of vitamins for rpoA, rps12, tufA, 23S rRNA, psbD, and atpB. No signal could be detected from the nuclear gene Rpl37 because under those conditions chloroplast transcripts are preferentially labeled (Guertin and Bellemare, 1979). A similar run-on experiment was performed with DPF1 grown for 144 h corresponding to a time point when the levels of rpoA and tufA RNA are increased. No corresponding increase of transcriptional activity was detected, indicating that the increased RNA level is due to increased stability. As control, we tested that there was no difference in transcriptional activity of the nuclear rRNA genes at the two time points (see Supplemental Figure 5B online).

The RpoA protein was tagged with a FLAG epitope, and its level was determined by immunoblotting with a FLAG antiserum. The decline of this protein followed closely that of its mRNA (Figures 5C and 7). A similar decline was observed using an RpoA antiserum (Figure 7). It is surprising that the decrease of both RpoA and TufA was faster upon inhibition of chloroplast transcription than of translation, whereas for other chloroplast-encoded proteins, such as Rps12 and D2, the decrease was slower as expected because of the time required for first reducing the level of RpoA (Figures 4 and 6). As in the case of rps12 mRNA depletion, the level of the cytosolic Rpl37 protein was unaffected, indicating that the accumulation of cytosolic ribosomes is insensitive to a block in chloroplast transcription and translation.

Figure 7. — Immunoblot Analysis of RpoA and Other Chloroplast and Cytoplasmic Proteins in DPF1.

Proteins were examined at different times after addition of the vitamins using antibodies against FLAG and the indicated proteins.

In land plants, inhibition of chloroplast protein synthesis leads to decreased accumulation of transcripts of nuclear genes encoding proteins involved in photosynthesis (Sullivan and Gray, 1999). To test whether a similar decline occurs in C. reinhardtii upon repression of chloroplast transcription and translation, the mRNA levels of several photosynthetic proteins were determined by quantitative RT-PCR (qRT-PCR) (Figure 8). A large decrease was observed for Lhca1, Lhcbm1, and PsaD mRNAs in the light in DPF1 and RR5 compared with the control A31 in the presence of vitamins. The decrease was larger for RR5 than for DPF1 in the light but the opposite occurred in the dark for Lhca1 and Lhcbm1. By contrast, Lhcbm9, which is known to be induced under stress conditions, was increased in the light and in the dark in both RR5 and DPF1 (Nguyen et al., 2008).

Figure 8. — Expression of Nuclear LHC Genes in A31, DPF1, and RR5.

The levels of the indicated mRNAs were determined by qRT-PCR from cells grown for 6 d in the light or 12 d in the dark in the presence of vitamins.

DISCUSSION

Conditional Vitamin-Mediated Repression of Chloroplast Gene Expression

Chloroplast genomes contain a set of genes that appear to have essential functions based on the fact that attempts to disrupt these genes through chloroplast transformation have failed, resulting in a heteroplasmic state with both wild-type and mutant gene copies. In this study, we established a method that allows for the conditional repression of any of these genes in the unicellular alga C. reinhardtii. Although long-term inactivation of these genes leads to cell death, it is possible to study the primary events that occur before the cells die and thereby to obtain important clues not only about the role of these genes by observing changes in transcript and protein levels but also on how plastid gene expression is integrated within cellular metabolism and signaling.

We took advantage of a repressible vitamin-dependent gene expression system that uses the TPP-responsive riboswitch located in the 5′UTR of the Thi4 transcript. This element can be repressed by inclusion of thiamine in the growth medium (Croft et al., 2005, 2007). Fusion of the Thi4 5′UTR to the Nac2 coding sequence resulted in expression of Nac2 in the absence of thiamine but not in its presence. In turn, Nac2 expression stabilizes the chloroplast psbD mRNA and allows for D2 expression and PSII activity. Because the psbD 5′UTR contains the major target site for Nac2, the system can be used for any chloroplast gene by fusing the psbD 5′UTR to its coding sequence. Although the design of this system is similar to the one we established previously with the copper-repressible Cyc6 promoter (Surzycki et al., 2007), it is easier to use, more versatile, and makes it possible to study the role of essential plastid genes.

Loss of Chloroplast Translation Leads to an Arrest in Cell Growth

Several cases have been described for land plants in which plastid translation is abolished. The albostrians mutant of barley has a variegated phenotype with green and white leaf sectors (Hess et al., 1994b). The latter lack plastid ribosomes and are deficient in plastid translation although transcription still occurs. In maize, the iojap mutation results in plastids lacking ribosomes and albino stripes in the leaves (Walbot and Coe, 1979). In this case, plastid growth and division are maintained in the absence of plastid protein synthesis. Moreover, many mutations that disrupt the splicing of chloroplast tRNAs or ribosomal protein mRNAs in maize lead to a complete loss of plastid ribosomes without affecting embryo development (Bryant et al., 2011; Lloyd and Meinke, 2012). Many additional albino mutants of maize and rice (Oryza sativa) lack plastid ribosomes, although the underlying mechanisms are not yet clear. A loss of plastid ribosomes has also been observed in heat-bleached rye (Secale cereale) and barley (Falk et al., 1993). In contrast with the grasses, embryo development is compromised in Arabidopsis mutants deficient in plastid translation. This difference between monocotyledonous and dicotyledonous plants appears to be due to the chloroplast accD locus involved in fatty acid biosynthesis, which is essential in Arabidopsis but not in maize, and rapeseed where nuclear genes compensate for its absence (Bryant et al., 2011).

Extensive genetic studies of C. reinhardtii did not reveal mutants completely deficient in chloroplast protein synthesis, although mutants affected in plastid ribosome assembly and impaired in chloroplast translation have been identified and characterized (Boynton et al., 1972; Harris et al., 1974). Analysis of some of these mutants showed that under conditions of reduced chloroplast protein synthesis, ribosomal proteins are preferentially translated in comparison to proteins involved in photosynthesis (Liu et al., 1989). As one-third of chloroplast ribosomal proteins are synthesized on chloroplast ribosomes, this response may slow the permanent loss of chloroplast ribosomes. Most likely, chloroplast translation is essential in this alga because its plastid genome contains several genes besides those of the components of the genetic system that are essential for cell growth. Examples include ClpP (Huang et al., 1994) and ORF1995 (Boudreau et al., 1997). It is also possible that chloroplast translation is intimately coupled to the cell cycle in C. reinhardtii and that arrest of chloroplast translation perturbs this cycle (Blamire et al., 1974).

To gain further insights into the role of plastid translation in C. reinhardtii and to avoid the use of bacterial translation inhibitors that have side effects on mitochondrial gene expression, we wanted to study the effect of the progressive and specific inactivation of chloroplast ribosomes via vitamin-induced repression of the rps12 gene through our riboswitch-Nac2 system. The Rps12 protein plays a key role in the decoding center of the ribosome. From earlier studies on in vitro ribosome assembly of Escherichia coli, it is known that Rps12 is assembled late after binding of the ribosomal proteins Rps4, Rps8, and Rps17 to 16S rRNA (Held et al., 1974). Thus, it is possible that ribosomal subparticles lacking Rps12 are still assembled after 48 h. However, the loss of Rps12 leads to a general block of chloroplast translation and is thereby expected to prevent the synthesis of 16 additional ribosomal proteins that are encoded by the chloroplast genome (Maul et al., 2002). The absence of Rps12 also leads to the gradual decrease of other chloroplast-encoded proteins with a time course that depends on the half-life of the proteins. It is also possible that competition for binding of mRNAs to ribosomes plays a role when ribosomes become limiting. Based on the immunoblot analysis of RR5 grown in the presence of the vitamins, the half-life of chloroplast ribosomes is at least several days (Figure 4B).

This repressible system should allow the testing of whether specific ribosomal proteins are essential for cell growth and the examination of their role in ribosome assembly and translational activity in vivo. In this respect, reverse genetic analysis in tobacco revealed that inactivation of the chloroplast ribosomal protein genes rps15 and rpl36 gave rise to homoplasmic transplastomic lines, indicating that these proteins are nonessential (Fleischmann et al., 2011). However, photosynthetic activity and growth were strongly impaired in the absence of Rpl36 and affected to a lesser extent in the absence of Rps15, indicating that translational activity is decreased in the absence of either of these two proteins.

Repression of Chloroplast Transcription and Translation Reveals Negative Regulatory Feedback Loops

Loss of Rps12 and RpoA has some unexpected effects on many chloroplast transcripts. Whereas mRNAs of the subunits of the photosynthetic complexes decreased or increased slightly after prolonged vitamin treatment of RR5, the RNA levels of a large set of chloroplast genes increased strongly (more than fivefold). It includes the mRNAs of subunits of the plastid RNA polymerase, most ribosomal proteins, some tRNAs, and TufA. Moreover, this group also comprises ClpP1, the PSI assembly factors Ycf3 and Ycf4, and the subunits of the enzyme involved in light-independent chlorophyll synthesis (chlB, chlL, and chlN) and heme ligation (ccsA), plastid proteins of unknown function (ORF59, ORF1995, and ORF2971), and transcripts from introns and the Wendy element (Figure 6; see Supplemental Figure 6 online). The inverse correlation between the progressive decrease of ClpP1 protein (Figure 4C) and the concomitant increase of its mRNA (Figure 5A) suggests a negative feedback mechanism exerted through ClpP1 itself. This is further confirmed by the increase of clpP1 mRNA in a mutant in which translation of ClpP1 is specifically attenuated (Figure 5B). In this context, it is interesting to note that the chloroplast RNA helicase RH3 is strongly upregulated in Arabidopsis mutants lacking Clp complex subunits (Asakura et al., 2012), pointing to a possible link between the Clp complex and chloroplast RNA metabolism. Interestingly, although the ClpP1 protein is also downregulated upon conditional inactivation of rpoA, we do not observe overaccumulation of its transcript in this condition (Figure 5C). It is possible that the ClpP1 protein has a longer half-life than its own mRNA; thus, in absence of active transcription, the presumed inhibitory effect of this protein on its own transcript further contributes to reduce the steady state level of the ClpP1 mRNA.

In the case of tufA, a significant increase of its RNA is already detectable after 12 h of vitamin treatment in RR5 when the decrease of Rps12 protein is only modest, which is followed by a massive increase of tufA RNA upon prolonged vitamin treatment (Figure 5A). Similarly, a marked increase of tufA RNA is already detectable after 12 h when transcription is repressed, although at this time point RpoA is only diminished 50% and Rps12 is barely affected (Figure 5C). This suggests that TufA may act as a sensor for both transcription and translation elongation and is in agreement with earlier findings that tufA RNA levels increase upon treatment of C. reinhardtii cells with rifampicin or chloramphenicol (Zicker et al., 2007). However, the chloroplast RNAs, which are strongly upregulated after 96 h, are apparently not translated as Rps12 could not be detected and ClpP1 level was strongly reduced at these time points (Figures 4B and 4C).

As expected, the transcripts of many genes involved in photosynthesis are downregulated in the DPF1 strain after prolonged vitamin treatment (Figure 5B; see Supplemental Figures 5 and 6 online). However, as in the case of RR5, the mRNAs of the plastid RNA polymerase and of the PSI assembly factors Ycf3 and Ycf4 are upregulated (Figure 5C; see Supplemental Figure 6 online). It should be noticed that the latter two mRNAs are cotranscribed with the ribosomal protein gene rps9, which is upregulated upon repression of rps12 and rpoA. Therefore, it is possible that the increased RNA levels of ycf3 and ycf4 are a simple consequence of their genomic location. Although the underlying molecular mechanisms are still unknown, this suggests the existence of negative feedback loops for many chloroplast transcripts that might compensate for a decrease in their transcription and or translation with an increase of their stability or a change in RNA processing (Figure 9). Alternatively a ribosome-associated nucleolytic activity could degrade chloroplast mRNAs once they are translated but no longer when translation or transcription is compromised. Increased mRNA levels have also been observed in several cases in mutants of C. reinhardtii deficient in translation of psbA (Kuchka et al., 1988), atpA (Drapier et al., 1992), and psaB mRNA (Xu et al., 1993). Similar feedback loops have also been detected in bacteria for ribosomal operons, including tufA (Young and Furano, 1981), suggesting that these regulatory mechanisms have been conserved during the evolution of chloroplasts. Finally, we note that loss of chloroplast transcription or translation does not affect the level of cytosolic 80S ribosomes as shown by the unchanged levels of the cytosolic ribosomal protein Rpl37 and 25 S rRNA (Figure 5).

Figure 9. — Negative Regulatory Feedback Loops in the Chloroplast Gene Circuitry.

Left, chloroplast compartment; right, nucleo-cytosol. A selected set of chloroplast mRNAs examined in this work is shown. They comprise mRNAs of ClpP1, chloroplast ribosomal proteins (r-prot), subunits of chloroplast RNA polymerase (rpo), elongation factor TufA (tufA), and subunits of the light-independent protochlorophyllide reductase (chl). Nucleus-encoded mRNA genes coding for chloroplast and cytoplasmic proteins are shown in red and blue, respectively, on the right. Negative regulatory feedback loops are revealed through repression of transcription (green lines) or translation (red lines). Factors involved are still unknown (X, Y, and Z) except for the ClpP1 protein, which represses accumulation of its own mRNA directly or indirectly (green circular line). The feedback loops act mostly at the level of RNA accumulation in contrast with CES (for Control of Epistasy of Synthesis), an assembly-dependent feedback process in which unassembled CES subunits inhibit directly or indirectly their own translation (green circular lines) (reviewed in Choquet and Wollman, 2002).

Loss of Chloroplast Gene Transcription and Translation Affects Nuclear Gene Expression

In land plants, inhibition of chloroplast protein synthesis triggers retrograde signaling from the chloroplast to the nucleus and leads to the decrease of mRNA levels of several nucleus-encoded chloroplast proteins (Nott et al., 2006). Several mutants, such as the constitutively photomorphogenetic cop1-4 from Arabidopsis and lip1 from pea (Pisum sativum), as well as the Arabidopsis gun1 mutant are deficient in this plastid gene expression-dependent pathway (Susek et al., 1993; Sullivan and Gray, 1999; Gray et al., 2003).

Here, we have shown that repression of rpoA and rps12 affects not only chloroplast transcripts but also the level of several mRNAs from nuclear-encoded proteins related to photosynthesis, suggesting that a similar retrograde signaling pathway is operating in C. reinhardtii as in land plants (Figure 8). We identified a restricted set of nucleus-encoded chloroplast proteins, including the LHCII proteins, which are downregulated (Figure 8), and Vipp1, Alb3.2, and several Hsp proteins, which are upregulated (Figures 6 and 7; see Supplemental Figure 6 online). Vipp1 is involved in some unknown way in thylakoid membrane biogenesis and probably also in chloroplast lipid trafficking (Kroll et al., 2001). It is possible that under conditions of limited plastid protein synthesis, Vipp1 is implicated in remodeling of the thylakoid membrane, a process that may be part of a stress response that also involves the chloroplast chaperones Hsp70B and Hsp90C and the cochaperone Cdj1 (Figure 4D). Moreover, the levels of mRNAs of some cytosolic proteins are also affected, in particular ubiquitin mRNA (Figures 5D). The upregulation of ubiquitin and chaperone proteins raises the possibility that under conditions in which chloroplast protein synthesis or any chloroplast downstream process is compromised, cytoplasmic protein degradation is enhanced. Whether this response participates in a salvage pathway in which proteins targeted to the plastid are specifically degraded or extracted from the organelle for degradation remains to be explored. It is interesting to note that a mitochondrial-associated degradation pathway has recently been reported in which proteins from the inner mitochondrial membrane are extracted and degraded by the cytoplasmic proteasome through the action of Cdc48 (Heo et al., 2010), the mRNA of which is upregulated upon repression of rps12 and rpoA in C. reinhardtii (Figure 6; see Supplemental Figure 6 online). Moreover, it is possible that impairment of plastid protein homeostasis upon abrogation of chloroplast translation might activate specific plastid-to-nucleus signals, resulting in a preferential upregulation of chaperone proteins targeted to this compartment similar to the mitochondrial unfolding response (Zhao et al., 2002).

The Chloroplast of C. reinhardtii Contains a Single E. coli–Like RNA Polymerase

In land plants, it is well established that chloroplasts contain at least two distinct RNA polymerases. The core of the E. coli–like plastid-encoded RNA polymerase (PEP), which initiates transcription from sequences resembling E. coli σ⁷⁰-type promoters, consists of two α-subunits, and the β-, β’-, and β’’-subunits encoded by the plastid genes rpoA, rpoB, rpoC1, and rpoC2, respectively (Igloi and Kössel, 1992). In C. reinhardtii, the rpoB and rpoC1 genes are further split in two parts (Fong and Surzycki, 1992; Maul et al., 2002). The α-subunit plays a role in the assembly of the polymerase complex. Although the sequence of this subunit is clearly similar to that of the bacterial alpha subunit, it is considerably larger in size because of an insertion that is unique to the C. reinhardtii protein (Klein, 2009). In addition, there is a nucleus-encoded plastid RNA polymerase (NEP) that transcribes a subset of chloroplast genes and recognizes promoters of a different type (Hajdukiewicz et al., 1997). However, the latter appears to exist only in flowering plants but not in lower plants and algae (Klein, 2009). Deletion of the rpoB gene was obtained in a homoplasmic form in tobacco and led to the absence of transcription from σ ⁷⁰-type promoters and to the loss of expression of the plastid genes involved in photosynthesis (Allison et al., 1996). However, transcription of a subset of plastid genes was maintained by NEP in the absence of PEP, and it was sufficient for plastid maintenance and plant development. It is interesting to note that in the parasitic plant Epiphagus virginiana, which lacks the photosynthetic apparatus, the rpo RNA polymerase genes have been lost during evolution (Wolfe et al., 1992).

In C. reinhardtii, current evidence suggests that NEP is missing and that chloroplast transcription is mainly performed by PEP (Smith and Purton, 2002). It was therefore important to reinvestigate this problem by avoiding chloroplast gene disruption through repressible chloroplast gene expression. Using the vitamin-repressible riboswitch-Nac2 system, we have shown that the loss of rpoA leads to the loss of cell growth on TAP medium, indicating that there is no additional major transcription system in the chloroplast of C. reinhardtii and that the PEP enzyme is most likely involved in the transcription of the entire chloroplast genome of this alga. There is thus a striking difference between unicellular algae and land plants in this respect. It is probably linked to plastid development in plants, which involves multiple steps from the proplastid with its rudimentary internal membrane system to the fully differentiated mature chloroplast. By contrast, in C. reinhardtii, there is no comparable developmental pathway for plastids as algal cell division is tightly coupled with the division of their differentiated chloroplast, at least in vegetative cells. Although it is likely that plastid differentiation must occur during the meiotic cycle of C. reinhardtii, it is not clear to what extent it resembles the proplastid to plastid differentiation in land plants. Mutants of C. reinhardtii exist that are unable to synthesize chlorophyll in the dark and therefore do not accumulate chlorophyll-protein complexes in their thylakoid membranes, but their plastids correspond to etioplasts rather than to proplastids (Ohad et al., 1967).

METHODS

Strains, Growth Conditions, and Media

The Chlamydomonas reinhardtii wild type, nac2-26, and the other strains generated were maintained on TAP or minimal (HSM) medium plates supplemented with 1.5% Bacto-agar (Gorman and Levine, 1966; Harris, 1989) at 25°C under constant light (60 to 40 μmol m⁻² s⁻¹)/dim light (10 μmol m⁻² s⁻¹) or in the dark depending on their phototropic growth abilities.

Medium with vitamins was prepared in the following way: Stock solutions 1000× of Thiamine-HCl (Sigma-Aldrich) and vitamin B12 (Sigma-Aldrich) were prepared in sterile MilliQ water and stored at 4°C in the dark. Upon sterilization, the medium was cooled and 1/1000 volume of each vitamin stock solution was added. Medium was stored at room temperature until ready to use.

At the beginning of each experiment, cells were preinoculated from fresh plates into liquid TAP media (Harris, 1989), unless indicated otherwise, and allowed to grow under continuous light at 25°C on a rotary shaker at 150 rpm to a density of 2 to 4 × 10⁶ cells/mL. Subsequently, they were diluted to a density of 0.5 to 10⁶ cells/mL and allowed to grow to the desired cell concentration.

In case of repression/derepression experiments, cells were allowed to grow to a density of 0.5 to 10⁷ cells/mL in medium with/without vitamins and were subsequently diluted (10 to 20 times) to a density of 0.5 to 10⁶ cells/mL in medium with/without vitamins. Although not absolutely necessary, in case of derepression, cells were pelleted by centrifugation at 2500g for 5 min at room temperature and washed two to three times in medium without vitamins prior to inoculation. Unless indicated otherwise, vitamins were supplied at the following concentration: 20 μM thiamine-HCl and 20 μg/L B12.

For growth tests, 7 μL of 1 × 10⁴, 5 × 10⁴, 1 × 10⁵, 5 × 10⁴, and 1 × 10⁶ cells were spotted on solid TAP or HSM medium with or without vitamins and placed in continuous light (60 μmol m⁻² s⁻¹) unless indicated otherwise.

Characterization of Suppressor Strains

When the RR5 and DPF1 strains were plated on TAP and HSM plates supplemented with vitamins, individual colonies appeared at a frequency of 10⁻⁴ and 10⁻³, respectively, after several days (Figure 3). Analysis of these suppressor strains revealed that the amount of Rps12 and RpoA was restored to the same or to an even higher level than in the cells grown without vitamins (see Supplemental Figure 7A online). The observation that ThiC was still repressed in these suppressors indicates that their appearance cannot be explained by a deficiency in thiamine delivery or in the synthesis of TPP (see Supplemental Figure 7B online). To test whether the absence of vitamin repression in these strains was due to a constitutive expression of Nac2 or to a change in the psbD 5′UTR sequence that would make the stabilization of psbD mRNA independent of Nac2, the level of Nac2 was determined by immunoblotting and the psbD 5′UTR of the suppressors was sequenced. Nac2 was found to be expressed constitutively in the presence of vitamins at even higher levels than in RR5 and DPF1 grown without vitamins (see Supplemental Figure 7B online). qRT-PCR revealed that while the transcript levels of Thi4 and ThiC were decreased in the presence of vitamins, the amount of Nac2 RNA was the same in the absence and presence of vitamins (see Supplemental Figure 7C online). Moreover, no change in the psbD 5′UTR sequence could be detected, raising the possibility that the riboswitch regulating Nac2 was altered. Indeed among five suppressors examined, two had changes in the riboswitch sequence, one with a single base deletion in the P4 helix and the other with the insertion of a retrotransposon in helix 5 (see Supplemental Figure 7D online). The other suppressors might have an epigenetic change or might be affected in an unknown factor interacting with the riboswitch. In contrast with RR5 and DPF1, no vitamin-resistant suppressors of Rep112 were found.

Construction of the Strains Rep112, A31, RR5, and DPF1

Plasmids used for generating the strains A31, Rep112, RR5, and DPF1 are described in Supplemental Methods 1 online. These strains were generated through nuclear and chloroplast transformation as described (Shimogawara et al., 1998; Surzycki et al., 2007).

Nuclear and Chloroplast Transformation

Nuclear transformation of the nac2-26 strain with pRAM23.1 was performed by the electroporation transformation method as described (Shimogawara et al., 1998). Briefly, nac2-26 cells were grown in TAP medium/dim light, harvested in mid-log phase (2 to 4 × 10⁶ cells/mL), and treated with gamete autolysin for 1 h, then resuspended in TAP medium plus 40 mM Suc. For each electroporation, 10⁸ treated cells were incubated with 0.5 to 2.5 μg of SapI-linearized pRAM23.1 and then transformed by electroporation in a 0.2-cm electroporation cuvette (Bio-Rad) using the Bio-Rad Gene Pulser II set to 0.75 kV, 25 μF, and no resistance. The resulting transformants were recovered in 1 mL fresh TAP plus 40 mM Suc plus 0.4% polyethylene glycol 8000 plus 20% starch medium for 10 min, followed by plating on HSM medium at 25°C in constant light (40 μmol m⁻² s⁻¹). The resultant photoautotrophic transformants were tested to have a wild-type fluorescence transient and were then screened for the inability to grow in HSM medium supplemented with vitamins (10 μM thiamine-HCl/10 μg B12/L). Their vitamin-repressible PSII activity was further analyzed by fluorescence measurements (F_v/F_m). On average, only 10% of the recovered transformants showed vitamin-repressible photoautotrophic growth.

Chloroplast biolistic transformation of Rep112 and A31 strain was performed with a helium-driven particle gun adapted from the design of Finer et al. (1992) according to the protocol developed by Boynton and Gillham (1993). For transformation of Rep112, 3 × 10⁷ cells grown in TAP medium were plated on solid agar TAP supplemented with 100 μg/mL spectinomycin (Sigma-Aldrich S9007) and bombarded with 550-nm-diameter gold particles (Seashell Technology S550d) coated with 300 ng of pRAM56.2 (rcy_aadA_psaA5′UTRpsbD) plasmid DNA. After 1 week of growth at 25°C in constant light (60 μmol m⁻² s⁻¹), single colonies were picked, replated four times on TAP supplemented with 100 μg/μL spectinomycin, and screened for photoautotrophic growth on HSM medium + 20 μM thiamine-HCl + 20 μg B12/L. The homoplasmic state of the selected transformants was verified by PCR analysis of chloroplast genomic DNA and through fluorescence measurements (F_v/F_m).

In the case of the A31 strain, a similar protocol was used for chloroplast biolistic transformation with pRAM61.1 (aadA_psbD 5′UTR_rps12) and pRAM68.13 (aadA_psbD 5′UTR_rpoAFLAG) plasmid DNA except that the transformants were replated under a more stringent selective pressure using TAP medium supplemented with 750 μg/mL spectinomycin. Single colonies were tested for growth on TAP + 10 μM thiamine-HCl + 20 μg B12/L.

Fluorescence Measurements

Maximum quantum efficiency of PSII (F_v/F_m) was measured with a plant efficiency analyzer (Handy PEA; Hansatech Instruments). The parameters were set according to the instructions of the manufacturer. Prior to each measurement, cells were adapted for ∼1 to 5 min in the dark.

Protein Extraction and Immunoblot Analysis

Total protein extraction and immunoblot analysis was performed as described with minor modifications (Rochaix et al., 1988). Total protein extracts were obtained by collecting the desired amount of cells (typically from 0.1 to 3 × 10⁸ cells). Pellets were frozen in liquid nitrogen and stored at −70°C until use. Cells were lysed by resuspending the pellet in the desired volume (typically 200 μL) of a 2× solution of Sigma-Aldrich protease inhibitor cocktail and mixing it with an equal volume of cell lysis buffer (100 mM Tris-HCl, pH 6.8, 4% SDS, and 20 mM EDTA). Cell were then incubated at 37°C for 30 min and unbroken membranes and other debris were removed by centrifugation at 16,000g for 30 min at 4°C in a tabletop centrifuge. Supernatants were used as total protein extracts. To determine protein concentration, 5 μL of protein extract were assayed by colorimetric measurements with bicinchonic acid (Sigma-Aldrich). During time-course experiments and for preparation of samples for immunoblot analysis with the Nac2 and ThiC antibody, RNA and proteins were extracted from the same pellet using the RNeasy Plant Mini Kit according to manufacturer’s instructions (Qiagen). For immunoblot analysis of samples, total protein extracts were separated by SDS-PAGE (acrylamide 4K [40%] 29:1; AppliChem) (Laemmli, 1970; Sambrook et al., 1989) and transferred to Protran 0.45-μm nitrocellulose membrane (Schleicher and Schuell) with a wet transfer cell. Membranes were blocked in Tris-buffered saline solution containing 5% nonfat dry milk and 0.1% Tween 20 (TBS-T). For primary antibody conjugation, dilutions of the antibody in TBS-T containing 5% nonfat milk were prepared as follows: D2, D1, PsaA, PsaD, CytF (gift of Y. Choquet), Rps12, RpoA, TufA, PRK, Nac2, Rpl37, Hsp70, Vipp1, DnaJ, Alb3.2, ClpP1 (gift of Olivier Vallon), CF1 antibodies (1:10,000 dilution), FLAG (Cell Signaling) and ThiC (gift of T. Fiztpatrick) antibodies (1:1000), Rps20 (gift of N. Gillham), rps21 (gift of N. Gillham), Rieske protein (gift of C. de Vitry), and Hsp90 (gift of M. Schroda) antibodies (1:3000), Rubisco-Holo, and Rubisco-ssu antibodies (1:50,000). Incubation was performed for 2 to 4 h at room temperature or overnight at 4°C. Subsequently, the membrane was washed three times for 10 to 15 min in TBS-T containing 1 or 5% nonfat milk. For secondary antibody conjugation, the membrane was incubated for 1 to 2 h at room temperature using anti-rabbit IgG (H+L), horseradish peroxidase conjugate (Promega) in TBS-T containing 1 or 5% nonfat milk at a final antibody dilution of 1:10,000. Anti-mouse IgG (H+L) horseradish peroxidase conjugate (Promega) were used in case of immunoblot with FLAG antibody. Anti-rabbit IgG alkaline-phosphatase conjugate (Sigma-Aldrich) was used in case of immunoblot with ThiC, Alb3.2, Nac2, and DnaK antibodies. In the case of horseradish peroxidase conjugate, the signal was visualized by enhanced chemiluminescence ECL (Durrant, 1990). In the case of alkaline-phosphatase conjugate, the signal was visualized by colorimetric detection with 5-bromo-4-chloro-3-indolyl-phosphate/nitro blue tetrazolium color development substrate (Promega S3771) following the manufacturer’s instructions.

Production of Polyclonal Antiserum of Rps12, Rpl37, RpoA, and TufA

For production of recombinant Rps12, Rpl37, RpoA, and TufA proteins, plasmids pRAM62.2, pRAM70.2, pRAM84.8, and pRAM86, respectively, were introduced into Escherichia coli BL21 (Novagen, EMD Biosciences). Expression of each fusion protein was induced by adding 1 mM isopropyl-β-d-thiogalactopyranoside (Invitrogen Life Technologies). Purification of each His-tagged protein was performed under denaturing conditions using a nickel-charged resin (nickel-nitrilotriacetic acid agarose; Qiagen) following the manufacturer’s instructions. The eluted proteins were concentrated using ultra-4 centrifugal filter units with ultracel-3 membrane (Amicon UFC800324) and subjected to further purification by SDS-PAGE (16 × 18 cm, 3-mm spacers, and 10 or 15% acrylamide). The gel was stained with 20% methanol and 0.25% Coomassie Brilliant Blue and destained with 10% methanol without acetic acid. The band of each recombinant protein was cut in little gel pieces and transferred in a small bag of dialysis tubing (SpectrumLabs Spectra/Por 3) in protein elution buffer (14g/L Gly, 3 g/L Tris-HCl, and 0.005% SDS). Electroelution of the proteins was performed in a horizontal electrophoresis apparatus with protein elution buffer at 4°C for ∼2 to 3 h at 200 V. The eluted proteins were reconcentrated using ultra-4 centrifugal filter units, and the final concentration was assayed using the Bradford method (Bio-Rad protein assay) following manufacturer’s instructions. Proteins were injected into rabbits five times at 3-week intervals.

RNA Extraction, RNA Gel Blot Analysis, and DNA Preparation

Isolation of total RNA from C. reinhardtii strains was achieved using the RNeasy plant mini kit according to the manufacturer’s instructions (Qiagen). In case of RNA samples taken during time-course experiments, 0.1 to 10⁸ cells were sedimented by centrifugation at 3000g for 5 min at 4°C, snap frozen in liquid nitrogen, and stored at −70°C until use.

RNA gel blot analysis was performed according to standard procedures. In brief, depending on the abundance of the transcript of interest (see Supplemental Tables 2 and 3 online), 3 to 15 μg of total RNA was electrophoresed in a 1.2 to 1.8% agarose to 4% formaldehyde gel in 1× MOPS buffer and transferred to a Hybond N+ nylon membrane (Amersham) in 10× SSC (1× SSC is 0.15 M NaCl and 0.015 M sodium citrate) buffer overnight. The membrane was rinsed with MilliQ water and the RNA was UV cross-linked using a Stratalinker cross-linking oven (Stratalinker) (set: autocross-linking 1200). Prehybridization (at least 2 h) and hybridization (12 to 60 h) of the membrane with each ³²P-labeled DNA probe was performed at 65°C in modified Church’s hybridization solution (0.5 M phosphate buffer, pH 7.2, 7% SDS [w/v], 10 mM EDTA, and 1 % BSA) (Church and Gilbert, 1984). After hybridization, the membranes were washed at least two times at 50 to 55°C for 10 min with high stringency washing buffer. Visualization of ³²P signal was achieved by autoradiography or with a phosphor imager (Molecular Dynamics). Each membrane was subjected to mild stripping 1 h at 60°C in 0.5% SDS and rehybridized with a loading control probe for a transcript with a different size (depending on the case, RPL37, 25S rRNA, or ORM). Based on the G+C content of the transcript of interest, probes were labeled with [α-³²P]dATP or [α-³²P]dCTP using random priming (Feinberg and Vogelstein, 1983) (see Supplemental Tables 2 and 3 online). The DNA templates necessary for this reaction were produced by PCR using primers corresponding to the full length or a fragment of the gene of interest (see Supplemental Tables 2 and 3 online). The same PCR fragments were used as probes during chloroplast run-on assays.

For isolation of genomic DNA, 0.1 to 1 × 10⁸ cells were centrifuged 5 min at 3200g at 4°C, and the pellet was incubated at 65°C in 600 μL of cetyltrimethylammoniumbromide extraction buffer (2% [w/v] cetyltrimethylammoniumbromide, 1.4 M NaCl, 20 mM EDTA, pH 8, 2% [v/v] β-mercaptoethanol, 100 mM Tris-HCl, pH 8.0) for 1 h. Genomic DNA was isolated by phenol/chloroform/isoamylalcohol (25:24:1) extraction followed by isopropanol precipitation. Any RNA contamination was removed through DNase-free RNase A treatment. Concentration and quality of each DNA preparation were determined by Nanodrop prior to use. To test the homoplasmicity of the transformants, PCR analysis was carried using 1 ng (1%) or 100 ng (100%) of genomic DNA and performing 40 PCR cycles.

For cDNA synthesis and quantitative PCR analyses, 5 to 10 μg of total RNA were treated with DNase I (Roche) for 20°min at 37°C, repurified using the RNeasy mini kit (Qiagen), and resuspended in Nuclease-Free Water (not diethylpyrocarbonate [DEPC] treated) (Ambion AM9932). Reverse transcription was performed using PrimeScript first-strand cDNA synthesis (Takara) following the manufacturer’s instructions. Quantitative PCR reactions were performed using LightCycler480 SYBR Green I Master (Roche) following the manufacturer’s instructions. Water controls were included in each 96-well PCR reaction, and dissociation analysis was performed at the end of each run to confirm the specificity of the reaction. Cycle threshold (Ct) values were obtained through LightCycler480 software release 1.5.0, and relative changes in gene expression were calculated using the 2^−ΔΔCt method. Oligonucleotides used are indicated in Supplemental Table 4 online.

Cell Vitality Test and Light Microscopy

Trypan Blue was added in each sample of cell culture to a final concentration of 0.02%. Cells were observed with a microscope (Eclipse 80i, Nikon), ×60/1.4 Plan Apochromat lens, 1.4–numerical aperture condenser lens, and differential interference contrast optics. Images were recorded using a digital camera, and contrast was optimized using Nis Elements F3.0 software.

NanoString nCounter Expression Analysis

Five nanograms of RNA from the A31, RR5, and DPF1 C. reinhardtii strains were hybridized with multiplexed Nanostring probes after experimentally comparing the signal detected upon hybridization of 500, 200, 100, 10, 5, and 1 ng of DNA-free total RNA extracted from three independent biological replicates. Samples were processed according to published procedures (Geiss et al., 2008). Barcodes were counted for 1150 fields of view per sample. Background correction was done by subtracting the mean + 2 sd of the negative controls for each sample. Values <1 were fixed to 1. Positive controls were used as quality assessment: We checked that the ratio between the highest and the lowest positive controls average among samples was below 3. Then, counts for target genes were normalized with the geometric mean of the four reference genes (Rpl37, Cblp, Tab2, and cytb), which were selected as the most stable using the geNorm algorithm (Vandesompele et al., 2002). Fold changes were calculated as ratios of the geometric mean of the counts in experimental conditions for RR5 and DPF1 at 0, 12, 48, and 146 h of vitamin treatment over that of control strain A31 at the same time points. Fold changes are expressed in log₂. Analysis of the hybridization results and preparation of heat maps were performed using Partek Genomic Suite 6.6 beta. Total RNAs used for nCounter analysis were extracted with fresh TRIzol reagent (Life Technologies) and further purified with acid-phenol:chloroform, (with indole-3-acetic acid, 125:24:1), pH 4.5, extraction (Life Technologies; AM9720) and chloroform ultrapure (AppliChem; A3633). Subsequently, they were precipitated with isopropanol at room temperature for 10 min, centrifuged 20 min at 8,000g at 4°C, washed twice with cold ethanol 75%, dried for 5 to 10 min at room temperature, and resuspended in 80 μL of nuclease-free water (not DEPC treated) (Ambion AM9932). Only RNA preparations with OD ratios (260/230 and 260/280) equal or higher than 2 were processed further. To remove any trace of genomic DNA contamination, 5 μg of each RNA sample was subjected to one up to three subsequent DNase I treatments (Roche) in presence of ribonuclease inhibitor (RNASE-OUT; Life Technologies). RNAs were then purified further using RNeasy MinElute cleanup kit and tested for effective removal of genomic DNA through a PCR reaction with the primer pair SR277/SR278 (which amplifies the genomic 23S rRNA locus in the inverted repeat region of the plastid genome). Their quality was further certified by the Qubit fluorometer/Quant-iT (Invitrogen) and by the 2100 Bioanalyzer (Agilent Technologies).

Statistical Analysis

Three biological replicates were used for the Nanostring hybridizations. The error bars are ±se of the population (n = 3). Standard errors were calculated using the square root of an unbiased estimator of the variance of the population in Matlab.

Chloroplast Run-on Transcription Assay

The reaction was performed as described by Gagné and Guertin (1992) with few modifications. Briefly, 2 × 10⁸ cells from a culture in exponential phase were pelleted at 4°C for 5 min at 2500g, washed with cold run-on washing buffer (10 mM HEPES, pH 7.5, 250 mM Suc, 150 mM KCl, 1 mM EDTA, and 0.4 mM PMSF freshly supplemented), frozen, and thawed to allow for cell wall permeabilization. Subsequently, 80 μL of transcription buffer (50 mM HEPES, pH 7.5, 1 M Suc, 60 mM MgCl₂, 15 mM DTT, 50 mM NaF, 240 units of RNase inhibitors, 250 μCi of [α-³²P]UTP, 400 nmol rATP, 200 nmol rGTP, and 200 nmol rCTP, freshly supplemented) was added and the cells were incubated for 15 min at 26°C. RNA was extracted with 1 mL of fresh TRIzol reagent (Life Technologies) and further purified with another acid-phenol:chloroform (with indole-3-acetic acid, 125:24:1), pH 4.5, extraction (Life Technologies). Subsequently, it was precipitated with cold ethanol at −20°C for 48 h, washed with cold ethanol 75%, dried for 5 to 10 min at room temperature, resuspended in 200 μL of nuclease-free water (not DEPC treated) (Ambion AM9932), quantified by Nanodrop, stored at −20°C, and diluted to the same concentration in 10 mL of hybridization buffer (5× SSPE [1× SSPE is 0.115 M NaCl, 10 mM sodium phosphate, and 1 mM EDTA, pH 7.4], 50% [v/v] deionized formamide, 5× Denhardt’s solution, 1% SDS, and 100 µg/mL of sheared denatured salmon sperm DNA) prior to use. DNA probes were obtained by PCR amplification of the gene of interest as during preparation of DNA probes for RNA gel blot analysis (Supplemental Table 3). One microgram of each PCR product was purified by gel extraction, precipitated by isopropanol, resuspended in a solution containing 0.4 M NaOH, 10 mM EDTA, pH 8, and a drop of Red Safe (Intron Biotechnology). It was denatured at 90°C for 10 min, cooled in ice for 10 min, and deposited onto a Hybond N⁺ membrane (Amersham) by pipetting or using hybridization manifold dot blot following the manufacturer’s instructions (Fisher Scientific). The membrane was quickly rinsed in MilliQ water and allowed to dry. DNA probes were then UV-cross-linked to the membrane using a Stratalinker cross-linking oven (Stratalinker) (set: autocross-linking 1200). Prehybridization (3 h) and hybridization (48 to 60 h) of the membrane with the labeled RNAs was performed at 42°C using the hybridization buffer described above. Membranes were washed twice or more, each time for 10 to 15 min at 42°C with 2× SSC containing 0.1% SDS. Visualization of ³²P signal was achieved by autoradiography as described above.

Polysome Preparation

The procedure for preparation of polysomes is a modified protocol of the methods described by Barkan (1988) and Minai et al. (2006). Chloramphenicol (100 μg/μL) was added to 100 mL of cell cultures in exponential phase 10 min prior to harvest by gentle centrifugation. Pellets were frozen, ground to powder in liquid nitrogen using mortar and pestle, and finally resuspended in 2 mL of polysome extraction buffer (200 mM Tris-HCl, pH 8.0, 20 mM KCl, 25 mM MgCl₂, 25 mM EGTA, 0.2 M Suc, 1% Triton X-100, 2% polyethelene-10-tridecyl-ether, and 0,5% sodium deoxycholate) supplemented with inhibitors (0.5 mg/mL heparin, 50 mM β-mercaptoethanol, 100 µg/mL chloramphenicol, 1 mM PMSF, 1 mM 1,10-phenanthroline, and 0.5% [v/v] protease inhibitor cocktail [Sigma-Aldrich]). The lysate was vortexed and centrifuged immediately at 4°C for 20 min at 8000g to remove unbroken cells and large cell debris. The supernatant was loaded on an 8-mL continuous Suc gradient prepared by an automatic gradient maker mixing 4 mL of a 15% Suc (w/v) solution (40 mM Tris-HCl, pH 8.0, 20 mM KCl, 30 mM MgCl₂, 5 mM EGTA, 50 mM β-mercaptoethanol, 0.5 mg/mL heparin, and 0.5% [v/v] protease inhibitor cocktail) with 4 mL of a 55% Suc (w/v) solution. The Suc gradients were centrifuged at 4°C for 3 h at 250,000g in a prechilled Beckmann SW41 rotor. Ten fractions of 1 mL were collected. Total RNA extraction was performed by precipitating each fraction overnight at −20°C through addition of 400 μL of 2 M KCl and 3 mL of ethanol. The following day, the RNAs were pelleted by centrifugation for 15 min at 8000g, resuspended in 400 μL of 0.1% SDS/5 mM EDTA, extracted once with an equal volume of phenol/chloroform/isoamyl alcohol (25:24:1), and reprecipitated with an equal volume of isopropanol at room temperature for 15 min. Purified RNAs were pelleted by centrifugation for 15 min at 8,000g at room temperature, washed with 75% ethanol, dried under vacuum for 5 min, resuspended in 5 mM EDTA and 0.1% SDS, and stored at −70°C. Five microliters of each RNA fraction was analyzed by RNA gel blotting.

Accession Numbers

Sequence data from this article can be found in the GenBank/EMBL data libraries under accession numbers M29284 (rps12) and EF587487 (rpoA).

Supplemental Data

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

Supplemental Figure 1. Identification of the Splicing Isoforms Generated upon Fusion of the Thi4 5′UTR with Nac2.
Supplemental Figure 2. Accumulation of Nac2 in Rep112 and A31 in the Presence and Absence of Vitamins.
Supplemental Figure 3. Effect of Vitamin Concentration on Maximum Photosynthetic Yield in Rep112.
Supplemental Figure 4. Homoplasmicity of the A31, RR5, and DPF1 Strains.
Supplemental Figure 5. RNA Analysis of DPF1.
Supplemental Figure 6. Kinetics of RNA Accumulation upon Repression of Chloroplast Translation (RR5) and Transcription (DPF1).
Supplemental Figure 7. Analysis of the Suppressors of RR5 and DPF1.
Supplemental Table 1. Growth Properties of the Vitamin-Repressible Strains of Chlamydomonas reinhardtii.
Supplemental Table 2. List of Primers.
Supplemental Table 3. List of DNA Probes Used for RNA Blot Analyses.
Supplemental Table 4. Primers Used for qPCR Analyses.
Supplemental Data Set 1. Time Course of RNA Quantification in RR5 and DPF1 upon Repression of Chloroplast Translation and Transcription (Nanostring nCounter Data).
Supplemental Methods 1.

Acknowledgments

We thank the Genomic Platform of the University of Geneva for performing the transcriptomic analysis, Emine Dinc for her valuable help during the preparation of the RNA samples, Martin Croft and Alison Smith (University of Cambridge, UK) for the plasmid containing the Thi4 riboswitch fused to the upstream region of the METE gene, Michael Moulin for sharing information about the Thi4 alternative transcripts and for HPLC measurements of thiamine in the suppressors strains, Teresa Fitzpatrick for the ThiC antibody, Olivier Vallon for the ClpP1 antibody, the ClpP-AUU strain, and his personal communication on the accumulation of the ClpP1 transcript in this strain, N. Roggli for photography and drawings, and M. Goldschmidt-Clermont for critical reading of the article. This work was supported by Grant 31003A_133089/1 from the Swiss National Foundation.

AUTHOR CONTRIBUTIONS

S.R. designed and performed the research, analyzed the data, and compiled the Methods. M.R. performed the research. O.S. performed the statistical analysis of the Nanostring nCounter expression data. J.-D.R. designed the research, analyzed the data, and wrote the article.

Glossary

PSII: photosystem II
PSI: photosystem I
Rubisco: ribulose-1,5-bis-phosphate carboxylase/oxygenase
5′UTR: 5′-untranslated region
TPP: thiamine pyrophosphate
F_v/F_m: the ratio between variable and maximum fluorescence
TAP: Tris-acetate phosphate
HSM: high-salt minimal
qRT-PCR: quantitative RT-PCR
PEP: plastid-encoded RNA polymerase
NEP: nucleus-encoded plastid RNA polymerase
TBS-T: 5% nonfat dry milk and 0.1% Tween 20
DEPC: diethylpyrocarbonate

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PERMALINK

Repression of Essential Chloroplast Genes Reveals New Signaling Pathways and Regulatory Feedback Loops in Chlamydomonas[W]

Silvia Ramundo

Michèle Rahire

Olivier Schaad

Jean-David Rochaix