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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 and Jean-David Rochaixa,1
a Department
of Molecular Biology and Plant Biology, University of Geneva, 1211 Geneva, Switzerland
Departments Platform, National Center of Competence in Research Frontiers in Genetics and Department of
Biochemistry, University of Geneva, 1211 Geneva 4, Switzerland
b Genomics
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 a-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
1 Address
correspondence to [email protected].
The author responsible for distribution of materials integral to the findings
presented in this article in accordance with the policy described in the
Instructions for Authors (www.plantcell.org) is: Jean-David Rochaix
([email protected]).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.112.103051
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
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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 59untranslated region (59UTR) 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 59UTR (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 59UTR, 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 a-subunit of the chloroplast DNAdependent 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 59UTR, 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 59UTR 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, Fv/Fm, 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 Fv/Fm
represents the repression of Nac2 expression. In Rep112, Fv/Fm
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, Fv/Fm 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
Repression of Essential Plastid Genes
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Figure 1. Reversible Vitamin-Controlled Repression of Chloroplast Gene Expression.
(A) Scheme of the vitamin-controlled repressible chloroplast gene expression system. The Nac2 gene, which is specifically required for the accumulation of the chloroplast psbD mRNA, is fused to the Thi4 59UTR containing the TPP-responsive riboswitch, and the MetE upstream region is used as
promoter. Addition of thiamine causes alternative splicing in the riboswitch region, which results in translation termination due to the inclusion of a stop
codon (red flag) (Croft et al., 2007). The yellow box below the second exon indicates the genomic location of the TPP riboswitch. Change of the box
color represents in a schematic way the conformational change of the riboswitch upon binding of TPP (green, no TPP binding; red, TPP binding).
Because Nac2 acts specifically on the psbD 59UTR, it is possible to render the expression of any chloroplast gene dependent on Nac2 by fusing its
coding sequence to the psbD 59UTR. To allow for phototrophic growth in the presence of the vitamins, the psbD 59UTR of the psbD gene was replaced
by the psaA 59UTR, thus making psbD expression independent of Nac2.
(B) Growth patterns of the wild-type (WT), nac2-26, Rep112, and A31 strains. Cells were spotted on TAP and HSM (minimal medium) plates in the
absence or presence of vitamins B12 (20 mg/L) and thiamine (20 mM) under an illumination of 60 mmol m22 s21.
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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 psaApsbD 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).
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 Fv/Fm 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 59UTR to rps12 encoding the ribosomal protein Rps12 and to the rpoA gene encoding the a-subunit of the chloroplast RNA polymerase. The
aadA cassette was inserted upstream of the psbD59-rpoA or
psbD59-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 psbD59-rpoA chimeric gene and
the RR transformants (translation inhibited) contain the psbD59rps12 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
Figure 1. (continued).
(C) Decrease of PSII activity of Rep112 after addition of vitamins. Fv/Fm was measured for the indicated strains at different times after addition of
vitamins (20 mM thiamine and 20 mg/L B12). The experiment was repeated three times with similar results.
(D) Growth curves of the indicated strains after addition of vitamins. To maintain cells in exponential growth during the entire time course, they were
diluted to 0.5 3 106 cells/mL when they reached a concentration between 2 and 4 3 106 cells/mL. The experiment was repeated three times with similar
results. div, divisions.
(E) Recovery of photosynthetic activity (Fv/Fm) upon transfer of Rep112 from vitamin-containing medium to vitamin-free medium.
(F) Growth of Rep112 under the same conditions as in (E). To maintain cells in exponential growth during the time course, they were diluted to 0.5 3 106
cells/mL when they reached a concentration between 2 and 4 3 106 cells/mL. The experiment was repeated three times with similar results.
Repression of Essential Plastid Genes
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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 (2) or presence of thiamine (T) and vitamin B12 (B12) in the nac2 and Rep112 strains.
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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 mg/L)
and thiamine (20 mM) under an illumination of 60 mmol m22 s21.
(B) Growth curves and Fv/Fm measurements in the presence of vitamins. Cell concentration and Fv/Fm 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 3 106 cells/mL when
they reached a concentration between 2 and 4 3 106 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 (2) of vitamins.
(D) Light microscopy of cells from A31, DPF1, and RR5 in the presence (+) or absence (2) of vitamins. Cells were stained with Trypan blue after 6 d in
the light (60 mmol m22 s21) or 12 d in the dark.
1024 and 1023, respectively, after several days (Figure 3A) (see
Methods for further characterization of these suppressors).
Of the transformants obtained, two of each type were used for
further studies: DPF1 and DPF9 (containing psbD59-rpoA) and
RR5 and RR20 (containing psbD59-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 mmol m22 s21 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 runoff 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
Repression of Essential Plastid Genes
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Figure 4. Polysome Profiles and Accumulation of Chloroplast Proteins in RR5 upon Repression of rps12 with Vitamins.
(A) Polysome isolation. Polysomes were isolated from untreated RR5 cells or RR5 cells treated with the vitamins for 48 h and subjected to Suc density
gradient centrifugation. Gradients containing EDTA to disrupt the polysomes are indicated. RNA was isolated from each fraction and hybridized to the
chloroplast psaB, psbD, and tufA probes. ORM encoding a cytosolic protein was used as control.
(B) Immunoblot analysis of Rps12, ribosomal proteins, and TufA in RR5 and A31. Proteins were examined at different time points after addition of the
vitamins using antibodies against the indicated proteins.
(C) Immunoblot analysis of ClpP1 and proteins involved in photosynthesis. Conditions were as in (B).
(D) Immunoblot analysis of Vipp1, Alb3.2, Hsp90C, DnaJ, and Hsp70B. Conditions were as in (B).
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, a- and b-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
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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 psbD59-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.
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
Repression of Essential Plastid Genes
9 of 20
Figure 5. RNA Analysis of RR5 and DPF1.
(A) RNA samples were examined at the indicated times after addition of vitamins. The size difference of rps12 mRNA in RR5 and A31 is due to the fusion
of the psbD 59UTR to the rps12 coding sequence in RR5. The hybridization probes for detecting other chloroplast RNAs are indicated. Cytoplasmic 25S
rRNA was not affected by the vitamin treatment and used as a loading control.
(B) RNA samples from A31, RR5, and clpP-AUU were examined after 120 h of vitamin treatment and hybridized with clpP1 and rpoA probes. Equal
loading was checked by rRNA staining with ethidium bromide.
(C) and (D) RNA gel blot analysis of rpoA mRNA in DPF1. RNA samples were examined at different times after addition of the vitamins using the
indicated hybridization probes.
(E) and (F) Chloroplast transcriptional activity is reduced in the DPF1 strain grown with vitamins. Cells were grown for 48 (E) and 144 h (F) in the
absence or presence of the vitamins, permeabilized through one freezing-defreezing cycle, and pulse labeled with 32P-UTP for 15 min. RNA was
isolated and hybridized with a filter containing the indicated gene probes.
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The Plant Cell
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.
Repression of Essential Plastid Genes
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 chloroplastencoded 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.
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
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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.
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).
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
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The Plant Cell
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
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.
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 59UTR 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 59UTR 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 59UTR contains the major target
site for Nac2, the system can be used for any chloroplast gene
by fusing the psbD 59UTR 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
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
Repression of Essential Plastid Genes
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 chloroplastencoded proteins with a time course that depends on the halflife 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 lightindependent 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,
13 of 20
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).
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 nucleusencoded 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
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The Plant Cell
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 assemblydependent feedback process in which unassembled CES subunits inhibit
directly or indirectly their own translation (green circular lines) (reviewed
in Choquet and Wollman, 2002).
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 s70-type promoters,
consists of two a-subunits, and the b-, b’-, and b’’-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 a-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 s 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
Repression of Essential Plastid Genes
15 of 20
METHODS
of 1024 and 1023, 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 59UTR sequence that
would make the stabilization of psbD mRNA independent of Nac2, the
level of Nac2 was determined by immunoblotting and the psbD 59UTR 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 59UTR 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.
Strains, Growth Conditions, and Media
Construction of the Strains Rep112, A31, RR5, and DPF1
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 mmol m22 s21)/dim light (10
mmol m22 s21) or in the dark depending on their phototropic growth abilities.
Medium with vitamins was prepared in the following way: Stock
solutions 10003 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 3 106 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 mM thiamineHCl and 20 mg/L B12.
For growth tests, 7 mL of 1 3 104, 5 3 104, 1 3 105, 5 3 104, and 1 3
106 cells were spotted on solid TAP or HSM medium with or without
vitamins and placed in continuous light (60 mmol m22 s21) unless indicated otherwise.
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).
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).
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
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 3 106 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 mg of SapI-linearized pRAM23.1 and then
transformed by electroporation in a 0.2-cm electroporation cuvette (BioRad) using the Bio-Rad Gene Pulser II set to 0.75 kV, 25 mF, 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 mmol m22 s21). 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 mM thiamine-HCl/10 mg B12/L). Their
vitamin-repressible PSII activity was further analyzed by fluorescence
measurements (Fv/Fm). 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 3 107 cells grown in TAP
medium were plated on solid agar TAP supplemented with 100 mg/mL
spectinomycin (Sigma-Aldrich S9007) and bombarded with 550-nmdiameter gold particles (Seashell Technology S550d) coated with 300 ng
of pRAM56.2 (rcy_aadA_psaA59UTRpsbD) plasmid DNA. After 1 week of
growth at 25°C in constant light (60 mmol m22 s21), single colonies were
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The Plant Cell
picked, replated four times on TAP supplemented with 100 mg/mL
spectinomycin, and screened for photoautotrophic growth on HSM
medium + 20 mM thiamine-HCl + 20 mg B12/L. The homoplasmic state of
the selected transformants was verified by PCR analysis of chloroplast
genomic DNA and through fluorescence measurements (Fv/Fm).
In the case of the A31 strain, a similar protocol was used for chloroplast
biolistic transformation with pRAM61.1 (aadA_psbD 59UTR_rps12) and
pRAM68.13 (aadA_psbD 59UTR_rpoAFLAG) plasmid DNA except that the
transformants were replated under a more stringent selective pressure using
TAP medium supplemented with 750 mg/mL spectinomycin. Single colonies
were tested for growth on TAP + 10 mM thiamine-HCl + 20 mg B12/L.
Fluorescence Measurements
Maximum quantum efficiency of PSII (Fv/Fm) 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 3 108 cells). Pellets were frozen in liquid nitrogen and stored
at 270°C until use. Cells were lysed by resuspending the pellet in the
desired volume (typically 200 mL) of a 23 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 mL 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-mm 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 isopropylb-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 SDSPAGE (16 3 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 270°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 mg of total RNA was
electrophoresed in a 1.2 to 1.8% agarose to 4% formaldehyde gel in 13
MOPS buffer and transferred to a Hybond N+ nylon membrane (Amersham) in 103 SSC (13 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 [a-32P]dATP
or [a-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 3 108 cells were centrifuged 5
min at 3200g at 4°C, and the pellet was incubated at 65°C in 600 mL of
Repression of Essential Plastid Genes
cetyltrimethylammoniumbromide extraction buffer (2% [w/v] cetyltrimethylammoniumbromide, 1.4 M NaCl, 20 mM EDTA, pH 8, 2% [v/v]
b-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 mg 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 22DDCt 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), 360/1.4 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 mL
of nuclease-free water (not DEPC treated) (Ambion AM9932). Only RNA
17 of 20
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 mg 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 6SE 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 3 108 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 mL 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 mCi of [a-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 220°C for 48 h, washed with cold ethanol
75%, dried for 5 to 10 min at room temperature, resuspended in 200 mL of
nuclease-free water (not DEPC treated) (Ambion AM9932), quantified by
Nanodrop, stored at 220°C, and diluted to the same concentration in 10
mL of hybridization buffer (53 SSPE [13 SSPE is 0.115 M NaCl, 10 mM
sodium phosphate, and 1 mM EDTA, pH 7.4], 50% [v/v] deionized
formamide, 53 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 23 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
18 of 20
The Plant Cell
(100 mg/mL) 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 b-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 b-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 220°C through addition of 400 mL 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 mL 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 270°C. Five microliters of each RNA fraction was analyzed by
RNA gel blotting.
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.
Received July 20, 2012; revised November 12, 2012; accepted December 11, 2012; published January 4, 2013.
REFERENCES
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 59UTR 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).
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Repression of Essential Chloroplast Genes Reveals New Signaling Pathways and Regulatory
Feedback Loops in Chlamydomonas
Silvia Ramundo, Michèle Rahire, Olivier Schaad and Jean-David Rochaix
Plant Cell; originally published online January 4, 2013;
DOI 10.1105/tpc.112.103051
This information is current as of June 17, 2017
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