Advertisement
JBC

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Originally published In Press as doi:10.1074/jbc.M408477200 on September 23, 2004

J. Biol. Chem., Vol. 279, Issue 48, 50366-50374, November 26, 2004
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/48/50366    most recent
M408477200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Schönfeld, C.
Right arrow Articles by Kruse, O.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Schönfeld, C.
Right arrow Articles by Kruse, O.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

The Nucleus-encoded Protein MOC1 Is Essential for Mitochondrial Light Acclimation in Chlamydomonas reinhardtii*

Christine Schönfeld{ddagger}, Lutz Wobbe{ddagger}, Rüdiger Borgstädt{ddagger}, Alexandra Kienast{ddagger}, Peter J. Nixon§, and Olaf Kruse{ddagger}

From the {ddagger}Molecular Cell Physiology Group, Department of Biology, University of Bielefeld, 33501 Bielefeld, Germany and the §Department of Biological Sciences, Wolfson Biochemistry Building, Imperial College London, South Kensington Campus, London, SW7 2AZ, United Kingdom

Received for publication, July 27, 2004 , and in revised form, September 14, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Mitochondrial respiration plays an important role in optimizing photosynthetic efficiency in plants. As yet, the mechanisms by which plant mitochondria sense and respond to changes in the environment are unclear, particularly when exposed to light. Here we describe the characterization of the Chlamydomonas reinhardtii mutant stm6, which was identified on the basis of impaired state transitions, a mechanism that regulates light harvesting in the chloroplast. The gene disrupted in stm6, termed Moc1, encodes a homologue of the human mitochondrial transcription termination factor (mTERF). MOC1 is targeted to the mitochondrion, and its expression is up-regulated in response to light. Loss of MOC1 causes a high light-sensitive phenotype and disrupts the transcription and expression profiles of the mitochondrial respiratory complexes causing, as compared with wild type, light-mediated changes in the expression levels of nuclear and mitochondrial encoded cytochrome c oxidase subunits and ubiquinone-NAD subunits. The absence of MOC1 leads to a reduction in the levels of cytochrome c oxidase and of rotenone-insensitive external NADPH dehydrogenase activities of the mitochondrial respiratory electron transfer chain. Overall, we have identified a novel mitochondrial factor that regulates the composition of the mitochondrial respiratory chain in the light so that it can act as an effective sink for reductant produced by the chloroplast.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
In plant cells, respiration within the mitochondrion not only provides ATP but also serves an important role in the maintenance of an appropriate reduction/oxidation (redox) state within the chloroplast (1). Through the use of metabolite shuttles within the inner envelope membrane of the chloroplast, reducing equivalents produced by photosynthetic electron transport activity can be exported and consumed within the mitochondrion through respiration (2). The respiratory chain in plant mitochondria possesses a variety of dehydrogenases and oxidases. Besides a typical rotenone-sensitive type I NADH:Q1 oxidoreductase (complex I), the inner membrane also contains rotenone-insensitive internal and external facing type II NAD(P)H:Q oxidoreductases (3–5). Two types of oxidases catalyze reduction of oxygen: a typical cyanide-sensitive cytochrome c oxidase and a cyanide-insensitive family of alternative oxidases. Cytochrome oxidase is coupled to ATP synthesis, whereas the alternative oxidase is not and possibly acts as a valve to allow oxidation of the ubiquinone pool (6).

A key signaling molecule in the chloroplast thylakoid membrane is plastoquinone (PQ), which according to its redox state appears to control gene transcription in the chloroplast (7) and to regulate light harvesting, which in turn appears to control the ratio of linear to cyclic electron flow in algal chloroplasts (8). Because the redox state of the plastoquinone pool is affected by mitochondrial respiration in the dark (9), and possibly in the light, the mitochondrion has the capacity to influence both chloroplast function and development. Important evidence for the role of the respiratory chain in optimizing photosynthesis (10–13) and the development of the chloroplast (14) has come from analysis of mutants lacking various respiratory complexes, as well as through the use of various mitochondrial-specific inhibitors (15).

It is already known that the mitochondrial proteome changes in the light. For instance, in potato, expression of a nucleus-encoded internal rotenone-insensitive NADH dehydrogenase (NDA1) is switched on by light, probably to oxidize NADH produced during photorespiration (16). However, there is no information about how expression of the mitochondrial genome is coordinated with the rest of the cell in response to light stress.

One approach to identify components that are important for the redox-controlled regulation of photosynthesis, including factors possibly involved in mitochondrial gene expression, is to screen for mutants that are perturbed in state transitions. This is a redox-controlled mechanism that balances the excitation of the two photosynthetic reaction centers involved in linear electron flow, photosystems I and II (17, 18). When the PQ pool, which is a component of the electron transport chain linking PSII and PSI, becomes more reduced, mobile light-harvesting antennae detach from PSII, following redox-activated phosphorylation, and dock with PSI, thus increasing the excitation of PSI and driving the PQ pool to become more oxidized (into so-called state 2). The reverse process occurs when the PQ becomes more oxidized (to achieve state 1) (19, 20). We and others have developed a rapid chlorophyll fluorescence-based screen to identify colonies of the model organism, the green alga Chlamydomonas reinhardtii, that are blocked in state transitions (the interconversion of state 1 and state 2) (21, 22). Mutants identified in such screens are beginning to identify the key genes involved in regulating state transitions (23). Here we identify another gene that is important for state transitions. Surprisingly, the gene product is targeted to the mitochondrion, not the chloroplast, and is involved in regulating mitochondrial gene expression.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Strains and Culture Conditions—The C. reinhardtii strain CC 1618 (arg7 cw15, mt-) was obtained from the Chlamydomonas Genetics Center Collection (Duke University, Raleigh, NC) and used as WT control in all experiments. WT13(mt+) is derived from the original WT 137c strain (Jacqueline Girard-Bascou, Paris, provided by Dr. Saul Purton, University College London, UK) and was used for segregation experiments.

All strains used were cultivated mixotrophically in TAP medium (Tris acetate-phosphate, pH 7.0) by illumination with 40 µmol m-2 s-1 white light at 20 °C (24) in a 12-h light-dark cycle to a cell density of 2 x 106 cells/ml. When required (for the arginine auxotroph strain CC1618) the medium was supplemented with 110 µg of arginine/ml.

Mutant Construction and Genetic Analysis—Nuclear transformation was performed as described following the methods of Refs. 25 and 26. Plasmids pARG7.8, containing a 7.8-kb genomic DNA fragment of the C. reinhardtii argininosuccinate lyase gene (27) and a 0.4-kb fragment of bacteriophage {varphi}X174 DNA (28), were used for transformation experiments. Prior to use, pARG7.8 was linearized by digestion with BamHI.

Genetic crosses were performed with WT13 and the mutant as described by Ref. 24. DNA sequences flanking the tag were cloned by ligation-mediated suppression-PCR (29) and by plasmid rescue (30).

Complementation—For complementation, a co-transformation strategy was established using the Cry1 gene as a dominant selectable marker conferring resistance to emetine (plasmid p613) (31) together with a 37-kb Moc1-containing cosmid. The cosmid was isolated from a cosmid library (kind gift of Dr. Saul Purton) by hybridization experiments using a specific probe derived from the 500-bp non-coding DNA region located between Moc1 and Toc1.

Cells were grown in TAP medium, harvested in the late logarithmic growth phase (OD750 nm = 0.8) and resuspended in TAP-1/10 N. The suspension was used in glass bead transformations, and transformed cells were collected in TAP-1/10 N medium and incubated for 4 days. Cells were then resuspended in TAP medium and incubated for another 8 days. Again, cells were pelleted, resuspended in TAP, spread over TAP agar plates, and grown for 12 days. Successful complementation was confirmed by PCR using Moc1-specific primers and by the recovery of state transitions as assayed by fluorescence video imaging.

Fluorescence Video Imaging, Pulse Amplitude-modulated Fluorescence Spectroscopy, and 77 K Fluorescence Spectroscopy—Screening of the mutants for state transitions defects was performed as described earlier (22) by video imaging with a FluorCam 700 MF apparatus (Photon System Instruments) at room temperature and at 77 K with a fluorescence/luminescence spectrometer (PerkinElmer Life Sciences LS50B). Samples were measured following illumination with 40 µmol m-2 s-1 620 nm light to gain state 2. For state 1 images, pictures were taken in the same white light briefly after an illumination period of 15 min entirely with 80 µmol m-2 s-1 710 nm (Schott filter, Germany) of PSI light. Alternatively, cells were adjusted to state 2 by incubation in the dark for 30 min under nitrogen atmosphere according to Ref. 9. Photochemical quenching was determined using pulse amplitude-modulated room temperature fluorescence spectroscopy (Heinz Walz GmbH) and fluorescence video imaging (Fluor Cam 700 MTM) and calculated as 1 - qp = 1 - (Fm' - Fs)/(Fm' - Fo') according to Ref. 32.

Chloroplast and Mitochondria Isolation—Chloroplasts were isolated according to Ref. 33. Mitochondria were isolated according to Ref. 34 using the supernatant of the chloroplast isolation after breaking the cells with a BioNebulizer (kindly provided by J. Nickelsen, Bochum, Germany). The purity of the preparations was confirmed by immunoblotting with anti-D2 (for possible chloroplast contamination) and anti-Cyt c (for possible mitochondria contamination).

Isolation of Nucleic Acids, Northern Analysis, and RT-PCR—Northern analysis was performed according to Ref. 35. RT-PCR was used for semiquantitative transcript analysis using the GeneCraftTM Invitrogen procedure.

Prior to RNA isolation, cells were cultivated in the dark until OD750 nm = 0.6 and, if desired, incubated for a further 4–6 h under different light conditions. RNA isolation was performed following a protocol of C. Admundsen (biosci.cbs.umn.edu/~amundsen/chlamy/methods/RNA_prrep.html) as described earlier (29) and DNA isolation using the DNeasy plant mini kit (Qiagen).

Immunoblotting—Protein analysis by immunoblotting was performed as described earlier by electroblotting onto nitrocellulose membranes (22). The polyclonal anti-phosphothreonine antibodies were obtained from Zymed Laboratories Inc.. Anti-Cyt c, -AOX1, and -COX90 are Chlamydomonas antisera provided by F. A. Wollman (Unité Propre de Recherche-CNRS Paris, France), Sabeeha Merchant (UCLA, CA), and Saul Purton, respectively. Anti-LCHII is a spinach antiserum provided by S. Jansson (Umea, Sweden).

Anti-MOC1 was produced by GenoSys from a keyhole limpet hemocyanin-conjugated 13-amino-acid synthetic peptide identical to the MOC1 C terminus. All antisera were polyclonal and used in conjunction with the anti-rabbit-IgG (Sigma)/anti-mouse-IgG (Sigma) alkaline phosphatase reaction or the CDP Star (Roche Applied Science) reaction, respectively.

Protein Phosphorylation—Phosphorylated proteins were detected by Western analysis using polyclonal anti-phosphothreonine (Zymed Laboratories Inc.) in state 1- and state 2-adapted cells as described earlier (22). States were fixed by the addition of p-benzoquinone according to Ref. 9.

Lipid Peroxidation—Lipid peroxidation was measured indirectly by the concentration of malonyldialdehyde and other carbonyl byproducts of lipid peroxidation, which form a colored complex in reaction with thiobarbituric acid that can be monitored at 532 nm (36).

Enzyme Activity Measurements—Cytochrome oxidase activity was measured in isolated mitochondria using a cytochrome c oxidase assay kit (Sigma) following the technical instructions. NADH:Q oxidoreductase activity and external NADPH oxidation activity were measured in isolated mitochondria according to Ref. 16. Mitochondria were osmotically burst in 1 mM MOPS, pH 7.2, 0.1 mM EGTA for 6 min at room temperature to permeabilize the membrane.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Isolation of State Transition Mutants—To identify genes involved in state transitions within C. reinhardtii, we screened a library of potentially tagged nuclear mutants that was generated through the random insertion of plasmid pArg7.8, carrying the Arg7 gene, into the nuclear genome of the arginine auxotrophic strain, CC1618 (27, 28). By selecting on medium lacking arginine, only colonies containing Arg7 inserted into the nuclear genome were able to grow. A plate-based fluorescence video-imaging screen, which involves the recording of chlorophyll fluorescence from individual colonies before and after illumination with light that induces state transitions in WT, was used to identify potential state transition mutants (stm) (22). Of the 2 x 104 colonies screened, seven possible stm mutants were identified. The characterization of mutant stm6 is described here.

stm6 Is Blocked in State 1 and Impaired in PSII-LHC Phosphorylation in the Light—Chlorophyll fluorescence assays conducted by fluorescence emission spectroscopy at 77 K (Fig. 1a) and chlorophyll video imaging with actinic 620 nm light at room temperature (Fig. 1b) revealed that stm6 was blocked in state 1. In contrast to the WT, light preferentially absorbed by PSII was unable to drive the cells into state 2, which is monitored by an increase in the fluorescence coming from PSI at 720 nm (Fig. 1a). It should be noted that a transition to state 2 was blocked to a far less extent when the PQ pool was driven reduced in the dark through anaerobiosis (data not shown). This result indicated that the observed block in state transitions in stm6 was primarily a defect observed in the light.



View larger version (27K):
[in this window]
[in a new window]
  		FIG. 1.
Analysis of light-induced short-term adaptation mechanisms in the chloroplast. a, state transitions analyzed by 77 K fluorescence emission spectroscopy in WT, B13, and stm6. b, fluorescence video image of WT, B13, and stm6 on TAP agar plates in state 2 and state 1 (illumination with actinic light at 620 nm ± 15 min illumination with 710 nm PSI light). c, in vivo phosphorylation protein pattern of thylakoid membranes isolated from state 2 (S2)- and state 1(S1)- (dark-) adapted WT and stm6 cells. Arrows indicate P9 and P11 proteins

 
Since state 2 is associated with the phosphorylation of threonine residues in the light-harvesting (LHC) antenna proteins of PSII (19), anti-phosphothreonine antibodies were used to assess the level of phosphorylation in immunoblots. Stm6 showed a marked reduction in the phosphorylation levels of CP29 (P9) and LHC-P11 proteins upon illumination conditions (state 2) that promote phosphorylation of these proteins in WT (Fig. 1c). Interestingly, the phosphorylation levels of the P13 and P17 proteins in stm6 in state 2 were comparable with WT levels, indicating regulatory differences in the phosphorylation mechanism between the various LHC proteins. Furthermore, no differences between WT and mutant were obtained when cultivated for 12 h in the dark (state 1). Control immunoblots confirmed that stm6 still contained WT levels of LHC proteins (Fig. 1c).

As transition to state 2 in the light is controlled by the reduction state of the PQ pool, it was important to confirm that the PQ pool could be driven to a reduced state in the light. To test this, chlorophyll fluorescence measurements were performed to determine the photochemical quenching parameter, qp, from which 1 - qp, which is a measure of the reduction state of the PQ pool, can be calculated (32). Under growth light conditions, the PQ pool was in a more reduced state in stm6 as compared with WT (1 - qp of 0.77 in stm6 and 0.59 in WT) (Table I).


View this table:
[in this window]
[in a new window]
  		TABLE I
Redox state of plastoquinone (1-qp), lipid peroxidation rates (malonyldialdehyde (MDA)), respiration rates, and external NADPH oxidation activity (NDB1) of WT, stm6, and B13 strains cultivated in standard growth light

 
Disruption of a Single mterf-type Gene Encoding for a Mitochondrial Protein Is Responsible for the Phenotype of stm6 —To test whether the Arg marker carried by plasmid pArg7.8-{varphi}X174 was tightly linked to stm6, crosses were performed between WT13 (mt+) and stm6 and the progeny scored for the existence of {varphi}X174 DNA (by PCR) and the ability to do state transitions as assessed by video imaging. Of the 50 progeny that contained {varphi}X174 DNA, all showed impaired state transitions. In contrast, all progeny lacking {varphi}X174 DNA showed normal state transitions (data not shown). These data suggested that stm6 was an Arg-tagged mutant.

A combination of plasmid rescue and ligation-mediated suppression-PCR led to the identification of the site of insertion of the Arg marker in stm6 (Fig. 2a). Sequencing and PCR analysis revealed that stm6 contained an insertion of two pArg7.8 plasmids at a single site (Fig. 2a) and the deletion of ~2 kb of genomic DNA (Fig. 2a). Subsequent sequence analysis and homology searches revealed that two genes were affected by the random integration of the two pArg7.8 plasmids. One gene was described previously as a nuclear transposon (Toc1) (37); the second gene had not been described previously and was subsequently designated Moc1 (mterf-like gene of Chlamydomonas) (AF531421 [GenBank] ).



View larger version (31K):
[in this window]
[in a new window]
  		FIG. 2.
Identification and localization of MOC1 in C. reinhardtii. a, identification of the genomic DNA site in stm6 affected by the tandem nuclear insertion of two pArg7.8 plasmids by sequence analysis after chromosome walking (ligation-mediated suppression-PCR) and by PCR analysis with specific moc1, toc1, and pArg7.8 primers. The two black arrow sets mark the primer pairs used for PCR. The two gray arrows mark the integration sites of the nuclear insertion in WT. b, RT-PCR with a specific cDNA probe of Moc1 to demonstrate that the Moc1 gene is not transcribed in stm6. c, localization of MOC1 as a mitochondrial (mito) protein by Western blotting with anti-MOC1. Mitochondrial Cyt c and chloroplast (chloro) D2 were used as controls (antisera dilution 1:1000).

 
PCR analysis in stm6 and WT using a 5'-specific Moc1-primer and a second primer derived from the Arg marker resulted in the amplification of a 1005-bp PCR product in stm6. This confirmed that the insertion caused a partial deletion of Moc1, leaving the 5' region of the gene untouched. The precise site of insertion of the pArg7.8 plasmids was confirmed through sequencing PCR fragments spanning the insertion sites. Overall, ~2 kb of the genome was deleted in stm6, extending from the 3' region of Moc1 to the 5' region of Toc1 (Fig. 2a).

To confirm that disruption of Moc1 caused the phenotype of stm6, complementation experiments were performed using a cosmid containing the Moc1 gene, which had been isolated from a cosmid library (kind gift of Dr. Saul Purton). DNA sequencing and PCR analysis revealed that this cosmid contained a 12-kb insert consisting of the entire Moc1 gene together with 2 and 5 kb of upstream and downstream flanking sequences, respectively. No other large open reading frames were identified in the insert of the cosmid. Complementation was performed using a co-transformation approach using the Moc1-containing cosmid in combination with a second vector containing the Cry1 gene conferring emetine resistance as a dominant selectable marker (31). Of ~200 emetine-resistant colonies assessed, four were found to have incorporated the intact Moc1 gene into the genome. These four colonies were able to express MOC1 and perform WT-like state transitions as assessed by fluorescence video imaging. According to previous slightly different two-plasmid co-transformation approaches (65, 66), a success rate of 2% with both transformed genes functionally expressed was expected. One of these colonies, B13, was examined in more detail and found to exhibit a clear WT phenotype with no perturbations in light acclimation (see Figs. 1, a and b, 4, and 6a and Table 1). In contrast, 9 out of the 200 emetine-resistant colonies that contained only fragments of the Moc1 gene as assessed by PCR were unable to perform state transitions. From these data, we conclude that Moc1 is required for functional light acclimation in C. reinhardtii.



View larger version (45K):
[in this window]
[in a new window]
  		FIG. 4.
stm6 is sensitive to light stress, and Moc1 is light-regulated. a, WT and stm6 cells cultivated for 200 h on HSM agar plates in different light intensities. b, WT, B13, and stm6 strains, cultivated to their mid-logarithmic growth phase in TAP medium, exposed to 1000 µmol m-2 s-1 white light for 4 h, subsequently inoculated on TAP agar plates and grown for 7 d in 40 µmol m-2 s-1 white light. High light cell rates for cells surviving high light treatment over a period of 4 h in WT, B13, and stm6 strains are shown. Cells were diluted to 1 x 104 cells/ml prior to light treatment. c, evidence for light regulation of Moc1: Moc1-Northern blots of isolated RNA from dark- and light-incubated (200 µmol m-2 s-1) WT. Shown are anti-MOC1 immunoblots of mitochondria isolated from dark cultivated WT cells (D) exposed to moderate high light (200 µmol m-2 s-1) for 0 and 6 h and from mitochondria isolated from standard light-cultivated WT cells (L) (96 h, 40 µmol m-2 s-1). M, marker protein.

 



View larger version (30K):
[in this window]
[in a new window]
  		FIG. 6.
ATP level, cytochrome c, oxidase and NADH:Q oxidoreductase activity measurements in WT, stm6, and B13. a, ATP levels in cells of WT, stm6, and B13 grown in the light and dark. b, specific activities of cytochrome c oxidase and NADH:Q oxidoreductase in isolated mitochondria from WT and stm6 cells cultivated in the dark (D) and subsequently exposed to 6 h 200 µmol m-2 s-1 light (L). Error bars indicate six different independent measurements.

 
The Moc1 gene encodes a 35-kDa protein containing, somewhat surprisingly, a putative mitochondrial transit sequence and two mitochondrial transcription termination factor domains (mTerf) with leucine zipper motifs characteristic of DNA-binding proteins (38).

RT-PCR experiments confirmed that Moc1 was not expressed in stm6 (Fig. 2b). Anti-peptide antibodies raised against MOC1 were used to confirm that MOC1 was targeted to the mitochondrion rather than the chloroplast (Fig. 2c).

Overall, there were striking similarities to the 35-kDa human mTERF protein (34% based on alignments covering the full-length sequences) (Fig. 3) and their homologues in Drosophila melanogaster (DmTTF (39)) and sea urchin (mtDBP (40)). Human mTERF binds downstream of tRNA genes and is thought to be involved in controlling the amount of tRNA and rRNA synthesized within the mitochondrion as well as the expression of other mitochondrial genes and consequently the functionality of the mitochondrial respiratory chain (41–43). Interestingly, nine homologues to MOC1 with mTerf domains have been identified in the genome of Arabidopsis thaliana, four of which are predicted to be targeted to the mitochondrion (At1g61980, At2g44020, At4g02990, At2g03050) and one of which is predicted to be targeted to the chloroplast (At5g55580). Analysis of the Chlamydomonas nuclear genome data base, plus DNA hybridization experiments (data not shown), suggest that there is only one copy of Moc1 in C. reinhardtii.



View larger version (64K):
[in this window]
[in a new window]
  		FIG. 3.
Protein sequence of MOC1 and alignment with the human transcription termination factor mTERF. The model and the boxes mark the locations of the mitochondrial transit sequences and the identified mTERF domain structures in MOC1. Gray frame, potential transit sequence; black frame, potential mTERF motifs. Black-labeled nucleotides are homologous or identical.

 
Growth of stm6 Is Sensitive to High Light, and MOC1 Expression Is Light-regulated—Besides effects on state transitions, a striking phenotype of the stm6 mutant was its sensitivity to light stress. In contrast to WT, stm6 cells did not survive photoautotrophic long term growth experiments on high salt medium agar plates incubated for 200 h at a constant irradiance of 200 µmol m-2 s-1 (Fig. 4a).

After 4 h of illumination by 1000 µmol m-2 s-1 of visible light, cells of the mutant showed a marked decline in viability, whereas the WT and the complemented strain, B13, were nearly unaffected (Fig. 4b). stm6 also showed a dramatic increase in singlet oxygen inside the cell upon illumination as assessed by the accumulation of lipid hydroperoxides (Table I). Importantly, expression of MOC1 in WT was up-regulated at both the RNA and the protein level upon a dark-to-light transition (Fig. 4c), suggesting a particular function of this protein in the light.

Mitochondrial Respiratory Complexes Are Expressed Aberrantly in Light-grown Cultures of stm6 —Since most mitochondrial genes in C. reinhardtii encode subunits of respiratory complexes, experiments were performed to assess whether the absence of MOC1 had affected the expression and the activity of mitochondrial respiratory complexes. Immunoblotting analysis of cells grown in the light revealed that accumulation of the nuclear encoded subunit 90 of cytochrome oxidase (44) (COX90) was heavily reduced in stm6 in light-grown cultures (Fig. 5a), whereas levels of a 30-kDa homologue of the NAD9 subunit of plant complex I (64), identified on Scaffold 96, contig5 in the C. reinhardtii genome data base (genome.jgi-psf.org/chlre1/chlre1.home.html) and the alternative oxidase(s) (AOX1, AOX2 (45)), were much higher in the mutant as compared with WT.



View larger version (35K):
[in this window]
[in a new window]
  		FIG. 5.
Evidence for the role of MOC1 in light-induced control of mitochondrial respiration. a, immunoblot of dark- and light-grown WT and stm6 cells using antibodies specific for cytochrome oxidase subunit COX90, the Q-oxidoreductase subunit NAD9, and the alternative oxidase AOX. b, semiquantitative RT-PCR of Nad1, Nad2, Nad6, and Cox1 in dark-grown WT (D) and stm6 cultures exposed to 200 µmol m-2 s-1 light (L) for 6 h. mRNA levels were normalized to that of actin. Different PCR cycles of Cox1 and Actin are shown in three independent approaches to give evidence that the assay is quantitative. Column bars represent changes in expression levels (in percentage) upon light incubation in dark-grown WT and stm6 (100% = no changes). c, Northern analysis of a Ndb1 homologue in light-grown WT and stm6 cultures.

 
Effect of Moc1 Deletion on a Homologue of the Rotenone-insensitive NADPH Dehydrogenase NDB1—Two rotenone-insensitive NAD(P)H dehydrogenases of the mitochondrial respiratory chain have been characterized recently in potato and A. thaliana by Rasmusson and colleagues (4, 5, 16). NDA1 functions as an internal NADH dehydrogenase, which is switched on by light, probably to oxidize matrix NADH generated through photorespiration. NDB1 is an external facing NADPH dehydrogenase and appears to be expressed constitutively in both the light and the dark. Whether there are homologues of these two enzymes in C. reinhardtii mitochondria has not been investigated previously.

In BLAST searches of the Chlamydomonas genome data base (genome.jgi-psf.org/chlre1/chlre1.home.html), we could identify a homologue of NDB1 on JGI-Scaffold 595. RT-PCR experiments identified mRNA of Ndb1 in WT and stm6 (not shown). Therefore we conclude that a homologue of NDB1 is indeed expressed in C. reinhardtii. Northern analyses revealed that, as compared with WT, the transcript levels of NDB1 were heavily reduced in standard light-grown cultures of stm6 (Fig. 5c).

Activity measurements in isolated mitochondria confirmed an up to 70% reduction in the activity of the external NADPH oxidation (Table I). Overall, these data provide strong support for the idea that stm6 is compromised in its ability to oxidize external sources of NADPH.

Absence of MOC1 Perturbs Regulation of the Transcript Profile of the Mitochondrial Genome upon Exposure to Light— Given that MOC1 encodes a transcription factor homologous to human mTERF, experiments were directed at detecting possible perturbations to the transcript profile of the mitochondrial genome, which for C. reinhardtii consists of a 15.8-kb linear genome (46). A number of respiratory subunits are encoded by the genome including subunit 1 of cytochrome oxidase (Cox1 gene product), five subunits of complex I (Nad1–2, Nad4–6 gene products), and apocytochrome b (Cob gene product). Nad transcripts in Chlamydomonas are transcribed bidirectionally with Nad1, Nad2, and Nad6 together with Cox1 in one direction.

Against the background that mitochondrial RNA makes only 1% of the total RNA (47), highly sensitive semiquantitative RT-PCR analysis was performed to analyze differences in the transcription profile of certain mitochondrial genes. The experiments confirmed differences in transcript levels of the Nad1, Nad2, Nad6, and Cox1 genes between WT and stm6 following exposure of dark-grown cells to light. In WT, comparison between light- and dark-treated cells revealed only very moderate light-induced (10–25%) deviations of Nad and Cox1 transcript levels (Fig. 5b). In stm6, differences in the RNA abundance of mitochondrial genes between dark-cultivated and light-treated cultures were far more dramatic. stm6 showed a ~60% reduced level of the Cox1 transcript in the light, a 80–90% decrease of the Nad1 and Nad6 transcripts and a 210% increase of Nad2 transcripts (Fig. 5b).

Together, these results suggest an important role for MOC1 in influencing the expression of both complex I (NADH:Q oxidoreductase) and complex IV (cytochrome oxidase) at the level of transcription of the mitochondrial Nad genes and of Cox1. Since it is likely that COX1 is the first subunit to be assembled into the cytochrome oxidase complex, the level of COX1 expression is likely to be an important determinant of the amount of enzyme synthesized (48).

Acclimation of the Mitochondrial Respiratory Chain to Light Is Compromised in stm6 —Enzyme activity measurements were also performed to investigate whether there were differences in the activities of the respiratory complexes upon acclimation of dark-grown cells to 6 h of light. Following the light treatment, there was a dramatic decrease in cytochrome oxidase activity in stm6 (Fig. 6b). The NADH:Q-oxidoreductase activities of both WT and stm6 declined in the light. However, here, in contrast to the obtained differences in complex IV activity, the light-induced decrease of NADH:Q-oxidoreductase activity was greater for WT than for stm6 (Fig. 6b). Levels of ATP in light-grown stm6 cells were also determined to be at about 50% of the levels of the WT and B13 (Fig. 6a), consistent with a role for cytochrome oxidase in the generation of ATP from photosynthetically derived reductant, in line with earlier suggestions on higher plants (1). In contrast, after the light treatment, the cyanide-insensitive respiration rate in stm6, assigned to the activity of the alternative oxidases, was much higher than the WT level (Table I).


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
Mutants Affecting State Transitions in the Chloroplast— State transitions represent an elegant mechanism to regulate photosynthetic electron transport within the chloroplast at the level of light harvesting. By changing the size of the antenna associated with either PSI or PSII, the state transition mechanism maintains optimal electron transport through PSI and PSII under varying light conditions. In addition, in the case of Chlamydomonas, and possibly vascular plants, state 2, in which LHC is associated with PSI, promotes cyclic electron transfer around PSI and the production of ATP (49). State transitions are therefore also in principle important for regulating the ratio of ATP/NADPH produced during the light reactions of photosynthesis (50). The results presented here indicate that abnormal LHC state transitions can also be used as a marker to detect general perturbations in cellular light acclimation processes.

The functional control of state transitions is complex. It is clear that the chloroplast Cyt b6f complex plays an important role in regulating the LHC kinase activity (51–54), and more recently, good candidates for this kinase have been identified in A. thaliana (55) and C. reinhardtii (23).

Defects in state transition mutants can be caused directly by deletion of functional genes such as LHC kinases. However, state transitions seem to be mainly controlled by the redox state of the plastoquinone pool (19). The PQH2:PQ ratio is influenced not only by linear and cyclic photosynthetic electron transport but also by alternative proton/electron feeding mechanisms (56). Consequently, perturbations in the control of the cellular redox homeostasis could indirectly cause defects in redox-regulated short term light acclimation of the chloroplast.

In this work, we have now identified an additional gene product, MOC1, which is needed for functional state transitions. Most surprisingly, we have shown that MOC1 is targeted to the mitochondrion, not the chloroplast, and so its effect on state transitions must be indirect.

MOC1 Is a Member of a Family of Proteins with Transcription Termination Motifs—MOC1 encodes a homologue of the human mitochondrial transcription termination factor mTERF, which in humans plays a central role in an attenuation phenomenon at the border between the 16 S rRNA and tRNA genes (57) and is reported to be important for controlling and coordinating mitochondrial gene transcription (42). Non-functional binding of mTERF to the mitochondrial DNA results in stress-related diseases such as mitochondrial encephalomyopathy, lactic acidosis, stroke (MELAS) encephalomyopathies (43) and special forms of diabetes and can cause cell death. Detailed molecular characterizations have shown that human mTERF is a multizipper protein and binds to mitochondrial DNA as a monomer (41). Although a number of homologues have now been identified in other systems such as sea urchin (mtDBP (40)) and D. melanogaster (DmTTF (39)), the binding and activation sites have not been conserved, possibly reflecting the differences in the gene order of mitochondrial genomes.

The identification of MOC1 in C. reinhardtii and the in silico identification of several potential mitochondrial, and even one possible chloroplast-targeted protein, with mTerf domains in A. thaliana suggest that this family of transcription factors might play an important role in the expression of plant organelle genomes as well. Analysis of the genome sequence for Chlamydomonas plus Southern hybridization analysis indicates that there is only one copy of Moc1. Previous studies had identified cDNA fragments of Moc1 in three independent Chlamydomonas expressed sequence tag libraries obtained from two different stress-treated cell cultures (www.chlamy.org, www.kazusa.or.jp/en/plant/chlamy/EST) so it is possible that MOC1 has a general role in regulating mitochondrial gene expression in response to stress conditions.

Electron microscopy performed with WT and stm6 cells grown under different light conditions did not reveal differences in the number of mitochondria per cell between WT and mutant under the same growth conditions (data not shown). Thus it is unlikely that MOC1 has a role in determining the number of mitochondria per cell, although effects on the copy number of the mitochondrial genome cannot be excluded at this stage.

MOC1 Is Required for Appropriate Expression of Mitochondrial Respiratory Complexes in the Light—It has been well established that the mitochondrion is important for optimal photosynthesis (10, 11, 58). Current ideas have emphasized the movement of metabolites between the two organelles so that perturbation of respiration would have a deleterious effect on chloroplast activity because of inappropriate redox conditions within the chloroplast. Our characterization of MOC1 builds on these ideas, and in addition, suggests that changes in the transcription of the mitochondrial genome in the light, mediated by MOC1, is important for acclimation to high light.

In higher plants, the onset of photosynthetic activity is known to lead to changes, some dramatic, in the relative content of the respiratory complexes in the mitochondrion. For instance, expression of the internal NADH dehydrogenase, NDA1, in potato is switched on in light, possibly to maintain high rates of glycine oxidation in the photorespiratory pathway and to prevent over-reduction of the matrix (16), and the external NADPH dehydrogenase NDB1 is reported to play an important role in regulating cellular redox homoostasis by transferring redox equivalents from cytosolic NADPH to the mitochondrial ubiquinone pool (5).

For WT C. reinhardtii, our work has also indicated marked differences in the composition of the respiratory chain in light-grown and dark-grown cultures. The reciprocal relationship in the expression of the type I and type II NAD(P)H dehydrogenases upon changing illumination conditions is quite similar to recent findings in potato and might reflect the need to increase rates of NAD(P)H oxidation but at the same time maintain an appropriate redox state for the Q pool (see Ref. 16, page 79, Fig. 8).

In stm6, there is a major perturbation in the relative content of the respiratory complexes as compared with the WT: most dramatically, less cytochrome oxidase and rotenone-insensitive NADPH dehydrogenase but higher levels of the alternative oxidases as compared with WT (Fig. 5). Clearly, the wide-ranging effect of the Moc1 mutation on the expression of both nuclear and mitochondrial-encoded proteins would indicate some indirect effects on gene expression.

A question that will be addressed in future work is the mechanism by which MOC1 controls transcription in the mitochondrion. The 15.8-kb mitochondrial genome of C. reinhardtii is transcribed bidirectionally to give two primary transcripts, which are then processed into smaller transcripts (46). Our RT-PCR and Northern data as well as activity measurements presented in Figs. 5 and 6 indicate that MOC1 is needed in the light to adjust the respiratory chain and to maintain in particular complex IV activity as a functional site for ATP production and the activity of the external NADPH-DH when chloroplast photophosphorylation is perturbed.

This is furthermore relevant given that rotenone-insensitive mitochondrial NAD(P)H dehydrogenases might be induced in the light (4, 5, 16) so that there would be a need for the synthesis of more cytochrome oxidase. Initial RT-PCR experiments have yet to provide evidence to support this model; thus it remains feasible that MOC1 binds at other sites (39), possibly even to RNA, and that its effects on transcription are indirect.

Chloroplast/Mitochondrial Interactions Are Perturbed in the Moc1 Mutant—We have shown here that MOC1 is needed for appropriate transcription of the mitochondrial genome in the light and that in its absence, there is a marked light-induced reduction of cytochrome oxidase activity and, as compared with WT, a more moderate reduction of NADH:Q oxidoreductase activity. Importantly, we have also identified for the first time the presence of the NDB1 type II NADPH dehydrogenase and have shown that the activities of the external rotenone-insensitive NADPH dehydrogenase are drastically reduced in stm6. These data strongly suggest that the mitochondria in stm6 are less able to function as electron sinks for the oxidation of reductant that is formed during photosynthesis. ATP levels are also dramatically reduced.

The influence of ATP levels for redox-controlled plastidic light adaptation mechanisms such as state transitions has already been described (9, 15, 59). It has been suggested that lower levels of ATP relieve the ATP-induced inhibition of phosphofructokinase activity in glycolysis, which then causes an increase in stromal NAD(P)H levels because of the activity of the downstream glyceraldehyde-3-phosphate dehydrogenase. Impaired oxidation of NAD(P)H by the mitochondrion in stm6 would also enhance NAD(P)H levels in the chloroplast. Levels of reduced PQ in the thylakoid could increase via reduction through a type-2 NAD(P)H:PQ oxidoreductase. Overall, ATP depletion would be predicted to result in a shift toward state 2, as observed recently for mitochondrial mutants lacking one or more of the respiratory complexes (13). In, contrast stm6 is shifted toward state 1 in the light, despite a reduced PQ pool. Recent evidence has pointed out that the activity of the LHC kinase is under two types of redox control in plants (60). Besides activation by the PQ pool, the kinase is switched off when levels of reduced thioredoxin in the stroma are high. Consequently, the inability of stm6 to undergo a transition to state 2 in the light could be explained by the presence of an over-reduced stroma. This suggestion would also nicely explain why stm6 can be driven into state 2 in the dark upon anaerobiosis; under these conditions, the stroma will be less reduced than in the light. It still remains possible, however, that state transitions might be regulated by other pathways that are dependent on the activity of the mitochondrion. It should be noted that under anaerobic conditions in standard growth light, stm6 is unable to switch from linear to cyclic electron transfer, most likely due to the block in state 1 (8, 50).

Current models would suggest that the over-reduced stroma in stm6 in the light could be due to a combination of low ATP levels, which promote glycolysis and the formation of NAD(P)H (9), and the reduced mitochondrial capacity to oxidize chloroplast stromal reductant (through the malate valve and the di-hydroxy-acetone-phosphate shuttle) (56). An inability to keep the stroma more oxidized would tend to enhance the production of reactive oxygen species in the chloroplast, as suggested from the measurement of lipid hydroperoxides (Table I). In addition, perturbations to mitorespiration in stm6 might also inhibit photorespiration (61), which again could cause sensitivity to photoinhibition. The reduced levels of ATP in stm6 would also impair the normal ATP-dependent repair processes leading to the observed sensitivity of growth to high light.

It should be noted that a reduced cytochrome oxidase activity and an increased AOX activity as shown for stm6 is similar to that reported by Ref. 62 for stress-induced nitric oxide signaling in plants. However, 2-(4-carboxylphenyl)-4,4,5,5-tetramethylimida-zoline-1-oxyl-3-oxide (cPTIO), which is known to prevent the killing effect of nitric oxide (63), did not prevent light-induced cell death in stm6, indicating that nitric oxide signaling is not involved.

In conclusion, we have identified a nuclear-encoded protein, MOC1, which is important for regulating mitochondrial respiration in response to light and possibly other stresses. As such, the identification of MOC1 is an important step in the search for factors that coordinate the redox poise between organelles.


   FOOTNOTES
 
The nucleotide sequence(s) reported in this paper has been submitted to the GenBankTM/EBI Data Bank with accession number(s) AF531421 [GenBank] .

* This work was supported by Grant FOR387 the Deutsche Forschungsgemeinschaft for financial support. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

To whom correspondence should be addressed. Tel.: +49-521-1065611; Fax: +49-521-1066410; E-mail: olaf.kruse{at}uni-bielefeld.de.

1 The abbreviations used are: Q, ubiquinone; PQ, plastoquinone; mTERF, mitochondrial transcription termination factor; PSI, photosystem I; PSII, photosystem II; WT, wild type; RT, reverse transcription; LHC, light-harvesting complex; COX, cytochrome c oxidase; AOX, alternative oxidase; MOPS, 4-morpholinepropanesulfonic acid; Cyt, cytochrome. Back


   ACKNOWLEDGMENTS
 
We thank F.-A. Wollman, Sabeeha Merchant, Saul Purton, Stefan Jansson, and J. M. Grienenberger for providing us with antibodies Cyt c, AOX1, COX90, LHCII, and NAD9.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Krömer, S., and Heldt, H. W. (1991) Plant Physiol. 95, 1270-1276[Abstract/Free Full Text]
  2. Flügge, U.-I. (1998) Curr. Opin. Plant Biol. 1, 201-206[CrossRef][Medline] [Order article via Infotrieve]
  3. Rasmusson, A. G., and Møller, I. M. (1991) Physiol. Plant. 83, 357-365[CrossRef]
  4. Michalecka, A. M., Svensson, A. S., Johansson, F. I., Agius, S. C., Johanson, U., Brennicke, A., Binder, S., and Rasmusson, A. G. (2003) Plant Physiol. 133, 642-652[Abstract/Free Full Text]
  5. Michalecka, A. M., Agius, S. C., Møller, I. M., and Rasmusson, A. G. (2004) Plant J. 37, 415-425[CrossRef][Medline] [Order article via Infotrieve]
  6. Millenaar, F. F., and Lambers, H. (2003) Plant Biol. (N. Y.) 5, 2-15
  7. Pfannschmidt, T., Nilsson, A., and Allen, J. (1999) Nature 397, 625-628[CrossRef]
  8. Finazzi, G., Rappaport, F., Furia, A., Fleischmann, M., Rochaix, J.-D., Zito, F., and Forti, G. (2002) EMBO Rep. 3, 280-285[CrossRef][Medline] [Order article via Infotrieve]
  9. Bulté, L., Gans, P., Rebéille, F., and Wollman, F.-A. (1990) Biochim. Biophys. Acta 1020, 72-80
  10. Dutilleul, C., Driscoll, S., Cornic, G., DePaepe, R., Foyer, C. H., and Noctor, G. (2003) Plant Physiol. 131, 264-275[Abstract/Free Full Text]
  11. Duby, F., and Matagne, R. F. (1999) Plant Cell 11, 115-125[Abstract/Free Full Text]
  12. Huertas, I. E., Colman, B., and Espie, G. S. (2002) Plant Physiol. 130, 284-291[Abstract/Free Full Text]
  13. Cardol, P., Gloire, G. Havaux, M., Remacle, C., Matagne, R., and Franck, F. (2003) Plant Physiol. 133, 2010-2020[Abstract/Free Full Text]
  14. Gu, D. M., and Newton, K. J. (1993) Plant Cell 5, 963-971[Abstract/Free Full Text]
  15. Gans, P., and Wollman, F.-A. (1995) Biochim. Biophys. Acta 1228, 51-57
  16. Svensson, A. S., and Rasmusson, A. G. (2001) Plant J. 28, 73-82[CrossRef][Medline] [Order article via Infotrieve]
  17. Murata, N. (1969) Biochim. Biophys. Acta 172, 242-251[Medline] [Order article via Infotrieve]
  18. Bonaventura, C., and Myers, J. (1969) Biochim. Biophys. Acta 189, 366-386[Medline] [Order article via Infotrieve]
  19. Allen, J. F., Bennett, J., Steinback, K. E., and Arntzen, C. J. (1981) Nature 291, 21-25[CrossRef]
  20. Horton, P., Allen, J. F., Black, M. T., and Bennett, J. (1981) FEBS Lett. 125, 193-196
  21. Fleischmann, M. M., Ravanel, S., Delosme, R., Olive, J., Zito, F., Wollman, F. A., and Rochaix, J.-D. (1999) J. Biol. Chem. 274, 30987-30994[Abstract/Free Full Text]
  22. Kruse, O., Nixon, P. J., Schmid, G. H., and Mullineaux, C. W. (1999) Photosynth. Res. 61, 43-51
  23. Depège, N., Bellafiore, S., and Rochaix, J.-D. (2003) Science 299, 1572-1575[Abstract/Free Full Text]
  24. Harris, E. (1989) The Chlamydomonas Sourcebook: A Comprehensive Guide to Biology and Laboratory Use, Academic Press, San Diego, CA
  25. Kindle, K. L., Schnell, R. A. Fernández, E., and Levebvre, P. A. (1989) J. Cell Biol. 109, 2589-2601[Abstract/Free Full Text]
  26. Purton, S., and Rochaix, J.-D. (1995) Eur. J. Phycol. 30, 141-148[CrossRef]
  27. Debuchy, R., Purton, S., and Rochaix, J.-D. (1989) EMBO J. 8, 2803-2809[Medline] [Order article via Infotrieve]
  28. Gumpel, N. J., and Purton, S. (1994) Trends Cell Biol. 4, 299-301[CrossRef][Medline] [Order article via Infotrieve]
  29. Strauss, C., Mussgnug, J. H., and Kruse, O. (2001) Photosynth. Res. 70, 311-320[CrossRef][Medline] [Order article via Infotrieve]
  30. Tam, L. W., and Lefebvre, P. A. (1993) Genetics 135, 375-384[Abstract]
  31. Nelson, J. A. E., Savereide, P. B., and Lefebvre, P. A. (1994) Mol. Cell. Biol. 14, 4011-4019[Abstract/Free Full Text]
  32. Lunde, C., Jensen, P. E., Haldrup, A., Knoetzel, J., and Scheller, H. V. (2000) Nature 408, 613-615[CrossRef][Medline] [Order article via Infotrieve]
  33. Zerges, W., and Rochaix, J.-D. (1998) J. Cell Biol. 140, 101-110[Abstract/Free Full Text]
  34. Nurani, G., Erikson, M., Knorpp, C., Glaser, E., and Franzen, L.-G. (1997) Plant Mol. Biol. 35, 973-980[CrossRef][Medline] [Order article via Infotrieve]
  35. Michel, K.-P., Krüger, F., Pühler, A., and Pistorius, E. K. (1999) Microbiology (N.Y.) 145, 1473-1484[Abstract/Free Full Text]
  36. Singh, D. P., and Varna, K. (1995) Photochem. Photobiol. 62, 314-319
  37. Day, A., Schirmer-Rahire, M., Kuchka, M. R., Mayfield, S. P., and Rochaix, J.-D. (1988) EMBO J. 7, 1917-1927[Medline] [Order article via Infotrieve]
  38. Daga, A., Micol, V., Hess, D., Aebersold, R., and Attardi, G. (1993) J. Biol. Chem. 268, 8124-8130
  39. Roberti, M., Polosa, P. L., Bruni, F., Musicco, C., Gadaleta, M. N., and Cantatore, P. (2003) Nucleic Acids Res. 31, 1597-1604[Abstract/Free Full Text]
  40. Loguericio, P. P., Roberti, M., Musicco, C., Gadaleta, M. N., Quagliariello, E., and Cantatore, P. (1999) Nucleic Acids Res. 27, 1890-1899[Abstract/Free Full Text]
  41. Fernandez-Silva, P., Martinez-Azorin, F., Mico l, V., and Attardi, G. (1997) EMBO J. 16, 1066-1079[CrossRef][Medline] [Order article via Infotrieve]
  42. Selwood, S. P., Chrzanowska-Lightowlers, Z. M. A., and Lightowlers, R. N. (2000) Biochem. Soc. Trans. 28, 154-159[Medline] [Order article via Infotrieve]
  43. Hess, J. F., Parasi, M. A., Bennett, J. L., and Clayton, D. A. (1991) Nature 351, 236-239[CrossRef][Medline] [Order article via Infotrieve]
  44. Lown, F. J., Watson, A. T., and Purton, S. (2001) Biochem. Soc. Trans. 29, 452-455[CrossRef][Medline] [Order article via Infotrieve]
  45. Baurain, D., Dinant, M., Coosemans, N., and Matagne, R. (2003) Plant Physiol. 131, 1418-1430[Abstract/Free Full Text]
  46. Gray, M. W., and Boer, P. H. (1988) Philos. Trans. R. Soc. Lond. B Biol. Sci. 319, 135-147[Abstract/Free Full Text]
  47. Dyer, T. A., and Leaver, C. J. (1981) in The Biochemistry of Plants (Stumpf, P. K., Conn, E., and Marcus, A., eds) p. 111, Academic Press New York
  48. Nijtmans, L. G. J., Taanman, J.–W., Muijsers, A. O., Speijer, D., and Van Den Bogert, C. (1998) Eur. J. Biochem. 254, 389-394[Medline] [Order article via Infotrieve]
  49. Wollman, F.-A. (2000) EMBO J. 20, 3623-3630[CrossRef]
  50. Finazzi, G., Furia, A., Barbagallo, R. P., and Forti, G. (1999) Biochim. Biophys. Acta 1413, 117-129[Medline] [Order article via Infotrieve]
  51. De Vitry, C., Finazzi, G., Baymann, F., and Kallas, T. (1999) Plant Cell 11, 2031-2044[Abstract/Free Full Text]
  52. Zito, F., Finazzi, G., Delosme, R., Nitschke, W., Picot, D., and Wollman, F.-A. (1999) EMBO J. 18, 2961-2969[CrossRef][Medline] [Order article via Infotrieve]
  53. Vener, A. V., van Kann, P. J., Rich, P. R., Andersson, B., and Ohad, I. (1997) Proc. Natl. Acad. Sci. U. S. A. 94, 1585-1590[Abstract/Free Full Text]
  54. Vener, A. V., Rokka, A., Fulgosi, H., Andersson, B., and Herrmann, R. G. (1999) Biochemistry 38, 14955-14965[CrossRef][Medline] [Order article via Infotrieve]
  55. Snyders, S., and Kohorn, B. D. (2001) J. Biol. Chem. 276, 32169-32176[Abstract/Free Full Text]
  56. Hoefnagel, M. H. N., Atkin, O. K., and Wiskich, J. T. (1998) Biochim. Biophys. Acta 1366, 235-255
  57. Kruse, B., Narasimhan, N., and Atardi, G. (1989) Cell 58, 391-397[CrossRef][Medline] [Order article via Infotrieve]
  58. Raghavendra, A. S., and Padmasree, K. (2003) Trends Plant Sci. 8, 546-553[CrossRef][Medline] [Order article via Infotrieve]
  59. Rebeille, F., and Gans, P. (1988) Plant Physiol. 88, 973-975[Abstract/Free Full Text]
  60. Rintamäki, E., Martinsuo, P., Pursiheimo, S., and Aro, E. M. (2000) Proc. Natl. Acad. Sci. U. S. A. 97, 11644-11649[Abstract/Free Full Text]
  61. Husic, D. W., and Tolbert, N. E. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 1555-1559[Abstract/Free Full Text]
  62. Millar, A. H., and Day, D. A. (1996) FEBS Lett. 398, 155-158[CrossRef][Medline] [Order article via Infotrieve]
  63. Zottini, M., Formentin, E., Scattolin, M., Carimi, F., Lo Schiavo, F., and Terzi, M. (2002) FEBS Lett. 515, 1-3[CrossRef][Medline] [Order article via Infotrieve]
  64. Lamattina, L., Gonzalez, D., Gualberto, J. M., and Grienenberger, J. M. (1993) Eur. J. Biochem. 217, 831-838[Medline] [Order article via Infotrieve]
  65. Stevens, D. R., Rochaix, J. D., and Purton, S. (1996) Mol. Gen. Genet. 251, 23-30[Medline] [Order article via Infotrieve]
  66. Hippler, M., Drepper, F., Haehnel, W., and Rochaix, J. D. (1998) Proc. Natl. Acad. Sci. U. S. A. 95, 7339-7344[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Complore Complore   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us   Add to Digg Digg   Add to Reddit Reddit   Add to Technorati Technorati    What's this?


This article has been cited by other articles:


Home page
 J Exp BotHome page
M. Timmins, S. R. Thomas-Hall, A. Darling, E. Zhang, B. Hankamer, U. C. Marx, and P. M. Schenk
Phylogenetic and molecular analysis of hydrogen-producing green algae
J. Exp. Bot., April 2, 2009; (2009) erp052v1.
[Abstract] [Full Text] [PDF]

Home page
 Plant CellHome page
J. H. Mussgnug, L. Wobbe, I. Elles, C. Claus, M. Hamilton, A. Fink, U. Kahmann, A. Kapazoglou, C. W. Mullineaux, M. Hippler, et al.
NAB1 Is an RNA Binding Protein Involved in the Light-Regulated Differential Expression of the Light-Harvesting Antenna of Chlamydomonas reinhardtii
PLANT CELL, December 1, 2005; 17(12): 3409 - 3421.
[Abstract] [Full Text] [PDF]

Home page
 J. Biol. Chem.Home page
O. Kruse, J. Rupprecht, K.-P. Bader, S. Thomas-Hall, P. M. Schenk, G. Finazzi, and B. Hankamer
Improved Photobiological H2 Production in Engineered Green Algal Cells
J. Biol. Chem., October 7, 2005; 280(40): 34170 - 34177.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
279/48/50366    most recent
M408477200v1
Right arrow Submit a Letter to Editor
Right arrow Alert me when this article is cited
Right arrow Alert me when eLetters are posted
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowRequest Permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Schönfeld, C.
Right arrow Articles by Kruse, O.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Schönfeld, C.
Right arrow Articles by Kruse, O.
Social Bookmarking
Add to CiteULike   Add to Complore   Add to Connotea   Add to Del.icio.us   Add to Digg   Add to Reddit   Add to Technorati  
What's this?

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   ASBMB Today 
Copyright © 2004 by the American Society for Biochemistry and Molecular Biology.
Advertisement
spacer
Advertisement
Advertisement