G. C. Ferreira and H. A. Dailey, Mouse protoporphyrinogen oxidase. Kinetic parameters and demonstration of inhibition by bilirubin, Biochem. J, vol.250, pp.597-603, 1988.

T. A. Dailey and H. A. Dailey, Human protoporphyrinoagen oxidase: Expression, purification, and characterization of the cloned enzyme, Protein Sci, vol.5, pp.98-105, 1996.

R. Van-lis, A. Atteia, L. A. Nogaj, and S. I. Beale, Subcellular localization and light-regulated expression of protoporphyrinogen IX oxidase and ferrochelatase in Chlamydomonas reinhardtii, Plant Physiol, vol.139, pp.1946-1958, 2005.
URL : https://hal.archives-ouvertes.fr/hal-00347332

N. Watanabe, Dual targeting of spinach protoporphyrinogen oxidase II to mitochondria and chloroplasts by alternative use of two in-frame initiation codons, J. Biol. Chem, vol.276, pp.20474-20481, 2001.

I. Lermontova, E. Kruse, H. P. Mock, and B. Grimm, Cloning and characterization of a plastidal and a mitochondrial isoform of tobacco protoporphyrinogen IX oxidase, Proc. Natl Acad. Sci. USA, vol.94, pp.8895-8900, 1997.

P. Bennoun, Evidence for a respiratory chain in the chloroplast, Proc. Natl Acad. Sci. USA, vol.79, pp.4352-4356, 1982.

C. Desplats, Characterization of Nda2, a plastoquinone-reducing type II NAD(P)H dehydrogenase in Chlamydomonas chloroplasts, J. Biol. Chem, vol.284, pp.4148-4157, 2009.

D. A. Berthold and P. Stenmark, Membrane-bound diiron carboxylate proteins, Annu. Rev. Plant. Biol, vol.54, pp.497-517, 2003.

L. Houille-vernes, F. Rappaport, F. Wollman, J. Alric, and X. Johnson, Plastid terminal oxidase 2 (PTOX2) is the major oxidase involved in chlororespiration in Chlamydomonas, Proc. Natl Acad. Sci. USA, vol.108, pp.20820-20825, 2011.

S. Buschlen, Y. Choquet, R. Kuras, and F. A. Wollman, Nucleotide sequences of the continuous and separated petA, petB and petD chloroplast genes in Chlamydomonas reinhardtii, FEBS Lett, vol.284, pp.257-262, 1991.

J. M. Jacobs and N. J. Jacobs, Porphyrin accumulation and export by isolated barley (Hordeum vulgare) plastids-effect of diphenyl ether herbicides, Plant Physiol, vol.101, pp.1181-1187, 1993.

H. J. Lee, M. V. Duke, and S. O. Duke, Cellular-localization of protoporphyrinogen-oxidizing activities of etiolated barley (Hordeum vulgare L.) leaves-relationship to mechanism of action of protoporphyrinogen oxidase-inhibiting herbicides, Plant Physiol, vol.102, pp.881-889, 1993.

H. Matsumoto, Y. Kashimoto, and E. Warabi, Basis for common chickweed (Stellaria media) tolerance to oxyfluorfen. Pestic, Biochem. Physiol, vol.64, pp.47-53, 1999.

S. O. Duke, J. M. Becerril, T. D. Sherman, and H. Matsumoto, Photosensitizing porphyrins as herbicides, Acs Symp. Ser, vol.449, pp.371-386, 1991.

R. Kuras and F. A. Wollman, The assembly of cytochrome b 6 /f complexes: an approach using genetic transformation of the green alga Chlamydomonas reinhardtii, EMBO J, vol.13, pp.1019-1027, 1994.

F. Jans, A type II NAD(P)H dehydrogenase mediates light-independent plastoquinone reduction in the chloroplast of Chlamydomonas, Proc. Natl Acad Sci, vol.105, pp.20546-20551, 2008.

A. Krieger-liszkay and A. W. Rutherford, Influence of herbicide binding on the redox potential of the quinone acceptor in photosystem-II. Relevance to photodamage and phytotoxicity, Biochemistry, vol.37, pp.17339-17344, 1998.

P. E. Jensen, L. C. Gibson, and C. N. Hunter, ATPase activity associated with the magnesium-protoporphyrin IX chelatase enzyme of Synechocystis PCC6803: evidence for ATP hydrolysis during Mg 2+ insertion, and the MgATP-dependent interaction of the ChlI and ChlD subunits, Biochem. J, vol.339, pp.127-134, 1999.

P. E. Jensen, J. D. Reid, and C. N. Hunter, Modification of cysteine residues in the ChlI and ChlH subunits of magnesium chelatase results in enzyme inactivation, Biochem. J, vol.352, pp.435-441, 2000.

A. Ikegami, The CHLI1 subunit of Arabidopsis thaliana magnesium chelatase is a target protein of the chloroplast thioredoxin, J. Biol. Chem, vol.282, pp.19282-19291, 2007.

A. Sawicki, S. Zhou, K. Kwiatkowski, M. Luo, and R. D. Willows, 1-N-histidine phosphorylation of ChlD by the AAA(+) ChlI2 stimulates magnesium chelatase activity in chlorophyll synthesis, Biochem. J, vol.474, pp.2095-2105, 2017.

K. Lohrig, B. Muller, J. Davydova, D. Leister, and D. A. Wolters, Phosphorylation site mapping of soluble proteins: bioinformatical filtering reveals potential plastidic phosphoproteins in Arabidopsis thaliana, Planta, vol.229, pp.1123-1134, 2009.

S. Reiland, Large-scale Arabidopsis phosphoproteome profiling reveals novel chloroplast kinase substrates and phosphorylation networks, Plant Physiol, vol.150, pp.889-903, 2009.

N. Sugiyama, Large-scale phosphorylation mapping reveals the extent of tyrosine phosphorylation in Arabidopsis, Mol. Syst. Biol, vol.4, p.193, 2008.

J. P. Woessner, Molecular and genetic analysis of the chloroplast ATPase of chlamydomonas, Plant Mol. Biol, vol.3, pp.177-190, 1984.

C. Lemaire, F. A. Wollman, and P. Bennoun, Restoration of phototrophic growth in a mutant of Chlamydomonas reinhardtii in which the chloroplast atpB gene of the ATP synthase has a deletion: an example of mitochondriadependent photosynthesis, Proc. Natl Acad Sci, vol.85, pp.1344-1348, 1988.

P. Brzezowski, Mg chelatase in chlorophyll synthesis and retrograde signaling in Chlamydomonas reinhardtii: CHLI2 cannot substitute for CHLI1, J. Exp. Bot, vol.67, pp.3925-3938, 2016.

J. Papenbrock, Impaired expression of the plastidic ferrochelatase by antisense RNA synthesis leads to a necrotic phenotype of transformed tobacco plants, Plant J, vol.28, pp.41-50, 2001.

I. Lermontova and B. Grimm, Reduced activity of plastid protoporphyrinogen oxidase causes attenuated photodynamic damage during high-light compared to low-light exposure, Plant J, vol.48, pp.499-510, 2006.

A. Fu, S. Park, and S. Rodermel, Sequences required for the activity of PTOX (IMMUTANS), a plastid terminal oxidase: in vitro and in planta mutagenesis of iron-binding sites and a conserved sequence that corresponds to Exon 8, J. Biol. Chem, vol.280, pp.42489-42496, 2005.

D. A. Berthold, M. E. Andersson, and P. Nordlund, New insight into the structure and function of the alternative oxidase, Biochim. Biophys. Acta, vol.1460, pp.241-254, 2000.

J. N. Siedow, A. L. Umbach, and A. L. Moore, The active site of the cyanideresistant oxidase from plant mitochondria contains a binuclear iron center, FEBS Lett, vol.362, pp.10-14, 1995.

A. L. Moore, A. L. Umbach, and J. N. Siedow, Structure-function relationships of the alternative oxidase of plant mitochondria: a model of the active site, J. Bioenerg. Biomembr, vol.27, pp.367-377, 1995.

D. A. Berthold, N. Voevodskaya, P. Stenmark, A. Graslund, and P. Nordlund, EPR studies of the mitochondrial alternative oxidase. Evidence for a diiron carboxylate center, J. Biol. Chem, vol.277, pp.43608-43614, 2002.

V. Pinta, M. Picaud, F. Reiss-husson, and C. Astier, Rubrivivax gelatinosus acsF (previously orf358) codes for a conserved, putative binuclear-ironcluster-containing protein involved in aerobic oxidative cyclization of Mgprotoporphyrin IX monomethylester, J. Bacteriol, vol.184, pp.746-753, 2002.

C. J. Walker, P. A. Castelfranco, and B. J. Whyte, Synthesis of divinyl protochlorophyllide. Enzymological properties of the Mg-protoporphyrin IX monomethyl ester oxidative cyclase system, Biochem. J, vol.276, pp.691-697, 1991.

J. Moseley, J. Quinn, M. Eriksson, and S. Merchant, The Crd1 gene encodes a putative di-iron enzyme required for photosystem I accumulation in copper deficiency and hypoxia in Chlamydomonas reinhardtii, EMBO J, vol.19, pp.2139-2151, 2000.

V. Steccanella, M. Hansson, and P. E. Jensen, Linking chlorophyll biosynthesis to a dynamic plastoquinone pool, Plant Physiol. Biochem, vol.97, pp.207-216, 2015.

M. Matringe, J. M. Camadro, P. Labbe, and R. Scalla, Protoporphyrinogen oxidase as a molecular target for diphenyl ether herbicides, Biochem. J, vol.260, pp.231-235, 1989.

S. Yamato, T. Ida, M. Katagiri, and H. Ohkawa, A tobacco soluble protoporphyrinogen-oxidizing enzyme similar to plant peroxidases in their amino acid sequences and immunochemical reactivity, Biosci. Biotechnol. Biochem, vol.59, pp.558-559, 1995.

J. M. Becerril and S. O. Duke, Protoporphyrin IX content correlates with activity of photobleaching herbicides, Plant Physiol, vol.90, pp.1175-1181, 1989.

G. Sandmann and P. Böger, Accumulation of protoporphyrin IX in the presence of peroxidizing herbicides, Z. Naturforsch, vol.43, pp.699-704, 1988.

J. J. Lee, H. Matsumoto, and K. Ishizuka, Light involvement in oxyfluorfeninduced protoporphyrin IX accumulation in several species of intact plants, Pestic. Biochem. Physiol, vol.44, pp.119-125, 1992.

G. A. Hunter, M. P. Sampson, and G. C. Ferreira, Metal ion substrate inhibition of ferrochelatase, J. Biol. Chem, vol.283, pp.23685-23691, 2008.

S. R. Norris, T. R. Barrette, and D. Dellapenna, Genetic dissection of carotenoid synthesis in Arabidopsis defines plastoquinone as an essential component of phytoene desaturation, Plant Cell, vol.7, pp.2139-2149, 1995.

P. Carol, Mutations in the Arabidopsis gene IMMUTANS cause a variegated phenotype by inactivating a chloroplast terminal oxidase associated with phytoene desaturation, Plant Cell, vol.11, pp.57-68, 1999.

F. S. Che, Molecular characterization and subcellular localization of protoporphyrinogen oxidase in spinach chloroplasts, Plant Physiol, vol.124, pp.59-70, 2000.

S. Narita, Molecular cloning and characterization of a cDNA that encodes protoporphyrinogen oxidase of Arabidopsis thaliana, Gene, vol.182, pp.169-175, 1996.

M. S. Manohara and B. C. Tripathy, Regulation of protoporphyrin IX biosynthesis by intraplastidic compartmentalization and adenosine triphosphate, Planta, vol.212, pp.52-59, 2000.

A. S. Richter, Phosphorylation of GENOMES UNCOUPLED 4 alters stimulation of Mg chelatase activity in angiosperms, Plant Physiol, vol.172, pp.1578-1595, 2016.

K. Mobius, Heme biosynthesis is coupled to electron transport chains for energy generation, Proc. Natl Acad. Sci. USA, vol.107, pp.10436-10441, 2010.
DOI : 10.1073/pnas.1000956107

URL : http://www.pnas.org/content/107/23/10436.full.pdf

N. Mochizuki, The cell biology of tetrapyrroles: a life and death struggle, Trends. Plant. Sci, vol.15, pp.488-498, 2010.

H. Schlicke, Function of tetrapyrroles, regulation of tetrapyrrole metabolism and methods for analyses of tetrapyrroles, Conjunction with International Conference on Natural Sciences, vol.14, pp.171-175, 2014.

B. Halliwell, Redox biology is a fundamental theme of aerobic life, Plant Physiol, vol.141, pp.312-322, 2006.

K. Apel and H. Hirt, Reactive oxygen species: metabolism, oxidative stress, and signal transduction, Annu. Rev. Plant. Biol, vol.55, pp.373-399, 2004.
DOI : 10.1146/annurev.arplant.55.031903.141701

Y. Balmer, Proteomics gives insight into the regulatory function of chloroplast thioredoxins, Proc. Natl Acad. Sci. USA, vol.100, pp.370-375, 2003.

C. Marchand, P. Le-marechal, Y. Meyer, and P. Decottignies, Comparative proteomic approaches for the isolation of proteins interacting with thioredoxin, Proteomics, vol.6, pp.6528-6537, 2006.
URL : https://hal.archives-ouvertes.fr/hal-00164223

A. S. Richter, Posttranslational influence of NADPH-dependent thioredoxin reductase C on enzymes in tetrapyrrole synthesis, Plant Physiol, vol.162, pp.63-73, 2013.

A. Stenbaek and P. E. Jensen, Redox regulation of chlorophyll biosynthesis, Phytochemistry, vol.71, pp.853-859, 2010.

S. D. Lemaire, L. Michelet, M. Zaffagnini, V. Massot, and E. Issakidis-bourguet, Thioredoxins in chloroplasts, Curr. Genet, vol.51, pp.343-365, 2007.
DOI : 10.1007/s00294-007-0128-z

G. Hanke and P. Mulo, Plant type ferredoxins and ferredoxin-dependent metabolism, Plant Cell Environ, vol.36, pp.1071-1084, 2013.

A. J. Serrato, J. M. Perez-ruiz, M. C. Spinola, and F. J. Cejudo, A novel NADPH thioredoxin reductase, localized in the chloroplast, which deficiency causes hypersensitivity to abiotic stress in Arabidopsis thaliana, J. Biol. Chem, vol.279, pp.43821-43827, 2004.

J. M. Perez-ruiz, Rice NTRC is a high-efficiency redox system for chloroplast protection against oxidative damage, Plant Cell, vol.18, pp.2356-2368, 2006.

A. Lepistö, Chloroplast NADPH-thioredoxin reductase interacts with photoperiodic development in Arabidopsis, Plant Physiol, vol.149, pp.1261-1276, 2009.

A. S. Richter and B. Grimm, Thiol-based redox control of enzymes involved in the tetrapyrrole biosynthesis pathway in plants, Front. Plant Sci, vol.4, p.371, 2013.

S. Hollingshead, Conserved chloroplast open-reading frame ycf54 is required for activity of the magnesium protoporphyrin monomethylester oxidative cyclase in Synechocystis PCC 6803, J. Biol. Chem, vol.287, pp.27823-27833, 2012.

C. A. Albus, LCAA, a novel factor required for magnesium protoporphyrin monomethylester cyclase accumulation and feedback control of aminolevulinic acid biosynthesis in tobacco, Plant Physiol, vol.160, pp.1923-1939, 2012.

J. Herbst, A. Girke, M. R. Hajirezaei, G. Hanke, and B. Grimm, Potential roles of YCF54 and ferredoxin-NADPH reductase for magnesium protoporphyrin monomethylester cyclase, Plant J, vol.94, pp.485-496, 2018.

T. A. Dailey and H. A. Dailey, Identification of an FAD superfamily containing protoporphyrinogen oxidases, monoamine oxidases, and phytoene desaturase. Expression and characterization of phytoene desaturase of Myxococcus xanthus, J. Biol. Chem, vol.273, pp.13658-13662, 1998.

A. G. Rasmusson, D. A. Geisler, and I. M. Moller, The multiplicity of dehydrogenases in the electron transport chain of plant mitochondria, Mitochondrion, vol.8, pp.47-60, 2008.

A. M. Melo, T. M. Bandeiras, and M. Teixeira, New insights into type II NAD (P)H:quinone oxidoreductases. Microbiol, Mol. Biol. Rev, vol.68, pp.603-616, 2004.

M. P. Mayer, P. Beyer, and H. Kleinig, Quinone compounds are able to replace molecular oxygen as terminal electron acceptor in phytoene desaturation in chromoplasts of Narcissus pseudonarcissus L, Eur. J. Biochem, vol.191, pp.359-363, 1990.

X. Johnson, R. Kuras, F. A. Wollman, and O. Vallon, Gene Hunting by Complementation of Pooled Chlamydomonas Mutants, pp.1093-1099, 2007.

N. Fischer and J. D. Rochaix, The flanking regions of PsaD drive efficient gene expression in the nucleus of the green alga Chlamydomonas reinhardtii, Mol. Genet. Genomics, vol.265, pp.888-894, 2001.

D. Mauzerall and S. Granick, The occurrence and determination of deltaamino-levulinic acid and porphobilinogen in urine, J. Biol. Chem, vol.219, pp.435-446, 1956.

J. D. Weinstein and S. I. Beale, Enzymatic conversion of glutamate to deltaaminolevulinate in soluble extracts of the unicellular green alga, Chlorella vulgaris, Arch. Biochem. Biophys, vol.237, pp.454-464, 1985.

X. Johnson, A new setup for in vivo fluorescence imaging of photosynthetic activity, Photosynth. Res, vol.102, pp.85-93, 2009.