Photocatalysis as the 'master switch' of photomorphogenesis in early plant development.
Journal
Nature plants
ISSN: 2055-0278
Titre abrégé: Nat Plants
Pays: England
ID NLM: 101651677
Informations de publication
Date de publication:
03 2021
03 2021
Historique:
received:
24
06
2020
accepted:
29
01
2021
pubmed:
10
3
2021
medline:
30
4
2021
entrez:
9
3
2021
Statut:
ppublish
Résumé
Enzymatic photocatalysis is seldom used in biology. Photocatalysis by light-dependent protochlorophyllide oxidoreductase (LPOR)-one of only a few natural light-dependent enzymes-is an exception, and is responsible for the conversion of protochlorophyllide to chlorophyllide in chlorophyll biosynthesis. Photocatalysis by LPOR not only regulates the biosynthesis of the most abundant pigment on Earth but it is also a 'master switch' in photomorphogenesis in early plant development. Following illumination, LPOR promotes chlorophyll production, plastid membranes are transformed and the photosynthetic apparatus is established. Given these remarkable, light-induced pigment and morphological changes, the LPOR-catalysed reaction has been extensively studied from catalytic, physiological and plant development perspectives, highlighting vital, and multiple, cellular roles of this intriguing enzyme. Here, we offer a perspective in which the link between LPOR photocatalysis and plant photomorphogenesis is explored. Notable breakthroughs in LPOR structural biology have uncovered the structural-mechanistic basis of photocatalysis. These studies have clarified how photon absorption by the pigment protochlorophyllide-bound in a ternary LPOR-protochlorophyllide-NADPH complex-triggers photocatalysis and a cascade of complex molecular and cellular events that lead to plant morphological changes. Photocatalysis is therefore the master switch responsible for early-stage plant development and ultimately life on Earth.
Identifiants
pubmed: 33686224
doi: 10.1038/s41477-021-00866-5
pii: 10.1038/s41477-021-00866-5
doi:
Substances chimiques
Plant Proteins
0
Oxidoreductases Acting on CH-CH Group Donors
EC 1.3.-
protochlorophyllide reductase
EC 1.3.1.33
Types de publication
Journal Article
Review
Langues
eng
Sous-ensembles de citation
IM
Pagination
268-276Commentaires et corrections
Type : ErratumIn
Références
Hörtensteiner, S. Update on the biochemistry of chlorophyll breakdown. Plant Mol. Biol. 82, 505–517 (2013).
pubmed: 22790503
doi: 10.1007/s11103-012-9940-z
Rudiger, W. Chlorophyll metabolism: from outer space down to the molecular level. Phytochemistry 46, 1151–1167 (1997).
doi: 10.1016/S0031-9422(97)80003-9
Masuda, T. & Fujita, Y. Regulation and evolution of chlorophyll metabolism. Photochem. Photobiol. Sci. 7, 1131–1149 (2008).
pubmed: 18846277
doi: 10.1039/b807210h
Stenbaek, A. & Jensen, P. E. Redox regulation of chlorophyll biosynthesis. Phytochemistry 71, 853–859 (2010).
pubmed: 20417532
doi: 10.1016/j.phytochem.2010.03.022
Mochizuki, N. et al. The cell biology of tetrapyrroles: a life and death struggle. Trends Plant Sci. 15, 488–498 (2010).
pubmed: 20598625
doi: 10.1016/j.tplants.2010.05.012
Czarnecki, O. & Grimm, B. Post-translational control of tetrapyrrole biosynthesis in plants, algae, and cyanobacteria. J. Exp. Bot. 63, 1675–1687 (2012).
pubmed: 22231500
doi: 10.1093/jxb/err437
Wang, P. & Grimm, B. Organization of chlorophyll biosynthesis and insertion of chlorophyll into the chlorophyll-binding proteins in chloroplasts. Photosynth. Res. 126, 189–202 (2015).
pubmed: 25957270
doi: 10.1007/s11120-015-0154-5
Kobayashi, K. & Masuda, T. Transcriptional regulation of tetrapyrrole biosynthesis in Arabidopsis thaliana. Front. Plant Sci. 7, 1811 (2016).
pubmed: 27990150
pmcid: 5130987
doi: 10.3389/fpls.2016.01811
Armarego-Marriott, T., Sandoval-Ibanez, O. & Kowalewska, L. Beyond the darkness: recent lessons from etiolation and de-etiolation studies. J. Exp. Bot. 71, 1215–1225 (2020).
pubmed: 31854450
doi: 10.1093/jxb/erz496
Chidgey, J. W., Jackson, P. J., Dickman, M. J. & Hunter, C. N. PufQ regulates porphyrin flux at the haem/bacteriochlorophyll branchpoint of tetrapyrrole biosynthesis via interactions with ferrochelatase. Mol. Microbiol. 106, 961–975 (2017).
pubmed: 29030914
pmcid: 5725709
doi: 10.1111/mmi.13861
Chen, G. E. et al. Complete enzyme set for chlorophyll biosynthesis in Escherichia coli. Sci. Adv. 4, eaaq1407 (2018).
pubmed: 29387799
pmcid: 5787379
doi: 10.1126/sciadv.aaq1407
Chen, X. et al. Structural insights into the catalytic mechanism of Synechocystis magnesium protoporphyrin IX O-methyltransferase (ChlM). J. Biol. Chem. 289, 25690–25698 (2014).
pubmed: 25077963
pmcid: 4162172
doi: 10.1074/jbc.M114.584920
Richter, A. S., Wang, P. & Grimm, B. Arabidopsis Mg-protoporphyrin IX methyltransferase activity and redox regulation depend on conserved cysteines. Plant Cell Physiol. 57, 519–527 (2016).
pubmed: 26759408
doi: 10.1093/pcp/pcw007
Hollingshead, S., Bliss, S., Baker, P. J. & Hunter, C. N. Conserved residues in Ycf54 are required for protochlorophyllide formation in Synechocystis sp. PCC 6803. Biochem. J. 474, 667–681 (2017).
pubmed: 28008132
doi: 10.1042/BCJ20161002
Chen, G. E., Canniffe, D. P. & Hunter, C. N. Three classes of oxygen-dependent cyclase involved in chlorophyll and bacteriochlorophyll biosynthesis. Proc. Natl Acad. Sci. USA 114, 6280–6285 (2017).
pubmed: 28559347
pmcid: 5474816
doi: 10.1073/pnas.1701687114
Heyes, D. J., Kruk, J. & Hunter, C. N. Spectroscopic and kinetic characterization of the light-dependent enzyme protochlorophyllide oxidoreductase (POR) using monovinyl and divinyl substrates. Biochem. J. 394, 243–248 (2006).
pubmed: 16274361
pmcid: 1386022
doi: 10.1042/BJ20051635
Hitchcock, A. et al. Biosynthesis of chlorophyll a in a purple bacterial phototroph and assembly into a plant chlorophyll–protein complex. ACS Synth. Biol. 5, 948–954 (2016).
pubmed: 27171912
doi: 10.1021/acssynbio.6b00069
Schoefs, B. & Franck, F. Protochlorophyllide reduction: mechanisms and evolution. Photochem. Photobiol. 78, 543–557 (2003).
pubmed: 14743862
doi: 10.1562/0031-8655(2003)078<0543:PRMAE>2.0.CO;2
Heyes, D. J. & Hunter, C. N. Making light work of enzyme catalysis: protochlorophyllide oxidoreductase. Trends Biochem. Sci. 30, 642–649 (2005).
pubmed: 16182531
doi: 10.1016/j.tibs.2005.09.001
Reinbothe, C. et al. Chlorophyll biosynthesis: spotlight on protochlorophyllide reduction. Trends Plant Sci. 15, 614–624 (2010).
pubmed: 20801074
doi: 10.1016/j.tplants.2010.07.002
Scrutton, N. S., Groot, M. L. & Heyes, D. J. Excited state dynamics and catalytic mechanism of the light-driven enzyme protochlorophyllide oxidoreductase. Phys. Chem. Chem. Phys. 14, 8818–8824 (2012).
pubmed: 22419074
doi: 10.1039/c2cp23789j
Gabruk, M. & Mysliwa-Kurdziel, B. Light-dependent protochlorophyllide oxidoreductase: phylogeny, regulation, and catalytic properties. Biochemistry 54, 5255–5262 (2015).
pubmed: 26230427
doi: 10.1021/acs.biochem.5b00704
Yang, J. & Cheng, Q. Origin and evolution of the light-dependent protochlorophyllide oxidase (LPOR) genes. Plant Biol. 6, 537–544 (2004).
pubmed: 15375724
doi: 10.1055/s-2004-821270
Armstrong, G. A. Greening in the dark: light independent chlorophyll biosynthesis from anoxygenic photosynthetic bacteria to gymnosperms. J. Photochem. Photobiol. B 43, 87–100 (1998).
doi: 10.1016/S1011-1344(98)00063-3
Vedalankar, P. & Tripathy, B. C. Evolution of light-independent protochlorophyllide oxidoreductase. Protoplasma 256, 293–312 (2019).
pubmed: 30291443
doi: 10.1007/s00709-018-1317-y
Sarma, R. et al. Crystal structure of the L protein of Rhodobacter sphaeroides light-independent protochlorophyllide reductase with MgADP bound: a homologue of the nitrogenase Fe protein. Biochemistry 47, 13004–13015 (2008).
pubmed: 19006326
doi: 10.1021/bi801058r
Bröcker, M. J. et al. Crystal structure of the nitrogenase-like dark operative protochlorophyllide oxidoreductase catalytic complex (ChlN/ChlB)
pubmed: 20558746
pmcid: 2930732
doi: 10.1074/jbc.M110.126698
Yamamoto, H., Kurumiya, S., Ohashi, R. & Fujita, Y. Oxygen sensitivity of a nitrogenase-like protochlorophyllide reductase from the cyanobacterium Leptolyngbya boryana. Plant Cell Physiol. 50, 1663–1673 (2009).
pubmed: 19643808
doi: 10.1093/pcp/pcp111
Kaschner, M. et al. Discovery of the first light dependent protochlorophyllide oxidoreductase in anoxygenic phototrophic bacteria. Mol. Microbiol. 93, 1066–1078 (2014).
pubmed: 25039543
doi: 10.1111/mmi.12719
Kasalicky, V. et al. Aerobic anoxygenic photosynthesis is commonly present within the genus Limnohabitans. Appl. Environ. Microbiol. 84, e02116–e02117 (2018).
pubmed: 29030444
doi: 10.1128/AEM.02116-17
Chernomor, O. et al. Complex evolution of light-dependent protochlorophyllide oxidoreductases in aerobic anoxygenic phototrophs: origin, phylogeny and function. Mol. Biol. Evol. https://doi.org/10.1093/molbev/msaa234 (2020).
Reinbothe, S., Quigley, F., Gray, J., Schemenewitz, A. & Reinbothe, C. Identification of plastid envelope proteins required for import of protochlorophyllide oxidoreductase A into the chloroplast of barley. Proc. Natl Acad. Sci. USA 101, 2197–2202 (2004).
pubmed: 14769934
pmcid: 380236
doi: 10.1073/pnas.0307284101
Reinbothe, S., Gray, J., Rustgi, S., von Wettstein, D. & Reinbothe, C. Cell growth defect factor 1 is crucial for the plastid import of NADPH:protochlorophyllide oxidoreductase A in Arabidopsis thaliana. Proc. Natl Acad. Sci. USA 112, 5838–5843 (2015).
pubmed: 25901327
pmcid: 4426407
doi: 10.1073/pnas.1506339112
Kauss, D., Bischof, S., Steiner, S., Apel, K. & Meskauskiene, R. FLU, a negative feedback regulator of tetrapyrrole biosynthesis, is physically linked to the final steps of the Mg
pubmed: 22212719
doi: 10.1016/j.febslet.2011.12.029
Zhang, W. et al. Characterization of ferredoxin-dependent biliverdin reductase PCYA1 reveals the dual function in retrograde bilin biosynthesis and interaction with light-dependent protochlorophyllide oxidoreductase LPOR in Chlamydomonas reinhardtii. Front. Plant Sci. 9, 676 (2018).
pubmed: 29875782
pmcid: 5974162
doi: 10.3389/fpls.2018.00676
Schottkowski, M., Ratke, J., Oster, U., Nowaczyk, M. & Nickelsen, J. Pitt, a novel tetratricopeptide repeat protein involved in light-dependent chlorophyll biosynthesis and thylakoid membrane biogenesis in Synechocystis sp. PCC 6803. Mol. Plant 2, 1289–1297 (2009).
pubmed: 19843617
doi: 10.1093/mp/ssp075
Hey, D. et al. LIL3, a light-harvesting complex protein, links terpenoid and tetrapyrrole biosynthesis in Arabidopsis thaliana. Plant Physiol. 174, 1037–1050 (2017).
pubmed: 28432258
pmcid: 5462053
doi: 10.1104/pp.17.00505
Smith, J. H. C. & Kupke, D. W. Some properties of extracted protochlorophyll holochrome. Nature 178, 751–752 (1956).
doi: 10.1038/178751a0
Solymosi, K. & Schoefs, B. Etioplast and etiochloroplast formation under natural conditions: the dark side of chlorophyll biosynthesis in angiosperms. Photosynth. Res. 105, 143–166 (2010).
pubmed: 20582474
doi: 10.1007/s11120-010-9568-2
Josse, E.-M. & Halliday, K. J. Skotomorphogenesis: the dark side of light signalling. Curr. Biol. 18, R1144–R1146 (2008).
pubmed: 19108774
doi: 10.1016/j.cub.2008.10.034
Adam, Z., Charuvi, D., Tsabari, O., Knopf, R. R. & Reich, Z. Biogenesis of thylakoid networks in angiosperms: knowns and unknowns. Plant Mol. Biol. 76, 221–234 (2011).
pubmed: 20859754
doi: 10.1007/s11103-010-9693-5
Grzyb, J. M., Solymosi, K., Strzalka, K. & Mysliwa-Kurdziel, B. Visualization and characterization of prolamellar bodies with atomic force microscopy. J. Plant Physiol. 170, 1217–1227 (2013).
pubmed: 23777838
doi: 10.1016/j.jplph.2013.04.017
Kowalewska, L., Mazur, R., Suski, S., Garstka, M. & Mostowska, A. Three-dimensional visualization of the tubular-lamellar transformation of the internal plastid membrane network during runner bean chloroplast biogenesis. Plant Cell 28, 875–891 (2016).
pubmed: 27002023
pmcid: 4863387
doi: 10.1105/tpc.15.01053
Gabruk, M., Mysliwa-Kurdziel, B. & Kruk, J. MGDG, PG and SQDG regulate the activity of light-dependent protochlorophyllide oxidoreductase. Biochem. J. 474, 1307–1320 (2017).
pubmed: 28188256
doi: 10.1042/BCJ20170047
Fujii, S., Kobayashi, K., Nagata, N., Masuda, T. & Wada, H. Monogalactosyldiacylglycerol facilitates synthesis of photoactive protochlorophyllide in etioplasts. Plant Physiol. 174, 2183–2198 (2017).
pubmed: 28655777
pmcid: 5543945
doi: 10.1104/pp.17.00304
Fujii, S., Kobayashi, K., Nagata, N., Masuda, T. & Wada, H. Digalactosyldiacylglycerol is essential for organization of the membrane structure in etioplasts. Plant Physiol. 177, 1487–1497 (2018).
pubmed: 29946018
pmcid: 6084665
doi: 10.1104/pp.18.00227
Fujii, S., Nagata, N., Masuda, T., Wada, H. & Kobayashi, K. Galactolipids are essential for internal membrane transformation during etioplast-to-chloroplast differentiation. Plant Cell Physiol. 60, 1224–1238 (2019).
pubmed: 30892620
pmcid: 6553665
doi: 10.1093/pcp/pcz041
Aronsson, H., Sundqvist, C. & Dahlin, C. POR hits the road: import and assembly of a plastid protein. Plant Mol. Biol. 51, 1–7 (2003).
pubmed: 12602886
doi: 10.1023/A:1020795415631
Selstam, E., Schelin, J., Brain, T. & Williams, W. P. The effects of low pH on the properties of protochlorophyllide oxidoreductase and the organization of prolamellar bodies of maize (Zea mays). Eur. J. Biochem. 269, 2336–2346 (2002).
pubmed: 11985616
doi: 10.1046/j.1432-1033.2002.02897.x
Selstam, E., Brain, A. P. R. & Williams, W. P. The relationship between different spectral forms of the protochlorophyllide oxidoreductase complex and the structural organisation of prolamellar bodies isolated from Zea mays. Photosynth. Res. 108, 47–59 (2011).
pubmed: 21505944
doi: 10.1007/s11120-011-9653-1
Masuda, S. et al. Prolamellar bodies formed by cyanobacterial protochlorophyllide oxidoreductase in Arabidopsis. Plant J. 58, 952–960 (2009).
pubmed: 19222806
doi: 10.1111/j.1365-313X.2009.03833.x
Yamamoto, H., Kojima-Ando, H., Ohki, K. & Fujita, Y. Formation of prolamellar-body-like ultrastructures in etiolated cyanobacterial cells overexpressing light-dependent protochlorophyllide oxidoreductase in Leptolyngbya boryana. J. Gen. Appl. Microbiol. 66, 129–139 (2020).
pubmed: 32238622
doi: 10.2323/jgam.2020.01.009
Pribil, M., Labs, M. & Leister, D. Structure and dynamics of thylakoids in land plants. J. Exp. Bot. 65, 1955–1972 (2014).
pubmed: 24622954
doi: 10.1093/jxb/eru090
Schoefs, B. & Franck, F. The photoenzymatic cycle of NADPH: protochlorophyllide oxidoreductase in primary bean leaves (Phaseolus vulgaris) during the first days of photoperiodic growth. Photosyn. Res. 96, 15–26 (2008).
doi: 10.1007/s11120-007-9274-x
Rumak, I. et al. 3-D modelling of chloroplast structure under (Mg
pubmed: 20621057
doi: 10.1016/j.bbabio.2010.07.001
Holtorf, H., Reinbothe, S., Reinbothe, C. & Bereza, B. Two routes of chlorophyllide synthesis that are differentially regulated by light in barley. Proc. Natl Acad. Sci. USA 92, 3254–3258 (1995).
pubmed: 7724548
pmcid: 42144
doi: 10.1073/pnas.92.8.3254
Su, Q., Frick, G., Armstrong, G. & Apel, K. POR C of Arabidopsis thaliana: a third light- and NADPH-dependent protochlorophyllide oxidoreductase that is differentially regulated by light. Plant Mol. Biol. 47, 805–813 (2001).
pubmed: 11785941
doi: 10.1023/A:1013699721301
Masuda, T. & Takamiya, K. Novel insights into the enzymology, regulation and physiological functions of light-dependent protochlorophyllide oxidoreductase in angiosperms. Photosyn. Res. 81, 1–29 (2004).
doi: 10.1023/B:PRES.0000028392.80354.7c
Gabruk, M. & Mysliwa-Kurdziel, B. The origin, evolution and diversification of multiple isoforms of light-dependent protochlorophyllide oxidoreductase (LPOR): focus on angiosperms. Biochem. J. 477, 2221–2236 (2020).
pubmed: 32568402
doi: 10.1042/BCJ20200323
Buhr, F. et al. Photoprotective role of NADPH:protochlorophyllide oxidoreductase A. Proc. Natl Acad. Sci. USA 105, 12629–12634 (2008).
pubmed: 18723681
pmcid: 2527962
doi: 10.1073/pnas.0803950105
Armstrong, G. A., Apel, K. & Rüdiger, W. Does a light-harvesting protochlorophyllide a/b-binding protein complex exist? Trends Plant Sci. 5, 40–44 (2000).
pubmed: 10637661
doi: 10.1016/S1360-1385(99)01513-7
Yuan, M. et al. Assembly of NADPH:protochlorophyllide oxidoreductase complex is needed for effective greening of barley seedlings. J. Plant Physiol. 169, 1311–1316 (2012).
pubmed: 22704664
doi: 10.1016/j.jplph.2012.05.010
Pattanayak, G. K. & Tripathy, B. C. Overexpression of protochlorophyllide oxidoreductase C regulates oxidative stress in Arabidopsis. PLoS ONE 6, e26532 (2011).
pubmed: 22031838
pmcid: 3198771
doi: 10.1371/journal.pone.0026532
Zhan, W. et al. An allele of ZmPORB2 encoding a protochlorophyllide oxidoreductase promotes tocopherol accumulation in both leaves and kernels of maize. Plant J. 100, 114–127 (2019).
pubmed: 31169939
doi: 10.1111/tpj.14432
Boddi, B., Lindsten, A., Ryberg, M. & Sundqvist, C. On the aggregational states of protochlorophyllide and its protein complexes in wheat etioplasts. Physiol. Plant. 76, 135–143 (1989).
doi: 10.1111/j.1399-3054.1989.tb05622.x
Schneidewind, J. et al. Consensus model of a cyanobacterial light-dependent protochlorophyllide oxidoreductase in its pigment-free apo-form and photoactive ternary complex. Commun. Biol. 2, 351 (2019).
pubmed: 31583285
pmcid: 6761149
doi: 10.1038/s42003-019-0590-4
Zhang, S. et al. Structural basis for enzymatic photocatalysis in chlorophyll biosynthesis. Nature 574, 722–725 (2019).
pubmed: 31645759
doi: 10.1038/s41586-019-1685-2
Gabruk, M. et al. Photoactive protochlorophyllide–enzyme complexes reconstituted with PORA, PORB and PORC proteins of A. thaliana: fluorescence and catalytic properties. PLoS ONE 10, e0116990 (2015).
pubmed: 25659137
pmcid: 4319759
doi: 10.1371/journal.pone.0116990
Gabruk, M. et al. Insight into the oligomeric structure of PORA from A. thaliana. Biochim. Biophys. Acta Proteins Proteom. 1864, 1757–1764 (2016).
doi: 10.1016/j.bbapap.2016.09.015
Zhang, S. et al. Dual role of the active site ‘lid’ regions of protochlorophyllide oxidoreductase in photocatalysis and plant development. FEBS J. 288, 175–189 (2021).
pubmed: 32866986
doi: 10.1111/febs.15542
Nguyen, H. C., Melo, A. A., Kruk, J., Frost, A. & Gabruk, M. Photocatalytic LPOR forms helical lattices that shape membranes for chlorophyll synthesis. Nat. Plants (in the press).
Aronsson, H., Sundqvist, C. & Dahlin, C. POR-import and membrane association of a key element in chloroplast development. Physiol. Plant. 118, 1–9 (2003).
pubmed: 12702007
doi: 10.1034/j.1399-3054.2003.00088.x
Heyes, D. J., Menon, B. R. K., Sakuma, M. & Scrutton, N. S. Conformational events during ternary enzyme–substrate complex formation are rate limiting in the catalytic cycle of the light-driven enzyme protochlorophyllide oxidoreductase. Biochemistry 47, 10991–10998 (2008).
pubmed: 18798649
doi: 10.1021/bi801521c
Heyes, D. J., Hardman, S. J. O., Mansell, D., Gardiner, J. M. & Scrutton, N. S. Mechanistic reappraisal of early stage photochemistry in the light-driven enzyme protochlorophyllide oxidoreductase. PLoS ONE 7, e45642 (2012).
pubmed: 23049830
pmcid: 3458894
doi: 10.1371/journal.pone.0045642
Kolossov, V. L., Kopetz, K. J. & Rebeiz, C. A. Chloroplast biogenesis 87: evidence of resonance excitation energy transfer between tetrapyrrole intermediates of the chlorophyll biosynthetic pathway and chlorophyll a. Photochem. Photobiol. 78, 184–196 (2003).
pubmed: 12945588
doi: 10.1562/0031-8655(2003)078<0184:CBEORE>2.0.CO;2
Sytina, O. A. et al. Conformational changes in an ultrafast light-driven enzyme determine catalytic activity. Nature 456, 1001–1004 (2008).
pubmed: 19092933
doi: 10.1038/nature07354
Stadler, A. M. et al. Ternary complex formation and photoactivation of a photoenzyme results in altered protein dynamics. J. Phys. Chem. B 123, 7372–7384 (2019).
pubmed: 31380636
doi: 10.1021/acs.jpcb.9b06608
Menon, B. R., Hardman, S. J., Scrutton, N. S. & Heyes, D. J. Multiple active site residues are important for photochemical efficiency in the light-activated enzyme protochlorophyllide oxidoreductase (POR). J. Photochem. Photobiol. B 161, 236–243 (2016).
pubmed: 27285815
pmcid: 4970445
doi: 10.1016/j.jphotobiol.2016.05.029
Gholami, S. et al. Theoretical model of the protochlorophyllide oxidoreductase from a hierarchy of protocols. J. Phys. Chem. B 122, 7668–7681 (2018).
pubmed: 29996651
doi: 10.1021/acs.jpcb.8b04231
Dong, C. S. et al. Crystal structures of cyanobacterial light-dependent protochlorophyllide oxidoreductase. Proc. Natl Acad. Sci. USA 117, 8455–8461 (2020).
pubmed: 32234783
pmcid: 7165480
doi: 10.1073/pnas.1920244117
Ho, M. Y., Shen, G., Canniffe, D. P., Zhao, C. & Bryant, D. A. Light-dependent chlorophyll f synthase is a highly divergent paralog of PsbA of photosystem II. Science 353, aaf9178 (2016).
pubmed: 27386923
doi: 10.1126/science.aaf9178
Aubert, C., Vos, M. H., Mathis, P., Eker, A. P. & Brettel, K. Intraprotein radical transfer during photoactivation of DNA photolyase. Nature 405, 586–590 (2000).
pubmed: 10850720
doi: 10.1038/35014644
Sorigue, D. et al. An algal photoenzyme converts fatty acids to hydrocarbons. Science 357, 903–907 (2017).
pubmed: 28860382
doi: 10.1126/science.aan6349
Schmermund, L. et al. Extending the library of light-dependent protochlorophyllide oxidoreductases and their solvent tolerance, stability in light and cofactor flexibility. ChemCatChem 12, 4044–4051 (2020).
doi: 10.1002/cctc.202000561
Dietzek, B. et al. Excited-state processes in protochlorophyllide a – a femtosecond time-resolved absorption study. Chem. Phys. Lett. 397, 110–115 (2004).
doi: 10.1016/j.cplett.2004.08.075
Dietzek, B., Kiefer, W., Hermann, G., Popp, J. & Schmitt, M. Solvent effects on the excited-state processes of protochlorophyllide: a femtosecond time-resolved absorption study. J. Phys. Chem. B 110, 4399–4406 (2006).
pubmed: 16509741
doi: 10.1021/jp0556456
Dietzek, B. et al. The excited-state chemistry of protochlorophyllide a: a time-resolved fluorescence study. ChemPhysChem 7, 1727–1733 (2006).
pubmed: 16841352
doi: 10.1002/cphc.200600172
Dietzek, B. et al. Protochlorophyllide a: a comprehensive photophysical picture. ChemPhysChem 10, 144–150 (2009).
pubmed: 18855967
doi: 10.1002/cphc.200800536
Dietzek, B. et al. Dynamics of charge separation in the excited-state chemistry of protochlorophyllide. Chem. Phys. Lett. 492, 157–163 (2010).
doi: 10.1016/j.cplett.2010.04.027
Sytina, O. A. et al. Protochlorophyllide excited-state dynamics in organic solvents studied by time-resolved visible and mid-infrared spectroscopy. J. Phys. Chem. B 114, 4335–4344 (2010).
pubmed: 20205376
doi: 10.1021/jp9089326
Heyes, D. J. et al. Excited-state properties of protochlorophyllide analogues and implications for light-driven synthesis of chlorophyll. J. Phys. Chem. B 121, 1312–1320 (2017).
pubmed: 28117585
doi: 10.1021/acs.jpcb.7b00528
Brandariz-de-Pedro, G. et al. Direct evidence of an excited-state triplet species upon photoactivation of the chlorophyll precursor protochlorophyllide. J. Phys. Chem. Lett. 8, 1219–81223 (2017).
pubmed: 28244763
doi: 10.1021/acs.jpclett.7b00200
Zhao, G. J. & Han, K. L. Site-specific solvation of the photoexcited protochlorophyllide a in methanol: formation of the hydrogen-bonded intermediate state induced by hydrogen-bond strengthening. Biophys. J. 94, 38–46 (2008).
pubmed: 17827245
doi: 10.1529/biophysj.107.113738
Heyes, D. J. et al. Excited state charge separation in the photochemical mechanism of the light-driven enzyme protochlorophyllide oxidoreductase. Angew. Chem. Int. Ed. 54, 1512–1515 (2015).
doi: 10.1002/anie.201409881
Heyes, D. J., Sakuma, M., de Visser, S. & Scrutton, N. S. Nuclear quantum tunneling in the light-activated enzyme protochlorophyllide oxidoreductase. J. Biol. Chem. 284, 3762–3767 (2009).
pubmed: 19073603
doi: 10.1074/jbc.M808548200
Heyes, D. J., Sakuma, M. & Scrutton, N. S. Solvent-slaved protein motions accompany proton but not hydride tunneling in light-activated protochlorophyllide oxidoreductase. Angew. Chem. Int. Ed. 48, 3850–3853 (2009).
doi: 10.1002/anie.200900086
Menon, B. R. K., Waltho, J. P., Scrutton, N. S. & Heyes, D. J. Cryogenic and laser photoexcitation studies identify multiple roles for active site residues in the light-driven enzyme protochlorophyllide oxidoreductase. J. Biol. Chem. 284, 18160–18166 (2009).
pubmed: 19439417
pmcid: 2709359
doi: 10.1074/jbc.M109.020719
Menon, B. R., Davison, P. A., Hunter, C. N., Scrutton, N. S. & Heyes, D. J. Mutagenesis alters the catalytic mechanism of the light-driven enzyme protochlorophyllide oxidoreductase. J. Biol. Chem. 285, 2113–2119 (2010).
pubmed: 19850924
doi: 10.1074/jbc.M109.071522
Heyes, D. J., Levy, C., Sakuma, M., Robertson, D. L. & Scrutton, N. S. A twin-track approach has optimized proton and hydride transfer by dynamically coupled tunneling during the evolution of protochlorophyllide oxidoreductase. J. Biol. Chem. 286, 11849–11854 (2011).
pubmed: 21317291
pmcid: 3064235
doi: 10.1074/jbc.M111.219626
Hoeven, R., Hardman, S. J. O., Heyes, D. J. & Scrutton, N. S. Cross-species analysis of protein dynamics associated with hydride and proton transfer in the catalytic cycle of the light-driven enzyme protochlorophyllide oxidoreductase. Biochemistry 55, 903–913 (2016).
pubmed: 26807652
doi: 10.1021/acs.biochem.5b01355
Archipowa, N., Kutta, R. J., Heyes, D. J. & Scrutton, N. S. Stepwise hydride transfer in a biological system: insights into the reaction mechanism of the light-dependent protochlorophyllide oxidoreductase. Angew. Chem. Int. Ed. 57, 2682–2686 (2018).
doi: 10.1002/anie.201712729
Heyes, D. J., Ruban, A. V., Wilks, H. M. & Hunter, C. N. Enzymology below 200 K: the kinetics and thermodynamics of the photochemistry catalyzed by protochlorophyllide oxidoreductase. Proc. Natl Acad. Sci. USA 99, 11145–11150 (2002).
pubmed: 12177453
pmcid: 123224
doi: 10.1073/pnas.182274199
Heyes, D. J., Ruban, A. V. & Hunter, C. N. Protochlorophyllide oxidoreductase: “dark” reactions of a light-driven enzyme. Biochemistry 42, 523–528 (2003).
pubmed: 12525180
doi: 10.1021/bi0268448
Heyes, D. J. & Hunter, C. N. Identification and characterization of the product release steps within the catalytic cycle of protochlorophyllide oxidoreductase. Biochemistry 43, 8265–8271 (2004).
pubmed: 15209523
doi: 10.1021/bi049576h
Heyes, D. J. et al. The first catalytic step of the light driven enzyme protochlorophyllide oxidoreductase proceeds via a charge transfer complex. J. Biol. Chem. 281, 26847–26853 (2006).
pubmed: 16867988
doi: 10.1074/jbc.M602943200
Durin, G. et al. Simultaneous measurements of solvent dynamics and functional kinetics in a light-activated enzyme. Biophys. J. 96, 1902–1910 (2009).
pubmed: 19254549
pmcid: 2717306
doi: 10.1016/j.bpj.2008.10.065
Garrone, A., Archipowa, N., Zipfel, P. F., Hermann, G. & Dietzek, B. Plant protochlorophyllide oxidoreductases A and B: catalytic efficiency and initial reaction steps. J. Biol. Chem. 290, 28530–28539 (2015).
pubmed: 26408201
pmcid: 4653708
doi: 10.1074/jbc.M115.663161
Raskin, V. I. & Schwartz, A. The charge-transfer complex between protochlorophyllide and NADPH: an intermediate in protochlorophyllide photoreduction. Photosyn. Res. 74, 181–186 (2002).
doi: 10.1023/A:1020955526882
Heyes, D. J., Sakuma, M. & Scrutton, N. S. Laser excitation studies of the product release steps in the catalytic cycle of the light-driven enzyme, protochlorophyllide oxidoreductase. J. Biol. Chem. 282, 32015–32020 (2007).
pubmed: 17848549
doi: 10.1074/jbc.M706098200
Levantino, M., Yorke, B. A., Monteiro, D. C., Cammarata, M. & Pearson, A. R. Using synchrotrons and XFELs for time-resolved X-ray crystallography and solution scattering experiments on biomolecules. Curr. Opin. Struct. Biol. 35, 41–48 (2015).
pubmed: 26342489
doi: 10.1016/j.sbi.2015.07.017