Conformational control over proton-coupled electron transfer in metalloenzymes.


Journal

Nature reviews. Chemistry
ISSN: 2397-3358
Titre abrégé: Nat Rev Chem
Pays: England
ID NLM: 101703631

Informations de publication

Date de publication:
02 Sep 2024
Historique:
accepted: 29 07 2024
medline: 3 9 2024
pubmed: 3 9 2024
entrez: 2 9 2024
Statut: aheadofprint

Résumé

From the reduction of dinitrogen to the oxidation of water, the chemical transformations catalysed by metalloenzymes underlie global geochemical and biochemical cycles. These reactions represent some of the most kinetically and thermodynamically challenging processes known and require the complex choreography of the fundamental building blocks of nature, electrons and protons, to be carried out with utmost precision and accuracy. The rate-determining step of catalysis in many metalloenzymes consists of a protein structural rearrangement, suggesting that nature has evolved to leverage macroscopic changes in protein molecular structure to control subatomic changes in metallocofactor electronic structure. The proton-coupled electron transfer mechanisms operative in nitrogenase, photosystem II and ribonucleotide reductase exemplify this interplay between molecular and electronic structural control. We present the culmination of decades of study on each of these systems and clarify what is known regarding the interplay between structural changes and functional outcomes in these metalloenzyme linchpins.

Identifiants

pubmed: 39223400
doi: 10.1038/s41570-024-00646-7
pii: 10.1038/s41570-024-00646-7
doi:

Types de publication

Journal Article Review

Langues

eng

Sous-ensembles de citation

IM

Informations de copyright

© 2024. Springer Nature Limited.

Références

Mitchell, P. Coupling of phosphorylation to electron and hydrogen transfer by a chemi-osmotic type of mechanism. Nature 191, 144–148 (1961).
pubmed: 13771349 doi: 10.1038/191144a0
Warren, J. J., Tronic, T. A. & Mayer, J. M. Thermochemistry of proton-coupled electron transfer reagents and its implications. Chem. Rev. 110, 6961–7001 (2010).
pubmed: 20925411 pmcid: 3006073 doi: 10.1021/cr100085k
Marcus, R. A. & Sutin, N. Electron transfers in chemistry and biology. BBA Rev. Bioenerg. 811, 265–322 (1985).
Reece, S. Y. & Nocera, D. G. Proton-coupled electron transfer in biology: results from synergistic studies in natural and model systems. Annu. Rev. Biochem. 78, 673–699 (2009).
pubmed: 19344235 pmcid: 4625787 doi: 10.1146/annurev.biochem.78.080207.092132
Soudackov, A. & Hammes-Schiffer, S. Derivation of rate expressions for nonadiabatic proton-coupled electron transfer reactions in solution. J. Chem. Phys. 113, 2385–2396 (2000).
doi: 10.1063/1.482053
Jencks, B. W. P. Binding energy, specificity, and enzymatic catalysis: the Circe effect. Adv. Enzym. 43, 220–410 (1975). The question of how enzymes accelerate reaction rates remains debated. This seminal work outlines key contributions to catalysis and provides experimental insights regarding the relative impacts of these different contributions. The term ‘Circe effect’ is coined to describe the way that enzymes use conformational changes to regulate activity.
Lumry, R. & Eyring, H. Conformation changes of proteins. J. Phys. Chem. 58, 110–120 (1954).
doi: 10.1021/j150512a005
Pratt, J. M. Metalloenzymes as molecular switches: the role of conformation changes in controlling activity. J. Inorg. Biochem. 28, 145–153 (1986).
pubmed: 3027253 doi: 10.1016/0162-0134(86)80078-2
Hammes-Schiffer, S. & Stuchebrukhov, A. A. Theory of coupled electron and proton transfer reactions. Chem. Rev. 110, 6939–6960 (2010).
pubmed: 21049940 pmcid: 3005854 doi: 10.1021/cr1001436
Migliore, A., Polizzi, N. F., Therien, M. J. & Beratan, D. N. Biochemistry and theory of proton-coupled electron transfer. Chem. Rev. 114, 3381–3465 (2014).
pubmed: 24684625 pmcid: 4317057 doi: 10.1021/cr4006654
Rutledge, H. L. & Tezcan, F. A. Electron transfer in nitrogenase. Chem. Rev. 120, 5158–5193 (2020).
pubmed: 31999100 pmcid: 7466952 doi: 10.1021/acs.chemrev.9b00663
Burgess, B. K. & Lowe, D. J. Mechanism of molybdenum nitrogenase. Chem. Rev. 96, 2983–3012 (1996).
pubmed: 11848849 doi: 10.1021/cr950055x
Hageman, R. V. & Burris, R. H. Nitrogenase and nitrogenase reductase associate and dissociate with each catalytic cycle. Proc. Natl Acad. Sci. USA 75, 2699–2702 (1978).
pubmed: 275837 pmcid: 392630 doi: 10.1073/pnas.75.6.2699
Garner, C. D. & Bristow, S. in Molybdenum Enzymes Vol. 7 (ed. Spiro T. G.) (Wiley, 1985).
Lowe, D. J. & Thorneley, R. N. The mechanism of Klebsiella pneumoniae nitrogenase action. Pre-steady-state kinetics of H
pubmed: 6395861 pmcid: 1144524 doi: 10.1042/bj2240877
Lowe, D. J. & Thorneley, R. N. F. The mechanism of Klebsiella pneumoniae nitrogenase action. The determination of rate constants required for the simulation of the kinetics of N
pubmed: 6395863 pmcid: 1144526 doi: 10.1042/bj2240895
Tezcan, F. A. et al. Nitrogenase complexes: multiple docking sites for a nucleotide switch protein. Science 309, 1377–1380 (2005).
pubmed: 16123301 doi: 10.1126/science.1115653
Schindelin, H., Kisker, C., Schlessman, J. L., Howard, J. B. & Rees, D. C. Structure of ADP·AlF
pubmed: 9163420 doi: 10.1038/387370a0
Schmid, B. et al. Biochemical and structural characterization of the cross-linked complex of nitrogenase: comparison to the ADP-AlF
pubmed: 12501184 doi: 10.1021/bi026642b
Chiu, H.-J. et al. MgATP-bound and nucleotide-free structures of a nitrogenase protein complex between the Leu 127Δ-Fe-protein and the MoFe-protein. Biochemistry 40, 641–650 (2001).
pubmed: 11170380 doi: 10.1021/bi001645e
Owens, C. P., Katz, F. E. H., Carter, C. H., Luca, M. A. & Tezcan, F. A. Evidence for functionally relevant encounter complexes in nitrogenase catalysis. J. Am. Chem. Soc. 137, 12704–12712 (2015).
pubmed: 26360912 pmcid: 4809638 doi: 10.1021/jacs.5b08310
Davidson, V. L. Protein control of true, gated, and coupled electron transfer reactions. Acc. Chem. Res. 41, 730–738 (2008).
pubmed: 18442271 pmcid: 4860822 doi: 10.1021/ar700252c
Danyal, K., Mayweather, D., Dean, D. R., Seefeldt, L. C. & Hoffman, B. M. Conformational gating of electron transfer from the nitrogenase Fe protein to MoFe protein. J. Am. Chem. Soc. 132, 6894–6895 (2010). This paper examines the rate of electron injection as a function of viscosity and osmolality, illuminating the nature of rate-limiting conformational changes in nitrogenase.
pubmed: 20429505 pmcid: 2892898 doi: 10.1021/ja101737f
Urbauer, J. L., Dorgan, L. J. & Schuster, S. M. Effects of deuterium on the kinetics of beef heart mitochondrial ATPase. Arch. Biochem. Biophys. 231, 498–502 (1984).
pubmed: 6329101 doi: 10.1016/0003-9861(84)90413-2
Jeuken, L. J. C. Conformational reorganisation in interfacial protein electron transfer. Biochim. Biophys. Acta Bioenerg. 1604, 67–76 (2003).
doi: 10.1016/S0005-2728(03)00026-4
Liang, Z.-X. et al. Dynamic docking and electron transfer between Zn-myoglobin and cytochrome b5. J. Am. Chem. Soc. 124, 6849–6859 (2002).
pubmed: 12059205 doi: 10.1021/ja0127032
Hazzard, J. T., Moench, S. J., Erman, J. E., Satterlee, J. D. & Tollin, G. Kinetics of intracomplex electron transfer and of reduction of the components of covalent and noncovalent complexes of cytochrome c and cytochrome c peroxidase by free flavin semiquinones. Biochemistry 27, 2002–2008 (1988).
pubmed: 2837280 doi: 10.1021/bi00406a029
Rutledge, H. L., Cook, B. D., Nguyen, H. P. M., Herzik, M. A. & Tezcan, F. A. Structures of the nitrogenase complex prepared under catalytic turnover conditions. Science 377, 865–869 (2022). In this paper, the catalytically active superstructure of nitrogenase is revealed by cryoEM, providing structural evidence for numerous conformational changes and the choreography of the dynamic protein–protein interactions that trigger them.
pubmed: 35901182 pmcid: 9949965 doi: 10.1126/science.abq7641
Ashby, G. A. & Thorneley, R. N. F. Nitrogenase of Klebsiella pneumoniae. Kinetic studies on the Fe protein involving reduction by sodium dithionite, the binding of MgADP and a conformation change that alters the reactivity of the 4Fe-4S centre. Biochem. J. 246, 455–465 (1987).
pubmed: 3318808 pmcid: 1148296 doi: 10.1042/bj2460455
Levitzki, A. & Koshland, D. E. Current Topics in Cellular Regulation Vol. 10 (eds. Horecker, B. L. & Stdtman, E. R.) 1–40 (Academic, 1976).
Nguyen, R. C., Stagliano, C. & Liu, A. Structural insights into the half-of-sites reactivity in homodimeric and homotetrameric metalloenzymes. Curr. Opin Chem. Biol. 75, 102332 (2023).
pubmed: 37269676 pmcid: 10528533 doi: 10.1016/j.cbpa.2023.102332
Howard, J. B. & Rees, D. C. Nitrogenase: a nucleotide-dependent molecular switch. Annu. Rev. Biochem. 63, 235–264 (1994).
pubmed: 7979238 doi: 10.1146/annurev.bi.63.070194.001315
Weston, M. F., Kotake, S. & Davis, L. C. Interaction of nitrogenase with nucleotide analogs of ATP and ADP and the effect of metal ions on ADP inhibition. Arch. Biochem. Biophys. 225, 809–817 (1983).
pubmed: 6354096 doi: 10.1016/0003-9861(83)90093-0
Danyal, K., Dean, D. R., Hoffman, B. M. & Seefeldt, L. C. Electron transfer within nitrogenase: evidence for a deficit-spending mechanism. Biochemistry 50, 9255–9263 (2011). Stopped-flow spectroscopic studies of a MoFeP variant support the conclusion that the immediate result of rate-limiting protein conformational changes is the internal ET from P-cluster to FeMoco, followed by backfill to P-cluster from FeP.
pubmed: 21939270 doi: 10.1021/bi201003a
Seefeldt, L. C. et al. Energy transduction in nitrogenase. Acc. Chem. Res. 51, 2179–2186 (2018).
pubmed: 30095253 pmcid: 6329455 doi: 10.1021/acs.accounts.8b00112
Chan, J. M., Christiansen, J., Dean, D. R. & Seefeldt, L. C. Spectroscopic evidence for changes in the redox state of the nitrogenase P-cluster during turnover. Biochemistry 38, 5779–5785 (1999).
pubmed: 10231529 doi: 10.1021/bi982866b
Peters, J. W. et al. Redox-dependent structural changes in the nitrogenase P-cluster. Biochemistry 36, 1181–1187 (1997).
pubmed: 9063865 doi: 10.1021/bi9626665
Lanzilotta, W. N. & Seefeldt, L. C. Changes in the midpoint potentials of the nitrogenase metal centers as a result of iron protein–molybdenum-iron protein complex formation. Biochemistry 36, 12976–12983 (1997).
pubmed: 9335558 doi: 10.1021/bi9715371
Kurnikov, I. V., Charnley, A. K. & Beratan, D. N. From ATP to electron transfer: electrostatics and free-energy transduction in nitrogenase. J. Phys. Chem. B 105, 5359–5367 (2001).
doi: 10.1021/jp002540o
Harris, D. F. et al. Mo-, V-, and Fe-nitrogenases use a universal eight-electron reductive-elimination mechanism to achieve N
pubmed: 31283201 doi: 10.1021/acs.biochem.9b00468
Davydov, R. et al. Exploring electron/proton transfer and conformational changes in the nitrogenase MoFe protein and FeMo-cofactor through cryoreduction/EPR measurements. Isr. J. Chem. 56, 841–851 (2016).
pubmed: 27777444 pmcid: 5074565 doi: 10.1002/ijch.201600026
Seefeldt, L. C. et al. Reduction of substrates by nitrogenases. Chem. Rev. 120, 5082–5106 (2020).
pubmed: 32176472 pmcid: 7703680 doi: 10.1021/acs.chemrev.9b00556
Durrant, M. C. Controlled protonation of iron–molybdenum cofactor by nitrogenase: a structural and theoretical analysis. Biochem. J. 355, 569–576 (2001).
pubmed: 11311117 pmcid: 1221770 doi: 10.1042/bj3550569
Morrison, C. N., Spatzal, T. & Rees, D. C. Reversible protonated resting state of the nitrogenase active site. J. Am. Chem. Soc. 139, 10856–10862 (2017).
pubmed: 28692802 pmcid: 5553094 doi: 10.1021/jacs.7b05695
Igarashi, R. Y. & Seefeldt, L. C. Nitrogen fixation: the mechanism of the Mo-dependent nitrogenase. Crit. Rev. Biochem. Mol. Biol. 38, 351–384 (2003).
pubmed: 14551236 doi: 10.1080/10409230391036766
Renger, G. The light reactions of photosynthesis. Curr. Sci. 98, 1305–1319 (2010).
Kok, B., Forbush, B. & McGloin, M. Cooperation of charges in photosynthetic O
pubmed: 5456273 doi: 10.1111/j.1751-1097.1970.tb06017.x
Vinyard, D. J., Ananyev, G. M. & Dismukes, G. C. Photosystem II: the reaction center of oxygenic photosynthesis. Annu. Rev. Biochem. 82, 577–606 (2013).
pubmed: 23527694 doi: 10.1146/annurev-biochem-070511-100425
Petrouleas, V. & Diner, B. A. Light-induced oxidation of the acceptor-side Fe(II) of photosystem II by exogenous quinones acting through the Q
doi: 10.1016/0005-2728(87)90032-6
Hermes, S. et al. A time-resolved iron-specific x-ray absorption experiment yields no evidence for an Fe
pubmed: 16401066 doi: 10.1021/bi0515725
He, W. Z., Newell, W. R., Haris, P. I., Chapman, D. & Barber, J. Protein secondary structure of the isolated photosystem II reaction center and conformational changes studied by Fourier transform infrared spectroscopy. Biochemistry 30, 4552–4559 (1991).
pubmed: 1850626 doi: 10.1021/bi00232a027
Klauss, A., Haumann, M. & Dau, H. Seven steps of alternating electron and proton transfer in photosystem II water oxidation traced by time-resolved photothermal beam deflection at improved sensitivity. J. Phys. Chem. B 119, 2677–2689 (2015).
pubmed: 25402235 doi: 10.1021/jp509069p
Nagy, G. et al. Dynamic properties of photosystem II membranes at physiological temperatures characterized by elastic incoherent neutron scattering. Increased flexibility associated with the inactivation of the oxygen evolving complex. Photosynth. Res. 111, 113–124 (2012).
pubmed: 22052408 doi: 10.1007/s11120-011-9701-x
Pieper, J. & Renger, G. Flash-induced structural dynamics in photosystem II membrane fragments of green plants. Biochemistry 48, 6111–6115 (2009).
pubmed: 19425568 doi: 10.1021/bi900414k
Zabret, J. et al. Structural insights into photosystem II assembly. Nat. Plants 7, 524–538 (2021).
pubmed: 33846594 pmcid: 8094115 doi: 10.1038/s41477-021-00895-0
Glöckner, C. et al. Structural changes of the oxygen-evolving complex in photosystem II during the catalytic cycle. J. Biol. Chem. 288, 22607–22620 (2013).
pubmed: 23766513 pmcid: 3829347 doi: 10.1074/jbc.M113.476622
Ibrahim, M. et al. Untangling the sequence of events during the S2 → S3 transition in photosystem II and implications for the water oxidation mechanism. Proc. Natl Acad. Sci. USA 117, 12624–12635 (2020).
pubmed: 32434915 pmcid: 7293653 doi: 10.1073/pnas.2000529117
Ishikita, H. & Knapp, E.-W. Induced conformational changes upon Cd
pubmed: 16254054 pmcid: 1283420 doi: 10.1073/pnas.0503826102
Rutherford, A. W. & Krieger-Liszkay, A. Herbicide-induced oxidative stress in photosystem II. Trends Biochem. Sci. 26, 648–653 (2001).
pubmed: 11701322 doi: 10.1016/S0968-0004(01)01953-3
Becker, K., Cormann, K. U. & Nowaczyk, M. M. Assembly of the water-oxidizing complex in photosystem II. J. Photochem. Photobiol. B 104, 204–211 (2011).
pubmed: 21382728 doi: 10.1016/j.jphotobiol.2011.02.005
Graige, M. S., Feher, G. & Okamura, M. Y. Conformational gating of the electron transfer reaction Q
pubmed: 9751725 pmcid: 21700 doi: 10.1073/pnas.95.20.11679
Renger, G., Gleiter, H. M., Haag, E. & Reifarth, F. Photosystem II: thermodynamics and kinetics of electron transport from Q
doi: 10.1515/znc-1993-3-419
Xu, Q. & Gunner, M. R. Trapping conformational intermediate states in the reaction center protein from photosynthetic bacteria. Biochemistry 40, 3232–3241 (2001).
pubmed: 11258940 doi: 10.1021/bi002326q
Renger, G. et al. Fluorescence and spectroscopic studies of exciton trapping and electron transfer in photosystem II of higher plants. Funct. Plant Biol. 22, 167–181 (1995).
doi: 10.1071/PP9950167
Kaminskaya, O., Renger, G. & Shuvalov, V. A. Effect of dehydration on light-induced reactions in photosystem II: photoreactions of cytochrome b559. Biochemistry 42, 8119–8132 (2003).
pubmed: 12846561 doi: 10.1021/bi020606v
Pieper, J. et al. Temperature- and hydration-dependent protein dynamics in photosystem II of green plants studied by quasielastic neutron scattering. Biochemistry 46, 11398–11409 (2007).
pubmed: 17867656 doi: 10.1021/bi700179s
Renger, G. & Renger, T. Photosystem II: the machinery of photosynthetic water splitting. Photosynth. Res. 98, 53–80 (2008).
pubmed: 18830685 doi: 10.1007/s11120-008-9345-7
de Wijn, R. & van Gorkom, H. J. Kinetics of electron transfer from Q
pubmed: 11570892 doi: 10.1021/bi010852r
Lupı́nková, L., Metz, J. G., Diner, B. A., Vass, I. & Komenda, J. Histidine residue 252 of the photosystem II D1 polypeptide is involved in a light-induced cross-linking of the polypeptide with the α subunit of cytochrome b-559: study of a site-directed mutant of Synechocystis PCC 6803. Biochim. Biophys. Acta Bioenerg. 1554, 192–201 (2002).
doi: 10.1016/S0005-2728(02)00243-8
Diner, B. A., Petrouleas, V. & Wendoloski, J. J. The iron-quinone electron-acceptor complex of photosystem II. Physiol. Plant. 81, 423–436 (1991).
doi: 10.1111/j.1399-3054.1991.tb08753.x
Kobayashi, T., Shimada, Y., Nagao, R. & Noguchi, T. pH-dependent regulation of electron flow in photosystem II by a histidine residue at the stromal surface. Biochemistry 61, 1351–1362 (2022). This research uses detection of stromal pH changes and modulation of the redox potential of Q
pubmed: 35686693 doi: 10.1021/acs.biochem.2c00150
Sigfridsson, K. G. V., Bernát, G., Mamedov, F. & Styring, S. Molecular interference of Cd
doi: 10.1016/j.bbabio.2004.07.003
Francia, F., Palazzo, G., Mallardi, A., Cordone, L. & Venturoli, G. Residual water modulates Q
pubmed: 14507738 pmcid: 1303499 doi: 10.1016/S0006-3495(03)74698-0
Pieper, J., Hauss, T., Buchsteiner, A. & Renger, G. The effect of hydration on protein flexibility in photosystem II of green plants studied by quasielastic neutron scattering. Eur. Biophys. J. 37, 657–663 (2008).
pubmed: 18351332 doi: 10.1007/s00249-008-0297-9
Vasil’ev, S., Bergmann, A., Redlin, H., Eichler, H.-J. & Renger, G. On the role of exchangeable hydrogen bonds for the kinetics of P680
doi: 10.1016/0005-2728(96)00027-8
Sugo, Y., Saito, K. & Ishikita, H. Conformational changes and H-bond rearrangements during quinone release in photosystem II. Biochemistry 61, 1836–1843 (2022).
pubmed: 35914244 doi: 10.1021/acs.biochem.2c00324
Sirohiwal, A. & Pantazis, D. A. Functional water networks in fully hydrated photosystem II. J. Am. Chem. Soc. 144, 22035–22050 (2022).
pubmed: 36413491 pmcid: 9732884 doi: 10.1021/jacs.2c09121
Alexov, E. G. & Gunner, M. R. Calculated protein and proton motions coupled to electron transfer: electron transfer from Q
pubmed: 10387071 doi: 10.1021/bi982700a
Saito, K., Rutherford, A. W. & Ishikita, H. Mechanism of proton-coupled quinone reduction in photosystem II. Proc. Natl Acad. Sci. USA 110, 954–959 (2013). Quantum mechanics/molecular mechanics analysis of the PT pathways and energetics involved in the two-step reduction of plastoquinone Q
pubmed: 23277574 doi: 10.1073/pnas.1212957110
Kulik, N., Kutý, M. & Řeha, D. The study of conformational changes in photosystem II during a charge separation. J. Mol. Model. 26, 75 (2020).
pubmed: 32152736 doi: 10.1007/s00894-020-4332-9
Paddock, M. L., Feher, G. & Okamura, M. Y. Proton transfer pathways and mechanism in bacterial reaction centers. FEBS Lett. 555, 45–50 (2003).
pubmed: 14630317 doi: 10.1016/S0014-5793(03)01149-9
Fufezan, C., Zhang, C., Krieger-Liszkay, A. & Rutherford, A. W. Secondary quinone in photosystem II of Thermosynechococcus elongatus: semiquinone–iron EPR signals and temperature dependence of electron transfer. Biochemistry 44, 12780–12789 (2005).
pubmed: 16171393 doi: 10.1021/bi051000k
Kimura, M., Kato, Y. & Noguchi, T. Protonation state of a key histidine ligand in the iron–quinone complex of photosystem II as revealed by light-induced ATR-FTIR spectroscopy. Biochemistry 59, 4336–4343 (2020).
pubmed: 33147961 doi: 10.1021/acs.biochem.0c00810
Stowell, M. H. B. et al. Light-induced structural changes in photosynthetic reaction center: implications for mechanism of electron-proton transfer. Science 276, 812–816 (1997).
pubmed: 9115209 doi: 10.1126/science.276.5313.812
Suga, M. et al. Light-induced structural changes and the site of O=O bond formation in PSII caught by XFEL. Nature 543, 131–135 (2017). The structural changes in PSII induced by two-flash illumination at room temperature are observed via time-resolved serial femtosecond crystallography with an X-ray free electron laser to 2.35 Å resolution.
pubmed: 28219079 doi: 10.1038/nature21400
Nabedryk, E. & Breton, J. Coupling of electron transfer to proton uptake at the Q
doi: 10.1016/j.bbabio.2008.06.012
Xu, Q., Baciou, L., Sebban, P. & Gunner, M. R. Exploring the energy landscape for Q
pubmed: 12146966 doi: 10.1021/bi025573y
Parak, F. et al. Evidence for a correlation between the photoinduced electron transfer and dynamic properties of the chromatophore membranes from Rhodospirillum rubrum. FEBS Lett. 117, 368–372 (1980).
pubmed: 6773810 doi: 10.1016/0014-5793(80)80982-3
Garbers, A., Reifarth, F., Kurreck, J., Renger, G. & Parak, F. Correlation between protein flexibility and electron transfer from to Q
pubmed: 9708974 doi: 10.1021/bi980296+
Koua, F. H. M. Structural changes in the acceptor site of photosystem II upon Ca
pubmed: 31416291 pmcid: 6722538 doi: 10.3390/biom9080371
Kargul, J. et al. Purification, crystallization and X-ray diffraction analyses of the T. elongatus PSII core dimer with strontium replacing calcium in the oxygen-evolving complex. Biochim. Biophys. Acta Bioenerg. 1767, 404–413 (2007).
doi: 10.1016/j.bbabio.2007.01.007
Koua, F. H. M., Umena, Y., Kawakami, K. & Shen, J.-R. Structure of Sr-substituted photosystem II at 2.1 Å resolution and its implications in the mechanism of water oxidation. Proc. Natl Acad. Sci. USA 110, 3889–3894 (2013).
pubmed: 23426624 pmcid: 3593836 doi: 10.1073/pnas.1219922110
Kato, Y. et al. Influence of the PsbA1/PsbA3, Ca
pubmed: 22721916 doi: 10.1016/j.bbabio.2012.06.006
Kato, Y. & Noguchi, T. Long-range interaction between the Mn
pubmed: 25029208 doi: 10.1021/bi500549b
Kato, Y., Ohira, A., Nagao, R. & Noguchi, T. Does the water-oxidizing Mn
pubmed: 31669461 doi: 10.1016/j.bbabio.2019.148082
Stubbe, J. & Nocera, D. G. Radicals in biology: your life is in their hands. J. Am. Chem. Soc. 143, 13463–13472 (2021).
pubmed: 34423635 pmcid: 8735831 doi: 10.1021/jacs.1c05952
Greene, B. L. et al. Ribonucleotide reductases: structure, chemistry, and metabolism suggest new therapeutic targets. Annu. Rev. Biochem. 89, 45–75 (2020).
pubmed: 32569524 pmcid: 7316142 doi: 10.1146/annurev-biochem-013118-111843
Ge, J., Yu, G., Ator, M. A. & Stubbe, J. A. Pre-steady-state and steady-state kinetic analysis of E. coli class I ribonucleotide reductase. Biochemistry 42, 10071–10083 (2003).
pubmed: 12939135 doi: 10.1021/bi034374r
Licht, S., Gerfen, G. J. & Stubbe, J. Thiyl radicals in ribonucleotide reductases. Science 271, 477–481 (1996).
pubmed: 8560260 doi: 10.1126/science.271.5248.477
Stubbe, J. Ribonucleotide reductases in the twenty-first century. Proc. Natl Acad. Sci. USA 95, 2723–2724 (1998).
pubmed: 9501154 pmcid: 34104 doi: 10.1073/pnas.95.6.2723
Kang, G., Taguchi, A. T., Stubbe, J. & Drennan, C. L. Structure of a trapped radical transfer pathway within a ribonucleotide reductase holocomplex. Science 368, 424–427 (2020). In this paper, the normally transient active RNR supercomplex is captured under catalytically relevant conditions and its structure is examined by cryoEM.
pubmed: 32217749 pmcid: 7774503 doi: 10.1126/science.aba6794
Climent, I., Sjöberg, B. M. & Huang, C. Y. Carboxyl-terminal peptides as probes for Escherichia coli ribonucleotide reductase subunit interaction: kinetic analysis of inhibition studies. Biochemistry 30, 5164–5171 (1991).
pubmed: 2036382 doi: 10.1021/bi00235a008
Mao, S. S. et al. A model for the role of multiple cysteine residues involved in ribonucleotide reduction: amazing and still confusing. Biochemistry 31, 9733–9743 (1992).
pubmed: 1382592 doi: 10.1021/bi00155a029
Ravichandran, K., Olshansky, L., Nocera, D. & Stubbe, J. Subunit interaction dynamics of class Ia ribonucleotide reductases: in search of a robust assay. Biochemistry 59, 1442–1453 (2020).
pubmed: 32186371 doi: 10.1021/acs.biochem.0c00001
Ando, N. et al. Structural interconversions modulate activity of Escherichia coli ribonucleotide reductase. Proc. Natl Acad. Sci. USA 108, 21046–21051 (2011).
pubmed: 22160671 pmcid: 3248520 doi: 10.1073/pnas.1112715108
Hofer, A., Crona, M., Logan, D. T. & Sjöberg, B.-M. DNA building blocks: keeping control of manufacture. Crit. Rev. Biochem. Mol. Biol. 47, 50–63 (2012).
pubmed: 22050358 doi: 10.3109/10409238.2011.630372
Minnihan, E. C. et al. Generation of a stable, aminotyrosyl radical-induced α2β2 complex of Escherichia coli class Ia ribonucleotide reductase. Proc. Natl Acad. Sci. USA 110, 3835–3840 (2013). This paper shows that trapping a radical along the RNR PCET pathway stalls the normally short-lived interactions between α
pubmed: 23431160 pmcid: 3593893 doi: 10.1073/pnas.1220691110
Minnihan, E. C., Seyedsayamdost, M. R., Uhlin, U. & Stubbe, J. Kinetics of radical intermediate formation and deoxynucleotide production in 3-aminotyrosine-substituted Escherichia coli ribonucleotide reductases. J. Am. Chem. Soc. 133, 9430–9440 (2011).
pubmed: 21612216 pmcid: 3125130 doi: 10.1021/ja201640n
Ravichandran, K. R. et al. Formal reduction potentials of difluorotyrosine and trifluorotyrosine protein residues: defining the thermodynamics of multistep radical transfer. J. Am. Chem. Soc. 139, 2994–3004 (2017).
pubmed: 28171730 pmcid: 5651514 doi: 10.1021/jacs.6b11011
Berry, B. W., Martinez-Rivera, M. C. & Tommos, C. Reversible voltammograms and a Pourbaix diagram for a protein tyrosine radical. Proc. Natl Acad. Sci. U A 109, 9739–9743 (2012).
doi: 10.1073/pnas.1112057109
Hay, S., Westerlund, K. & Tommos, C. Moving a phenol hydroxyl group from the surface to the interior of a protein: effects on the phenol potential and pK
pubmed: 16128591 doi: 10.1021/bi050901q
Ravichandran, K. R. et al. A >200 meV uphill thermodynamic landscape for radical transport in Escherichia coli ribonucleotide reductase determined using fluorotyrosine-substituted enzymes. J. Am. Chem. Soc. 138, 13706–13716 (2016).
pubmed: 28068088 pmcid: 5224885 doi: 10.1021/jacs.6b08200
Wörsdörfer, B. et al. Function of the diiron cluster of Escherichia coli class Ia ribonucleotide reductase in proton-coupled electron transfer. J. Am. Chem. Soc. 135, 8585–8593 (2013). Leveraging the ability to trap the active RNR complex, the researchers of this paper use Mössbauer spectroscopy to define the initial target of rate-limiting conformational changes in RNR as consisting of PT from the diiron metallocofactor to Y
pubmed: 23676140 doi: 10.1021/ja401342s
Seyedsayamdost, M. R., Chan, C. T. Y., Mugnaini, V., Stubbe, J. & Bennati, M. PELDOR spectroscopy with DOPA-β2 and NH
pubmed: 18047343 doi: 10.1021/ja076459b
Lebrette, H. et al. Structure of a ribonucleotide reductase R2 protein radical. Science 382, 109–113 (2023).
pubmed: 37797025 pmcid: 7615503 doi: 10.1126/science.adh8160
Yokoyama, K., Uhlin, U. & Stubbe, J. Site-specific incorporation of 3-nitrotyrosine as a probe of pK
pubmed: 20518462 pmcid: 2905227 doi: 10.1021/ja101097p
Ator, M., Salowe, S. P., Stubbe, J., Emptage, M. H. & Robins, M. J. 2′-Azido-2′-deoxynucleotide interaction with E. coli ribonucleotide reductase: generation of a new radical species. J. Am. Chem. Soc. 106, 1886–1887 (1984).
doi: 10.1021/ja00318a082
Sjöberg, B. M., Gräslund, A. & Eckstein, F. A substrate radical intermediate in the reaction between ribonucleotide reductase from Escherichia coli and 2′-azido-2′-deoxynucleoside diphosphates. J. Biol. Chem. 258, 8060–8067 (1983).
pubmed: 6305969 doi: 10.1016/S0021-9258(20)82027-4
Högbom, M. et al. Displacement of the tyrosyl radical cofactor in ribonucleotide reductase obtained by single-crystal high-field EPR and 1.4-Å x-ray data. Proc. Natl Acad. Sci. USA 100, 3209–3214 (2003).
pubmed: 12624184 pmcid: 404301 doi: 10.1073/pnas.0536684100
Offenbacher, A. R., Burns, L. A., Sherrill, C. D. & Barry, B. A. Redox-linked conformational control of proton-coupled electron transfer: Y122 in the ribonucleotide reductase β2 subunit. J. Phys. Chem. B 117, 8457–8468 (2013).
pubmed: 23822111 pmcid: 3757525 doi: 10.1021/jp404757r
Offenbacher, A. R., Minnihan, E. C., Stubbe, J. & Barry, B. A. Redox-linked changes to the hydrogen-bonding network of ribonucleotide reductase β2. J. Am. Chem. Soc. 135, 6380–6383 (2013).
pubmed: 23594029 pmcid: 3694779 doi: 10.1021/ja3032949
Watson, R. A., Offenbacher, A. R. & Barry, B. A. Detection of catalytically linked conformational changes in wild-type class Ia ribonucleotide reductase using reaction-induced FTIR spectroscopy. J. Phys. Chem. B 125, 8362–8372 (2021).
pubmed: 34289692 doi: 10.1021/acs.jpcb.1c03038
Yokoyama, K., Smith, A. A., Corzilius, B., Griffin, R. G. & Stubbe, J. Equilibration of tyrosyl radicals (Y356·, Y731·, Y730·) in the radical propagation pathway of the Escherichia coli class Ia ribonucleotide reductase. J. Am. Chem. Soc. 133, 18420–18432 (2011).
pubmed: 21967342 pmcid: 3236566 doi: 10.1021/ja207455k
Ravichandran, K. R., Minnihan, E. C., Wei, Y., Nocera, D. G. & Stubbe, J. A. Reverse electron transfer completes the catalytic cycle in a 2,3,5-trifluorotyrosine-substituted ribonucleotide reductase. J. Am. Chem. Soc. 137, 14387–14395 (2015).
pubmed: 26492582 pmcid: 4678968 doi: 10.1021/jacs.5b09189
Yokoyama, K., Uhlin, U. & Stubbe, J. A hot oxidant, 3-NO
pubmed: 20929229 pmcid: 3005585 doi: 10.1021/ja1069344
Offenbacher, A. R., Watson, R. A., Pagba, C. V. & Barry, B. A. Redox-dependent structural coupling between the α2 and β2 subunits in E. coli ribonucleotide reductase. J. Phys. Chem. B 118, 2993–3004 (2014).
pubmed: 24606240 doi: 10.1021/jp501121d
Srinivas, V. et al. Metal-free ribonucleotide reduction powered by a DOPA radical in mycoplasma pathogens. Nature 563, 416–420 (2018).
pubmed: 30429545 pmcid: 6317698 doi: 10.1038/s41586-018-0653-6
Blaesi, E. J. et al. Metal-free class Ie ribonucleotide reductase from pathogens initiates catalysis with a tyrosine-derived dihydroxyphenylalanine radical. Proc. Natl Acad. Sci. USA 115, 10022–10027 (2018).
pubmed: 30224458 pmcid: 6176560 doi: 10.1073/pnas.1811993115
Bollinger, J. M. Jr. et al. Mechanism of assembly of the tyrosyl radical-diiron(III) cofactor of E. coli ribonucleotide reductase. 3. Kinetics of the limiting Fe
doi: 10.1021/ja00097a009
Minnihan, E. C., Nocera, D. G. & Stubbe, J. Reversible, long-range radical transfer in E. coli class Ia ribonucleotide reductase. Acc. Chem. Res. 46, 2524–2535 (2013).
pubmed: 23730940 doi: 10.1021/ar4000407
Oyala, P. H. et al. Biophysical characterization of fluorotyrosine probes site-specifically incorporated into enzymes: E. coli ribonucleotide reductase as an example. J. Am. Chem. Soc. 138, 7951–7964 (2016).
pubmed: 27276098 pmcid: 4929525 doi: 10.1021/jacs.6b03605
Seyedsayamdost, M. R., Reece, S. Y., Nocera, D. G. & Stubbe, J. Mono-, di-, tri-, and tetra-substituted fluorotyrosines: new probes for enzymes that use tyrosyl radicals in catalysis. J. Am. Chem. Soc. 128, 1569–1579 (2006).
pubmed: 16448128 doi: 10.1021/ja055926r
Pizano, A. A. et al. Photo-ribonucleotide reductase β2 by selective cysteine labeling with a radical phototrigger. Proc. Natl Acad. Sci. USA 109, 39–43 (2012).
pubmed: 22171005 doi: 10.1073/pnas.1115778108
Cui, C., Song, D. Y., Drennan, C. L., Stubbe, J. & Nocera, D. G. Radical transport facilitated by a proton transfer network at the subunit interface of ribonucleotide reductase. J. Am. Chem. Soc. 145, 5145–5154 (2023).
pubmed: 36812162 pmcid: 10561588 doi: 10.1021/jacs.2c11483
Cui, C. et al. Gated proton release during radical transfer at the subunit interface of ribonucleotide reductase. J. Am. Chem. Soc. 143, 176–183 (2021).
pubmed: 33353307 doi: 10.1021/jacs.0c07879
Ravichandran, K. et al. Glutamate 350 plays an essential role in conformational gating of long-range radical transport in Escherichia coli class Ia ribonucleotide reductase. Biochemistry 56, 856–868 (2017).
pubmed: 28103007 doi: 10.1021/acs.biochem.6b01145
Zhong, J., Reinhardt, C. R. & Hammes-Schiffer, S. Direct proton-coupled electron transfer between interfacial tyrosines in ribonucleotide reductase. J. Am. Chem. Soc. 145, 4784–4790 (2023).
pubmed: 36802630 pmcid: 10344599 doi: 10.1021/jacs.2c13615
Nick, T. U. et al. Hydrogen bond network between amino acid radical intermediates on the proton-coupled electron transfer pathway of E. coli α2 ribonucleotide reductase. J. Am. Chem. Soc. 137, 289–298 (2015).
pubmed: 25516424 doi: 10.1021/ja510513z
Hecker, F., Stubbe, J. & Bennati, M. Detection of water molecules on the radical transfer pathway of ribonucleotide reductase by
pubmed: 33957040 pmcid: 8154519 doi: 10.1021/jacs.1c01359
Greene, B. L., Taguchi, A. T., Stubbe, J. & Nocera, D. G. Conformationally dynamic radical transfer within ribonucleotide reductase. J. Am. Chem. Soc. 139, 16657–16665 (2017).
pubmed: 29037038 pmcid: 5702266 doi: 10.1021/jacs.7b08192
Kasanmascheff, M., Lee, W., Nick, T. U., Stubbe, J. & Bennati, M. Radical transfer in E. coli ribonucleotide reductase: a NH
pubmed: 29899944 doi: 10.1039/C5SC03460D
Meyer, A. et al.
pubmed: 35652913 pmcid: 9248007 doi: 10.1021/jacs.2c02906
Stubbe, J. A., Ackles, D., Ator, M. & Krenitsky, T. Mechanism of ribonucleoside diphosphate reductase from Escherichia coli. Evidence for 3′-C-H bond cleavage. J. Biol. Chem. 255, 1625–1630 (1980).
doi: 10.1016/S0021-9258(19)70598-5
Levitz, T. S. et al. A rapid and sensitive assay for quantifying the activity of both aerobic and anaerobic ribonucleotide reductases acting upon any or all substrates. PLoS ONE 17, e0269572 (2022).
pubmed: 35675376 pmcid: 9176816 doi: 10.1371/journal.pone.0269572
Livoreil, A., Dietrich-Buchecker, C. O. & Sauvage, J. P. Electrochemically triggerred swinging of a [2]-catenate. J. Am. Chem. Soc. 116, 9399–9400 (1994).
pubmed: 27715033 doi: 10.1021/ja00099a095
Meylemans, H. A., Hewitt, J. T., Abdelhaq, M., Vallett, P. J. & Damrauer, N. H. Exploiting conformational dynamics to facilitate formation and trapping of electron-transfer photoproducts in metal complexes. J. Am. Chem. Soc. 132, 11464–11466 (2010).
pubmed: 20684515 doi: 10.1021/ja1055559
Lister, F. G. A., Le Bailly, B. A. F., Webb, S. J. & Clayden, J. Ligand-modulated conformational switching in a fully synthetic membrane-bound receptor. Nat. Chem. 9, 420–425 (2017).
doi: 10.1038/nchem.2736
Xie, X., Crespo, G. A., Mistlberger, G. & Bakker, E. Photocurrent generation based on a light-driven proton pump in an artificial liquid membrane. Nat. Chem. 6, 202–207 (2014).
pubmed: 24557134 doi: 10.1038/nchem.1858
Gemen, J. et al. Disequilibrating azobenzenes by visible-light sensitization under confinement. Science 381, 1357–1363 (2023).
pubmed: 37733864 doi: 10.1126/science.adh9059

Auteurs

Saman Fatima (S)

Department of Chemistry, College of Liberal Arts and Sciences, University of Illinois Urbana-Champaign, Urbana, IL, USA.

Lisa Olshansky (L)

Department of Chemistry, College of Liberal Arts and Sciences, University of Illinois Urbana-Champaign, Urbana, IL, USA. lolshans@illinois.edu.
Center for Biophysics and Quantitative Biology, University of Illinois Urbana-Champaign, Urbana, IL, USA. lolshans@illinois.edu.
Materials Research Laboratory, The Grainger College of Engineering, University of Illinois Urbana-Champaign, Urbana, IL, USA. lolshans@illinois.edu.
The Beckman Institute for Advanced Science and Technology, University of Illinois Urbana-Champaign, Urbana, IL, USA. lolshans@illinois.edu.

Classifications MeSH