Abstract
Lytic polysaccharide monooxygenases (LPMOs) have an essential role in global carbon cycle, industrial biomass processing and microbial pathogenicity by catalysing the oxidative cleavage of recalcitrant polysaccharides. Despite initially being considered monooxygenases, experimental and theoretical studies show that LPMOs are essentially peroxygenases, using a single copper ion and H2O2 for C–H bond oxygenation. Here, we examine LPMO catalysis, emphasizing key studies that have shaped our comprehension of their function, and address side and competing reactions that have partially obscured our understanding. Then, we compare this novel copper–peroxygenase reaction with reactions catalysed by haem iron enzymes, highlighting the different chemistries at play. We conclude by addressing some open questions surrounding LPMO catalysis, including the importance of peroxygenase and monooxygenase reactions in biological contexts, how LPMOs modulate copper site reactivity and potential protective mechanisms against oxidative damage.
This is a preview of subscription content, access via your institution
Access options
Access Nature and 54 other Nature Portfolio journals
Get Nature+, our best-value online-access subscription
$29.99 / 30 days
cancel any time
Subscribe to this journal
Receive 12 digital issues and online access to articles
$119.00 per year
only $9.92 per issue
Buy this article
- Purchase on Springer Link
- Instant access to full article PDF
Prices may be subject to local taxes which are calculated during checkout
Similar content being viewed by others
Change history
22 January 2024
A Correction to this paper has been published: https://doi.org/10.1038/s41570-024-00580-8
References
Koshland, D. E. Jr Stereochemistry and the mechanism of enzymatic reactions. Biol. Rev. Camb. Philos. Soc. 28, 416–436 (1953).
Vuong, T. V. & Wilson, D. B. Glycoside hydrolases: catalytic base/nucleophile diversity. Biotechnol. Bioeng. 107, 195–205 (2010).
Davies, G. & Henrissat, B. Structures and mechanisms of glycosyl hydrolases. Structure 3, 853–859 (1995).
Rye, C. S. & Withers, S. G. Glycosidase mechanisms. Curr. Opin. Chem. Biol. 4, 573–580 (2000).
Bissaro, B., Monsan, P., Fauré, R. & O’Donohue, M. J. Glycosynthesis in a waterworld: new insight into the molecular basis of transglycosylation in retaining glycoside hydrolases. Biochem. J. 467, 17–35 (2015).
Jongkees, S. A. K. & Withers, S. G. Unusual enzymatic glycoside cleavage mechanisms. Acc. Chem. Res. 47, 226–235 (2014).
Vaaje-Kolstad, G., Horn, S. J., Van Aalten, D. M. F., Synstad, B. & Eijsink, V. G. H. The non-catalytic chitin-binding protein CBP21 from Serratia marcescens is essential for chitin degradation. J. Biol. Chem. 280, 28492–29497 (2005).
Vaaje-Kolstad, G. et al. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330, 219–222 (2010).
Quinlan, R. J. et al. Insights into the oxidative degradation of cellulose by a copper metalloenzyme that exploits biomass components. Proc. Natl Acad. Sci. USA 108, 15079–15084 (2011).
Aachmann, F. L., Sørlie, M., Skjåk-Bræk, G., Eijsink, V. G. H. & Vaaje-Kolstad, G. NMR structure of a lytic polysaccharide monooxygenase provides insight into copper binding, protein dynamics, and substrate interactions. Proc. Natl Acad. Sci. USA 109, 18779–18784 (2012).
Phillips, C. M., Beeson, W. T., Cate, J. H. & Marletta, M. A. Cellobiose dehydrogenase and a copper-dependent polysaccharide monooxygenase potentiate cellulose degradation by Neurospora crassa. ACS Chem. Biol. 6, 1399–1406 (2011).
Wang, Y., Lan, D., Durrani, R. & Hollmann, F. Peroxygenases en route to becoming dream catalysts. What are the opportunities and challenges? Curr. Opin. Chem. Biol. https://doi.org/10.1016/j.cbpa.2016.10.007 (2017).
Denisov, I. G., Makris, T. M., Sligar, S. G. & Schlichting, I. Structure and chemistry of cytochrome P450. Chem. Rev. 105, 2253–2277 (2005).
Johansen, K. S. Discovery and industrial applications of lytic polysaccharide mono-oxygenases. Biochem. Soc. Trans. 44, 143–149 (2016).
Chylenski, P. et al. Lytic polysaccharide monooxygenases in enzymatic processing of lignocellulosic biomass. ACS Catal. 9, 4970–4991 (2019).
Vandhana, T. M. et al. On the expansion of biological functions of lytic polysaccharides monooxygenases. New Phytol. 233, 2380–2396 (2022).
Garcia-Santamarina, S. et al. A lytic polysaccharide monooxygenase-like protein functions in fungal copper import and meningitis. Nat. Chem. Biol. 16, 337–344 (2020).
Martinez-D’Alto, A. et al. Characterization of a unique polysaccharide monooxygenase from the plant pathogen Magnaporthe oryzae. Proc. Natl Acad. Sci. USA 120, e2215426120 (2023).
Drula, E. et al. The carbohydrate-active enzyme database: functions and literature. Nucleic Acids Res. 50, D571–D577 (2022).
Vu, V. V., Beeson, W. T., Span, E. A., Farquhar, E. R. & Marletta, M. A. A family of starch-active polysaccharide monooxygenases. Proc. Natl Acad. Sci. USA 111, 13822–13827 (2014).
Couturier, M. et al. Lytic xylan oxidases from wood-decay fungi unlock biomass degradation. Nat. Chem. Biol. 14, 306–310 (2018).
Sabbadin, F. et al. An ancient family of lytic polysaccharide monooxygenases with roles in arthropod development and biomass digestion. Nat. Commun. 9, 756 (2018).
Filiatrault-Chastel, C. et al. AA16, a new lytic polysaccharide monooxygenase family identified in fungal secretomes. Biotechnol. Biofuels 12, 55 (2019).
Sabbadin, F. et al. Secreted pectin monooxygenases drive plant infection by pathogenic oomycetes. Science 373, 774–779 (2021).
Hemsworth, G. R., Henrissat, B., Davies, G. J. & Walton, P. H. Discovery and characterization of a new family of lytic polysaccharide monooxygenases. Nat. Chem. Biol. 10, 122–126 (2014).
Ruscic, B. Active thermochemical tables: sequential bond dissociation enthalpies of methane, ethane, and methanol and the related thermochemistry. J. Phys. Chem. A 119, 7810–7837 (2015).
Gao, J., Thomas, D. A., Sohn, C. H. & Beauchamp, J. L. Biomimetic reagents for the selective free radical and acid-base chemistry of glycans: application to glycan structure determination by mass spectrometry. J. Am. Chem. Soc. 135, 10684–10692 (2013).
Luo, Y. R. Handbook of Bond Dissociation Energies in Organic Compounds 1st edn (CRC, 2002).
Hedegård, E. D. & Ryde, U. Targeting the reactive intermediate in polysaccharide monooxygenases. J. Biol. Inorg. Chem. 22, 1029–1037 (2017).
Bissaro, B. et al. Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2. Nat. Chem. Biol. 13, 1123–1128 (2017).
Bissaro, B., Várnai, A., Røhr, Å. K. & Eijsink, V. G. H. Oxidoreductases and reactive oxygen species in conversion of lignocellulosic biomass. Microbiol. Mol. Biol. Rev. 82, e00029 (2018).
Meier, K. K. et al. Oxygen activation by Cu LPMOs in recalcitrant carbohydrate polysaccharide conversion to monomer sugars. Chem. Rev. 118, 2593–2635 (2018).
Kovaleva, E. G. & Lipscomb, J. D. Crystal structures of Fe2+ dioxygenase superoxo, alkylperoxo, and bound product intermediates. Science 316, 453–457 (2007).
Williams, P. A. et al. Crystal structures of human cytochrome P450 3A4 bound to metyrapone and progesterone. Science 305, 683–686 (2004).
Ramirez-Escudero, M. et al. Structural insights into the substrate promiscuity of a laboratory-evolved peroxygenase. ACS Chem. Biol. 13, 3259–3268 (2018).
Tan, T.-C. et al. Structural basis for cellobiose dehydrogenase action during oxidative cellulose degradation. Nat. Commun. 6, 7542 (2015).
Hecht, H. J., Kalisz, H. M., Hendle, J., Schmid, R. D. & Schomburg, D. Crystal structure of glucose oxidase from Aspergillus niger refined at 2.3 Å resolution. J. Mol. Biol. 229, 153–172 (1993).
Bertrand, T. et al. Crystal structure of a four-copper laccase complexed with an arylamine: insights into substrate recognition and correlation with kinetics. Biochemistry 41, 7325–7333 (2002).
Carlsson, G. H., Nicholls, P., Svistunenko, D., Berglund, G. I. & Hajdu, J. Complexes of horseradish peroxidase with formate, acetate, and carbon monoxide. Biochemistry 44, 635–642 (2005).
Ciano, L., Davies, G. J., Tolman, W. B. & Walton, P. H. Bracing copper for the catalytic oxidation of C–H bonds. Nat. Catal. 1, 571–577 (2018).
Hagemann, M. M. & Hedegård, E. D. Molecular mechanism of substrate oxidation in lytic polysaccharide monooxygenases: insight from theoretical investigations. Chem. Eur. J. 29, e202202379 (2023).
Bissaro, B., Kommedal, E., Røhr, Å. K. & Eijsink, V. G. H. Controlled depolymerization of cellulose by light-driven lytic polysaccharide oxygenases. Nat. Commun. 11, 890 (2020).
Stepnov, A. A. et al. The impact of reductants on the catalytic efficiency of a lytic polysaccharide monooxygenase and the special role of dehydroascorbic acid. FEBS Lett. 596, 53–70 (2022).
Eijsink, V. G. H. et al. On the functional characterization of lytic polysaccharide monooxygenases (LPMOs). Biotechnol. Biofuels 12, 58 (2019).
Bissaro, B. et al. Fenton-type chemistry by a copper enzyme: molecular mechanism of polysaccharide oxidative cleavage. Preprint at https://doi.org/10.1101/097022 (2016).
Hangasky, J. A., Iavarone, A. T. & Marletta, M. A. Reactivity of O2 versus H2O2 with polysaccharide monooxygenases. Proc. Natl Acad. Sci. USA 115, 4915–4920 (2018).
Jones, S. M. et al. Kinetic analysis of amino acid radicals formed in H2O2-driven CuI LPMO reoxidation implicates dominant homolytic reactivity. Proc. Natl Acad. Sci. USA 117, 11916–11922 (2020).
Hemsworth, G. R. Revisiting the role of electron donors in lytic polysaccharide monooxygenase biochemistry. Essays Biochem. 18, 585–595 (2023).
Chang, H. et al. Investigating lytic polysaccharide monooxygenase-assisted wood cell wall degradation with microsensors. Nat. Commun. 13, 6258 (2022).
Hedison, T. M. et al. Insights into the H2O2-driven catalytic mechanism of fungal lytic polysaccharide monooxygenases. FEBS J. 288, 4115–4128 (2021).
Kuusk, S. et al. Kinetic insights into the role of the reductant in H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase. J. Biol. Chem. 294, 1516–1528 (2019).
Kracher, D. et al. Extracellular electron transfer systems fuel cellulose oxidative degradation. Science 352, 1098–1101 (2016).
Cordas, C. M. et al. Electrochemical characterization of a family AA10 LPMO and the impact of residues shaping the copper site on reactivity. J. Inorg. Biochem. 238, 112056 (2023).
Frommhagen, M. et al. Lytic polysaccharide monooxygenases from Myceliophthora thermophila C1 differ in substrate preference and reducing agent specificity. Biotechnol. Biofuels 9, 186 (2016).
Frommhagen, M., Westphal, A. H., van Berkel, W. J. H. & Kabel, M. A. Distinct substrate specificities and electron-donating systems of fungal lytic polysaccharide monooxygenases. Front. Microbiol. 9, 1080 (2018).
Dimarogona, M., Topakas, E. & Christakopoulos, P. Cellulose degradation by oxidative enzymes. Comput. Struct. Biotechnol. J. 2, e201209015 (2012).
Garajova, S. et al. Single-domain flavoenzymes trigger lytic polysaccharide monooxygenases for oxidative degradation of cellulose. Sci. Rep. 6, 28276 (2016).
Haddad Momeni, M. et al. Discovery of fungal oligosaccharide-oxidising flavo-enzymes with previously unknown substrates, redox-activity profiles and interplay with LPMOs. Nat. Commun. 12, 2132 (2021).
Langston, J. A. et al. Oxidoreductive cellulose depolymerization by the enzymes cellobiose dehydrogenase and glycoside hydrolase 61. Appl. Environ. Microbiol. 77, 7007–7015 (2011).
Várnai, A., Umezawa, K., Yoshida, M. & Eijsink, V. G. H. The pyrroloquinoline–quinone-dependent pyranose dehydrogenase from Coprinopsis cinerea drives lytic polysaccharide monooxygenase action. Appl. Environ. Microbiol. 84, e00156–e00618 (2018).
Cannella, D. et al. Light-driven oxidation of polysaccharides by photosynthetic pigments and a metalloenzyme. Nat. Commun. 7, 11134 (2016).
Kommedal, E. G., Sæther, F., Hahn, T. & Eijsink, V. G. H. Natural photoredox catalysts promote light-driven lytic polysaccharide monooxygenase reactions and enzymatic turnover of biomass. Proc. Natl Acad. Sci. USA 119, e2204510119 (2022).
Kracher, D. et al. Polysaccharide oxidation by lytic polysaccharide monooxygenase is enhanced by engineered cellobiose dehydrogenase. FEBS J. 287, 897–908 (2020).
Branch, J. et al. C-type cytochrome-initiated reduction of bacterial lytic polysaccharide monooxygenases. Biochem. J. 478, 2927 (2021).
Loose, J. S. M. et al. Activation of bacterial lytic polysaccharide monooxygenases with cellobiose dehydrogenase. Protein Sci. 25, 2175–2186 (2016).
Bissaro, B. et al. Molecular mechanism of the chitinolytic peroxygenase reaction. Proc. Natl Acad. Sci. USA 117, 1504–1513 (2020).
Rieder, L., Stepnov, A. A., Sørlie, M. & Eijsink, V. G. H. Fast and specific peroxygenase reactions catalyzed by fungal mono-copper enzymes. Biochemistry 60, 3633–3643 (2021).
Golten, O. et al. Reductants fuel lytic polysaccharide monooxygenase activity in a pH-dependent manner. FEBS Lett. 597, 1363–1374 (2023).
Forsberg, Z. & Courtade, G. On the impact of carbohydrate-binding modules (CBMs) in lytic polysaccharide monooxygenases (LPMOs). Essays Biochem. 67, 561–574 (2022).
Forsberg, Z. et al. Structural and functional characterization of a conserved pair of bacterial cellulose-oxidizing lytic polysaccharide monooxygenases. Proc. Natl Acad. Sci. USA 111, 8446–8451 (2014).
Frandsen, K. E. H. & Lo Leggio, L. Lytic polysaccharide monooxygenases: a crystallographer’s view on a new class of biomass-degrading enzymes. IUCrJ 3, 448–467 (2016).
Tandrup, T., Frandsen, K. E. H., Johansen, K. S., Berrin, J. G. & Leggio, L. L. Recent insights into lytic polysaccharide monooxygenases (LPMOs). Biochem. Soc. Trans. 46, 1431–1447 (2018).
Borisova, A. S. et al. Structural and functional characterization of a lytic polysaccharide monooxygenase with broad substrate specificity. J. Biol. Chem. 290, 22955–22969 (2015).
Forsberg, Z. et al. Comparative study of two chitin-active and two cellulose-active AA10-type lytic polysaccharide monooxygenases. Biochemistry 53, 1647–1656 (2014).
Courtade, G. et al. Mechanistic basis of substrate-O2 coupling within a chitin-active lytic polysaccharide monooxygenase: an integrated NMR/EPR study. Proc. Natl Acad. Sci. USA 117, 19178–19189 (2020).
Bissaro, B., Isaksen, I., Vaaje-Kolstad, G., Eijsink, V. G. H. & Røhr, Å. K. How a lytic polysaccharide monooxygenase binds crystalline chitin. Biochemistry 57, 1893–1906 (2018).
Wu, M. et al. Crystal structure and computational characterization of the lytic polysaccharide monooxygenase GH61D from the Basidiomycota fungus Phanerochaete chrysosporium. J. Biol. Chem. 57, 12828–12839 (2013).
Vaaje-Kolstad, G., Houston, D. R., Riemen, A. H. K., Eijsink, V. G. H. & Van Aalten, D. M. F. Crystal structure and binding properties of the Serratia marcescens chitin-binding protein CBP2. J. Biol. Chem. 280, 11313–11319 (2005).
Harris, P. V. et al. Stimulation of lignocellulosic biomass hydrolysis by proteins of glycoside hydrolase family 61: structure and function of a large, enigmatic family. Biochemistry 49, 3305–3316 (2010).
Danneels, B., Tanghe, M. & Desmet, T. Structural features on the substrate-binding surface of fungal lytic polysaccharide monooxygenases determine their oxidative regioselectivity. Biotechnol. J. 14, 1800211 (2019).
Forsberg, Z. et al. Structural determinants of bacterial lytic polysaccharide monooxygenase functionality. J. Biol. Chem. 293, 1397–1412 (2018).
Frandsen, K. E. H. et al. Identification of the molecular determinants driving the substrate specificity of fungal lytic polysaccharide monooxygenases (LPMOs). J. Biol. Chem. 296, 100086 (2021).
Jensen, M. S. et al. Engineering chitinolytic activity into a cellulose-active lytic polysaccharide monooxygenase provides insights into substrate specificity. J. Biol. Chem. 294, 19349–19364 (2019).
Isaksen, T. et al. A C4-oxidizing lytic polysaccharide monooxygenase cleaving both cellulose and cello-oligosaccharides. J. Biol. Chem. 289, 2632–2642 (2014).
Agger, J. W. et al. Discovery of LPMO activity on hemicelluloses shows the importance of oxidative processes in plant cell wall degradation. Proc. Natl Acad. Sci. USA 111, 6287–6292 (2014).
Frandsen, K. E. H. et al. The molecular basis of polysaccharide cleavage by lytic polysaccharide monooxygenases. Nat. Chem. Biol. 12, 298–303 (2016).
Bennati-Granier, C. et al. Substrate specificity and regioselectivity of fungal AA9 lytic polysaccharide monooxygenases secreted by Podospora anserina. Biotechnol. Biofuels 8, 90 (2015).
Kracher, D., Andlar, M., Furtmüller, P. G. & Ludwig, R. Active-site copper reduction promotes substrate binding of fungal lytic polysaccharide monooxygenase and reduces stability. J. Biol. Chem. 293, 1676–1687 (2018).
Kuusk, S. et al. Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase. J. Biol. Chem. 293, 523–531 (2018).
Rieder, L., Petrović, D., Väljamäe, P., Eijsink, V. G. H. & Sørlie, M. Kinetic characterization of a putatively chitin-active LPMO reveals a preference for soluble substrates and absence of monooxygenase activity. ACS Catal. 11, 11685–11695 (2021).
Li, X., Beeson, W. T., Phillips, C. M., Marletta, M. A. & Cate, J. H. D. Structural basis for substrate targeting and catalysis by fungal polysaccharide monooxygenases. Structure 20, 1051–1061 (2012).
Hangasky, J. A. & Marletta, M. A. A random-sequential kinetic mechanism for polysaccharide monooxygenases. Biochemistry 57, 3191–3199 (2018).
Schröder, G. C., O’Dell, W. B., Webb, S. P., Agarwal, P. K. & Meilleur, F. Capture of activated dioxygen intermediates at the copper-active site of a lytic polysaccharide monooxygenase. Chem. Sci. 13, 13303–13320 (2022).
O’Dell, W. B., Agarwal, P. K. & Meilleur, F. Oxygen activation at the active site of a fungal lytic polysaccharide monooxygenase. Angew. Chem. Int. Ed. Engl. 56, 767–770 (2017).
Kjaergaard, C. H. et al. Spectroscopic and computational insight into the activation of O2 by the mononuclear Cu center in polysaccharide monooxygenases. Proc. Natl Acad. Sci. USA 111, 8797–8802 (2014).
Kuusk, S. & Väljamäe, P. Kinetics of H2O2-driven catalysis by a lytic polysaccharide monooxygenase from the fungus Trichoderma reesei. J. Biol. Chem. 297, 101256 (2021).
Beeson, W. T., Phillips, C. M., Cate, J. H. D. & Marletta, M. A. Oxidative cleavage of cellulose by fungal copper-dependent polysaccharide monooxygenases. J. Am. Chem. Soc. 134, 890–892 (2012).
Kittl, R., Kracher, D., Burgstaller, D., Haltrich, D. & Ludwig, R. Production of four Neurospora crassa lytic polysaccharide monooxygenases in Pichia pastoris monitored by a fluorimetric assay. Biotechnol. Biofuels 5, 79 (2012).
Hegnar, O. A. et al. pH-dependent relationship between catalytic activity and hydrogen peroxide production shown via characterization of a lytic polysaccharide monooxygenase from Gloeophyllum trabeum. Appl. Environ. Microbiol. 85, e02612–e02618 (2019).
Kont, R., Bissaro, B., Eijsink, V. G. H. & Väljamäe, P. Kinetic insights into the peroxygenase activity of cellulose-active lytic polysaccharide monooxygenases (LPMOs). Nat. Commun. 11, 5786 (2020).
Filandr, F. et al. The H2O2-dependent activity of a fungal lytic polysaccharide monooxygenase investigated with a turbidimetric assay. Biotechnol. Biofuels 13, 37 (2020).
Wang, B. et al. QM/MM studies into the H2O2-dependent activity of lytic polysaccharide monooxygenases: evidence for the formation of a caged hydroxyl radical intermediate. ACS Catal. 8, 1346–1351 (2018).
Hedegård, E. D. & Ryde, U. Molecular mechanism of lytic polysaccharide monooxygenases. Chem. Sci. 9, 3866–3880 (2018).
Kim, S., Ståhlberg, J., Sandgren, M., Paton, R. S. & Beckham, G. T. Quantum mechanical calculations suggest that lytic polysaccharide monooxygenases use a copper-oxyl, oxygen-rebound mechanism. Proc. Natl Acad. Sci. USA 111, 149–154 (2014).
Wang, B., Wang, Z., Davies, G. J., Walton, P. H. & Rovira, C. Activation of O2 and H2O2 by lytic polysaccharide monooxygenases. ACS Catal. 10, 12760–12769 (2020).
Kim, B. et al. Fenton-like chemistry by a copper(I) complex and H2O2 relevant to enzyme peroxygenase C–H hydroxylation. J. Am. Chem. Soc. 145, 11735–11744 (2023).
Bertini, L. et al. Catalytic mechanism of fungal lytic polysaccharide monooxygenases investigated by first-principles calculations. Inorg. Chem. 57, 86–97 (2018).
Wang, B., Walton, P. H. & Rovira, C. Molecular mechanisms of oxygen activation and hydrogen peroxide formation in lytic polysaccharide monooxygenases. ACS Catal. 9, 4958–4969 (2019).
Singh, R. K. et al. Detection and characterization of a novel copper-dependent intermediate in a lytic polysaccharide monooxygenase. Chem. Eur. J. 26, 454–463 (2020).
Paradisi, A. et al. Formation of a copper(II)-tyrosyl complex at the active site of lytic polysaccharide monooxygenases following oxidation by H2O2. J. Am. Chem. Soc. 141, 18585–18599 (2019).
Mcevoy, A. et al. The role of the active site tyrosine in the mechanism of lytic polysaccharide monooxygenase. Chem. Sci. 12, 352 (2021).
Gray, H. B. & Winkler, J. R. Hole hopping through tyrosine/tryptophan chains protects proteins from oxidative damage. Proc. Natl Acad. Sci. USA 112, 10920–10925 (2015).
Bissaro, B. & Eijsink, V. G. H. Lytic polysaccharide monooxygenases: enzymes for controlled and site-specific Fenton-like chemistry. Essays Biochem. 18, 575–584 (2023).
Beeson, W. T., Vu, V. V., Span, E. A., Phillips, C. M. & Marletta, M. A. Cellulose degradation by polysaccharide monooxygenases. Annu. Rev. Biochem. 84, 923–946 (2015).
Westereng, B. et al. Simultaneous analysis of C1 and C4 oxidized oligosaccharides, the products of lytic polysaccharide monooxygenases acting on cellulose. J. Chromatogr. A 1445, 46–54 (2016).
Kojima, Y. et al. A lytic polysaccharide monooxygenase with broad xyloglucan specificity from the brown-rot fungus Gloeophyllum trabeum and its action on cellulose-xyloglucan complexes. Appl. Environ. Microbiol. 82, 6557–6572 (2016).
Vu, V. V., Beeson, W. T., Phillips, C. M., Cate, J. H. D. & Marletta, M. A. Determinants of regioselective hydroxylation in the fungal polysaccharide monooxygenases. J. Am. Chem. Soc. 136, 562–565 (2014).
Caldararu, O., Oksanen, E., Ryde, U. & Hedegård, E. D. Mechanism of hydrogen peroxide formation by lytic polysaccharide monooxygenase. Chem. Sci. 10, 576–586 (2019).
Span, E. A., Suess, D. L. M., Deller, M. C., Britt, R. D. & Marletta, M. A. The role of the secondary coordination sphere in a fungal polysaccharide monooxygenase. ACS Chem. Biol. 12, 1095–1103 (2017).
Sun, P. et al. AA16 oxidoreductases boost cellulose-active AA9 lytic polysaccharide monooxygenases from Myceliophthora thermophila. ACS Catal. 13, 4454–4467 (2023).
Li, F. et al. A lytic polysaccharide monooxygenase from a white-rot fungus drives the degradation of lignin by a versatile peroxidase. Appl. Environ. Microbiol. 85, e02803–e02818 (2019).
Breslmayr, E. et al. A fast and sensitive activity assay for lytic polysaccharide monooxygenase. Biotechnol. Biofuels 11, 79 (2018).
Loose, J. S. M. et al. Multipoint precision binding of substrate protects lytic polysaccharide monooxygenases from self-destructive off-pathway processes. Biochemistry 57, 4114–4124 (2018).
Petrović, D. M. et al. Methylation of the N-terminal histidine protects a lytic polysaccharide monooxygenase from auto-oxidative inactivation. Protein Sci. 27, 1636–1650 (2018).
Torbjörnsson, M., Hagemann, M. M., Ryde, U. & Donovan, H. E. Histidine oxidation in lytic polysaccharide monooxygenase. J. Biol. Inorg. Chem. 28, 317–328 (2023).
Winterbourn, C. C. & Metodiewa, D. Reactivity of biologically important thiol compounds with superoxide and hydrogen peroxide. Free Radic. Biol. Med. 27, 322–328 (1999).
Winterbourn, C. C. The biological chemistry of hydrogen peroxide. Methods Enzymol. 528, 3–25 (2013).
Fenton, H. J. H. Oxidation of tartaric acid in presence of iron. J. Chem. Soc. Trans. 65, 899–910 (1894).
Haber, F. & Weiss, J. Über die katalyse des hydroperoxydes. Naturwissenschaften 20, 948–950 (1932).
Hussain, S., Aneggi, E., Trovarelli, A. & Goi, D. Heterogeneous Fenton-like oxidation of ibuprofen over zirconia-supported iron and copper catalysts: effect of process variables. J. Water Process Eng. 44, 102343 (2021).
Burek, B. O., Bormann, S., Hollmann, F., Bloh, J. Z. & Holtmann, D. Hydrogen peroxide driven biocatalysis. Green Chem. 21, 3232–3249 (2019).
Picard, M. et al. Metal-free bacterial haloperoxidases as unusual hydrolases: activation of H2O2 by the formation of peracetic acid. Angew. Chem. Int. Ed. Engl. 36, 1196–1199 (1997).
Holtmann, D. & Hollmann, F. The oxygen dilemma: a severe challenge for the application of monooxygenases? ChemBioChem 17, 1391–1398 (2016).
Krest, C. M. et al. Reactive intermediates in cytochrome P450 catalysis. J. Biol. Chem. 288, 17074 (2013).
Hobisch, M. et al. Recent developments in the use of peroxygenases — exploring their high potential in selective oxyfunctionalisations. Biotechnol. Adv. 51, 107615 (2021).
Dawson, J. H. Probing structure-function relations in heme-containing oxygenases and peroxidases. Science 240, 433–439 (1988).
Matsunaga, I. & Shiro, Y. Peroxide-utilizing biocatalysts: structural and functional diversity of heme-containing enzymes. Curr. Opin. Chem. Biol. 8, 127–132 (2004).
Rydberg, P., Sigfridsson, E. & Ryde, U. On the role of the axial ligand in heme proteins: a theoretical study. J. Biol. Inorg. Chem. 9, 203–223 (2004).
Faiza, M., Huang, S., Lan, D. & Wang, Y. New insights on unspecific peroxygenases: superfamily reclassification and evolution. BMC Evol. Biol. 19, 76 (2019).
De Montellano, P. R. O. (ed.) Cytochrome P450: Structure, Mechanism, and Biochemistry 3rd edn (Springer, 2005).
Munro, A. W., Girvan, H. M., Mason, A. E., Dunford, A. J. & McLean, K. J. What makes a P450 tick? Trends Biochem. Sci. 38, 140–150 (2013).
Cirino, P. C. & Arnold, F. H. Regioselectivity and activity of cytochrome P450 BM-3 and mutant F87A in reactions driven by hydrogen peroxide. Adv. Synth. Catal. 344, 932–937 (2002).
Matsumura, H. et al. Modulation of redox potential and alteration in reactivity via the peroxide shunt pathway by mutation of cytochrome P450 around the proximal heme ligand. Biochemistry 47, 4834–4842 (2008).
Renneberg, R., Scheller, F., Ruckpaul, K., Pirrwitz, J. & Mohr, P. NADPH and H2O2-dependent reactions of cytochrome P-450LM compared with peroxidase catalysis. FEBS Lett. 96, 349–353 (1978).
Shumyantseva, V. V. et al. N-Terminal truncated cytochrome P450 2B4: catalytic activities and reduction with alternative electron sources. Biochem. Biophys. Res. Commun. 263, 678–680 (1999).
Walton, P. H., Davies, G. J., Diaz, D. E. & Franco-Cairo, J. P. The histidine brace: nature’s copper alternative to haem? FEBS Lett. 597, 485–494 (2023).
Poulos, T. L. & Kraut, J. The stereochemistry of peroxidase catalysis. J. Biol. Chem. 255, 8199–8205 (1980).
Derat, E. & Shaik, S. The Poulos–Kraut mechanism of compound I formation in horseradish peroxidase: a QM/MM. J. Phys. Chem. B 110, 10526–10533 (2006).
Wang, B., Zhang, X., Fang, W., Rovira, C. & Shaik, S. How do metalloproteins tame the Fenton reaction and utilize •OH radicals in constructive manners? Acc. Chem. Res. 55, 2280–2290 (2022).
Wang, B., Li, C., Dubey, K. D. & Shaik, S. Quantum mechanical/molecular mechanical calculated reactivity networks reveal how cytochrome P450cam and its T252A mutant select their oxidation pathways. J. Am. Chem. Soc. 137, 7379–7390 (2015).
Wang, B. et al. How do enzymes utilize reactive OH radicals? lessons from nonheme HppE and Fenton systems. J. Am. Chem. Soc. 138, 8489–8496 (2016).
Wang, C. et al. Evidence that the fosfomycin-producing epoxidase, HppE, is a non-heme-iron peroxidase. Science 342, 991–995 (2013).
Brander, S. et al. Biochemical evidence of both copper chelation and oxygenase activity at the histidine brace. Sci. Rep. 10, 16369 (2020).
Ipsen, J. et al. Copper binding and reactivity at the histidine brace motif: insights from mutational analysis of the Pseudomonas fluorescens copper chaperone CopC. FEBS Lett. 595, 1708–1720 (2021).
Quist, D. A., Diaz, D. E., Liu, J. J. & Karlin, K. D. Activation of dioxygen by copper metalloproteins and insights from model complexes. J. Biol. Inorg. Chem. 22, 253–288 (2017).
Castillo, I. et al. Cellulose depolymerization with LPMO-inspired Cu complexes. ChemCatChem 13, 4700–4704 (2021).
Bouchey, C. J., Shopov, D. Y., Gruen, A. D. & Tolman, W. B. Mimicking the Cu active site of lytic polysaccharide monooxygenase using monoanionic tridentate N-donor ligands. ACS Omega 7, 35217–35232 (2022).
Concia, A. L. et al. Copper complexes as bioinspired models for lytic polysaccharide monooxygenases. Inorg. Chem. 56, 1023–1026 (2017).
Costa, T. H. F. et al. Demonstration-scale enzymatic saccharification of sulfite-pulped spruce with addition of hydrogen peroxide for LPMO activation. Biofuels Bioprod. Bioref. 14, 734–745 (2020).
Kommedal, E. G. et al. Visible light-exposed lignin facilitates cellulose solubilization by lytic polysaccharide monooxygenases. Nat. Commun. 14, 1063 (2023).
Askarian, F. et al. The lytic polysaccharide monooxygenase CbpD promotes Pseudomonas aeruginosa virulence in systemic infection. Nat. Commun. 12, 1230 (2021).
Acknowledgements
We thank the current and past members of our research groups that have contributed to LPMO research. We also thank our colleagues from the LPMO community. We apologize in advance to all investigators whose research cannot be appropriately discussed and cited owing to space limitations. A.M. and B.B. disclose support for the research of this work from INRAE (EvoFun project PAF_02). V.G.H.E. discloses support for the research of this work from the European Research Council (grant number 856446).
Author information
Authors and Affiliations
Contributions
All authors contributed substantially to discussion of the content. A.M. and B.B. wrote the first draft of the article. All authors reviewed and edited the manuscript before submission.
Corresponding author
Ethics declarations
Competing interests
The authors declare no competing interests.
Peer review
Peer review information
Nature Reviews Chemistry thanks William DeGrado, Erik Hedegård and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.
Additional information
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Springer Nature or its licensor (e.g. a society or other partner) holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.
About this article
Cite this article
Munzone, A., Eijsink, V.G.H., Berrin, JG. et al. Expanding the catalytic landscape of metalloenzymes with lytic polysaccharide monooxygenases. Nat Rev Chem 8, 106–119 (2024). https://doi.org/10.1038/s41570-023-00565-z
Accepted:
Published:
Issue Date:
DOI: https://doi.org/10.1038/s41570-023-00565-z