A fungal family of lytic polysaccharide monooxygenase-like copper proteins


Lytic polysaccharide monooxygenases (LPMOs) are copper-containing enzymes that play a key role in the oxidative degradation of various biopolymers such as cellulose and chitin. While hunting for new LPMOs, we identified a new family of proteins, defined here as X325, in various fungal lineages. The three-dimensional structure of X325 revealed an overall LPMO fold and a His brace with an additional Asp ligand to Cu(II). Although LPMO-type activity of X325 members was initially expected, we demonstrated that X325 members do not perform oxidative cleavage of polysaccharides, establishing that X325s are not LPMOs. Investigations of the biological role of X325 in the ectomycorrhizal fungus Laccaria bicolor revealed exposure of the X325 protein at the interface between fungal hyphae and tree rootlet cells. Our results provide insights into a family of copper-containing proteins, which is widespread in the fungal kingdom and is evolutionarily related to LPMOs, but has diverged to biological functions other than polysaccharide degradation.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Fig. 1: Phylogeny of the X325 family.
Fig. 2: Structure of X325 and configuration of the copper active site.
Fig. 3: LaX325 structural and biochemical features do not support LPMO-type activity.
Fig. 4: Immunolocalization of LbX325 in L. bicolor.

Data availability

LaX325 nucleotide sequence was deposited in GenBank under accession number MK088083. The X-ray structures of LaX325 were deposited in the Protein Data Bank under accession numbers 6IBH, 6IBI and 6IBJ. Raw EPR data are available on request through the Research Data York (https://doi.org/10.15124/a034974e-2782-415e-8b02-2b6e4098760e).


  1. 1.

    Vaaje-Kolstad, G. et al. An oxidative enzyme boosting the enzymatic conversion of recalcitrant polysaccharides. Science 330, 219–222 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  2. 2.

    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).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. 3.

    Horn, S. J., Vaaje-Kolstad, G., Westereng, B. & Eijsink, V. G. Novel enzymes for the degradation of cellulose. Biotechnol. Biofuels 5, 45 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. 4.

    Bissaro, B. et al. Oxidative cleavage of polysaccharides by monocopper enzymes depends on H2O2. Nat. Chem. Biol. 13, 1123 (2017).

    CAS  PubMed  Google Scholar 

  5. 5.

    Johansen, K. S. Discovery and industrial applications of lytic polysaccharide mono-oxygenases. Biochem. Soc. Trans. 44, 143–149 (2016).

    CAS  PubMed  Google Scholar 

  6. 6.

    Bennati-Granier, C. et al. Substrate specificity and regioselectivity of fungal AA9 lytic polysaccharide monooxygenases secreted by Podospora anserina. Biotechnol. Biofuels 8, 90 (2015).

    PubMed  PubMed Central  Google Scholar 

  7. 7.

    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).

    PubMed  PubMed Central  Google Scholar 

  8. 8.

    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).

    CAS  Google Scholar 

  9. 9.

    Gilbert, H. J., Knox, J. P. & Boraston, A. B. Advances in understanding the molecular basis of plant cell wall polysaccharide recognition by carbohydrate-binding modules. Curr. Opin. Struc. Biol. 23, 669–677 (2013).

    CAS  Google Scholar 

  10. 10.

    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).

    CAS  PubMed  Google Scholar 

  11. 11.

    Lo Leggio, L. et al. Structure and boosting activity of a starch-degrading lytic polysaccharide monooxygenase. Nat. Commun. 6, 5961 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  12. 12.

    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).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. 13.

    Sabbadin, F. et al. An ancient family of lytic polysaccharide monooxygenases with roles in arthropod development and biomass digestion. Nat. Commun. 9, 756 (2018).

    PubMed  PubMed Central  Google Scholar 

  14. 14.

    Couturier, M. et al. Lytic xylan oxidases from wood-decay fungi unlock biomass degradation. Nat. Chem. Biol. 14, 306 (2018).

    CAS  PubMed  Google Scholar 

  15. 15.

    Filiatrault-Chastel, C. et al. AA16, a new lytic polysaccharide monooxygenase family identified in fungal secretomes. Biotech. Biofuels 12, 55 (2019).

    Google Scholar 

  16. 16.

    Berrin, J.-G., Rosso, M.-N. & Abou Hachem, M. Fungal secretomics to probe the biological functions of lytic polysaccharide monooxygenases. Carbohydr. Res. 448, 155–160 (2017).

    CAS  PubMed  Google Scholar 

  17. 17.

    Navarro, D. et al. Fast solubilization of recalcitrant cellulosic biomass by the basidiomycete fungus Laetisaria arvalis involves successive secretion of oxidative and hydrolytic enzymes. Biotechnol. Biofuels 7, 143 (2014).

    PubMed  PubMed Central  Google Scholar 

  18. 18.

    Kelley, L. A., Mezulis, S., Yates, C. M., Wass, M. N. & Sternberg, M. J. E. The Phyre2 web portal for protein modeling, prediction and analysis. Nat. Protoc. 10, 845–858 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. 19.

    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).

    CAS  PubMed  Google Scholar 

  20. 20.

    Low, M. & Saltiel, A. Structural and functional roles of glycosyl-phosphatidylinositol in membranes. Science 239, 268–275 (1988).

    CAS  PubMed  Google Scholar 

  21. 21.

    Berbee, M. L., James, T. Y. & Strullu-Derrien, C. Early diverging fungi: diversity and impact at the dawn of terrestrial life. Annu. Rev. Microbiol. 71, 41–60 (2017).

    CAS  PubMed  Google Scholar 

  22. 22.

    Tandrup, T., Frandsen, K. E. H., Johansen, K. S., Berrin, J.-G. & Lo Leggio, L. Recent insights into lytic polysaccharide monooxygenases (LPMOs). Biochem. Soc. Trans. 46, 1431–1447 (2018).

    CAS  PubMed  Google Scholar 

  23. 23.

    Holm, L. & Laakso, L. M. Dali server update. Nucleic Acids Res. 44, W351–W355 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  24. 24.

    Vu, V. V. & Ngo, S. T. Copper active site in polysaccharide monooxygenases. Coord. Chem. Rev. 368, 134–157 (2018).

    CAS  Google Scholar 

  25. 25.

    Chiu, E. et al. Structural basis for the enhancement of virulence by viral spindles and their in vivo crystallization. Proc. Natl Acad. Sci. USA 112, 3973–3978 (2015).

    CAS  PubMed  Google Scholar 

  26. 26.

    Tan, T.-C. et al. Structural basis for cellobiose dehydrogenase action during oxidative cellulose degradation. Nat. Commun. 6, 7542 (2015).

    PubMed  PubMed Central  Google Scholar 

  27. 27.

    Lawton, T. J., Kenney, G. E., Hurley, J. D. & Rosenzweig, A. C. The CopC family: structural and bioinformatic insights into a diverse group of periplasmic copper binding proteins. Biochemistry 55, 2278–2290 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. 28.

    Ross, M. O. et al. Particulate methane monooxygenase contains only mononuclear copper centers. Science 364, 566–570 (2019).

    CAS  PubMed  PubMed Central  Google Scholar 

  29. 29.

    Liew, E. F., Tong, D., Coleman, N. V. & Holmes, A. J. Mutagenesis of the hydrocarbon monooxygenase indicates a metal centre in subunit-C, and not subunit-B, is essential for copper-containing membrane monooxygenase activity. Microbiology 160, 1267–1277 (2014).

    CAS  PubMed  Google Scholar 

  30. 30.

    Cao, L., Caldararu, O., Rosenzweig, A. C. & Ryde, U. Quantum refinement does not support dinuclear copper sites in crystal structures of particulate methane monooxygenase. Angew. Chem. Int. Ed. Eng. 57, 162–166 (2018).

    CAS  Google Scholar 

  31. 31.

    Peisach, J. & Blumberg, W. E. Structural implications derived from the analysis of electron paramagnetic resonance spectra of natural and artificial copper proteins. Arch. Biochem. Biophys. 165, 691–708 (1974).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. 32.

    Kuusk, S. et al. Kinetics of H2O2-driven degradation of chitin by a bacterial lytic polysaccharide monooxygenase. J. Biol. Chem. 293, 523–531 (2018).

    CAS  PubMed  Google Scholar 

  33. 33.

    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).

    CAS  PubMed  Google Scholar 

  34. 34.

    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).

    CAS  PubMed  Google Scholar 

  35. 35.

    Bey, M. et al. Cello-oligosaccharide oxidation reveals differences between two lytic polysaccharide monooxygenases (family GH61) from Podospora anserina. Appl. Environ. Microbiol. 79, 488–496 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  36. 36.

    Chen, K. H. et al. Bacteriohemerythrin bolsters the activity of the particulate methane monooxygenase (pMMO) in Methylococcus capsulatus (Bath). J. Inorg. Biochem. 111, 10–17 (2012).

    CAS  PubMed  Google Scholar 

  37. 37.

    Bernardes et al. Carbohydrate binding modules enhance cellulose enzymatic hydrolysis by increasing access of cellulases to the substrate. Carbohydr. Polym. 211, 57–68 (2019).

    CAS  PubMed  Google Scholar 

  38. 38.

    Shah, F. et al. Ectomycorrhizal fungi decompose soil organic matter using oxidative mechanisms adapted from saprotrophic ancestors. New Phytol. 209, 1705–1719 (2016).

    CAS  PubMed  Google Scholar 

  39. 39.

    Balestrini R. M. & Kottke I. In Molecular Mycorrhizal Symbiosis (ed. Martin, F.) 47–62 (John Wiley & Sons, 2016).

  40. 40.

    Balestrini, R., Hahn, M. G., Faccio, A., Mendgen, K. & Bonfante, P. Differential localization of carbohydrate epitopes in plant cell walls in the presence and absence of arbuscular mycorrhizal fungi. Plant Physiol. 111, 203–213 (1996).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. 41.

    Balestrini, R. & Bonfante, P. Cell wall remodeling in mycorrhizal symbiosis: a way towards biotrophism. Front. Plant Sci. 5, 237 (2014).

    PubMed  PubMed Central  Google Scholar 

  42. 42.

    Yamazaki, H., Tanaka, A., Kaneko, J.-i, Ohta, A. & Horiuchi, H. Aspergillus nidulans ChiA is a glycosylphosphatidylinositol (GPI)-anchored chitinase specifically localized at polarized growth sites. Fungal Genet. Biol. 45, 963–972 (2008).

    CAS  PubMed  Google Scholar 

  43. 43.

    Nakajima, M., Yamashita, T., Takahashi, M., Nakano, Y. & Takeda, T. A novel glycosylphosphatidylinositol-anchored glycoside hydrolase from Ustilago esculenta functions in β-1,3-glucan degradation. Appl. Environ. Microbiol. 78, 5682–5689 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  44. 44.

    Smith, H. W. & Marshall, C. J. Regulation of cell signalling by uPAR. Nat. Rev. Mol. Cell Biol. 11, 23–36 (2010).

    CAS  PubMed  Google Scholar 

  45. 45.

    Gaggelli, E., Kozlowski, H., Valensin, D. & Valensin, G. Copper homeostasis and neurodegenerative disorders (Alzheimer’s, prion, and Parkinson’s diseases and amyotrophic lateral sclerosis). Chem. Rev. 106, 1995–2044 (2006).

    CAS  PubMed  Google Scholar 

  46. 46.

    García-Santamarina, S. & Thiele, D. J. Copper at the fungal pathogen–host axis. J. Biol. Chem. 290, 18945–18953 (2015).

    PubMed  PubMed Central  Google Scholar 

  47. 47.

    Bolchi, A. et al. Genome-wide inventory of metal homeostasis-related gene products including a functional phytochelatin synthase in the hypogeous mycorrhizal fungus Tuber melanosporum. Fungal Genet. Biol. 48, 573–584 (2011).

    CAS  PubMed  Google Scholar 

  48. 48.

    Lalucque, H. et al. IDC2 and IDC3, two genes involved in cell non-autonomous signaling of fruiting body development in the model fungus Podospora anserina. Dev. Biol. 421, 126–138 (2017).

    CAS  PubMed  Google Scholar 

  49. 49.

    Green, K. A. et al. SymB and SymC, two membrane associated proteins, are required for Epichloë festucae hyphal cell–cell fusion and maintenance of a mutualistic interaction with Lolium perenne. Mol. Microbiol. 103, 657–677 (2016).

    PubMed  Google Scholar 

  50. 50.

    García-Santamarina, S. et al. A lytic polysaccharide monooxygenase-like protein functions in copper import and fungal meningitis. Nat. Chem. Biol. https://doi.org/10.1038/s41589-019-0437-9.

  51. 51.

    Kohler, A. et al. Convergent losses of decay mechanisms and rapid turnover of symbiosis genes in mycorrhizal mutualists. Nat. Genet. 47, 410–415 (2015).

    CAS  PubMed  Google Scholar 

  52. 52.

    Altschul, S. F., Gish, W., Miller, W., Myers, E. W. & Lipman, D. J. Basic local alignment search tool. J. Mol. Biol. 215, 403–410 (1990).

    CAS  Google Scholar 

  53. 53.

    Edgar, R. C. MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Res. 32, 1792–1797 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. 54.

    Lemoine, F. et al. Renewing Felsenstein’s phylogenetic bootstrap in the era of big data. Nature 556, 452–456 (2018).

    CAS  PubMed  PubMed Central  Google Scholar 

  55. 55.

    Huson, D. H. & Scornavacca, C. Dendroscope 3: an interactive tool for rooted phylogenetic trees and networks. Syst. Biol. 61, 1061–1067 (2012).

    PubMed  Google Scholar 

  56. 56.

    Haon, M. et al. Recombinant protein production facility for fungal biomass-degrading enzymes using the yeast Pichia pastoris. Front. Microbiol. 6, 1002 (2015).

    PubMed  PubMed Central  Google Scholar 

  57. 57.

    Pitarch, A., Sánchez, M., Nombela, C. & Gil, C. Sequential fractionation and two-dimensional gel analysis unravels the complexity of the dimorphic fungus Candida albicans cell wall proteome. Mol. Cell. Proteom. 1, 967–982 (2002).

    CAS  Google Scholar 

  58. 58.

    Felten, J. et al. The ectomycorrhizal fungus Laccaria bicolor stimulates lateral root formation in poplar and Arabidopsis through auxin transport and signaling. Plant Physiol. 151, 1991–2005 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  59. 59.

    Martin, F. et al. The genome of Laccaria bicolor provides insights into mycorrhizal symbiosis. Nature 452, 88–92 (2008).

    CAS  PubMed  Google Scholar 

  60. 60.

    Gorrec, F. The MORPHEUS protein crystallization screen. J. Appl. Crystallogr. 42, 1035–1042 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  61. 61.

    Terwilliger, T. C. et al. Decision-making in structure solution using Bayesian estimates of map quality: the PHENIX AutoSol wizard. Acta Cryst. D Biol. Crystallogr. 65, 582–601 (2009).

    CAS  Google Scholar 

  62. 62.

    Kabsch, W. XDS. Acta Crystallogr. D Biol. Crystallogr. 66, 125–132 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. 63.

    Vagin, A. & Teplyakov, A. Molecular replacement with MOLREP. Acta Crystallogr. D Biol. Crystallogr. 66, 22–25 (2010).

    CAS  PubMed  Google Scholar 

  64. 64.

    Murshudov, G. N. et al. REFMAC5 for the refinement of macromolecular crystal structures. Acta Crystallogr. D Biol. Crystallogr. 67, 355–367 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  65. 65.

    Zeldin, O. B., Gerstel, M. & Garman, E. F. RADDOSE-3D: time- and space-resolved modelling of dose in macromolecular crystallography. J. Appl. Crystallogr. 46, 1225–1230 (2013).

    CAS  Google Scholar 

Download references


A.L. was funded by a Marie Curie Individual Fellowship within the Horizon 2020 Research and Innovation Framework Programme (748758). The Danish Ministry of Higher Education and Science through the Instrument Center DANSCATT funded travel to synchrotrons. We would also like to acknowledge MAX-IV, Lund, Sweden, the ESRF, Grenoble, France and DESY, Hamburg, Germany for synchrotron beamtime and related assistance. We thank J.-C. Poulsen for technical assistance; A. Kohler and D. Thiele for helpful discussions; F. Chaspoul for assistance with ICP-MS analyses and R. Balestrini (Institute for Sustainable Plant Protection, Italy) for providing gold-labeled WGA lectin. L.L.L. and T.T. thank the Novo Nordisk Foundation for funding through grant NF17SA0027704. K.E.H.F. thanks the Carlsberg Foundation through an Internationalization Postdoc Fellowship (grants CF16-0673 and CF17-0533) and the EU, framework of the Marie Curie FP7 COFUND People Programme (AgreenSkills+ fellowship 609398) for financial support. L.L.L., T.T. and K.E.H.F. are members of ISBUC Integrative Structural Biology at the University of Copenhagen (https://isbuc.ku.dk/). K.S.J. thanks the Novo Nordisk Foundation for funding through grant NNF17SA0027704. P.H.W. and L.C. thank the UK Biotechnology and Biological Sciences Research Council (BB/L021633/1) for funding. F.M. is funded by the French National Research Agency through the Laboratory of Excellence Advanced Research on the Biology of Tree and Forest Ecosystems (grant ANR-11-LABX 0002 01). A.Z. thanks the French Embassy in Greece, together with the French Ministry of Higher Education, Research and Innovation, for the scholarship ‘Séjour scientifique de haut niveau’ (SSHN). The electron microscopy experiments were performed on the PiCSL-FBI core facility (IBDM, Marseille), member of the France-BioImaging national research infrastructure (ANR-10-INBS-04).

Author information




A.L., D.N. and M.-N.R. identified the new proteins. A.L. and B.H. performed bioinformatic analyses. A.L., S.G. and M.H. performed recombinant protein production and purification. T.T. and K.E.H.F. crystallized LaX325, determined and analyzed the X-ray crystal structure. T.T. collected X-ray data. K.E.H.F. made relevant figures and tables. L.L.L. directed the crystallographic studies. K.E.H.F., K.S.J. and L.L.L. analyzed the structure. K.E.H.F. and L.L.L. drafted relevant parts of the manuscript. A.L., B.B. and A.Z. performed enzyme assays. M.F. and D.R. performed mass spectrometry analyses. L.C. and P.H.W. conceived and carried out the EPR study. A.L. and F.Z. performed confocal microscopy under the supervision of F.M. A.L. and N.B. performed transmission electron microscopy. J.-G.B. coordinated the work. A.L. and J.-G.B. organized the data and drafted the manuscript. All authors made comments on the manuscript and approved the final version.

Corresponding author

Correspondence to Jean-Guy Berrin.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Supplementary Information

Supplementary Tables 1–6 and Supplementary Figs. 1–9.

Reporting Summary

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Labourel, A., Frandsen, K.E.H., Zhang, F. et al. A fungal family of lytic polysaccharide monooxygenase-like copper proteins. Nat Chem Biol 16, 345–350 (2020). https://doi.org/10.1038/s41589-019-0438-8

Download citation

Further reading


Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing