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Structures of mouse DUOX1–DUOXA1 provide mechanistic insights into enzyme activation and regulation


DUOX1, an NADPH oxidase family member, catalyzes the production of hydrogen peroxide. DUOX1 is expressed in various tissues, including the thyroid and respiratory tract, and plays a crucial role in processes such as thyroid hormone biosynthesis and innate host defense. DUOX1 co-assembles with its maturation factor DUOXA1 to form an active enzyme complex. However, the molecular mechanisms for activation and regulation of DUOX1 remain mostly unclear. Here, I present cryo-EM structures of the mammalian DUOX1–DUOXA1 complex, in the absence and presence of substrate NADPH, as well as DUOX1–DUOXA1 in an unexpected dimer-of-dimers configuration. These structures reveal atomic details of the DUOX1-DUOXA1 interaction, a lipid-mediated NADPH-binding pocket and the electron transfer path. Furthermore, biochemical and structural analyses indicate that the dimer-of-dimers configuration represents an inactive state of DUOX1–DUOXA1, suggesting an oligomerization-dependent regulatory mechanism. Together, my work provides structural bases for DUOX1–DUOXA1 activation and regulation.

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Fig. 1: Structure of DUOX1–DUOXA1 in the absence of NADPH.
Fig. 2: Interaction between DUOX1 and DUOXA1.
Fig. 3: Heme- and FAD-binding sites.
Fig. 4: The NADPH-binding site and electron transfer path.
Fig. 5: Structure of DUOX1–DUOXA1 in the dimer-of-dimers configuration.
Fig. 6: The dimer-of-dimers conformation of DUOX1–DUOXA1 represents an inactivated state.

Data availability

Cryo-EM maps and atomic models for mouse DUOX1–DUOXA1 complexes have been deposited in the EMDB and wwPDB with the following accession numbers: EMD-21962 and PDB 6WXR (apo state); EMD-21963 and PDB 6WXU (dimer of dimers); EMD-21964 and PDB 6WXV (with NADPH).


  1. 1.

    Devasagayam, T. P. et al. Free radicals and antioxidants in human health: current status and future prospects. J. Assoc. Physicians India 52, 794–804 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  2. 2.

    Bedard, K. & Krause, K. H. The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology. Physiol. Rev. 87, 245–313 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. 3.

    Vlahos, R. et al. Inhibition of Nox2 oxidase activity ameliorates influenza A virus-induced lung inflammation. PLoS Pathog. 7, e1001271 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  4. 4.

    Khomich, O. A., Kochetkov, S. N., Bartosch, B. & Ivanov, A. V. Redox biology of respiratory viral infections. Viruses 10, 392 (2018).

    Google Scholar 

  5. 5.

    Imai, Y. et al. Identification of oxidative stress and Toll-like receptor 4 signaling as a key pathway of acute lung injury. Cell 133, 235–249 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  6. 6.

    De Deken, X., Corvilain, B., Dumont, J. E. & Miot, F. Roles of DUOX-mediated hydrogen peroxide in metabolism, host defense and signaling. Antioxid. Redox Signal. 20, 2776–2793 (2014).

    PubMed  Google Scholar 

  7. 7.

    Brandes, R. P., Weissmann, N. & Schroder, K. Nox family NADPH oxidases: molecular mechanisms of activation. Free Radic. Biol. Med. 76, 208–226 (2014).

    CAS  PubMed  Google Scholar 

  8. 8.

    Dupuy, C. et al. Purification of a novel flavoprotein involved in the thyroid NADPH oxidase. Cloning of the porcine and human cDNAs. J. Biol. Chem. 274, 37265–37269 (1999).

    CAS  PubMed  Google Scholar 

  9. 9.

    Leseney, A. M. et al. Biochemical characterization of a Ca2+/NAD(P)H-dependent H2O2 generator in human thyroid tissue. Biochimie 81, 373–380 (1999).

    CAS  PubMed  Google Scholar 

  10. 10.

    De Deken, X. et al. Cloning of two human thyroid cDNAs encoding new members of the NADPH oxidase family. J. Biol. Chem. 275, 23227–23233 (2000).

    PubMed  Google Scholar 

  11. 11.

    Ameziane-El-Hassani, R., Schlumberger, M. & Dupuy, C. NADPH oxidases: new actors in thyroid cancer? Nat. Rev. Endocrinol. 12, 485–494 (2016).

    CAS  PubMed  Google Scholar 

  12. 12.

    Boots, A. W. et al. ATP-mediated activation of the NADPH oxidase DUOX1 mediates airway epithelial responses to bacterial stimuli. J. Biol. Chem. 284, 17858–17867 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  13. 13.

    Koff, J. L., Shao, M. X., Ueki, I. F. & Nadel, J. A. Multiple TLRs activate EGFR via a signaling cascade to produce innate immune responses in airway epithelium. Am. J. Physiol. Lung Cell Mol. Physiol. 294, L1068–L1075 (2008).

    CAS  PubMed  Google Scholar 

  14. 14.

    Grasberger, H. & Refetoff, S. Identification of the maturation factor for dual oxidase. Evolution of an eukaryotic operon equivalent. J. Biol. Chem. 281, 18269–18272 (2006).

    CAS  PubMed  Google Scholar 

  15. 15.

    Luxen, S. et al. Heterodimerization controls localization of Duox–DuoxA NADPH oxidases in airway cells. J. Cell Sci. 122, 1238–1247 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  16. 16.

    Korzeniowska, A., Donko, A. P., Morand, S. & Leto, T. L. Functional characterization of DUOX enzymes in reconstituted cell models. Methods Mol. Biol. 1982, 173–190 (2019).

    CAS  PubMed  Google Scholar 

  17. 17.

    Ameziane-El-Hassani, R. et al. Dual oxidase-2 has an intrinsic Ca2+-dependent H2O2-generating activity. J. Biol. Chem. 280, 30046–30054 (2005).

    CAS  PubMed  Google Scholar 

  18. 18.

    Rigutto, S. et al. Activation of dual oxidases Duox1 and Duox2: differential regulation mediated by cAMP-dependent protein kinase and protein kinase C-dependent phosphorylation. J. Biol. Chem. 284, 6725–6734 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  19. 19.

    Deme, D., Virion, A., Hammou, N. A. & Pommier, J. NADPH-dependent generation of H2O2 in a thyroid particulate fraction requires Ca2+. FEBS Lett. 186, 107–110 (1985).

    CAS  PubMed  Google Scholar 

  20. 20.

    Meitzler, J. L., Hinde, S., Banfi, B., Nauseef, W. M. & Ortiz de Montellano, P. R. Conserved cysteine residues provide a protein-protein interaction surface in dual oxidase (DUOX) proteins. J. Biol. Chem. 288, 7147–7157 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. 21.

    Singh, P. K. et al. Structure of bovine lactoperoxidase with a partially linked heme moiety at 1.98 Å resolution. Biochim. Biophys. Acta Proteins Proteom. 1865, 329–335 (2017).

    CAS  PubMed  Google Scholar 

  22. 22.

    Meitzler, J. L. & Ortiz de Montellano, P. R. Caenorhabditis elegans and human dual oxidase 1 (DUOX1) ‘peroxidase’ domains: insights into heme binding and catalytic activity. J. Biol. Chem. 284, 18634–18643 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. 23.

    Magnani, F. et al. Crystal structures and atomic model of NADPH oxidase. Proc. Natl Acad. Sci. USA 114, 6764–6769 (2017).

    CAS  PubMed  Google Scholar 

  24. 24.

    Varela, V. et al. Three mutations (p.Q36H, p.G418fsX482 and g.IVS19-2A>C) in the dual oxidase 2 gene responsible for congenital goiter and iodide organification defect. Clin. Chem. 52, 182–191 (2006).

    CAS  PubMed  Google Scholar 

  25. 25.

    Carre, A. et al. When an intramolecular disulfide bridge governs the interaction of DUOX2 with its partner DUOXA2. Antioxid. Redox Signal. 23, 724–733 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  26. 26.

    Louzada, R. A. et al. Conformation of the N-terminal ectodomain elicits different effects on DUOX function: a potential impact on congenital hypothyroidism caused by a H2O2 production defect. Thyroid 28, 1052–1062 (2018).

    CAS  PubMed  Google Scholar 

  27. 27.

    Ganasen, M. et al. Structural basis for promotion of duodenal iron absorption by enteric ferric reductase with ascorbate. Commun. Biol. 1, 120 (2018).

    PubMed  PubMed Central  Google Scholar 

  28. 28.

    Lu, P. et al. Structure and mechanism of a eukaryotic transmembrane ascorbate-dependent oxidoreductase. Proc. Natl Acad. Sci. USA 111, 1813–1818 (2014).

    CAS  PubMed  Google Scholar 

  29. 29.

    Hanukoglu, I. Proteopedia: Rossmann fold: a β-α-β fold at dinucleotide binding sites. Biochem Mol. Biol. Educ. 43, 206–209 (2015).

    CAS  PubMed  Google Scholar 

  30. 30.

    Balabin, I. A., Hu, X. & Beratan, D. N. Exploring biological electron transfer pathway dynamics with the Pathways plugin for VMD. J. Comput. Chem. 33, 906–910 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  31. 31.

    Beratan, D. N., Betts, J. N. & Onuchic, J. N. Protein electron transfer rates set by the bridging secondary and tertiary structure. Science 252, 1285–1288 (1991).

    CAS  PubMed  PubMed Central  Google Scholar 

  32. 32.

    Chovancova, E. et al. CAVER 3.0: a tool for the analysis of transport pathways in dynamic protein structures. PLoS Comput Biol. 8, e1002708 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  33. 33.

    Ueyama, T. et al. The extracellular A-loop of dual oxidases affects the specificity of reactive oxygen species release. J. Biol. Chem. 290, 6495–6506 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  34. 34.

    Zeng, J. & Fenna, R. E. X-ray crystal structure of canine myeloperoxidase at 3-Å resolution. J. Mol. Biol. 226, 185–207 (1992).

    CAS  Google Scholar 

  35. 35.

    Jurrus, E. et al. Improvements to the APBS biomolecular solvation software suite. Protein Sci. 27, 112–128 (2018).

    CAS  PubMed  Google Scholar 

  36. 36.

    Sun, J. & MacKinnon, R. Structural basis of human KCNQ1 modulation and gating. Cell 180, 340–347 (2020).

    CAS  Google Scholar 

  37. 37.

    Goehring, A. et al. Screening and large-scale expression of membrane proteins in mammalian cells for structural studies. Nat. Protoc. 9, 2574–2585 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  38. 38.

    Kirchhofer, A. et al. Modulation of protein properties in living cells using nanobodies. Nat. Struct. Mol. Biol. 17, 133–138 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  39. 39.

    Mastronarde, D. N. Automated electron microscope tomography using robust prediction of specimen movements. J. Struct. Biol. 152, 36–51 (2005).

    PubMed  PubMed Central  Google Scholar 

  40. 40.

    Tang, G. et al. EMAN2: an extensible image processing suite for electron microscopy. J. Struct. Biol. 157, 38–46 (2007).

    CAS  PubMed  Google Scholar 

  41. 41.

    Zheng, S. Q. et al. MotionCor2: anisotropic correction of beam-induced motion for improved cryo-electron microscopy. Nat. Methods 14, 331–332 (2017).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. 42.

    Zhang, K. Gctf: real-time CTF determination and correction. J. Struct. Biol. 193, 1–12 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  43. 43.

    Zivanov, J. et al. New tools for automated high-resolution cryo-EM structure determination in RELION-3. Elife 7, e42166 (2018).

    PubMed  PubMed Central  Google Scholar 

  44. 44.

    Scheres, S. H. RELION: implementation of a Bayesian approach to cryo-EM structure determination. J. Struct. Biol. 180, 519–530 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. 45.

    Punjani, A., Rubinstein, J. L., Fleet, D. J. & Brubaker, M. A. cryoSPARC: algorithms for rapid unsupervised cryo-EM structure determination. Nat. Methods 14, 290–296 (2017).

    CAS  Google Scholar 

  46. 46.

    Emsley, P., Lohkamp, B., Scott, W. G. & Cowtan, K. Features and development of Coot. Acta Crystallogr. D Biol. Crystallogr. 66, 486–501 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  47. 47.

    Yang, J. et al. The I-TASSER Suite: protein structure and function prediction. Nat. Methods 12, 7–8 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  48. 48.

    Yang, J. & Zhang, Y. I-TASSER server: new development for protein structure and function predictions. Nucleic Acids Res. 43, W174–W181 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  49. 49.

    Afonine, P. V., Grosse-Kunstleve, R. W., Adams, P. D. & Urzhumtsev, A. Bulk-solvent and overall scaling revisited: faster calculations, improved results. Acta Crystallogr. D Biol. Crystallogr. 69, 625–634 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  50. 50.

    Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D Biol. Crystallogr. 66, 12–21 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  51. 51.

    Pettersen, E. F. et al. UCSF Chimera—a visualization system for exploratory research and analysis. J. Comput. Chem. 25, 1605–1612 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

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I thank members of the Cryo-electron Microscopy and Tomography Center of St Jude Children’s Research Hospital for help with cryo-EM data collection; P. Hixson and R. Kalathur (Protein Technology Center) for help with mammalian cell culture; A. Myasnikov, M. Halic and C. Lee for helpful discussions; Z. Luo for help with bio-illustration; and C. Kalodimos, M. Halic and M. Babu for critical reading of the manuscript. J.S. is funded by the NIH (HL143037) and American Lebanese Syrian Associated Charities (ALSAC).

Author information




J.S. designed and performed all the experiments, analyzed the results and prepared the manuscript.

Corresponding author

Correspondence to Ji Sun.

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The author declares no competing interests.

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Peer review information Inês Chen was the primary editor on this article and managed its editorial process and peer review in collaboration with the rest of the editorial team.

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Extended data

Extended Data Fig. 1 Structure determination of the DUOX1–DUOXA1 complex.

a, Construct design of DUOX1 and DUOXA1 used for structural studies and size exclusion chromatography profile of the DUOX1–DUOXA1 complex. Fractions of second peak (PK2) in the red box are concentrated and used for single-particle analysis. b, Flowchart of DUOX1–DUOXA1 structure determination. The steps in blue are carried out in cryoSPARC, ones in green in RELION. In the 2D classification, dimer-of-dimer classes are indicated by red dashed cycles. c, Interaction between the S3-S4 linker and PHD of DUOX1. The out leaflet of the membrane bilayer is indicated by a gray line. d, Disulfide bonds and glycosylation sites of the DUOX1–DUOXA1 complex. Sugars and their linked Asn side chains are shown as sticks and balls. Disulfide bonds are colored in green.

Extended Data Fig. 2 Structure of the PHD of DUOX1.

a, Superimposition between LPO and PHD of DUOX1. LPO and DUOX1 are colored in gray and blue, respectively. b, Putative heme-binding site. The possible heme-binding pocket indicated by a red oval. Positions of Ser326 and Ser108 (colored in brown) are where the heme-coordinating histidines are located. c, The putative calcium-binding site and surrounding residues. Calcium is indicated by a green sphere. d, A potential ion binding site and surrounding residues. Cryo-EM density is contoured by gray meshes.

Extended Data Fig. 3 Heme- and FAD-binding sites.

a, Structural comparison of heme-binding sites between DUOX1 (blue) and csNOX5 (gray). The transmembrane helices are labeled with S1–S6. b, Structural conservation of heme coordination in ferric oxidoreductases. DUOX1, csNOX5, Cyto b561 and Dcytb are colored in blue, gray, light blue and cyan, respectively. c, FAD-binding site in 2D representation. Hydrogen bonds, hydrophobic interactions and cation-π interactions are indicated by dashes, spokes and vertical dashes, respectively. Residues from FBD, NBD, TMD are colored in light pink, gray and blue, respectively. d, The putative oxygen-binding site. Oxygen is represented by a dashed red oval. e, The possible oxygen entering and hydrogen peroxide exiting paths.

Extended Data Fig. 4 NADPH-binding site.

a, Cryo-EM density of DUOX1–DUOXA1 with and without NADPH. The potential NADPH-binding site is indicated by red dashes. b, Structure of an NADPH molecule. c, The NADPH-binding site. Residues from TMD and NBD are colored in blue and gray, respectively. The “invisible” nicotinamide group is cycled in a cyan dashed oval. d, Cartoon and structure of the NADPH-binding site. The conserved glycines on the “GXGXG” motif are shown as magenta balls. e, The lipid-binding pocket of csNOX5 and DUOX1. Lipid or alkyl chains are colored in red. f, Functional analyses of F1097 mutations. F1097 is mutated to Tyr, Ala, Ile and Val, and the activity of the mutations are normalized to the wild-type protein (Data shown are mean and s.d. of n = 4 independent experiments).

Extended Data Fig. 5 Formation of the dimer-of-dimer interface.

a, b, Structural comparison of DUOX1 and DUOXA1 in heterodimeric and dimer-of-dimer states. c, Interaction between DUOX1 and DUOXA1. Left: interactions between DUOX1 and the N-terminal loop and glycan chain linked to N109 of DUOXA1. Right: cryo-EM density and cartoon of the glycan chain on N109. d, Interactions between transmembrane domains of DUOX1 and DUOXA1 mediated by a lipid molecule. Left: interaction details. Right: density of the lipid molecule. e, PHD-PHD interactions in the dimer-of-dimer configuration. Left: interactions between PHDs of DUOX1. Right: comparison between MPO dimers and PHD dimers of DUOX1.

Extended Data Fig. 6 The interface between DUOX1–DUOXA1 heterodimers.

a, Modeling of two DUOX1–DUOXA1 dimers into the dimer-of-dimer state. Structural crashes are indicated by red arrows. b, The potential oxygen entering/hydrogen peroxide exiting paths in the DUOX1–DUOXA1 dimer of dimers. c, FSEC curves of mouse DUOX1–DUOXA1 (blue), human DUOX1–DUOXA1 (gray) and human DUOX2DUOXA2 (orange). d, FSEC curves of mouse DUOX1–DUOXA1 (gray), mouse DUOX1–DUOXA1 with NADPH (green) and mouse DUOX1–DUOXA1 with FAD (orange). e, Modeling of DUOX2DUOXA2 complex and mapping of the hypothyroidism disease mutations. f, Accessibility of the outer heme of csNOX5 to extracellular space. The heme molecule is colored in green, indicated by a red arrow. The csNOX5 is shown as gray surface. g, The positively charged environment surrounding the heme molecule (heme #1).

Supplementary information

Reporting Summary

Supplementary Video 1

Flexibility of cytoplasmic domains of DUOX1–DUOXA1 in the dimer-of-dimers conformation.

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Sun, J. Structures of mouse DUOX1–DUOXA1 provide mechanistic insights into enzyme activation and regulation. Nat Struct Mol Biol 27, 1086–1093 (2020).

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