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Structural basis for GTP hydrolysis and conformational change of MFN1 in mediating membrane fusion

Abstract

Fusion of the outer mitochondrial membrane is mediated by the dynamin-like GTPase mitofusin (MFN). Here, we determined the structure of the minimal GTPase domain (MGD) of human MFN1 in complex with GDP-BeF3. The MGD folds into a canonical GTPase fold with an associating four-helix bundle, HB1, and forms a dimer. A potassium ion in the catalytic core engages GDP and BeF3 (GDP-BeF3). Enzymatic analysis has confirmed that efficient GTP hydrolysis by MFN1 requires potassium. Compared to previously reported MGD structures, the HB1 structure undergoes a major conformational change relative to the GTPase domains, as they move from pointing in opposite directions to point in the same direction, suggesting that a swing of the four-helix bundle can pull tethered membranes closer to achieve fusion. The proposed model is supported by results from in vitro biochemical assays and mitochondria morphology rescue assays in MFN1-deleted cells. These findings offer an explanation for how Charcot–Marie–Tooth neuropathy type 2 A (CMT2A)-causing mutations compromise MFN-mediated fusion.

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Fig. 1: Crystal structure of MFN1MGD dimer.
Fig. 2: GTP hydrolysis of MFN1.
Fig. 3: Dimerization of MFN1.
Fig. 4: Conformational changes in MFN1.
Fig. 5: Tests of CMT2A-causing mutations.
Fig. 6: FRET and membrane tethering assay of MFN1.
Fig. 7: Models of MFN-mediated fusion.

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References

  1. Hoppins, S. & Nunnari, J. The molecular mechanism of mitochondrial fusion. Biochim. Biophys. Acta 1793, 20–26 (2009).

    Article  CAS  PubMed  Google Scholar 

  2. Lackner, L. L. & Nunnari, J. M. The molecular mechanism and cellular functions of mitochondrial division. Biochim. Biophys. Acta 1792, 1138–1144 (2009).

    Article  CAS  PubMed  Google Scholar 

  3. Youle, R. J. & van der Bliek, A. M. Mitochondrial fission, fusion, and stress. Science 337, 1062–1065 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  4. Mishra, P. & Chan, D. C. Metabolic regulation of mitochondrial dynamics. J. Cell Biol. 212, 379–387 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  5. Pernas, L. & Scorrano, L. Mito-morphosis: mitochondrial fusion, fission, and cristae remodeling as key mediators of cellular function. Annu. Rev. Physiol. 78, 505–531 (2016).

    Article  CAS  PubMed  Google Scholar 

  6. Santel, A. & Fuller, M. T. Control of mitochondrial morphology by a human mitofusin. J. Cell Sci. 114, 867–874 (2001).

    CAS  PubMed  Google Scholar 

  7. Rojo, M., Legros, F., Chateau, D. & Lombès, A. Membrane topology and mitochondrial targeting of mitofusins, ubiquitous mammalian homologs of the transmembrane GTPase Fzo. J. Cell Sci. 115, 1663–1674 (2002).

    CAS  PubMed  Google Scholar 

  8. Chen, H. et al. Mitofusins Mfn1 and Mfn2 coordinately regulate mitochondrial fusion and are essential for embryonic development. J. Cell Biol. 160, 189–200 (2003).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  9. Hales, K. G. & Fuller, M. T. Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase. Cell 90, 121–129 (1997).

    Article  CAS  PubMed  Google Scholar 

  10. Hermann, G. J. et al. Mitochondrial fusion in yeast requires the transmembrane GTPase Fzo1p. J. Cell Biol. 143, 359–373 (1998).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  11. Rapaport, D., Brunner, M., Neupert, W. & Westermann, B. Fzo1p is a mitochondrial outer membrane protein essential for the biogenesis of functional mitochondria in Saccharomyces cerevisiae. J. Biol. Chem. 273, 20150–20155 (1998).

    Article  CAS  PubMed  Google Scholar 

  12. Züchner, S. et al. Mutations in the mitochondrial GTPase mitofusin 2 cause Charcot-Marie-Tooth neuropathy type 2A. Nat. Genet. 36, 449–451 (2004).

    Article  PubMed  Google Scholar 

  13. Harrison, S. C. Viral membrane fusion. Nat. Struct. Mol. Biol. 15, 690–698 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  14. Jahn, R. & Scheller, R. H. SNAREs–engines for membrane fusion. Nat. Rev. Mol. Cell Biol. 7, 631–643 (2006).

    Article  CAS  PubMed  Google Scholar 

  15. Wickner, W. & Schekman, R. Membrane fusion. Nat. Struct. Mol. Biol. 15, 658–664 (2008).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  16. Martens, S. & McMahon, H. T. Mechanisms of membrane fusion: disparate players and common principles. Nat. Rev. Mol. Cell Biol. 9, 543–556 (2008).

    Article  CAS  PubMed  Google Scholar 

  17. Südhof, T. C. & Rothman, J. E. Membrane fusion: grappling with SNARE and SM proteins. Science 323, 474–477 (2009).

    Article  PubMed  PubMed Central  Google Scholar 

  18. Wang, Y. et al. SNARE-mediated membrane fusion in autophagy. Semin. Cell Dev. Biol. 60, 97–104 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  19. Hu, J. et al. A class of dynamin-like GTPases involved in the generation of the tubular ER network. Cell 138, 549–561 (2009).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  20. Orso, G. et al. Homotypic fusion of ER membranes requires the dynamin-like GTPase atlastin. Nature 460, 978–983 (2009).

    Article  CAS  PubMed  Google Scholar 

  21. Anwar, K. et al. The dynamin-like GTPase Sey1p mediates homotypic ER fusion in S. cerevisiae. J. Cell Biol. 197, 209–217 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  22. Yan, L. et al. Structures of the yeast dynamin-like GTPase Sey1p provide insight into homotypic ER fusion. J. Cell Biol. 210, 961–972 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  23. Zhang, M. et al. ROOT HAIR DEFECTIVE3 family of dynamin-like GTPases mediates homotypic endoplasmic reticulum fusion and is essential for Arabidopsis development. Plant Physiol. 163, 713–720 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  24. Liu, T. Y. et al. Cis and trans interactions between atlastin molecules during membrane fusion. Proc. Natl Acad. Sci. USA 112, E1851–E1860 (2015).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  25. Hu, J. & Rapoport, T. A. Fusion of the endoplasmic reticulum by membrane-bound GTPases. Semin. Cell Dev. Biol. 60, 105–111 (2016).

    Article  CAS  PubMed  Google Scholar 

  26. Ishihara, N., Eura, Y. & Mihara, K. Mitofusin 1 and 2 play distinct roles in mitochondrial fusion reactions via GTPase activity. J. Cell Sci. 117, 6535–6546 (2004).

    Article  CAS  PubMed  Google Scholar 

  27. Brandt, T., Cavellini, L., Kühlbrandt, W. & Cohen, M. M. A mitofusin-dependent docking ring complex triggers mitochondrial fusion in vitro. eLife 5, e14618 (2016).

    Article  PubMed  PubMed Central  Google Scholar 

  28. Qi, Y. et al. Structures of human mitofusin 1 provide insight into mitochondrial tethering. J. Cell Biol. 215, 621–629 (2016).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  29. Cao, Y. L. et al. MFN1 structures reveal nucleotide-triggered dimerization critical for mitochondrial fusion. Nature 542, 372–376 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Bian, X. et al. Structures of the atlastin GTPase provide insight into homotypic fusion of endoplasmic reticulum membranes. Proc. Natl Acad. Sci. USA 108, 3976–3981 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  31. Byrnes, L. J. & Sondermann, H. Structural basis for the nucleotide-dependent dimerization of the large G protein atlastin-1/SPG3A. Proc. Natl Acad. Sci. USA 108, 2216–2221 (2011).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  32. Byrnes, L. J. et al. Structural basis for conformational switching and GTP loading of the large G protein atlastin. EMBO J. 32, 369–384 (2013).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  33. Wu, F., Hu, X., Bian, X., Liu, X. & Hu, J. Comparison of human and Drosophila atlastin GTPases. Protein Cell 6, 139–146 (2015).

    Article  CAS  PubMed  Google Scholar 

  34. Low, H. H. & Löwe, J. A bacterial dynamin-like protein. Nature 444, 766–769 (2006).

    Article  CAS  PubMed  Google Scholar 

  35. Low, H. H., Sachse, C., Amos, L. A. & Löwe, J. Structure of a bacterial dynamin-like protein lipid tube provides a mechanism for assembly and membrane curving. Cell 139, 1342–1352 (2009).

    Article  PubMed  PubMed Central  Google Scholar 

  36. Chappie, J. S., Acharya, S., Leonard, M., Schmid, S. L. & Dyda, F. G domain dimerization controls dynamin’s assembly-stimulated GTPase activity. Nature 465, 435–440 (2010).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  37. Scrima, A. & Wittinghofer, A. Dimerisation-dependent GTPase reaction of MnmE: how potassium acts as GTPase-activating element. EMBO J. 25, 2940–2951 (2006).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  38. Chen, Y. et al. Conformational dynamics of dynamin-like MxA revealed by single-molecule FRET. Nat. Commun. 8, 15744 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Huang, X. et al. Sequences flanking the transmembrane segments facilitate mitochondrial localization and membrane fusion by mitofusin. Proc. Natl Acad. Sci. USA 114, E9863–E9872 (2017).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  40. Daumke, O. & Roux, A. Mitochondrial homeostasis: how do dimers of mitofusins mediate mitochondrial fusion? Curr. Biol. 27, R353–R356 (2017).

    Article  CAS  PubMed  Google Scholar 

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

  42. Afonine, P. V. et al. Towards automated crystallographic structure refinement with phenix.refine. Acta Crystallogr. D Biol. Crystallogr. 68, 352–367 (2012).

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  43. Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D Biol. Crystallogr. 60, 2126–2132 (2004).

    Article  PubMed  Google Scholar 

  44. Yin, J., Lin, A. J., Golan, D. E. & Walsh, C. T. Site-specific protein labeling by Sfp phosphopantetheinyl transferase. Nat. Protoc. 1, 280–285 (2006).

    Article  CAS  PubMed  Google Scholar 

Download references

Acknowledgements

We thank J. Shaw (University of Utah) and D. Chan (Caltech) for providing the yeast strains and MEF cell lines, Dr. A. M. Prater for proofreading, L. Wu from BL17B for help with structure determination, and the Tsinghua University Branch of China National Center for Protein Sciences (Beijing) for technical support. J.H. is supported by the National Key Research and Development Program (Grant No. 2016YFA0500201), the National Natural Science Foundation of China (Grant No. 31225006), and an International Early Career Scientist grant from Howard Hughes Medical Institute. L.Y. is supported by the National Natural Science Foundation of China (Grant No. 31700659). Z.L. is supported by the National Natural Science Foundation of China (Grant No. 81322023) and the National Basic Research Program of China (973 Program, Grants 2013CB911103, 2014CBA02003 and 2014CB542802).

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Contributions

Z.L. and J.H. contributed to the overall study design. L.Y., Y.Q., X.H., and C.Y. performed most experiments, with help from L.L. and X.G. Data were analyzed by L.Y., Y.Q., X.H., C.Y., L.L., X.G., Z.R., Z.L., and J.H. The manuscript was written primarily by J.H., with contributions from the other authors.

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Correspondence to Junjie Hu or Zhiyong Lou.

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Integrated supplementary information

Supplementary Figure 1 Comparison of different MFN1 structures.

(A) Crystal packing of the MFN1 MGD in complex with GDP and BeF3-. Different views of molecule packing in the crystal are shown. Three MFN1 molecules (in lime, orange and blue) in the unit cell (green lines) and three from symmetry operation (purple, green and cyan) are shown in cartoon representation. (B) Superposition of the non-crystallographic dimer with the crystallographic dimer. (C) Superposition of the new MGD dimer (in pink) with previous structures (5GOM in cyan and 5GNT in green). (D) Comparison of the dimer interface of the two MGD dimers. Residues at the interface are highlighted and labeled. Buried surface areas and dimensions of the dimers are indicated.

Supplementary Figure 2 Comparison of MFN1 and dynamin superfamily proteins.

(A) Dimers of dynamin superfamily GTPases are shown in surface representation, with two protomers colored in pink and green, respectively. Nucleotide states and conformational states are labeled. BSE, bundle signaling element. (B) Monomers of dynamin superfamily GTPases are shown in cartoon representation and colored as in Fig. 1B. PDB codes and key structural elements are labeled.

Supplementary Figure 3 Catalytic cores of MFN1 and other dynamin superfamily proteins.

(A) Cutaway views of the catalytic core of MFN1, dynamin and ATL1. The nucleotides are shown in sticks and ions in spheres. (B) Catalytic core of MFN1, dynamin and MnmE. Monovalent ions and their ligands are shown. (C) GTPase activity of MFN1 measured with varied [K+]. 5 μM protein was used for each sample. The activities were measured by phosphate release at saturating GTP concentrations (0.5 mM) using the MGD. GTPase activity of MFN1 measured with varied [GTP] is shown on the right. Each point is the mean and SD of eight measurements. (D) As in C, but with 150 mM K+ and varied [Mg2+] and or [Ca2+]. (E) As in C, but with 150 mM K+, 4 mM Mg2+, and varied [Na+]. (F) As in C, but with 150 mM K+, 4 mM Mg2+, and varied pH. (G) As in C, but with various amounts of MFN1-MGD, 150 mM K+ and 4 mM Mg2+. (H) As in C, but with 2 μM proteins, 150 mM K+ or Na+, and 4 mM Mg2+.

Supplementary Figure 4 Nucleotide binding and dimerization of MFN1.

(A) Stopped-flow analysis of mGDP-BeF3- binding to MGD in the presence of different salts. Preincubated mGDP-MGD (10μM mGDP and 60μM MGD) was mixed in a 1:1 ratio with BeF3- in a stopped-flow apparatus. The mant-group was excited at 360nm and fluorescence change was monitored through a 395nm cutoff filter with time. (B) The binding of indicated nucleotides to S85A MFN1 (0.03 mM) was determined by ITC by stepwise injection of a 0.45 mM nucleotide solution. The estimated dissociation constants KD are given. See Supplementary Table 1 for more details. (C) The sizes of wild-type MFN1-MGD (theoretical molecular mass 49.5 kDa) were determined by MALS coupled with gel filtration in the presence of indicated nucleotides. The estimated molecular masses are shown by the right axis. The data are representative of at least three repetitions. (D) The sizes of wild-type MFN1-MGD (theoretical molecular mass 49.5 kDa) in complex with indicated nucleotides were determined by AUC. The estimated molecular masses are shown on top of the peaks. (E) As in B, but with mutants of the dimer interface in the presence of GDP and BeF3-. (F) As in B, but with mutants of the catalytic core in the presence of GDP and BeF3-. (G) HA-tagged MFN1 H147A or Q255A was transfected into MFN1-deleted MEF cells. Its localization was determined by anti-HA antibodies (green) and compared to MitoDsRed using confocal microscopy. The right images show enlargements of the boxed regions. The mitochondria morphology of indicated samples was categorized and shown on the right with data for “wt” and “vector” from Fig. 2F for comparison. A total of 100-120 cells were counted for each sample. All graphs are representative of at least three repetitions. Scale bars, 10 μm.

Supplementary Figure 5 Sequence conservation of MFN, Fzo1p and BDLP.

(A) Sequence alignment of MFNs, Fzo1p and BDLP. Predicted and observed secondary structural elements and GTPase motifs are highlighted (predicted α helices, yellow; predicted β strands, light green; observed α helices, orange; observed β strands, green). The α helices are shown in bars and β strands with arrows. Heptad repeat 1 (HR1) of MFN is underlined in brown, HR2 in purple, and TMs in red. (B) GTPase activity of disease-causing mutants measured with 150 mM K+ and 4 mM Mg2+. 5 μM protein was used for each sample. The activities were measured by phosphate release at saturating GTP concentrations (0.5 mM) using the MGD. Each point is the mean and SD of eight measurements. (C) Expression levels of HA-Fzo1p used in Fig. S5D are shown with PGK as a loading control. (D) Serial dilution assay to analyze the growth of fzo1Δ yeast expressing the indicated alleles of FZO1. Growth on rich dextrose (YPD) and glycerol (YPG) plates is shown. These strains were generated through a plasmid-shuffle strategy in which pRS416-FZO1 is replaced by a pRS315-2XHA-FZO1 plasmid containing the indicated mutations.

Supplementary Figure 6 Controls for FRET-based assay and tethering assay of MFN1.

(A) Structural elements in “HB-open” and “HB-closed” states are superimposed. Key components involved in conformational changes are shown. (B) Purified MGD, with or without YBBR tag, was labelled with CoA-Cy3, and separated by SDS-PAGE, and analysed by fluorescent imaging (top) or Coomassie staining (bottom). (C) GTPase activity of MGD, with or without YBBR tag, was measured by phosphate release at saturating GTP concentrations (0.5 mM). (D) Mitochondrial morphology rescue in MFN1-deleted MEF cells was tested using wt or MFN1 with YBBR tag insertion. E, MFN1-MGD, containing a YBBR tag at either GTPase site or HB1 site, was purified and labelled with CoA-Cy3 or CoA-Cy5 fluorescent dyes using Sfp phosphopantetheinyl transferase. Cy3- and Cy5-labeled proteins were mixed 1:1 (0.5 μM each), and the indicated nucleotide (nt), 50 μM unlabeled MGD, or buffer was added at the indicated time point. FRET was determined by exciting the donor fluorophore (Cy3) at 537 nm and measuring the emission of the donor and acceptor dyes (Cy5) at 570 nm and 667 nm, respectively. FRET is expressed as I A /(I A + I D ), where I A and I D are fluorescence intensities of the acceptor and donor emission, respectively. (F) Purified TM-containing MGDs were analyzed by SDS-PAGE and Coomassie staining.

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Supplementary Figures 1–6 and Supplementary Table 1

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Supplementary Video 1

Conformational changes of MFN1–MGD from open to closed state. MFN1–MGD structures are shown in cartoon representation. Key residues that stabilize each state are labelled. Buried surface areas of the dimers are indicated.

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Yan, L., Qi, Y., Huang, X. et al. Structural basis for GTP hydrolysis and conformational change of MFN1 in mediating membrane fusion. Nat Struct Mol Biol 25, 233–243 (2018). https://doi.org/10.1038/s41594-018-0034-8

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