Abstract
The mechanochemical protein dynamin is the prototype of the dynamin superfamily of large GTPases, which shape and remodel membranes in diverse cellular processes1. Dynamin forms predominantly tetramers in the cytosol, which oligomerize at the neck of clathrin-coated vesicles to mediate constriction and subsequent scission of the membrane1. Previous studies have described the architecture of dynamin dimers2,3, but the molecular determinants for dynamin assembly and its regulation have remained unclear. Here we present the crystal structure of the human dynamin tetramer in the nucleotide-free state. Combining structural data with mutational studies, oligomerization measurements and Markov state models of molecular dynamics simulations, we suggest a mechanism by which oligomerization of dynamin is linked to the release of intramolecular autoinhibitory interactions. We elucidate how mutations that interfere with tetramer formation and autoinhibition can lead to the congenital muscle disorders Charcot–Marie–Tooth neuropathy4 and centronuclear myopathy5, respectively. Notably, the bent shape of the tetramer explains how dynamin assembles into a right-handed helical oligomer of defined diameter, which has direct implications for its function in membrane constriction.
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Acknowledgements
This project was supported by grants from the Deutsche Forschungsgemeinschaft (MA1081/8-2 to D.J.M.; SFB740/D7 and SFB958/A04 to F.N.; SFB740/C8 and SFB 958/A7 to V.H.; SFB 740/C7 and SFB958/A12 to O.D.; and ES410/2-1 to S.E.), an ERC consolidator grant (ERC-2013-CoG-616024 to O.D.), an ERC starting grant (pcCell to F.N.) and a grant from the Einstein Foundation Berlin (SOoPiC to N.P.). T.F.R. acknowledges partial financial support by the Cluster of Excellence REBIRTH (DFG EXC 62/1). We thank B. Purfürst for help with electron microscopy; S. Hertel, L. Litz, P. Straub and S. Wohlgemuth for experimental assistance; and the staff at beamlines X06SA (PXI) and X06DA (PXIII) at the Swiss Light Source (Villigen, Switzerland) for help during data collection. We thank Y.-W. Liu for discussions, and A. Wittinghofer for his support and discussions in the initial stages of the project.
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Contributions
T.F.R. grew the crystals and collected data; K.F. solved the structure; T.F.R., K.F. and S.E. refined the structure; T.F.R. and R.A. purified protein for crystallization and monomeric dynamin; K.F. and J.S. purified all other proteins, performed liposome co-sedimentation, EM and GTPase assays; U.C. performed and analysed analytical ultracentrifugation experiments; Y.P. and K.K. performed transferrin uptake assays; N.P. and. F.N. conducted and analysed molecular modelling and molecular dynamics simulations. N.P. and Y.P. contributed equally to this work. T.F.R., K.F., F.N., V.H., O.D. and S.E. interpreted structural data. T.F.R., K.F., Y.P., N.P., U.C., D.J.M., F.N., V.H., O.D. and S.E. designed the research. T.F.R., K.F., F.N., O.D. and S.E. wrote the manuscript.
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Extended data figures and tables
Extended Data Figure 1 Characterization of the dynamin 3 construct.
a, Top: the domain structure of dynamin 3. The previously used sequence-derived domain nomenclature is shown below. Bottom: a dynamin 3 monomer colour-coded according to the domain architecture. b, SDS–PAGE representing a typical purification of dynamin 3. M, marker proteins; NI, whole-cell-lysate non-induced; SN, supernatant of cleared lysate; E, elution peak of the Talon-Co2+ column; CL, after cleavage with TEV protease; P, pool after gel filtration. c, Representative electron density map (stereo view). Two stalk helices are shown as stick models, the 2Fo − Fc map is contoured at 1.0σ. Raw data for b is available in Supplementary Information.
Extended Data Figure 2 Dimerization of dynamin 3.
a, Superposition of dynamin 1 (grey; PDB code: 3SNH) and dynamin 3 (magenta and green) dimers, colour-coded as in Fig. 1. The stalk arrangement in dynamin 3 is essentially the same as in dynamin 1. b, Interface 2 in dynamin 3. The view is rotated by 90° with respect to a. The zoom shows the side chains of residues involved in interface formation. Residues, whose mutation render dynamin 1 and dynamin 3 monomeric, are marked with an asterisk. c, Top: stalks of the dynamin 3 tetramer, as seen in the crystal structure (left). Dynamin dimers (dark and light blue) are formed via interface 2 (I2) and assemble into the tetramer via interfaces 1 and 3 (I1 and I3, respectively). In alternative dimerization models (middle and right)52, dynamin monomers assemble via interface 1 (middle) or interface 3 (right) to form elongated dimers of different shapes. Bottom: arrangement of stalks of a dynamin 1 as fitted into a cryo-EM density map of a super-constricted dynamin 1 helix (PDB code: 4UUD)26. d, Oligomeric state of dimer interface mutants, as assayed by analytical ultracentrifugation at a protein concentration of 20 µM. The following molecular masses were obtained from c(s) analyses: dynamin 1(R361S/R399A) (176 kDa, dimeric) in dark blue; dynamin 1(I481D/H687D/L688S) (84 kDa, monomeric) in light blue; dynamin 3(K361S/R399A) (165 kDa, dimeric) in red, dynamin 3(I481D/H677D/L678S) (83 kDa, monomeric) in black.
Extended Data Figure 3 Dynamin assembly via interface 3.
a, Schematic overview of the interactions in interface 3. b, Details of loop L1NS. The 2Fo − Fc electron density is contoured at 1.0σ. c, The Charcot–Marie–Tooth-related mutation G358R is located at the C-terminal end of loop L1NS. It probably disturbs the structural integrity of this loop and therefore might interfere with oligomerization. d, Clathrin-coated pit dynamics in HeLa cells expressing interface 3 mutants of dynamin 2. HeLa cells treated with dynamin 2 siRNA were co-transfected with plasmids encoding eGFP or siRNA-resistant dynamin 2–eGFP and mRFP–clathrin-light-chain, and live cells were imaged at 37 °C by TIRF microscopy. Shown are representative time-resolved line scans (kymographs) from at least ten time-lapse recordings of individual cells. Attenuated clathrin-coated pit dynamics upon depletion of endogenous dynamin 2 are only rescued by re-expression of wild-type but not mutant dynamin 2–eGFP. Note that the dynamin 2 mutants tested displayed a more diffuse subcellular distribution although they were still recruited to clathrin-coated pits.
Extended Data Figure 4 Localization of the PH domain in the tetramer.
a, Superposition of an outer dynamin molecule (magenta) and an inner molecule (green) of the dynamin 3 tetramer. The comparison reveals an ∼40° rotation of the G domains and BSEs. Furthermore, the PH domain is visible only in the outer molecule. b, Superposition of the stalk and PH domain in dynamin 1 (grey) and dynamin 3 (magenta). c, Connectivity of PH domain and stalk in the outer molecule. Shown are the stalk and PH domain of an outer molecule (magenta) and the stalk of the corresponding inner molecule (green) from a dimer. Since the gap of ∼58 Å between V629 of the PH domain and P643 of the inner stalk is too large to be spanned by the missing 13 residues (grey dashed line), we can unambiguously assign the PH domains in dynamin 3 to the outer stalks (black dashed lines). All other potential connections including molecules from the second dimer or symmetry-related tetramers span even larger distances (not shown). In the crystal structures of dynamin 1, an unequivocal assignment of the PH domain to a specific stalk was not possible, due to the long unresolved linker regions between the stalk and the PH domains. Concomitantly, the impact of the interface between stalk and PH domain has not been generally recognized53. d, The outer PH domains are clearly defined in the electron density (left panel), whereas no density for a PH domain is observed in the equivalent position at the inner stalks (right panel). The density visible in the right panel corresponds mainly to a G domain from a symmetry-related molecule. The 2Fo − Fc electron density is contoured at 1.0σ. e, Modelling of a PH domain (grey) relative to an inner stalk (green) in the same geometry as seen in the outer molecules leads to steric clashes (black oval) with an adjacent stalk (blue).
Extended Data Figure 5 The PH domains regulate oligomerization of dynamin.
a, In the absence of liposomes, a dynamin 3 variant lacking the PH domain (ΔPH) was sedimented more efficiently than wild-type dynamin 3 (WT). Both ΔPH and wild type lacked the PRD. The proteins were sedimented by ultracentrifugation after 20 h of incubation at low salt concentrations (60 mM NaCl) in the presence of the non-hydrolysable GTP analogue GMPPCP. S, supernatant; P, pellet fraction. b, c, Representative negative-stain electron micrographs of wild type (b) and ΔPH (c) under the same conditions as in a. For each protein, at least eight micrographs were recorded. Both constructs showed oligomeric ring structures, similar to structures seen for full-length dynamin7. Our data indicate that oligomerization of dynamin does not require membrane binding, but membrane binding requires oligomerization (Fig. 2). d, In liposome co-sedimentation assays, dynamin 3 bound to Folch liposomes independently of their size. Not extr., not extruded. e, At physiological salt concentrations (150 mM NaCl), dynamin 3 efficiently tubulated unfiltered Folch liposomes. In contrast, ΔPH did not decorate the liposome surface and did not induce liposome tubulation. For each setup at least 12 micrographs were recorded. f, When expressed in HeLa cells, dynamin 2(ΔPH) formed large cytosolic aggregates that did not co-localize with mRFP–clathrin. Arrowheads indicate co-localization for wild-type dynamin 2. Shown are magnified insets of representative images from at least 20 individual cells, acquired by TIRF microscopy. g, Dynamin 2(ΔPH) was dominant-negative in transferrin uptake assays. Data shown represent mean ± s.e.m., the number of independent experiments is indicated in the bar. Raw data for a and d is available in Supplementary Information.
Extended Data Figure 6 Mutational analysis of the interface between PH domain and stalk.
a, Analytical gel filtration analysis for wild-type dynamin 3 and the mutant R518D. The proteins were pre-incubated for 10 min at 22 °C or 37 °C. When pre-incubated at 37 °C, only R518D showed a higher molecular weight species. AU, arbitrary units. b, Intrinsic GTPase activity of wild-type dynamin 3 and the mutant R518D at 37 °C in the absence of liposomes. The lines represent linear fits of GTP hydrolysis versus time. For R518D, a biphasic behaviour of the GTPase activity was apparent (for wild type: kobs = 0.5 min−1; for R518D: kobs1 = 2.2 min−1 and kobs2 = 13.3 min−1). This biochemical behaviour is reminiscent of dynamin 1 mutants in the PH-domain–stalk interface that show increased oligomerization and GTPase rates when incubated at 37 °C20. Perturbations in this interface appear to promote oligomerization of dynamin, pointing to an autoinhibitory function of this interface for oligomerization. The average of two independent measurements is shown with deviations ranging from 0% to 0.05% for wild type and 0% to 0.62% for R518D. c, Analytical ultracentrifugation experiments for the indicated dynamin 3 variants, as in Fig. 2a. For the mutant K361S that sediments as a single species, a molecular mass of 164 kDa could be obtained from c(s) analysis, indicating that this mutant forms dimers in solution. d, Liposome co-sedimentation analysis for the indicated mutants. S, supernatant; P, pellet fraction. e, GTPase activity of the indicated mutants in the absence and presence of liposomes. Shown is the average of two independent measurements, with deviations ranging from 1% to 11%. f, Ability of dynamin 2 mutants to rescue defective CME of transferrin in absence of endogenous dynamin 2. The assay was performed as described in Fig. 2d. R518 in dynamin 3 corresponds to R522 in dynamin 2 and the R522H mutation in dynamin 2 is implicated in centronuclear myopathy. Data shown represent mean ± s.e.m., the number of independent experiments is indicated in the bar. Note, we generally observed that the GTPase experiments were the most sensitive indicators of structural perturbations induced by mutations. Compared to membrane binding assays, GTPase assays appear to be more sensitive to the actual architecture of the dynamin oligomer and alterations induced by point mutations. Transferrin uptake assays could be influenced by cellular factors, such as BAR-domain protein that may stabilize mutant dynamin forms with deficits in oligomerization. Raw data for d is available in Supplementary Information.
Extended Data Figure 7 Molecular dynamics simulations and Markov models.
a, The PH-domain–stalk interaction is characterized by a number of mainly polar interactions. The represented conformation is one of the starting structures (setup 2) for the MD simulations and quickly converts into one of the metastable conformations shown in Fig. 3. b, Relaxation timescales of different constructs as a function of lag time computed from Markov models. The timescales of all models (black) have converged at a lag time of about 20 ns within statistical uncertainty (colour-shaded regions), indicating approximate Markovianity. The grey area indicates the region with lag times larger than relaxation timescales. c, Top: intrinsic conformation dynamics of the L1NS loop shown for the wild type (black) and the mutant K361S (red). Bottom: six metastable conformations and their equilibrium probabilities of the L1NS loop (setup 3) for the wild type (black) and mutant K361S (red) computed from the Markov model. d, Residue pairs used to characterize the L1NS loop and stalk–PH domain interactions.
Extended Data Figure 8 Interactions of the G domain, stalk and BSE in the tetramer.
a, Two views on a fitting of the dynamin 3 tetramer crystal structure into the EM density of non-constricted oligomerized dynamin 1 (ref 24). The positions of the inner G domains are shown in all four molecules since the outer G domains in our crystals are stabilized by crystal contacts. Apparently, membrane binding and oligomerization is associated with major movements of the G domain, BSE and the PH domain (indicated by arrows). b, A loop of the outer PH domain and an inner G domain are in close proximity. c, The outer G domains (left), but not the inner G domains (right), are well defined in the electron density. The 2Fo − Fc electron density is contoured at 1.0σ. The weak electron density for the inner G domains and the resulting uncertainty in determining the contact sites prevented us from analysing this interaction in more detail. d, The BSE of an inner monomer (grey) interacts with the stalk of an outer monomer (magenta). This contact involves R465 which is mutated to tryptophan in some centronuclear myopathy patients. The R465W mutation leads to hyperactive dynamin4 that fragments the T tubule network in mouse-myoblast-derived myotubes and Drosophila body wall muscle (Y.-W. Liu, personal communication).
Extended Data Figure 9 Disease-relevant mutations in dynamin.
Localizations of mutations leading to Charcot–Marie–Tooth neuropathy (black balls) and centronuclear myopathy (pink balls) are plotted onto a dynamin 3 monomer. Colour code as in Fig. 1.
Supplementary information
Supplementary Information
This file contains Supplementary Figure 1, sequence alignment of the dynamin superfamily. The figure contains a structure-based alignment of protein sequences of important members of the dynamin superfamily. Amino acid residues mutated in this study are highlighted. It also contains gel images for Figure 2 and Extended Data Figures 5 and 6. (PDF 2723 kb)
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Reubold, T., Faelber, K., Plattner, N. et al. Crystal structure of the dynamin tetramer. Nature 525, 404–408 (2015). https://doi.org/10.1038/nature14880
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DOI: https://doi.org/10.1038/nature14880
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