Synaptotagmin, complexin, and neuronal SNARE (soluble N-ethylmaleimide sensitive factor attachment protein receptor) proteins mediate evoked synchronous neurotransmitter release, but the molecular mechanisms mediating the cooperation between these molecules remain unclear. Here we determine crystal structures of the primed pre-fusion SNARE–complexin–synaptotagmin-1 complex. These structures reveal an unexpected tripartite interface between synaptotagmin-1 and both the SNARE complex and complexin. Simultaneously, a second synaptotagmin-1 molecule interacts with the other side of the SNARE complex via the previously identified primary interface. Mutations that disrupt either interface in solution also severely impair evoked synchronous release in neurons, suggesting that both interfaces are essential for the primed pre-fusion state. Ca2+ binding to the synaptotagmin-1 molecules unlocks the complex, allows full zippering of the SNARE complex, and triggers membrane fusion. The tripartite SNARE–complexin–synaptotagmin-1 complex at a synaptic vesicle docking site has to be unlocked for triggered fusion to start, explaining the cooperation between complexin and synaptotagmin-1 in synchronizing evoked release on the sub-millisecond timescale.
Subscribe to Journal
Get full journal access for 1 year
only $3.90 per issue
All prices are NET prices.
VAT will be added later in the checkout.
Rent or Buy article
Get time limited or full article access on ReadCube.
All prices are NET prices.
Südhof, T. C. Neurotransmitter release: the last millisecond in the life of a synaptic vesicle. Neuron 80, 675–690 (2013)
Rothman, J. E. The principle of membrane fusion in the cell (Nobel lecture). Angew. Chem. Int. Ed. 53, 12676–12694 (2014)
Sutton, R. B., Fasshauer, D., Jahn, R. & Brunger, A. T. Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 Å resolution. Nature 395, 347–353 (1998)
Weber, T. et al. SNAREpins: minimal machinery for membrane fusion. Cell 92, 759–772 (1998)
Geppert, M. et al. Synaptotagmin I: a major Ca2+ sensor for transmitter release at a central synapse. Cell 79, 717–727 (1994)
Fernández-Chacón, R. et al. Synaptotagmin I functions as a calcium regulator of release probability. Nature 410, 41–49 (2001)
Perin, M. S., Fried, V. A., Mignery, G. A., Jahn, R. & Südhof, T. C. Phospholipid binding by a synaptic vesicle protein homologous to the regulatory region of protein kinase C. Nature 345, 260–263 (1990)
Perin, M. S. et al. Structural and functional conservation of synaptotagmin (p65) in Drosophila and humans. J. Biol. Chem. 266, 615–622 (1991)
Xu, J., Mashimo, T. & Südhof, T. C. Synaptotagmin-1, -2, and -9: Ca2+ sensors for fast release that specify distinct presynaptic properties in subsets of neurons. Neuron 54, 567–581 (2007)
Wen, H. et al. Distinct roles for two synaptotagmin isoforms in synchronous and asynchronous transmitter release at zebrafish neuromuscular junction. Proc. Natl Acad. Sci. USA 107, 13906–13911 (2010)
Bacaj, T. et al. Synaptotagmin-1 and synaptotagmin-7 trigger synchronous and asynchronous phases of neurotransmitter release. Neuron 80, 947–959 (2013)
Brose, N., Petrenko, A. G., Südhof, T. C. & Jahn, R. Synaptotagmin: a calcium sensor on the synaptic vesicle surface. Science 256, 1021–1025 (1992)
Davletov, B. A. & Südhof, T. C. A single C2 domain from synaptotagmin I is sufficient for high affinity Ca2+/phospholipid binding. J. Biol. Chem. 268, 26386–26390 (1993)
Chapman, E. R. & Davis, A. F. Direct interaction of a Ca2+ binding loop of synaptotagmin with lipid bilayers. J. Biol. Chem. 273, 13995–14001 (1998)
Fernandez, I. et al. Three-dimensional structure of the synaptotagmin 1 C2B-domain: synaptotagmin 1 as a phospholipid binding machine. Neuron 32, 1057–1069 (2001)
Kuo, W., Herrick, D. Z., Ellena, J. F. & Cafiso, D. S. The calcium-dependent and calcium-independent membrane binding of synaptotagmin 1: two modes of C2B binding. J. Mol. Biol. 387, 284–294 (2009)
Choi, U. B. et al. Single-molecule FRET-derived model of the synaptotagmin 1–SNARE fusion complex. Nat. Struct. Mol. Biol. 17, 318–324 (2010)
Kochubey, O. & Schneggenburger, R. Synaptotagmin increases the dynamic range of synapses by driving Ca2+-evoked release and by clamping a near-linear remaining Ca2+ sensor. Neuron 69, 736–748 (2011)
Vrljic, M. et al. Post-translational modifications and lipid binding profile of insect cell-expressed full-length mammalian synaptotagmin 1. Biochemistry 50, 9998–10012 (2011)
Brewer, K. D. et al. Dynamic binding mode of a Synaptotagmin-1–SNARE complex in solution. Nat. Struct. Mol. Biol. 22, 555–564 (2015)
Zhou, Q. et al. Architecture of the synaptotagmin–SNARE machinery for neuronal exocytosis. Nature 525, 62–67 (2015)
Pérez-Lara, Á. et al. PtdInsP2 and PtdSer cooperate to trap synaptotagmin-1 to the plasma membrane in the presence of calcium. eLife 5, 115–118 (2016)
Wang, S., Li, Y. & Ma, C. Synaptotagmin-1 C2B domain interacts simultaneously with SNAREs and membranes to promote membrane fusion. eLife 5, 209–217 (2016)
Mohrmann, R., Dhara, M. & Bruns, D. Complexins: small but capable. Cell. Mol. Life Sci. 72, 4221–4235 (2015)
Maximov, A., Tang, J., Yang, X., Pang, Z. P. & Südhof, T. C. Complexin controls the force transfer from SNARE complexes to membranes in fusion. Science 323, 516–521 (2009)
Xu, J., Pang, Z. P., Shin, O.-H. & Südhof, T. C. Synaptotagmin-1 functions as a Ca2+ sensor for spontaneous release. Nat. Neurosci. 12, 759–766 (2009)
Jorquera, R. A., Huntwork-Rodriguez, S., Akbergenova, Y., Cho, R. W. & Littleton, J. T. Complexin controls spontaneous and evoked neurotransmitter release by regulating the timing and properties of synaptotagmin activity. J. Neurosci. 32, 18234–18245 (2012)
Dhara, M. et al. Complexin synchronizes primed vesicle exocytosis and regulates fusion pore dynamics. J. Cell Biol. 204, 1123–1140 (2014)
Chen, X. et al. Three-dimensional structure of the complexin/SNARE complex. Neuron 33, 397–409 (2002)
Wu, D. et al. Postsynaptic synaptotagmins mediate AMPA receptor exocytosis during LTP. Nature 544, 316–321 (2017)
Lee, J., Guan, Z., Akbergenova, Y. & Littleton, J. T. Genetic analysis of synaptotagmin C2 domain specificity in regulating spontaneous and evoked neurotransmitter release. J. Neurosci. 33, 187–200 (2013)
Ernst, J. A. & Brunger, A. T. High resolution structure, stability, and synaptotagmin binding of a truncated neuronal SNARE complex. J. Biol. Chem. 278, 8630–8636 (2003)
Bracher, A., Kadlec, J., Betz, H. & Weissenhorn, W. X-ray structure of a neuronal complexin-SNARE complex from squid. J. Biol. Chem. 277, 26517–26523 (2002)
Kümmel, D. et al. Complexin cross-links prefusion SNAREs into a zigzag array. Nat. Struct. Mol. Biol. 18, 927–933 (2011)
Shin, O.-H., Xu, J., Rizo, J. & Südhof, T. C. Differential but convergent functions of Ca2+ binding to synaptotagmin-1 C2 domains mediate neurotransmitter release. Proc. Natl Acad. Sci. USA 106, 16469–16474 (2009)
Yang, X., Kaeser-Woo, Y. J., Pang, Z. P., Xu, W. & Südhof, T. C. Complexin clamps asynchronous release by blocking a secondary Ca2+ sensor via its accessory α helix. Neuron 68, 907–920 (2010)
Schonn, J.-S., Maximov, A., Lao, Y., Südhof, T. C. & Sørensen, J. B. Synaptotagmin-1 and -7 are functionally overlapping Ca2+ sensors for exocytosis in adrenal chromaffin cells. Proc. Natl Acad. Sci. USA 105, 3998–4003 (2008)
Gustavsson, N. & Han, W. Calcium-sensing beyond neurotransmitters: functions of synaptotagmins in neuroendocrine and endocrine secretion. Biosci. Rep. 29, 245–259 (2009)
Cao, P., Yang, X. & Südhof, T. C. Complexin activates exocytosis of distinct secretory vesicles controlled by different synaptotagmins. J. Neurosci. 33, 1714–1727 (2013)
Lai, Y. et al. N-terminal domain of complexin independently activates calcium-triggered fusion. Proc. Natl Acad. Sci. USA 113, E4698–E4707 (2016)
Choi, U. B., Zhao, M., Zhang, Y., Lai, Y. & Brunger, A. T. Complexin induces a conformational change at the membrane-proximal C-terminal end of the SNARE complex. eLife 5, e16886 (2016)
Tang, J. et al. A complexin/synaptotagmin 1 switch controls fast synaptic vesicle exocytosis. Cell 126, 1175–1187 (2006)
Gipson, P. et al. Morphologies of synaptic protein membrane fusion interfaces. Proc. Natl Acad. Sci. USA http://dx.doi.org/10.1073/pnas.1708492114 (2017)
Cipriano, D. J. et al. Processive ATP-driven substrate disassembly by the N-ethylmaleimide-sensitive factor (NSF) molecular machine. J. Biol. Chem. 288, 23436–23445 (2013)
Studier, F. W. Protein production by auto-induction in high density shaking cultures. Protein Expr. Purif. 41, 207–234 (2005)
Kabsch, W. Xds. Acta Crystallogr. D 66, 125–132 (2010)
Winn, M. D. et al. Overview of the CCP4 suite and current developments. Acta Crystallogr. D 67, 235–242 (2011)
McCoy, A. J. et al. Phaser crystallographic software. J. Appl. Crystallogr. 40, 658–674 (2007)
Emsley, P. & Cowtan, K. Coot: model-building tools for molecular graphics. Acta Crystallogr. D 60, 2126–2132 (2004)
Adams, P. D . et al. PHENIX: Building new software for automated crystallographic structure determination. Acta Crystallogr. D 58, 1948–1954 (2002)
Chen, V. B. et al. MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr. D 66, 12–21 (2010)
Krissinel, E. & Henrick, K. Inference of macromolecular assemblies from crystalline state. J. Mol. Biol. 372, 774–797 (2007)
Choi, H.-J. et al. αE-catenin is an autoinhibited molecule that coactivates vinculin. Proc. Natl Acad. Sci. USA 109, 8576–8581 (2012)
Radhakrishnan, A., Stein, A., Jahn, R. & Fasshauer, D. The Ca2+ affinity of synaptotagmin 1 is markedly increased by a specific interaction of its C2B domain with phosphatidylinositol 4,5-bisphosphate. J. Biol. Chem. 284, 25749–25760 (2009)
Evans, C. S. et al. Functional analysis of the interface between the tandem C2 domains of synaptotagmin-1. Mol. Biol. Cell 27, 979–989 (2016)
Maximov, A., Pang, Z. P., Tervo, D. G. R. & Südhof, T. C. Monitoring synaptic transmission in primary neuronal cultures using local extracellular stimulation. J. Neurosci. Methods 161, 75–87 (2007)
Pang, Z. P. & Südhof, T. C. Cell biology of Ca2+-triggered exocytosis. Curr. Opin. Cell Biol. 22, 496–505 (2010)
Zhou, P. et al. Syntaxin-1 N-peptide and Habc-domain perform distinct essential functions in synaptic vesicle fusion. EMBO J. 32, 159–171 (2013)
We thank P. Gipson, J. Leitz, A. Lyubimov, and W. I. Weis for discussions, S. Muennich and S. Pokutta for assistance with ITC, and the National Institutes of Health (NIH) for support (R37 MH63105 to A.T.B.; P50 MH086403 to T.C.S.). Crystal diffraction screening and data collection were performed at synchrotron facilities provided by the Advanced Photon Source at Argonne National Laboratory, the Stanford Synchrotron Radiation Lightsource, California, and the Advanced Light Source, Berkeley, California, funded by Department of Energy (DOE) under contracts DE-AC02-06CH11357 (Advanced Photon Source), DE-AC02-76SF00515 (Stanford Synchrotron Radiation Lightsource), and DE-AC02-05CH11231 (ALS). We thank the staff at these beamlines for help with diffraction data collection. The Northeastern Collaborative Access Team (NECAT) beamlines are funded by the National Institute of General Medical Sciences (NIGMS) from the NIH (P41 GM103403). The Pilatus 6M detector at the 24-ID-C beam line is funded by an NIH-ORIP HEI grant (S10 RR029205). The Stanford Synchrotron Radiation Lightsource Structural Molecular Biology Program including beam line BL12-2 is supported by the DOE Office of Biological and Environmental Research, and by the NIH NIGMS (P41 GM103393). The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of NIGMS or NIH.
The authors declare no competing financial interests.
Reviewer Information Nature thanks R. Heidelberger, C. Montecucco and the other anonymous reviewer(s) for their contribution to the peer review of this work.
Publisher's note: Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Extended data figures and tables
a, b, Views of the crystal lattice of the Syt1–SNARE–Cpx–Syt1 C2AB (a) and Syt1–SNARE–Cpx–Syt1 C2B (b) crystal structures. The red rectangles highlight the C-terminal ends of the SNARE complexes. They are unstructured and form no crystal contacts in either crystal form. c, d, B-factor coloured cartoon representations of the Syt1–SNARE–Cpx–Syt1 C2AB (c) and Syt1–SNARE–Cpx–Syt1 C2B (d) crystal structures (the second Syt1 molecule is related to the first by crystallographic symmetry). Both the primary and tripartite interfaces have low B-factors. e, f, Views of the molecular packing arrangements around the SNARE (coloured) and Syt1 C2B (orange) components that form that primary interface in the Syt1–SNARE–Cpx–Syt1 C2AB crystal structure (e) and the SNARE–Syt1 C2AB crystal structure (f, PDB accession number 5CCH). The molecular packing arrangements around the SNARE and Syt1 C2B components that form that primary interface are vastly different in these crystal structures, illustrating that the formation of the primary interface is entirely independent of crystal packing environments. We also note that the protein constructs and crystallization conditions were very different for these structures.
Extended Data Figure 2 Additional details of the Syt1–SNARE–Cpx–Syt1 tripartite interface and primary sequence alignment.
a, Close-up view of the contact interface between a pseudo-helix in Syt1 C2B (including the 310 helix T3 situated between β5 and β6, and the loop between α-helices β7 and HB), syntaxin-1A, and SNAP-25_N. Syt1 Lys354 and SNAP-25 Glu24 as well as Syt1 Glu350 and SNAP-25 Arg17 interact via salt bridges. Syt1 Gln353 and syntaxin-1A His199, Syt1 Glu350, and SNAP-25 Gln20 form separate hydrogen bonds. b, Close-up view of the contact site between the HB α-helix of the Syt1 C2B domain and the Cpx central α-helix. Interacting residues are shown as sticks and are labelled, while water molecules are shown as red balls. Dashed lines indicate hydrogen bonds or salt bridges. c, Sequence alignment of rat synaptotagmin isoforms, rat Doc2A/B, and rabphilin3A, for residues around the SNARE–Cpx–Syt1 tripartite interface. The arrows show the residues involved in specific sidechain interactions in the tripartite interface (see also discussion the text). The red stars show the mutated residues used in this study. The alignment was performed using Clustal Omega (http://www.ebi.ac.uk/Tools/msa/clustalo/). The figure was prepared with Boxshade3.21 (http://www.ch.embnet.org/software/BOX_form.html). d, Cartoon representation of the known crystal structures of C2A and C2B domains of synaptotagmin isoforms, Doc2 isoforms, rabphilin3A (Rph), and Munc13-1. The black arrows indicate the presence of the HA or T3 helices in these structures.
a, Top: circular dichroism spectra of WT and mutant Syt1 C2B domains in the absence of Ca2+. Bottom: circular dichroism thermal melting curves, monitored at 216 nm in the absence of Ca2+ (black) and in the presence of 5 mM Ca2+ (red). The specified melting temperatures were estimated by the mid-point of the melting curves (Methods). b, Circular dichroism spectra of WT SNARE complex (left) and SNAREQ (right). c, Circular dichroism thermal melting curves of WT SNARE complex (black circle) and SNAREQ complex (red triangle), monitored at 220 nm.
a–n, Differential power traces and heats of injection traces of the specified samples in the syringe and the cell of the ITC instrument. The experimental conditions are described in the Methods. For j–l, three independent experimental repeats were performed, and all ITC data curves are shown; shown are means ± s.e.m. for three independent repeat experiments. For all other ITC experiments, the error bars were obtained from a fit of the data points of the particular ITC experiments. The schemas in the insets summarize the mutations used in the particular experiments. The ITC experiments produce well-determined n values for the following experiments: C2BKA titrated into the SNARE complex (n = 0.94 ± 0.02), Cpx(48–73) titrated into the SNARE complex (n = 0.97 ± 0.02) or the SNAREQ complex (n = 0.98 ± 0.01). For the other ITC experiments, it was difficult to achieve high enough concentrations of the injected sample to obtain optimal conditions for reliable determination of n. ND, not detectable. Experimental conditions are described in the Methods.
Extended Data Figure 5 Superposition of observed interactions between the Syt1 C2B domain and the SNARE complex in solution and in crystal structures.
a, Close-up view of a small contact between the polybasic region of the Syt1 C2B domain and the SNARE complex in the Syt1–SNARE–Cpx–Syt1 crystal structures. Interacting residues are shown as sticks and are labelled. Dashed lines indicate hydrogen bonds or salt bridges. b, The primary interface (PDB accession number 5CCG21, and structures in this work), the small contact shown in a, and the deposited five conformers derived from solution NMR experiments involving the polybasic region of Syt1 C2B (sticks and balls indicate residues of the polybasic region of Syt1 C2B). These NMR studies revealed that there are dynamic binding modes between the polybasic region of Syt1 C2B and the SNARE complex in solution20. Although other interfaces between Syt1 and the SNARE complex have been observed (for example, secondary and tertiary interfaces in the crystal structure PDB accession number 5CCG), the ITC data (Extended Data Fig. 4) suggest that these are the only interactions and interfaces that occur in solution (see text).
a, Quantification of IPSC charge transfer (from the same neurons as in Fig. 4a). b, Recordings of mIPSCs from cultured cortical neurons with Syt1 conditional KO and Syt7 constitutive KO infected with lentiviruses expressing ΔCre/Cre recombinase and WT Syt1 or Syt1 mutants. Left to right: sample traces, cumulative probability plot of inter-event intervals, and quantification of event frequency, amplitude, and 10–90% rise time. c, Quantification of IPSC charge transfer (from the same neurons as in Fig. 4b). d, e, mIPSCs (d) and mEPSCs (e) from cultured cortical neurons infected with lentiviruses expressing Syt1 mutants. Left to right: sample traces, cumulative probability plot of inter-event intervals, and quantification of event frequency, amplitude, and 10–90% rise time. f, Quantification of IPSC charge transfer (from the same neurons as in Fig. 4d). g, h, Recordings of mIPSCs and mEPSCs from cultured cortical neurons infected with lentiviruses expressing Syt1WT or Syt1DA, without or with lentiviruses expressing Cpx1/2 shRNAs (Cpx1/2 DKD). Left to right: sample traces, cumulative probability plot of inter-event intervals, and quantification of event frequency, amplitude, and 10–90% rise time. Shown are means ± s.e.m.; the numbers of neurons/independent cultures are indicated. Statistical significance was assessed by Student’s t-test (*P < 0.05; **P < 0.01; ***P < 0.001; NS, no significant difference) with respect to either the Cre (red) or the Cre + Syt1 group (black) in a and b, either the control (black) or the Syt1DA group (red) in c–e, and between Syt1WT and Syt1DA with or without Cpx1/2 DKD in f–h.
Recordings of mIPSCs from cultured cortical neurons infected with lentiviruses expressing Syt1WT or Syt1DA, without or with 10 μM BAPTA-AM preincubated for 30 min at 37 °C. Sample traces (left), cumulative probability plot of inter-event intervals (middle), and quantification of event frequency, amplitude, and 10–90% rise time (right) of mIPSCs. Shown are means ± s.e.m.; the numbers of neurons/independent cultures are indicated. Statistical significance was assessed by Student’s t-test (**P < 0.01; ***P < 0.001; NS, no significant difference) with respect to either the Syt1WT (black) or the Syt1DA group (red). The absence of a dominant-negative effect of the Syt1DA group on spontaneous release in the presence of BAPTA-AM is consistent with the notion that spontaneous release largely depends on a different Ca2+ sensor. We speculate that this mini-release Ca2+ sensor may compete with Syt1 in binding to the tripartite interface. Elimination of the primary interface or presence of the Syt1DA mutant would affect the binding equilibrium, possibly explaining the increase of spontaneous release. We note that the effect of the Syt1DA mutant on spontaneous release (as assessed by mIPSCs and mEPSCs) is opposite to the effect on evoked release (as assessed by IPSCs and EPSCs) (Fig. 4).
Extended Data Figure 8 Superpositions of the Syt1–SNARE–Cpx–Syt1 crystal structures and other known crystal structures.
a–c, Conserved and variable regions of the SNARE–Cpx subcomplex in the Syt1–SNARE–Cpx–Syt1 C2B crystal structure (coloured), and crystal structures of the SNARE–Cpx subcomplex (PDB accession number 1KIL29 (black) and PDB accession number 3RK3 (ref. 34) (white)). The interaction between the central α-helix of Cpx and the SNARE complex is essentially identical in all crystal structures, while the angle at which the accessory helix protrudes away from the SNARE complex is variable. Such variability was also observed by single-molecule fluorescence resonance transfer experiments41. d, The superposition of the Syt1 C2B domain of the Syt1–SNARE–Cpx–Syt1 C2AB crystal structure with crystal structures of uncomplexed C2AB fragments (PDB accession numbers are indicated in the figure) illustrates variability of the position of the C2A domain relative to the C2B domain of Syt1.
Extended Data Figure 9 A possible supramolecular arrangement of one Syt1 C2B molecule bridging two SNARE complexes.
Orthogonal views (cartoon presentation, left; schema, right) of an arrangement where one Syt1 C2B domain bridges two SNARE complexes via the primary and tripartite interfaces, respectively. Directions (N terminus to C terminus) of the α-helices of the SNARE complex and Cpx are indicated by crosses (pointing into the page) and dots (pointing out of the page). The bridged complex would be sandwiched between two membranes as suggested in the schema in the lower right panel.
During the first 35 sec, a rotating view of the cartoon representation of the Syt1-SNARE-Cpx-Syt1 complex is shown. Subsequently, the interacting residues of the two interfaces are shown, and then, at 58 sec, the residues of the Ca2+ binding sites (from 58 to 1 min 12 sec) and polybasic regions (from 1 min 13 sec to 1 min 28 sec) of two Syt1 C2B domains are also shown in stick-and-ball representation. At 1 min 29 sec, the molecular surfaces of the molecules are shown. For clarity, we omitted the Syt1 C2A domains after 35 sec. (MOV 13954 kb)
Rotating view of the cartoon representation of the Syt1-SNARE-Cpx-Syut1 complex with mutated residues shown in stick-and-ball representation. For clarity, we omitted the Syt1 C2A domains. (MOV 3798 kb)
About this article
Cite this article
Zhou, Q., Zhou, P., Wang, A. et al. The primed SNARE–complexin–synaptotagmin complex for neuronal exocytosis. Nature 548, 420–425 (2017). https://doi.org/10.1038/nature23484
Physiological Reviews (2020)
Neurobiology of Aging (2020)
CNS & Neurological Disorders - Drug Targets (2020)
Coupling of Conformational Switches in Calcium Sensor Unraveled with Local Markov Models and Transfer Entropy
Journal of Chemical Theory and Computation (2020)
Proceedings of the National Academy of Sciences (2020)