Review Article | Published:

The control of release probability at nerve terminals

Nature Reviews Neurosciencevolume 20pages177186 (2019) | Download Citation

Abstract

Exocytosis is a fundamental membrane fusion process by which the soluble or membrane-associated cargoes of a secretory vesicle are delivered to the extracellular milieu or the cell surface. While essential for all organs, the brain relies on a specialized form of exocytosis to mediate information flow throughout its vast circuitry. Neurotransmitter-laden synaptic vesicles fuse with the plasma membrane on cue with astonishing speed in a probabilistic process that is both tightly regulated and capable of a fascinating array of plasticities. Here, we examine progress in the molecular understanding of synaptic vesicle fusion and its control.

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References

  1. 1.

    Feng, T. P. The Chinese Journal of Physiology Vol. 11 437–450 (Chinese Physiological Society, 1937).

  2. 2.

    Katz, B. & Miledi, R. The effect of calcium on acetylcholine release from motor nerve terminals. Proc. R. Soc. Lond. B 161, 496–503 (1965).

  3. 3.

    Dodge, F. A. Jr & Rahamimoff, R. Co-operative action a calcium ions in transmitter release at the neuromuscular junction. J. Physiol. 193, 419–432 (1967).

  4. 4.

    Augustine, G. J., Charlton, M. P. & Smith, S. J. Calcium entry and transmitter release at voltage-clamped nerve terminals of squid. J. Physiol. 367, 163–181 (1985).

  5. 5.

    Bollmann, J. H., Sakmann, B. & Borst, J. G. Calcium sensitivity of glutamate release in a calyx-type terminal. Science 289, 953–957 (2000).

  6. 6.

    Schneggenburger, R. & Neher, E. Intracellular calcium dependence of transmitter release rates at a fast central synapse. Nature 406, 889–893 (2000).

  7. 7.

    Wang, L. Y., Neher, E. & Taschenberger, H. Synaptic vesicles in mature calyx of Held synapses sense higher nanodomain calcium concentrations during action potential-evoked glutamate release. J. Neurosci. 28, 14450–14458 (2008).

  8. 8.

    Nakamura, Y. et al. Nanoscale distribution of presynaptic Ca(2+) channels and its impact on vesicular release during development. Neuron 85, 145–158 (2015). This paper — a technical tour du force — utilized a combination of SDS-FRL, electrophysiology, presynaptic calcium imaging and computational modelling at the calyx of Held to investigate the nanoscale organization of presynaptic calcium channels. The authors conclude that release sites are organized in a stereotyped topography they term ‘perimeter release’, such that a small cluster of calcium channels is surrounded by one or more SVs. Thus, vesicle fusion at a single release site is driven by the activity of a small number of nearby (~10–20 nm) calcium channels at the calyx.

  9. 9.

    Heidelberger, R., Heinemann, C., Neher, E. & Matthews, G. Calcium dependence of the rate of exocytosis in a synaptic terminal. Nature 371, 513–515 (1994).

  10. 10.

    Pang, Z. P., Sun, J., Rizo, J., Maximov, A. & Südhof, T. C. Genetic analysis of synaptotagmin 2 in spontaneous and Ca2+-triggered neurotransmitter release. EMBO J. 25, 2039–2050 (2006).

  11. 11.

    Sun, J. et al. A dual-Ca2+-sensor model for neurotransmitter release in a central synapse. Nature 450, 676–682 (2007).

  12. 12.

    Chen, C., Arai, I., Satterfield, R., Young, S. M. & Jonas, P. Synaptotagmin 2 is the fast Ca2+ sensor at a central synapse. Cell Rep. 18, 723–736 (2017).

  13. 13.

    Papahadjopoulos, D., Vail, W. J., Pangborn, W. A. & Poste, G. Studies on membrane fusion. II. Induction of fusion in pure phospholipid membranes by calcium ions and other divalent metals. Biochim. Biophys. Acta 448, 265–283 (1976).

  14. 14.

    Feigenson, G. W. On the nature of calcium ion binding between phosphatidylserine lamellae. Biochemistry 25, 5819–5825 (1986).

  15. 15.

    Feigenson, G. W. Calcium ion binding between lipid bilayers: the four-component system of phosphatidylserine, phosphatidylcholine, calcium chloride, and water. Biochemistry 28, 1270–1278 (1989).

  16. 16.

    Wilschut, J., Düzgünes, N., Fraley, R. & Papahadjopoulos, D. Studies on the mechanism of membrane fusion: kinetics of calcium ion induced fusion of phosphatidylserine vesicles followed by a new assay for mixing of aqueous vesicle contents. Biochemistry 19, 6011–6021 (1980).

  17. 17.

    Bai, J., Tucker, W. C. & Chapman, E. R. PIP2 increases the speed of response of synaptotagmin and steers its membrane-penetration activity toward the plasma membrane. Nat. Struct. Mol. Biol. 11, 36–44 (2004).

  18. 18.

    Südhof, T. C. Calcium control of neurotransmitter release. Cold Spring Harb. Perspect. Biol. 4, a011353 (2012).

  19. 19.

    Rizo, J. & Xu, J. The synaptic vesicle release machinery. Annu. Rev. Biophys. 44, 339–367 (2015).

  20. 20.

    Hui, E. et al. Three distinct kinetic groupings of the synaptotagmin family: candidate sensors for rapid and delayed exocytosis. Proc. Natl Acad. Sci. USA 102, 5210–5214 (2005).

  21. 21.

    Takamori, S. et al. Molecular anatomy of a trafficking organelle. Cell 127, 831–846 (2006).

  22. 22.

    Chapman, E. R. How does synaptotagmin trigger neurotransmitter release? Annu. Rev. Biochem. 77, 615–641 (2008).

  23. 23.

    Brenowitz, S. D. & Regehr, W. G. Reliability and heterogeneity of calcium signaling at single presynaptic boutons of cerebellar granule cells. J. Neurosci. 27, 7888–7898 (2007).

  24. 24.

    Xu-Friedman, M. A., Harris, K. M. & Regehr, W. G. Three-dimensional comparison of ultrastructural characteristics at depressing and facilitating synapses onto cerebellar Purkinje cells. J. Neurosci. 21, 6666–6672 (2001).

  25. 25.

    Luo, F., Dittrich, M., Stiles, J. R. & Meriney, S. D. Single-pixel optical fluctuation analysis of calcium channel function in active zones of motor nerve terminals. J. Neurosci. 31, 11268–11281 (2011).

  26. 26.

    Li, L., Bischofberger, J. & Jonas, P. Differential gating and recruitment of P/Q, N, and R-type Ca2+ channels in hippocampal mossy fiber boutons. J. Neurosci. 27, 13420–13429 (2007).

  27. 27.

    Rollenhagen, A. & Lübke, J. H. The mossy fiber bouton: the “common” or the “unique” synapse? Front. Synapt. Neurosci. 2, 2 (2010).

  28. 28.

    Fujimoto, K. Freeze-fracture replica electron microscopy combined with SDS digestion for cytochemical labeling of integral membrane proteins. Application to the immunogold labeling of intercellular junctional complexes. J. Cell Sci. 108, 3443–3449 (1995). This paper describes the original breakthrough whereby judicious use of detergents allowed antibody labelling of membrane proteins to be combined with freeze–fracture electron microscopy and thus enabled the identification of specific molecules.

  29. 29.

    Althof, D. et al. Inhibitory and excitatory axon terminals share a common nano-architecture of their Cav2.1 (P/Q-type) Ca(2+) channels. Front. Cell Neurosci. 9, 315 (2015).

  30. 30.

    Miki, T. et al. Numbers of presynaptic Ca2+ channel clusters match those of functionally defined vesicular docking sites in single central synapses. Proc. Natl Acad. Sci. USA 114, E5246–E5255 (2017). This paper describes detailed comparisons of estimates of the number of release sites at a small cerebellar synapse with the number of calcium channels present based on the technique described in reference 28.

  31. 31.

    Holderith, N. et al. Release probability of hippocampal glutamatergic terminals scales with the size of the active zone. Nat. Neurosci. 15, 988–997 (2012).

  32. 32.

    Dolphin, A. C. The α2δ subunits of voltage-gated calcium channels. Biochim. Biophys. Acta 1828, 1541–1549 (2013).

  33. 33.

    Dolphin, A. C. Voltage-gated calcium channels and their auxiliary subunits: physiology and pathophysiology and pharmacology. J. Physiol. 594, 5369–5390 (2016).

  34. 34.

    Campbell, K., Leung, A. & Sharp, A. The biochemistry and molecular-biology of the dihydropyridine-sensitive caclium-channel. Trends Neurosci. 11, 425–430 (1988).

  35. 35.

    Wu, J. et al. Structure of the voltage-gated calcium channel Ca(v)1.1 at 3.6Å resolution. Nature 537, 191–196 (2016).

  36. 36.

    Hoppa, M. B., Lana, B., Margas, W., Dolphin, A. C. & Ryan, T. A. α2δ expression sets presynaptic calcium channel abundance and release probability. Nature 486, 122–125 (2012). This paper demonstrates that the calcium channel subunit α2δ serves as a molecular gatekeeper, determining how many calcium channels are present at small CNS nerve terminals, thereby influencing the release probability.

  37. 37.

    Llinás, R. R. Depolarization release coupling: an overview. Ann. NY Acad. Sci. 635, 3–17 (1991).

  38. 38.

    Neher, E. Vesicle pools and Ca2+ microdomains: new tools for understanding their roles in neurotransmitter release. Neuron 20, 389–399 (1998).

  39. 39.

    Scimemi, A. & Diamond, J. S. The number and organization of Ca2+ channels in the active zone shapes neurotransmitter release from Schaffer collateral synapses. J. Neurosci. 32, 18157–18176 (2012).

  40. 40.

    Stanley, E. Activation of quantal transmitter release by single calcium domains at a cholinergic presynaptic nerve-terminal. Biophys. J. 64, A115–A115 (1993).

  41. 41.

    Stanley, E. F. Single calcium channels and acetylcholine release at a presynaptic nerve terminal. Neuron 11, 1007–1011 (1993).

  42. 42.

    Kim, M., Li, G. & von Gersdorff, H. Single Ca2+ channels and exocytosis at sensory synapses. J. Physiol. 591, 3167–3178 (2013).

  43. 43.

    Nakamura, Y., Reva, M. & DiGregorio, D. A. Variations in Ca2+ influx can alter Ca2+-chelator-based estimates of Ca2+ channel-synaptic vesicle coupling distance. J. Neurosci. 38, 3971–3987 (2018). This study develops a detailed quantitative analysis of how to estimate the coupling distance of calcium channels and release sites using an exogenous calcium chelator. The approach emphasizes the use of diverse sets of experimentally measured parameters to constrain computational models of excitation–secretion coupling.

  44. 44.

    Bucurenciu, I., Kulik, A., Schwaller, B., Frotscher, M. & Jonas, P. Nanodomain coupling between Ca2+ channels and Ca2+ sensors promotes fast and efficient transmitter release at a cortical GABAergic synapse. Neuron 57, 536–545 (2008).

  45. 45.

    Arai, I. & Jonas, P. Nanodomain coupling explains Ca2+ independence of transmitter release time course at a fast central synapse. eLife 3, e04057 (2014).

  46. 46.

    Baur, D. et al. Developmental tightening of cerebellar cortical synaptic influx-release coupling. J. Neurosci. 35, 1858–1871 (2015).

  47. 47.

    Schmidt, H. et al. Nanodomain coupling at an excitatory cortical synapse. Curr. Biol. 23, 244–249 (2013).

  48. 48.

    Kawaguchi, S. & Sakaba, T. Fast Ca2+ buffer-dependent reliable but plastic transmission at small CNS synapses revealed by direct bouton recording. Cell Rep. 21, 3338–3345 (2017).

  49. 49.

    Vyleta, N. P. & Jonas, P. Loose coupling between Ca2+ channels and release sensors at a plastic hippocampal synapse. Science 343, 665–670 (2014).

  50. 50.

    Kittel, R. J. et al. Bruchpilot promotes active zone assembly, Ca2+ channel clustering, and vesicle release. Science 312, 1051–1054 (2006).

  51. 51.

    Fouquet, W. et al. Maturation of active zone assembly by Drosophila Bruchpilot. J. Cell Biol. 186, 129–145 (2009).

  52. 52.

    Böhme, M. A. et al. Active zone scaffolds differentially accumulate Unc13 isoforms to tune Ca(2+) channel-vesicle coupling. Nat. Neurosci. 19, 1311–1320 (2016). This study employs super-resolution imaging at the fly NMJ to analyse the molecular organization of the active zone. The authors discover that a developmental switch between two Unc13 isoforms of different lengths correspond to a change in the organization and calcium coupling of the synapse. The results suggest that two distinct types of calcium coupling may coexist at mature active zones, determined in part by expression of specific Unc13 isoforms.

  53. 53.

    Liu, C. et al. The active zone protein family ELKS supports Ca2+ influx at nerve terminals of inhibitory hippocampal neurons. J. Neurosci. 34, 12289–12303 (2014).

  54. 54.

    Han, Y., Kaeser, P. S., Sudhof, T. C. & Schneggenburger, R. RIM determines Ca(2) + channel density and vesicle docking at the presynaptic active zone. Neuron 69, 304–316 (2011).

  55. 55.

    Grauel, M. K. et al. RIM-binding protein 2 regulates release probability by fine-tuning calcium channel localization at murine hippocampal synapses. Proc. Natl Acad. Sci. USA 113, 11615–11620 (2016).

  56. 56.

    Tong, X. J. et al. Retrograde synaptic inhibition is mediated by α-neurexin binding to the α2δ subunits of N-type calcium channels. Neuron 95, 326–340 (2017).

  57. 57.

    Zamponi, G. & Currie, K. Regulation of Ca(v)2 calcium channels by G protein coupled receptors. Biochim. Biophys. Acta 1828, 1629–1643 (2013).

  58. 58.

    Kim, S. H. & Ryan, T. A. Balance of calcineurin Aalpha and CDK5 activities sets release probability at nerve terminals. J. Neurosci. 33, 8937–8950 (2013).

  59. 59.

    Lee, A. et al. Ca2+/calmodulin binds to and modulates P/Q-type calcium channels. Nature 399, 155–159 (1999).

  60. 60.

    Ben-Johny, M. & Yue, D. T. Calmodulin regulation (calmodulation) of voltage-gated calcium channels. J. Gen. Physiol. 143, 679–692 (2014).

  61. 61.

    Rowan, M. J., Tranquil, E. & Christie, J. M. Distinct Kv channel subtypes contribute to differences in spike signaling properties in the axon initial segment and presynaptic boutons of cerebellar interneurons. J. Neurosci. 34, 6611–6623 (2014).

  62. 62.

    Rowan, M. J., DelCanto, G., Yu, J. J., Kamasawa, N. & Christie, J. M. Synapse-level determination of action potential duration by K(+) channel clustering in axons. Neuron 91, 370–383 (2016).

  63. 63.

    Hoppa, M. B., Gouzer, G., Armbruster, M. & Ryan, T. A. Control and plasticity of the presynaptic action potential waveform at small CNS nerve terminals. Neuron 84, 778–789 (2014).

  64. 64.

    Sabatini, B. L. & Regehr, W. G. Control of neurotransmitter release by presynaptic waveform at the granule cell to Purkinje cell synapse. J. Neurosci. 17, 3425–3435 (1997).

  65. 65.

    Cho, I. H., Panzera, L. C., Chin, M. & Hoppa, M. B. Sodium channel β2 subunits prevent action potential propagation failures at axonal branch points. J. Neurosci. 37, 9519–9533 (2017).

  66. 66.

    Dobrunz, L. E., Huang, E. P. & Stevens, C. F. Very short-term plasticity in hippocampal synapses. Proc. Natl Acad. Sci. USA 94, 14843–14847 (1997).

  67. 67.

    Brody, D. L. & Yue, D. T. Release-independent short-term synaptic depression in cultured hippocampal neurons. J. Neurosci. 20, 2480–2494 (2000).

  68. 68.

    Calloway, N., Gouzer, G., Xue, M. & Ryan, T. A. The active-zone protein Munc13 controls the use-dependence of presynaptic voltage-gated calcium channels. eLife 4, e07728 (2015).

  69. 69.

    Lipstein, N. et al. Dynamic control of synaptic vesicle replenishment and short-term plasticity by Ca2+-calmodulin-Munc13-1 signaling. Neuron 79, 82–96 (2013).

  70. 70.

    Ermolyuk, Y. S. et al. Differential triggering of spontaneous glutamate release by P/Q, N and R-type Ca2+ channels. Nat. Neurosci. 16, 1754–1763 (2013).

  71. 71.

    Landò, L., Giovannini, J. & Zucker, R. S. Cobalt blocks the decrease in MEPSP frequency on depolarization in calcium-free hypertonic media. J. Neurobiol. 17, 707–712 (1986).

  72. 72.

    Schotten, S. et al. Additive effects on the energy barrier for synaptic vesicle fusion cause supralinear effects on the vesicle fusion rate. eLife 4, e05531 (2015).

  73. 73.

    Atasoy, D. et al. Spontaneous and evoked glutamate release activates two populations of NMDA receptors with limited overlap. J. Neurosci. 28, 10151–10166 (2008).

  74. 74.

    Cathala, L., Brickley, S., Cull-Candy, S. & Farrant, M. Maturation of EPSCs and intrinsic membrane properties enhances precision at a cerebellar synapse. J. Neurosci. 23, 6074–6085 (2003).

  75. 75.

    Grafmüller, A., Shillcock, J. & Lipowsky, R. The fusion of membranes and vesicles: pathway and energy barriers from dissipative particle dynamics. Biophys. J. 96, 2658–2675 (2009).

  76. 76.

    Marrink, S. J. & Mark, A. E. The mechanism of vesicle fusion as revealed by molecular dynamics simulations. J. Am. Chem. Soc. 125, 11144–11145 (2003).

  77. 77.

    Smirnova, Y. G., Marrink, S. J., Lipowsky, R. & Knecht, V. Solvent-exposed tails as prestalk transition states for membrane fusion at low hydration. J. Am. Chem. Soc. 132, 6710–6718 (2010).

  78. 78.

    Kasson, P. M. et al. Ensemble molecular dynamics yields submillisecond kinetics and intermediates of membrane fusion. Proc. Natl Acad. Sci. USA 103, 11916–11921 (2006).

  79. 79.

    François-Martin, C., Rothman, J. E. & Pincet, F. Low energy cost for optimal speed and control of membrane fusion. Proc. Natl Acad. Sci. USA 114, 1238–1241 (2017). This study resurrects a classic physicochemical approach to measure the activation energy of spontaneous vesicle fusion. The authors monitor liposome fusion in a bulk fusion assay conducted at a range of temperatures. Fits to the Arrhenius rate law indicate an energy barrier of approximately 30k B T , consistent with spontaneous fusion rates measured at the synapse and much smaller than the 100–150k B T energy barrier proposed in past theoretical studies.

  80. 80.

    Rothman, J. E., Krishnakumar, S. S., Grushin, K. & Pincet, F. Hypothesis — buttressed rings assemble, clamp, and release SNAREpins for synaptic transmission. FEBS Lett. 591, 3459–3480 (2017).

  81. 81.

    Oelkers, M., Witt, H., Halder, P., Jahn, R. & Janshoff, A. SNARE-mediated membrane fusion trajectories derived from force-clamp experiments. Proc. Natl Acad. Sci. USA 113, 13051–13056 (2016).

  82. 82.

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

  83. 83.

    Jahn, R. & Fasshauer, D. Molecular machines governing exocytosis of synaptic vesicles. Nature 490, 201–207 (2012).

  84. 84.

    Lai, Y. et al. Molecular mechanisms of synaptic vesicle priming by Munc13 and Munc18. Neuron 95, 591–607 (2017).

  85. 85.

    Ma, C., Su, L., Seven, A. B., Xu, Y. & Rizo, J. Reconstitution of the vital functions of Munc18 and Munc13 in neurotransmitter release. Science 339, 421–425 (2013). This study uses in vitro fusion assays in the presence of numerous crucial presynaptic fusion proteins to support a new perspective on SNARE-mediated fusion. The authors propose that MUNC13 and MUNC18 orchestrate a particular trajectory for SNARE assembly, protecting bound SNARE complexes from disassembly in the face of constitutive NSF activity. This viewpoint helps explain the well-known observation that vesicle fusion is largely eliminated when either MUNC13 or MUNC18 is absent.

  86. 86.

    Dhara, M., Mohrmann, R. & Bruns, D. v-SNARE function in chromaffin cells. Pflugers Arch. 470, 169–180 (2018).

  87. 87.

    Wu, Z., Thiyagarajan, S., O’Shaughnessy, B. & Karatekin, E. Regulation of exocytotic fusion pores by SNARE protein transmembrane domains. Front. Mol. Neurosci. 10, 315 (2017).

  88. 88.

    Sudhof, T. C. Neurotransmitter release: the last millisecond in the life of a synaptic vesicle. Neuron 80, 675–690 (2013).

  89. 89.

    Wiederhold, K. & Fasshauer, D. Is assembly of the SNARE complex enough to fuel membrane fusion? J. Biol. Chem. 284, 13143–13152 (2009).

  90. 90.

    Li, F., Tiwari, N., Rothman, J. E. & Pincet, F. Kinetic barriers to SNAREpin assembly in the regulation of membrane docking/priming and fusion. Proc. Natl Acad. Sci. USA 113, 10536–10541 (2016).

  91. 91.

    Sørensen, J. B. et al. Sequential N to C-terminal SNARE complex assembly drives priming and fusion of secretory vesicles. EMBO J. 25, 955–966 (2006).

  92. 92.

    Mohrmann, R., de Wit, H., Verhage, M., Neher, E. & Sorensen, J. B. Fast vesicle fusion in living cells requires at least three SNARE complexes. Science 330, 502–505 (2010).

  93. 93.

    Imig, C. et al. The morphological and molecular nature of synaptic vesicle priming at presynaptic active zones. Neuron 84, 416–431 (2014). In this paper, detailed electron tomography of the active zone synaptic vesicle distribution is performed in hippocampal synapses across a variety of synaptic mutants, including SNAREs, MUNC13, synaptotagmin and complexin. The authors conclude that docking (vesicles within 2 nm of the PM) requires MUNC13 and the SNAREs. These results are consistent with the notion that partial assembly of the SNARE complex is necessary to dock and prime a synaptic vesicle.

  94. 94.

    Vasin, A., Volfson, D., Littleton, J. T. & Bykhovskaia, M. Interaction of the complexin accessory helix with synaptobrevin regulates spontaneous fusion. Biophys. J. 111, 1954–1964 (2016).

  95. 95.

    Bao, H. et al. Dynamics and number of trans-SNARE complexes determine nascent fusion pore properties. Nature 554, 260–263 (2018). In this study, SNARE-mediated membrane fusion is examined at an unprecedented time resolution using v-SNARE-containing nanodiscs fusing with t-SNARE planar lipid bilayers. The resulting fusion pores are highly dynamic on the millisecond to second timescale. Surprisingly, the pore flicker dynamics correspond to the metastable assembly of trans -SNAREs, suggesting that even after pore formation, trans -SNARE assembly remains dynamic and reversible.

  96. 96.

    Diao, J. et al. Synaptic proteins promote calcium-triggered fast transition from point contact to full fusion. eLife 1, e00109 (2012).

  97. 97.

    He, E. et al. Munc13-1 and Munc18-1 together prevent NSF-dependent de-priming of synaptic vesicles. Nat. Commun. 8, 15915 (2017).

  98. 98.

    Richmond, J. E., Weimer, R. M. & Jorgensen, E. M. An open form of syntaxin bypasses the requirement for UNC-13 in vesicle priming. Nature 412, 338–341 (2001).

  99. 99.

    Yang, X. et al. Syntaxin opening by the MUN domain underlies the function of Munc13 in synaptic-vesicle priming. Nat. Struct. Mol. Biol. 22, 547–554 (2015).

  100. 100.

    Wang, S. et al. Conformational change of syntaxin linker region induced by Munc13s initiates SNARE complex formation in synaptic exocytosis. EMBO J. 36, 816–829 (2017).

  101. 101.

    Baker, R. W. et al. A direct role for the Sec1/Munc18-family protein Vps33 as a template for SNARE assembly. Science 349, 1111–1114 (2015).

  102. 102.

    Reddy-Alla, S. et al. Stable positioning of Unc13 restricts synaptic vesicle fusion to defined release sites to promote synchronous neurotransmission. Neuron 95, 1350–1364 (2017).

  103. 103.

    Sakamoto, H. et al. Synaptic weight set by Munc13-1 supramolecular assemblies. Nat. Neurosci. 21, 41–49 (2018). In this paper, super-resolution optical imaging of presynaptic terminals is combined with optical quantal analysis of exocytosis to reveal that MUNC13, a protein essential to all known forms of neurotransmitter release, is clustered in small assemblies that likely correspond to release sites for SV fusion.

  104. 104.

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

  105. 105.

    Schluter, O. M., Basu, J., Sudhof, T. C. & Rosenmund, C. Rab3 superprimes synaptic vesicles for release: implications for short-term synaptic plasticity. J. Neurosci. 26, 1239–1246 (2006).

  106. 106.

    Basu, J., Betz, A., Brose, N. & Rosenmund, C. Munc13-1 C1 domain activation lowers the energy barrier for synaptic vesicle fusion. J. Neurosci. 27, 1200–1210 (2007).

  107. 107.

    Taschenberger, H., Woehler, A. & Neher, E. Superpriming of synaptic vesicles as a common basis for intersynapse variability and modulation of synaptic strength. Proc. Natl Acad. Sci. USA 113, E4548–E4557 (2016).

  108. 108.

    Lee, J. S., Ho, W. K., Neher, E. & Lee, S. H. Superpriming of synaptic vesicles after their recruitment to the readily releasable pool. Proc. Natl Acad. Sci. USA 110, 15079–15084 (2013).

  109. 109.

    Rhee, J. S. et al. Beta phorbol ester- and diacylglycerol-induced augmentation of transmitter release is mediated by Munc13s and not by PKCs. Cell 108, 121–133 (2002).

  110. 110.

    Lou, X., Korogod, N., Brose, N. & Schneggenburger, R. Phorbol esters modulate spontaneous and Ca2+-evoked transmitter release via acting on both Munc13 and protein kinase C. J. Neurosci. 28, 8257–8267 (2008).

  111. 111.

    Madison, J. M., Nurrish, S. & Kaplan, J. M. UNC-13 interaction with syntaxin is required for synaptic transmission. Curr. Biol. 15, 2236–2242 (2005).

  112. 112.

    Michelassi, F., Liu, H., Hu, Z. & Dittman, J. S. A C1-C2 module in Munc13 inhibits calcium-dependent neurotransmitter release. Neuron 95, 577–590 (2017).

  113. 113.

    Xu, J. et al. Mechanistic insights into neurotransmitter release and presynaptic plasticity from the crystal structure of Munc13-1 C1C2BMUN. eLife 6, e22567 (2017).

  114. 114.

    Südhof, T. C. The presynaptic active zone. Neuron 75, 11–25 (2012).

  115. 115.

    Augustin, I., Rosenmund, C., Sudhof, T. C. & Brose, N. Munc13-1 is essential for fusion competence of glutamatergic synaptic vesicles. Nature 400, 457–461 (1999).

  116. 116.

    Wu, Z. et al. Dilation of fusion pores by crowding of SNARE proteins. eLife 6, e22964 (2017).

  117. 117.

    Sharma, S. & Lindau, M. t-SNARE transmembrane domain clustering modulates lipid organization and membrane curvature. J. Am. Chem. Soc. 139, 18440–18443 (2017).

  118. 118.

    Chabrol, F. P., Arenz, A., Wiechert, M. T., Margrie, T. W. & DiGregorio, D. A. Synaptic diversity enables temporal coding of coincident multisensory inputs in single neurons. Nat. Neurosci. 18, 718–727 (2015).

  119. 119.

    Llano, I. et al. Presynaptic calcium stores underlie large-amplitude miniature IPSCs and spontaneous calcium transients. Nat. Neurosci. 3, 1256–1265 (2000).

  120. 120.

    de Juan-Sanz, J. et al. Axonal endoplasmic reticulum Ca2+ content controls release probability in CNS nerve terminals. Neuron 93, 867–881 (2017).

  121. 121.

    Neher, E. & Augustine, G. J. Calcium gradients and buffers in bovine chromaffin cells. J. Physiol. 450, 273–301 (1992).

  122. 122.

    Dittrich, M. et al. An excess-calcium-binding-site model predicts neurotransmitter release at the neuromuscular junction. Biophys. J. 104, 2751–2763 (2013).

  123. 123.

    Kawaguchi, S. Y. & Sakaba, T. Control of inhibitory synaptic outputs by low excitability of axon terminals revealed by direct recording. Neuron 85, 1273–1288 (2015).

  124. 124.

    Bucurenciu, I., Bischofberger, J. & Jonas, P. A small number of open Ca2+ channels trigger transmitter release at a central GABAergic synapse. Nat. Neurosci. 13, 19–21 (2010).

  125. 125.

    Wang, L. Y. & Augustine, G. J. Presynaptic nanodomains: a tale of two synapses. Front. Cell Neurosci. 8, 455 (2014).

  126. 126.

    Stern, M. D. Buffering of calcium in the vicinity of a channel pore. Cell Calcium 13, 183–192 (1992).

  127. 127.

    Naraghi, M. & Neher, E. Linearized buffered Ca2+ diffusion in microdomains and its implications for calculation of [Ca2+] at the mouth of a calcium channel. J. Neurosci. 17, 6961–6973 (1997).

  128. 128.

    Smith, G. D. Analytical steady-state solution to the rapid buffering approximation near an open Ca2+ channel. Biophys. J. 71, 3064–3072 (1996).

  129. 129.

    Lis, L. J., McAlister, M., Fuller, N., Rand, R. P. & Parsegian, V. A. Interactions between neutral phospholipid bilayer membranes. Biophys. J. 37, 657–665 (1982).

  130. 130.

    Chernomordik, L. V. & Kozlov, M. M. Mechanics of membrane fusion. Nat. Struct. Mol. Biol. 15, 675–683 (2008).

  131. 131.

    Leikin, S. L., Kozlov, M. M., Chernomordik, L. V., Markin, V. S. & Chizmadzhev, Y. A. Membrane fusion: overcoming of the hydration barrier and local restructuring. J. Theor. Biol. 129, 411–425 (1987).

  132. 132.

    Kozlovsky, Y. & Kozlov, M. M. Stalk model of membrane fusion: solution of energy crisis. Biophys. J. 82, 882–895 (2002).

  133. 133.

    Siegel, D. P. The modified stalk mechanism of lamellar/inverted phase transitions and its implications for membrane fusion. Biophys. J. 76, 291–313 (1999).

  134. 134.

    Kuzmin, P. I., Zimmerberg, J., Chizmadzhev, Y. A. & Cohen, F. S. A quantitative model for membrane fusion based on low-energy intermediates. Proc. Natl Acad. Sci. USA 98, 7235–7240 (2001).

  135. 135.

    Markin, V. S. & Albanesi, J. P. Membrane fusion: stalk model revisited. Biophys. J. 82, 693–712 (2002).

  136. 136.

    Chernomordik, L. V., Zimmerberg, J. & Kozlov, M. M. Membranes of the world unite! J. Cell Biol. 175, 201–207 (2006).

  137. 137.

    Dhara, M. et al. v-SNARE transmembrane domains function as catalysts for vesicle fusion. eLife 5, e17571 (2016).

  138. 138.

    Bao, H. et al. Exocytotic fusion pores are composed of both lipids and proteins. Nat. Struct. Mol. Biol. 23, 67–73 (2016).

  139. 139.

    Cohen, F. S. & Melikyan, G. B. The energetics of membrane fusion from binding, through hemifusion, pore formation, and pore enlargement. J. Membr. Biol. 199, 1–14 (2004).

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Acknowledgements

The authors thank D. DiGregorio (Institut Pasteur) and J. Feigenson (Cornell University) for useful discussions. This work was supported by grants MH085783 (T.A.R.) and GM095674 (J.S.D.).

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Nature Reviews Neuroscience thanks K. Hirose and the other anonymous reviewer(s) for their contribution to the peer review of this work.

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  1. Department of Biochemistry, Weill Cornell Medical College, New York, NY, USA

    • Jeremy S. Dittman
    •  & Timothy A. Ryan

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T.A.R. and J.S.D. made substantial contributions to the discussion of content, writing, review and editing of the manuscript before submission.

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Glossary

Isocalcium surfaces

Convenient graphical representations of the calcium concentration indicating regions that are at the same calcium levels similar to weather maps plotting regions at the same temperature (isotherms) or pressure (isobars).

Diffusion

The random migration of molecules or small particles arising from motion due to thermal energy.

Optical fluctuation analyses

Variant of noise analyses using a fluorescent reporter of calcium to monitor the trial-to-trial fluctuations in calcium levels following a stimulus-triggered calcium transient in order to count the number of calcium channels contributing to the fluorescence.

k B T

A natural unit of thermal energy available to drive nanoscale processes, defined by the product of the Boltzmann constant with absolute thermodynamic temperature (in Kelvin).

Arrhenius law

A rudimentary quantitative relationship between the microscopic rate constant of a transition between two states and the energy barrier (ΔE) separating the states: rate = A exp(−ΔE/kBT), where A is the pre-exponential factor, kB is the Boltzmann constant and T is the absolute thermodynamic temperature. The barrier height is commonly referred to as the activation energy.

SNARE

An acronym derived from soluble N-ethylmaleimide-sensitive factor attachment protein (SNAP) receptor. The SNARE proteins comprise a deeply conserved protein family involved in membrane fusion in eukaryotes. The neuronal SNAREs responsible for synaptic vesicle fusion are syntaxin 1, SNAP25 and VAMP2.

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https://doi.org/10.1038/s41583-018-0111-3