Phase separation at the synapse


Emerging evidence indicates that liquid–liquid phase separation, the formation of a condensed molecular assembly within another diluted aqueous solution, is a means for cells to organize highly condensed biological assemblies (also known as biological condensates or membraneless compartments) with very broad functions and regulatory properties in different subcellular regions. Molecular machineries dictating synaptic transmissions in both presynaptic boutons and postsynaptic densities of neuronal synapses may be such biological condensates. Here we review recent developments showing how phase separation can build dense synaptic molecular clusters, highlight unique features of such condensed clusters in the context of synaptic development and signaling, discuss how aberrant phase-separation-mediated synaptic assembly formation may contribute to dysfunctional signaling in psychiatric disorders, and present some challenges and opportunities of phase separation in synaptic biology.

Access options

Rent or Buy article

Get time limited or full article access on ReadCube.


All prices are NET prices.

Fig. 1: Basic principles of phase separation illustrated by a simple two-component system.
Fig. 2: Phase separation in neurons.
Fig. 3: Phase-separation-mediated formation of PSD assemblies.
Fig. 4: Phase separation in presynaptic boutons.
Fig. 5: Mutual exclusion of excitatory and inhibitory PSD condensates.


  1. 1.

    Ramon y Cajal, S. Un sencillo metodo de coloracion selectiva del reticulo protoplasmatico y sus efectos en los diversos organos nerviosos de vertebrados e invertebrados. Trab. Lab. Invest. Biol. Univ. Madrid 2, 129–221 (1903).

  2. 2.

    Banani, S. F., Lee, H. O., Hyman, A. A. & Rosen, M. K. Biomolecular condensates: organizers of cellular biochemistry. Nat. Rev. Mol. Cell Biol. 18, 285–298 (2017).

  3. 3.

    Shin, Y. & Brangwynne, C. P. Liquid phase condensation in cell physiology and disease. Science 357, eaaf4382 (2017).

  4. 4.

    Hyman, A. A., Weber, C. A. & Jülicher, F. Liquid-liquid phase separation in biology. Annu. Rev. Cell Dev. Biol. 30, 39–58 (2014).

  5. 5.

    Van Treeck, B. & Parker, R. Emerging roles for intermolecular RNA-RNA interactions in RNP assemblies. Cell 174, 791–802 (2018).

  6. 6.

    Harris, K. M. & Weinberg, R. J. Ultrastructure of synapses in the mammalian brain. Cold Spring Harb. Perspect. Biol. 4, a005587 (2012).

  7. 7.

    Chen, X. et al. Organization of the core structure of the postsynaptic density. Proc. Natl Acad. Sci. USA 105, 4453–4458 (2008).

  8. 8.

    Blomberg, F., Cohen, R. S. & Siekevitz, P. The structure of postsynaptic densities isolated from dog cerebral cortex. II. Characterization and arrangement of some of the major proteins within the structure. J. Cell Biol. 74, 204–225 (1977).

  9. 9.

    Couteaux, R. & Pécot-Dechavassine, M. [Synaptic vesicles and pouches at the level of “active zones” of the neuromuscular junction]. C. R. Acad. Sci. Hebd. Seances Acad. Sci. D 271, 2346–2349 (1970).

  10. 10.

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

  11. 11.

    Biederer, T., Kaeser, P. S. & Blanpied, T. A. Transcellular nanoalignment of synaptic function. Neuron 96, 680–696 (2017).

  12. 12.

    Rizzoli, S. O. & Betz, W. J. Synaptic vesicle pools. Nat. Rev. Neurosci. 6, 57–69 (2005).

  13. 13.

    Alabi, A. A. & Tsien, R. W. Synaptic vesicle pools and dynamics. Cold Spring Harb. Perspect. Biol. 4, a013680 (2012).

  14. 14.

    Zeng, M. et al. Phase transition in postsynaptic densities underlies formation of synaptic complexes and synaptic plasticity. Cell 166, 1163–1175.e12 (2016).

  15. 15.

    Zeng, M. et al. Reconstituted postsynaptic density as a molecular platform for understanding synapse formation and plasticity. Cell 174, 1172–1187.e16 (2018).

  16. 16.

    Milovanovic, D., Wu, Y., Bian, X. & De Camilli, P. A liquid phase of synapsin and lipid vesicles. Science 361, 604–607 (2018).

  17. 17.

    Wu, X. et al. RIM and RIM-BP form presynaptic active-zone-like condensates via phase separation. Mol. Cell 73, 971–984.e5 (2019).

  18. 18.

    Nedelsky, N. B. & Taylor, J. P. Bridging biophysics and neurology: aberrant phase transitions in neurodegenerative disease. Nat. Rev. Neurol. 15, 272–286 (2019).

  19. 19.

    Elbaum-Garfinkle, S. Matter over mind: liquid phase separation and neurodegeneration. J. Biol. Chem. 294, 7160–7168 (2019).

  20. 20.

    Taylor, J. P., Brown, R. H. Jr. & Cleveland, D. W. Decoding ALS: from genes to mechanism. Nature 539, 197–206 (2016).

  21. 21.

    Palay, S. L. Synapses in the central nervous system. J. Biophys. Biochem. Cytol. 2(Suppl), 193–202 (1956).

  22. 22.

    Gray, E. G. Axo-somatic and axo-dendritic synapses of the cerebral cortex: an electron microscope study. J. Anat. 93, 420–433 (1959).

  23. 23.

    Banker, G., Churchill, L. & Cotman, C. W. Proteins of the postsynaptic density. J. Cell Biol. 63, 456–465 (1974).

  24. 24.

    Cheng, D. et al. Relative and absolute quantification of postsynaptic density proteome isolated from rat forebrain and cerebellum. Mol. Cell. Proteomics 5, 1158–1170 (2006).

  25. 25.

    Roy, M. et al. Proteomic analysis of postsynaptic proteins in regions of the human neocortex. Nat. Neurosci. 21, 130–138 (2018).

  26. 26.

    Wilson, R. S. et al. Development of targeted mass spectrometry-based approaches for quantitation of proteins enriched in the postsynaptic density (PSD). Proteomes 7, 12 (2019).

  27. 27.

    Kennedy, M. B. The postsynaptic density at glutamatergic synapses. Trends Neurosci. 20, 264–268 (1997).

  28. 28.

    Sheng, M. & Hoogenraad, C. C. The postsynaptic architecture of excitatory synapses: a more quantitative view. Annu. Rev. Biochem. 76, 823–847 (2007).

  29. 29.

    Li, J. et al. Spatiotemporal profile of postsynaptic interactomes integrates components of complex brain disorders. Nat. Neurosci. 20, 1150–1161 (2017).

  30. 30.

    Cohen, R. S., Blomberg, F., Berzins, K. & Siekevitz, P. The structure of postsynaptic densities isolated from dog cerebral cortex. I. Overall morphology and protein composition. J. Cell Biol. 74, 181–203 (1977).

  31. 31.

    Kornau, H. C., Schenker, L. T., Kennedy, M. B. & Seeburg, P. H. Domain interaction between NMDA receptor subunits and the postsynaptic density protein PSD-95. Science 269, 1737–1740 (1995).

  32. 32.

    Kim, E. et al. GKAP, a novel synaptic protein that interacts with the guanylate kinase-like domain of the PSD-95/SAP90 family of channel clustering molecules. J. Cell Biol. 136, 669–678 (1997).

  33. 33.

    Naisbitt, S. et al. Shank, a novel family of postsynaptic density proteins that binds to the NMDA receptor/PSD-95/GKAP complex and cortactin. Neuron 23, 569–582 (1999).

  34. 34.

    Xiao, B. et al. Homer regulates the association of group 1 metabotropic glutamate receptors with multivalent complexes of homer-related, synaptic proteins. Neuron 21, 707–716 (1998).

  35. 35.

    Berry, K. P. & Nedivi, E. Spine dynamics: are they all the same? Neuron 96, 43–55 (2017).

  36. 36.

    Nishiyama, J. & Yasuda, R. Biochemical computation for spine structural plasticity. Neuron 87, 63–75 (2015).

  37. 37.

    Harris, K. M., Jensen, F. E. & Tsao, B. Three-dimensional structure of dendritic spines and synapses in rat hippocampus (CA1) at postnatal day 15 and adult ages: implications for the maturation of synaptic physiology and long-term potentiation. J. Neurosci. 12, 2685–2705 (1992).

  38. 38.

    Huganir, R. L. & Nicoll, R. A. AMPARs and synaptic plasticity: the last 25 years. Neuron 80, 704–717 (2013).

  39. 39.

    Borgdorff, A. J. & Choquet, D. Regulation of AMPA receptor lateral movements. Nature 417, 649–653 (2002).

  40. 40.

    Heine, M. et al. Surface mobility of postsynaptic AMPARs tunes synaptic transmission. Science 320, 201–205 (2008).

  41. 41.

    Makino, H. & Malinow, R. AMPA receptor incorporation into synapses during LTP: the role of lateral movement and exocytosis. Neuron 64, 381–390 (2009).

  42. 42.

    MacGillavry, H. D., Song, Y., Raghavachari, S. & Blanpied, T. A. Nanoscale scaffolding domains within the postsynaptic density concentrate synaptic AMPA receptors. Neuron 78, 615–622 (2013).

  43. 43.

    Blanpied, T. A., Kerr, J. M. & Ehlers, M. D. Structural plasticity with preserved topology in the postsynaptic protein network. Proc. Natl Acad. Sci. USA 105, 12587–12592 (2008).

  44. 44.

    Nair, D. et al. Super-resolution imaging reveals that AMPA receptors inside synapses are dynamically organized in nanodomains regulated by PSD95. J. Neurosci. 33, 13204–13224 (2013).

  45. 45.

    Bosch, M. et al. Structural and molecular remodeling of dendritic spine substructures during long-term potentiation. Neuron 82, 444–459 (2014).

  46. 46.

    Kuriu, T., Inoue, A., Bito, H., Sobue, K. & Okabe, S. Differential control of postsynaptic density scaffolds via actin-dependent and -independent mechanisms. J. Neurosci. 26, 7693–7706 (2006).

  47. 47.

    Kim, J. H., Liao, D., Lau, L. F. & Huganir, R. L. SynGAP: a synaptic RasGAP that associates with the PSD-95/SAP90 protein family. Neuron 20, 683–691 (1998).

  48. 48.

    Chen, H. J., Rojas-Soto, M., Oguni, A. & Kennedy, M. B. A synaptic Ras-GTPase activating protein (p135 SynGAP) inhibited by CaM kinase II. Neuron 20, 895–904 (1998).

  49. 49.

    Araki, Y., Zeng, M., Zhang, M. & Huganir, R. L. Rapid dispersion of SynGAP from synaptic spines triggers AMPA receptor insertion and spine enlargement during LTP. Neuron 85, 173–189 (2015).

  50. 50.

    Pena, V. et al. The C2 domain of SynGAP is essential for stimulation of the Rap GTPase reaction. EMBO Rep. 9, 350–355 (2008).

  51. 51.

    Vazquez, L. E., Chen, H.-J., Sokolova, I., Knuesel, I. & Kennedy, M. B. SynGAP regulates spine formation. J. Neurosci. 24, 8862–8872 (2004).

  52. 52.

    Zeng, M., Bai, G. & Zhang, M. Anchoring high concentrations of SynGAP at postsynaptic densities via liquid-liquid phase separation. Small GTPases 10, 296–304 (2017).

  53. 53.

    Clement, J. P. et al. Pathogenic SYNGAP1 mutations impair cognitive development by disrupting maturation of dendritic spine synapses. Cell 151, 709–723 (2012).

  54. 54.

    Kilinc, M. et al. Species-conserved SYNGAP1 phenotypes associated with neurodevelopmental disorders. Mol. Cell. Neurosci. 91, 140–150 (2018).

  55. 55.

    Zeng, M., Ye, F., Xu, J. & Zhang, M. PDZ ligand binding-induced conformational coupling of the PDZ-SH3-GK tandems in PSD-95 family MAGUKs. J. Mol. Biol. 430, 69–86 (2018).

  56. 56.

    McGee, A. W. et al. Structure of the SH3-guanylate kinase module from PSD-95 suggests a mechanism for regulated assembly of MAGUK scaffolding proteins. Mol. Cell 8, 1291–1301 (2001).

  57. 57.

    Li, P. et al. Phase transitions in the assembly of multivalent signalling proteins. Nature 483, 336–340 (2012).

  58. 58.

    Walkup, W. G. et al. A model for regulation by SynGAP-alpha1 of binding of synaptic proteins to PDZ-domain ‘Slots’ in the postsynaptic density. eLife 5, e16813 (2016).

  59. 59.

    Petralia, R. S., Sans, N., Wang, Y. X. & Wenthold, R. J. Ontogeny of postsynaptic density proteins at glutamatergic synapses. Mol. Cell. Neurosci. 29, 436–452 (2005).

  60. 60.

    Valtschanoff, J. G. & Weinberg, R. J. Laminar organization of the NMDA receptor complex within the postsynaptic density. J. Neurosci. 21, 1211–1217 (2001).

  61. 61.

    Dosemeci, A., Weinberg, R. J., Reese, T. S. & Tao-Cheng, J. H. The postsynaptic density: there is more than meets the eye. Front. Synaptic Neurosci. 8, 23 (2016).

  62. 62.

    Lowenthal, M. S., Markey, S. P. & Dosemeci, A. Quantitative mass spectrometry measurements reveal stoichiometry of principal postsynaptic density proteins. J. Proteome Res. 14, 2528–2538 (2015).

  63. 63.

    Ting, J. T., Peça, J. & Feng, G. Functional consequences of mutations in postsynaptic scaffolding proteins and relevance to psychiatric disorders. Annu. Rev. Neurosci. 35, 49–71 (2012).

  64. 64.

    Zhu, J., Shang, Y. & Zhang, M. Mechanistic basis of MAGUK-organized complexes in synaptic development and signalling. Nat. Rev. Neurosci. 17, 209–223 (2016).

  65. 65.

    Feng, W. & Zhang, M. Organization and dynamics of PDZ-domain-related supramodules in the postsynaptic density. Nat. Rev. Neurosci. 10, 87–99 (2009).

  66. 66.

    Zeng, M. et al. Phase separation-mediated TARP/MAGUK complex condensation and AMPA receptor synaptic transmission. Neuron 104, 529–543.e6 (2019).

  67. 67.

    Landis, D. M. D., Hall, A. K., Weinstein, L. A. & Reese, T. S. The organization of cytoplasm at the presynaptic active zone of a central nervous system synapse. Neuron 1, 201–209 (1988).

  68. 68.

    Rosahl, T. W. et al. Essential functions of synapsins I and II in synaptic vesicle regulation. Nature 375, 488–493 (1995).

  69. 69.

    Pieribone, V. A. et al. Distinct pools of synaptic vesicles in neurotransmitter release. Nature 375, 493–497 (1995).

  70. 70.

    Milovanovic, D. & De Camilli, P. Synaptic vesicle clusters at synapses: a distinct liquid phase? Neuron 93, 995–1002 (2017).

  71. 71.

    Siksou, L. et al. Three-dimensional architecture of presynaptic terminal cytomatrix. J. Neurosci. 27, 6868–6877 (2007).

  72. 72.

    Acuna, C., Liu, X. & Südhof, T. C. How to make an active zone: unexpected universal functional redundancy between RIMs and RIM-BPs. Neuron 91, 792–807 (2016).

  73. 73.

    Wang, S. S. H. et al. Fusion competent synaptic vesicles persist upon active zone disruption and loss of vesicle docking. Neuron 91, 777–791 (2016).

  74. 74.

    Benfenati, F., Bähler, M., Jahn, R. & Greengard, P. Interactions of synapsin I with small synaptic vesicles: distinct sites in synapsin I bind to vesicle phospholipids and vesicle proteins. J. Cell Biol. 108, 1863–1872 (1989).

  75. 75.

    Südhof, T. C. et al. Synapsins: mosaics of shared and individual domains in a family of synaptic vesicle phosphoproteins. Science 245, 1474–1480 (1989).

  76. 76.

    Shupliakov, O., Haucke, V. & Pechstein, A. How synapsin I may cluster synaptic vesicles. Semin. Cell Dev. Biol. 22, 393–399 (2011).

  77. 77.

    Esser, L. et al. Synapsin I is structurally similar to ATP-utilizing enzymes. EMBO J. 17, 977–984 (1998).

  78. 78.

    Hosaka, M. & Südhof, T. C. Homo- and heterodimerization of synapsins. J. Biol. Chem. 274, 16747–16753 (1999).

  79. 79.

    Hosaka, M., Hammer, R. E. & Südhof, T. C. A phospho-switch controls the dynamic association of synapsins with synaptic vesicles. Neuron 24, 377–387 (1999).

  80. 80.

    Cheetham, J. J. et al. Identification of synapsin I peptides that insert into lipid membranes. Biochem. J. 354, 57–66 (2001).

  81. 81.

    De Camilli, P., Harris, S. M. Jr., Huttner, W. B. & Greengard, P. Synapsin I (Protein I), a nerve terminal-specific phosphoprotein. II. Its specific association with synaptic vesicles demonstrated by immunocytochemistry in agarose-embedded synaptosomes. J. Cell Biol. 96, 1355–1373 (1983).

  82. 82.

    Chi, P., Greengard, P. & Ryan, T. A. Synapsin dispersion and reclustering during synaptic activity. Nat. Neurosci. 4, 1187–1193 (2001).

  83. 83.

    Benfenati, F. et al. Synaptic vesicle-associated Ca2+/calmodulin-dependent protein kinase II is a binding protein for synapsin I. Nature 359, 417–420 (1992).

  84. 84.

    Zhai, R. G. & Bellen, H. J. The architecture of the active zone in the presynaptic nerve terminal. Physiology (Bethesda) 19, 262–270 (2004).

  85. 85.

    Tang, A. H. et al. A trans-synaptic nanocolumn aligns neurotransmitter release to receptors. Nature 536, 210–214 (2016).

  86. 86.

    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).

  87. 87.

    Nakamura, Y. et al. Nanoscale distribution of presynaptic Ca(2+) channels and its impact on vesicular release during development. Neuron 85, 145–158 (2015).

  88. 88.

    Eggermann, E., Bucurenciu, I., Goswami, S. P. & Jonas, P. Nanodomain coupling between Ca2+ channels and sensors of exocytosis at fast mammalian synapses. Nat. Rev. Neurosci. 13, 7–21 (2011).

  89. 89.

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

  90. 90.

    Kaeser, P. S. et al. RIM proteins tether Ca2+ channels to presynaptic active zones via a direct PDZ-domain interaction. Cell 144, 282–295 (2011).

  91. 91.

    Acuna, C., Liu, X., Gonzalez, A. & Südhof, T. C. RIM-BPs mediate tight coupling of action potentials to Ca(2+)-triggered neurotransmitter release. Neuron 87, 1234–1247 (2015).

  92. 92.

    Wilhelm, B. G. et al. Composition of isolated synaptic boutons reveals the amounts of vesicle trafficking proteins. Science 344, 1023–1028 (2014).

  93. 93.

    Galkin, O., Chen, K., Nagel, R. L., Hirsch, R. E. & Vekilov, P. G. Liquid-liquid separation in solutions of normal and sickle cell hemoglobin. Proc. Natl Acad. Sci. USA 99, 8479–8483 (2002).

  94. 94.

    Brangwynne, C. P. et al. Germline P granules are liquid droplets that localize by controlled dissolution/condensation. Science 324, 1729–1732 (2009).

  95. 95.

    Zhu, J. et al. Synaptic targeting and function of SAPAPs mediated by phosphorylation-dependent binding to PSD-95 MAGUKs. Cell Rep. 21, 3781–3793 (2017).

  96. 96.

    Xiao, B., Tu, J. C. & Worley, P. F. Homer: a link between neural activity and glutamate receptor function. Curr. Opin. Neurobiol. 10, 370–374 (2000).

  97. 97.

    Sala, C. et al. Inhibition of dendritic spine morphogenesis and synaptic transmission by activity-inducible protein Homer1a. J. Neurosci. 23, 6327–6337 (2003).

  98. 98.

    Diering, G. H. et al. Homer1a drives homeostatic scaling-down of excitatory synapses during sleep. Science 355, 511–515 (2017).

  99. 99.

    de Vivo, L. et al. Ultrastructural evidence for synaptic scaling across the wake/sleep cycle. Science 355, 507–510 (2017).

  100. 100.

    Kubota, Y., Hatada, S., Kondo, S., Karube, F. & Kawaguchi, Y. Neocortical inhibitory terminals innervate dendritic spines targeted by thalamocortical afferents. J. Neurosci. 27, 1139–1150 (2007).

  101. 101.

    Villa, K. L. et al. Inhibitory synapses are repeatedly assembled and removed at persistent sites in vivo. Neuron 89, 756–769 (2016).

  102. 102.

    Kato, M. et al. Cell-free formation of RNA granules: low complexity sequence domains form dynamic fibers within hydrogels. Cell 149, 753–767 (2012).

  103. 103.

    Kim, H. J. et al. Mutations in prion-like domains in hnRNPA2B1 and hnRNPA1 cause multisystem proteinopathy and ALS. Nature 495, 467–473 (2013).

  104. 104.

    Toretsky, J. A. & Wright, P. E. Assemblages: functional units formed by cellular phase separation. J. Cell Biol. 206, 579–588 (2014).

  105. 105.

    Jiang, H. et al. Phase transition of spindle-associated protein regulate spindle apparatus assembly. Cell 163, 108–122 (2015).

  106. 106.

    Molliex, A. et al. Phase separation by low complexity domains promotes stress granule assembly and drives pathological fibrillization. Cell 163, 123–133 (2015).

  107. 107.

    Patel, A. et al. A liquid-to-solid phase transition of the ALS protein FUS accelerated by disease mutation. Cell 162, 1066–1077 (2015).

  108. 108.

    Wang, J. et al. A molecular grammar governing the driving forces for phase separation of prion-like RNA binding proteins. Cell 174, 688–699.e16 (2018).

  109. 109.

    Baldan, A. Progress in Ostwald ripening theories and their applications to nickel-base superalloys. J. Mater. Sci. 37, 2171–2202 (2002).

  110. 110.

    Guo, L. et al. Nuclear-import receptors reverse aberrant phase transitions of RNA-binding proteins with prion-like domains. Cell 173, 677–692.e20 (2018).

  111. 111.

    Qamar, S. et al. FUS phase separation is modulated by a molecular chaperone and methylation of arginine cation-π interactions. Cell 173, 720–734.e15 (2018).

Download references


Work in our laboratory is supported by grants from RGC of Hong Kong (AoE-M09-12 and C6004-17G) and a grant from Simons Foundation for Autism Research (510178). M.Z. receives support from a Kerry Holdings Professorship of Science and a Senior Fellowship of IAS at HKUST.

Author information

Correspondence to Mingjie Zhang.

Ethics declarations

Competing interests

The authors declare no competing interests.

Additional information

Peer review information Nature Neuroscience thanks Thomas Biederer 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.

Rights and permissions

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Chen, X., Wu, X., Wu, H. et al. Phase separation at the synapse. Nat Neurosci (2020).

Download citation