Abstract
Cell–cell interfaces are found throughout multicellular organisms, from transient interactions between motile immune cells to long-lived cell–cell contacts in epithelia. Studies of immune cell interactions, epithelial cell barriers, neuronal contacts and sites of cell–cell fusion have identified a core set of features shared by cell–cell interfaces that critically control their function. Data from diverse cell types also show that cells actively and passively regulate the localization, strength, duration and cytoskeletal coupling of receptor interactions governing cell–cell signalling and physical connections between cells, indicating that cell–cell interfaces have a unique membrane organization that emerges from local molecular and cellular mechanics. In this Review, we discuss recent findings that support the emerging view of cell–cell interfaces as specialized compartments that biophysically constrain the arrangement and activity of their protein, lipid and glycan components. We also review how these biophysical features of cell–cell interfaces allow cells to respond with high selectivity and sensitivity to multiple inputs, serving as the basis for wide-ranging cellular functions. Finally, we consider how the unique properties of cell–cell interfaces present opportunities for therapeutic intervention.
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References
Honig, B. & Shapiro, L. Adhesion protein structure, molecular affinities, and principles of cell–cell recognition. Cell 181, 520–535 (2020).
Rozbesky, D. & Jones, E. Y. Cell guidance ligands, receptors and complexes—orchestrating signalling in time and space. Curr. Opin. Struct. Biol. 61, 79–85 (2020).
Helle, S. C. J. et al. Organization and function of membrane contact sites. Biochim. Biophys. Acta Mol. Cell Res. 1833, 2526–2541 (2013).
Phillips, M. J. & Voeltz, G. K. Structure and function of ER membrane contact sites with other organelles. Nat. Rev. Mol. Cell Biol. 17, 69–82 (2016).
Prinz, W. A. Bridging the gap: membrane contact sites in signaling, metabolism, and organelle dynamics. J. Cell Biol. 205, 759–769 (2014).
Prinz, W. A., Toulmay, A. & Balla, T. The functional universe of membrane contact sites. Nat. Rev. Mol. Cell Biol. 21, 7–24 (2020).
Singer, S. J. & Nicolson, G. L. The fluid mosaic model of the structure of cell membranes. Science 175, 720–731 (1972).
Aimon, S. et al. Membrane shape modulates transmembrane protein distribution. Dev. Cell 28, 212–218 (2014).
Domanov, Y. A. et al. Mobility in geometrically confined membranes. Proc. Natl Acad. Sci. USA 108, 12605–12610 (2011).
Fenz, S. F., Merkel, R. & Sengupta, K. Diffusion and intermembrane distance: case study of avidin and E-cadherin mediated adhesion. Langmuir 25, 1074–1085 (2009).
Thoumine, O., Lambert, M., Mège, R.-M. & Choquet, D. Regulation of N-cadherin dynamics at neuronal contacts by ligand binding and cytoskeletal coupling. Mol. Biol. Cell 17, 862–875 (2006).
Cavey, M., Rauzi, M., Lenne, P.-F. & Lecuit, T. A two-tiered mechanism for stabilization and immobilization of E-cadherin. Nature 453, 751–756 (2008).
Nusrat, A. et al. Tight junctions are membrane microdomains. J. Cell Sci. 113, 1771–1781 (2000).
Shigetomi, K., Ono, Y., Inai, T. & Ikenouchi, J. Adherens junctions influence tight junction formation via changes in membrane lipid composition. J. Cell Biol. 217, 2373–2381 (2018). This study finds that cholesterol is enriched at tight junction membranes and, surprisingly, that the lipid compositional bias is not regulated by the tight junction but by the neighbouring adherens junction.
Lewis, J. D. et al. The desmosome is a mesoscale lipid raft-like membrane domain. Mol. Biol. Cell 30, 1390–1405 (2019). This study finds that the desmoglein 1 long transmembrane domain (24 amino acids), in cooperation with cholesterol, contributes to increased lipid thickness at desmosomes and that decreasing the height of the transmembrane domain causes severe dermatitis by compromising desmoglein 1 incorporation into desmosomes.
Freeman, S. A. et al. Integrins form an expanding diffusional barrier that coordinates phagocytosis. Cell 164, 128–140 (2016). This study uses a micropatterned IgG surface to examine macrophage interface (~6 nm) organization during Fcγ receptor-driven phagocytosis and finds that integrins actively set up around Fcγ receptor to augment and reinforce height differences between the interface and CD45.
Freeman, S. A. et al. Transmembrane pickets connect cyto- and pericellular skeletons forming barriers to receptor engagement. Cell 172, 305–317.e10 (2018). This study shows that CD44 in a macrophage complexes hyaluronic acid, a type of proteoglycan, through its Link domain in the extracellular space and creates a pericellular picket fence that restricts Fcγ receptor mobility.
Ostrowski, P. P., Grinstein, S. & Freeman, S. A. Diffusion barriers, mechanical forces, and the biophysics of phagocytosis. Dev. Cell 38, 135–146 (2016).
Bakalar, M. H. et al. Size-dependent segregation controls macrophage phagocytosis of antibody-opsonized targets. Cell 174, 131–142.e13 (2018). This study shows that macrophage phagocytosis scales with the size of the antibody–antigen complex compared with the size of CD45, with the most efficient engulfment seen for antigens that were <10 nm in size.
Felce, J. H. et al. CD45 exclusion- and cross-linking-based receptor signaling together broaden FcεRI reactivity. Sci. Signal. 11, eaat0756 (2018).
Goodridge, H. S. et al. Activation of the innate immune receptor Dectin-1 upon formation of a ‘phagocytic synapse’. Nature 472, 471–475 (2011).
James, J. R. & Vale, R. D. Biophysical mechanism of T-cell receptor triggering in a reconstituted system. Nature 487, 64–69 (2012).
Varma, R., Campi, G., Yokosuka, T., Saito, T. & Dustin, M. L. T cell receptor-proximal signals are sustained in peripheral microclusters and terminated in the central supramolecular activation cluster. Immunity 25, 117–127 (2006).
Schmid, E. M. et al. Size-dependent protein segregation at membrane interfaces. Nat. Phys. 12, 704–711 (2016). This study uses a series of binding and non-binding proteins of different heights to form adhesions between giant unilamellar vesicles or between a giant unilamellar vesicle and a supported lipid bilayer, and finds that protein height differences of 5 nm or more can drive switch-like exclusion of proteins from in vitro membrane interfaces.
Farquhar, M. G. & Palade, G. E. Junctional complexes in various epithelia. J. Cell Biol. 17, 375–412 (1963).
Franke, W. W. Discovering the molecular components of intercellular junctions—a historical view. Cold Spring Harb. Perspect. Biol. 1, a003061 (2009).
Hoffman, D. P. et al. Correlative three-dimensional super-resolution and block-face electron microscopy of whole vitreously frozen cells. Science 367, eaaz5357 (2020).
Weikl, T. R. & Lipowsky, R. in Advances in Planar Lipid Bilayers and Liposomes Vol. 5 Ch. 4 (ed. Leitmannova Liu, A.). 63–127 (Academic Press, 2006).
Paszek, M., Boettiger, D., Weaver, V. & Hammer, D. Integrin clustering is driven by mechanical resistance from the glycocalyx and the substrate. PLoS Comput. Biol. 5, e1000604 (2009).
Hakomori, S. Tumor malignancy defined by aberrant glycosylation and sphingo(glyco)lipid metabolism. Cancer Res. 56, 5309–5318 (1996).
Hollingsworth, M. A. & Swanson, B. J. Mucins in cancer: protection and control of the cell surface. Nat. Rev. Cancer 4, 45–60 (2004).
Horm, T. M. & Schroeder, J. A. MUC1 and metastatic cancer. Cell Adh. Migr. 7, 187–198 (2013).
Paszek, M. J. et al. The cancer glycocalyx mechanically primes integrin-mediated growth and survival. Nature 511, 319–325 (2014).
Woods, E. C. et al. A bulky glycocalyx fosters metastasis formation by promoting G1 cell cycle progression. eLife 6, e25752 (2017).
Chen, W., Lou, J. & Zhu, C. Forcing switch from short- to intermediate- and long-lived states of the αA domain generates LFA-1/ICAM-1 catch bonds. J. Biol. Chem. 285, 35967–35978 (2010).
Franziska Fenz, S., Smith, A.-S., Merkel, R. & Sengupta, K. Inter-membrane adhesion mediated by mobile linkers: effect of receptor shortage. Soft Matter 7, 952–962 (2011).
Dustin, M. L. & Colman, D. R. Neural and immunological synaptic relations. Science 298, 785–789 (2002).
Fenz, S. F. et al. Membrane fluctuations mediate lateral interaction between cadherin bonds. Nat. Phys. 13, 906–913 (2017). By interrogating E-cadherin bond formation between a giant unilamellar vesicle and a supported lipid bilayer, this paper finds that membrane fluctuations give rise to numerous adhesion morphologies by promoting a series of events — formation of trans dimers at an interface, cis aggregation over a lateral range of ~120–500 nm and E-cadherin clustering at an interface.
Steinkühler, J. et al. Membrane fluctuations and acidosis regulate cooperative binding of ‘marker of self’ protein CD47 with the macrophage checkpoint receptor SIRPα. J. Cell Sci. 132, jcs216770 (2019).
Biswas, K. H. et al. E-Cadherin junction formation involves an active kinetic nucleation process. Proc. Natl Acad. Sci. USA 112, 10932–10937 (2015).
Cai, E. et al. Visualizing dynamic microvillar search and stabilization during ligand detection by T cells. Science 356, eaal3118 (2017).
Bell, G. I. Models for the specific adhesion of cells to cells. Science 200, 618–627 (1978). Pioneering theoretical framework for bond formation at membrane interfaces.
Dustin, M. L., Ferguson, L. M., Chan, P. Y., Springer, T. A. & Golan, D. E. Visualization of CD2 interaction with LFA-3 and determination of the two-dimensional dissociation constant for adhesion receptors in a contact area. J. Cell Biol. 132, 465–474 (1996).
Pielak, R. M. et al. Early T cell receptor signals globally modulate ligand:receptor affinities during antigen discrimination. Proc. Natl Acad. Sci. USA 114, 12190–12195 (2017).
Wu, Y., Vendome, J., Shapiro, L., Ben-Shaul, A. & Honig, B. Transforming binding affinities from three dimensions to two with application to cadherin clustering. Nature 475, 510–513 (2011). This paper uses Monte Carlo simulations to calculate the 2D affinity constant for E-cadherin and N-cadherin trans interaction, and finds that domains enriched in cadherins emerge because the trans association alters the energetic landscape for the following cis interaction.
Özkan, E. et al. Extracellular architecture of the SYG-1/SYG-2 adhesion complex instructs synaptogenesis. Cell 156, 482–494 (2014).
Shapiro, L. & Weis, W. I. Structure and biochemistry of cadherins and catenins. Cold Spring Harb. Perspect. Biol. 1, a003053 (2009).
Brasch, J. et al. Visualization of clustered protocadherin neuronal self-recognition complexes. Nature 569, 280–283 (2019).
Schwartz, J. C., Zhang, X., Fedorov, A. A., Nathenson, S. G. & Almo, S. C. Structural basis for co-stimulation by the human CTLA-4/B7-2 complex. Nature 410, 604–608 (2001).
Schubert, D. et al. Autosomal dominant immune dysregulation syndrome in humans with CTLA4 mutations. Nat. Med. 20, 1410–1416 (2014).
Chan, A. C. et al. Activation of ZAP-70 kinase activity by phosphorylation of tyrosine 493 is required for lymphocyte antigen receptor function. EMBO J. 14, 2499–2508 (1995).
Choudhuri, K. et al. Polarized release of T-cell-receptor-enriched microvesicles at the immunological synapse. Nature 507, 118–123 (2014).
Campi, G., Varma, R. & Dustin, M. L. Actin and agonist MHC–peptide complex-dependent T cell receptor microclusters as scaffolds for signaling. J. Exp. Med. 202, 1031–1036 (2005).
Springer, T. A. Adhesion receptors of the immune system. Nature 346, 425–434 (1990).
Courtney, A. H. et al. CD45 functions as a signaling gatekeeper in T cells. Sci. Signal. 12, eaaw8151 (2019).
Cai, H. et al. Full control of ligand positioning reveals spatial thresholds for T cell receptor triggering. Nat. Nanotechnol. 13, 610–617 (2018).
Chang, V. T. et al. Initiation of T cell signaling by CD45 segregation at ‘close contacts’. Nat. Immunol. 17, 574–582 (2016).
Taylor, M. J., Husain, K., Gartner, Z. J., Mayor, S. & Vale, R. D. A DNA-based T cell receptor reveals a role for receptor clustering in ligand discrimination. Cell 169, 108–119.e20 (2017). This study precisely controls the binding affinities of TCR–pMHC using a DNA-conjugated chimeric TCR and MHC, and finds that the TCR–MHC clusters were formed because kon for new bindings increases directly adjacent to preformed ligated receptors, contributing both to ‘kinetic proofreading’ and ‘kinetic segregation’.
Bays, J. L., Campbell, H. K., Heidema, C., Sebbagh, M. & DeMali, K. A. Linking E-cadherin mechanotransduction to cell metabolism through force-mediated activation of AMPK. Nat. Cell Biol. 19, 724–731 (2017).
Finer, J. T., Simmons, R. M. & Spudich, J. A. Single myosin molecule mechanics: picoNewton forces and nanometre steps. Nature 368, 113–119 (1994).
Liu, B., Chen, W., Evavold, B. D. & Zhu, C. Accumulation of dynamic catch bonds between TCR and agonist peptide-MHC triggers T cell signaling. Cell 157, 357–368 (2014). This study uses a state-of-the-art pipette pulling assay to elucidate a catch bond behaviour exclusively for the TCRs bound to matching pMHCs.
Murugesan, S. et al. Formin-generated actomyosin arcs propel T cell receptor microcluster movement at the immune synapse. J. Cell Biol. 215, 383–399 (2016).
Huppa, J. B. et al. TCR–peptide-MHC interactions in situ show accelerated kinetics and increased affinity. Nature 463, 963–967 (2010).
Sibener, L. V. et al. Isolation of a structural mechanism for uncoupling T cell receptor signaling from peptide-MHC binding. Cell 174, 672–687.e27 (2018). This paper identifies a region within the peptide loaded in MHC that directly forms a catch bond with the TCR.
Luca, V. C. et al. Notch–Jagged complex structure implicates a catch bond in tuning ligand sensitivity. Science 355, 1320–1324 (2017).
Borghi, N. et al. E-Cadherin is under constitutive actomyosin-generated tension that is increased at cell–cell contacts upon externally applied stretch. Proc. Natl Acad. Sci. USA 109, 12568–12573 (2012). This paper demonstrates that membrane-bound E-cadherin is under tension by installing a tension sensor into the cytoplasmic domain of E-cadherin.
Yao, M. et al. Force-dependent conformational switch of α-catenin controls vinculin binding. Nat. Commun. 5, 4525 (2014).
Huang, D. L., Bax, N. A., Buckley, C. D., Weis, W. I. & Dunn, A. R. Vinculin forms a directionally asymmetric catch bond with F-actin. Science 357, 703–706 (2017). This study uses the state-of-the-art optical tweezer to show that vinculin exhibits catch bond behaviour towards F-actin.
Zhang, Y., Sivasankar, S., Nelson, W. J. & Chu, S. Resolving cadherin interactions and binding cooperativity at the single-molecule level. Proc. Natl Acad. Sci. USA 106, 109–114 (2009). This paper finds that the X-dimer conformation forms catch bonds in the presence of calcium, whereas the strand–swap dimer forms slip bonds, effectively allowing E-cadherin to increase affinity above the weak trans interaction observed in solution.
Manibog, K., Li, H., Rakshit, S. & Sivasankar, S. Resolving the molecular mechanism of cadherin catch bond formation. Nat. Commun. 5, 1–11 (2014).
Rakshit, S., Zhang, Y., Manibog, K., Shafraz, O. & Sivasankar, S. Ideal, catch, and slip bonds in cadherin adhesion. Proc. Natl Acad. Sci. USA 109, 18815–18820 (2012).
Manibog, K. et al. Molecular determinants of cadherin ideal bond formation: conformation-dependent unbinding on a multidimensional landscape. Proc. Natl Acad. Sci. USA 113, E5711–E5720 (2016).
Chugh, P. et al. Actin cortex architecture regulates cell surface tension. Nat. Cell Biol. 19, 689–697 (2017).
Manning, M. L., Foty, R. A., Steinberg, M. S. & Schoetz, E.-M. Coaction of intercellular adhesion and cortical tension specifies tissue surface tension. Proc. Natl Acad. Sci. USA 107, 12517–12522 (2010).
Toret, C. P., Collins, C. & Nelson, W. J. An Elmo–Dock complex locally controls Rho GTPases and actin remodeling during cadherin-mediated adhesion. J. Cell Biol. 207, 577–587 (2014). This study uses genome-wide RNAi screening to find that the ELMO–DOCK complex promotes formation of cadherin-based cell–cell contacts by locally regulating actin assembly.
Shilagardi, K. et al. Actin-propelled invasive membrane protrusions promote fusogenic protein engagement during cell–cell fusion. Science 340, 359–363 (2013). This work demonstrates that the fusogenic interface consists of two main adhesion proteins, Sns and Duf, that organize actin into podosome-like protrusive structures and thin sheaths of actin beneath the membrane, respectively.
Kim, J. H. et al. Mechanical tension drives cell membrane fusion. Dev. Cell 32, 561–573 (2015).
Kumari, S. et al. Actin foci facilitate activation of the phospholipase C-γ in primary T lymphocytes via the WASP pathway. eLife 4, e04953 (2015).
Gomez, T. S. et al. Formins regulate the actin-related protein 2/3 complex-independent polarization of the centrosome to the immunological synapse. Immunity 26, 177–190 (2007).
Belardi, B., Hamkins-Indik, T., Harris, A. R., Kim, J., Xu, K., Fletcher, D. A. A weak link with actin organizes tight junctions to control epithelial permeability. Dev. Cell 54, 792-804 (2020).
Indra, I., Troyanovsky, R. B., Shapiro, L., Honig, B. & Troyanovsky, S. M. Sensing actin dynamics through adherens junctions. Cell Rep. 30, 2820–2833.e3 (2020).
Yi, J., Balagopalan, L., Nguyen, T., McIntire, K. M. & Samelson, L. E. TCR microclusters form spatially segregated domains and sequentially assemble in calcium-dependent kinetic steps. Nat. Commun. 10, 1–13 (2019).
Yokosuka, T. & Saito, T. Dynamic regulation of T-cell costimulation through TCR–CD28 microclusters. Immunol. Rev. 229, 27–40 (2009).
Davis, S. J., Ikemizu, S., Wild, M. K. & van der Merwe, P. A. CD2 and the nature of protein interactions mediating cell–cell recognition. Immunol. Rev. 163, 217–236 (1998).
Demetriou, P et al. A dynamic CD2-rich compartment at the outer edge of the immunological synapse boosts and integrates signals. Nat. Immunol. 21, 1232–1243 (2020).
Weledji, E. P. & Assob, J. C. The ubiquitous neural cell adhesion molecule (N-CAM). Ann. Med. Surg. 3, 77–81 (2014).
Kiselyov, V. V., Soroka, V., Berezin, V. & Bock, E. Structural biology of NCAM homophilic binding and activation of FGFR. J. Neurochem. 94, 1169–1179 (2005).
Doherty, P., Fazeli, M. S. & Walsh, F. S. The neural cell adhesion molecule and synaptic plasticity. J. Neurobiol. 26, 437–446 (1995).
Rønn, L. C., Berezin, V. & Bock, E. The neural cell adhesion molecule in synaptic plasticity and ageing. Int. J. Dev. Neurosci. 18, 193–199 (2000).
Mace, E. M., Gunesch, J. T., Dixon, A. & Orange, J. S. Human NK cell development requires CD56-mediated motility and formation of the developmental synapse. Nat. Commun. 7, 12171 (2016).
Taouk, G. et al. CD56 expression in breast cancer induces sensitivity to natural killer-mediated cytotoxicity by enhancing the formation of cytotoxic immunological synapse. Sci. Rep. 9, 8756 (2019).
Ditlevsen, D. K., Povlsen, G. K., Berezin, V. & Bock, E. NCAM-induced intracellular signaling revisited. J. Neurosci. Res. 86, 727–743 (2008).
Case, L. B., Ditlev, J. A. & Rosen, M. K. Regulation of transmembrane signaling by phase separation. Annu. Rev. Biophys. 48, 465–494 (2019).
Banjade, S. & Rosen, M. K. Phase transitions of multivalent proteins can promote clustering of membrane receptors. eLife 3, e04123 (2014).
Su, X. et al. Phase separation of signaling molecules promotes T cell receptor signal transduction. Science 352, 595–599 (2016). This paper shows that purified, phosphorylated LAT of T cells can form a phase-separated cluster in the presence of GRB2 and SOS1, which promotes the MAPK signalling cascade in cells.
Case, L. B., Zhang, X., Ditlev, J. A. & Rosen, M. K. Stoichiometry controls activity of phase-separated clusters of actin signaling proteins. Science 363, 1093–1097 (2019).
Ditlev, J. A. et al. A composition-dependent molecular clutch between T cell signaling condensates and actin. eLife 8, e42695 (2019).
Huang, W. Y. C. et al. A molecular assembly phase transition and kinetic proofreading modulate Ras activation by SOS. Science 363, 1098–1103 (2019). This paper shows that the GEF domain of SOS requires long activation times (~50 s), and a biomolecular condensate, such as that formed between LAT–GRB2–SOS in 2D, extends the dwell times of SOS to release its autoinhibition and allow the protein to achieve full activation of its GEF activity for RAS — a necessary step in T cell activation.
Missler, M., Südhof, T. C. & Biederer, T. Synaptic cell adhesion. Cold Spring Harb. Perspect. Biol. 4, a005694 (2012).
Frank, R. A. & Grant, S. G. Supramolecular organization of NMDA receptors and the postsynaptic density. Curr. Opin. Neurobiol. 45, 139–147 (2017).
Broadhead, M. J. et al. PSD95 nanoclusters are postsynaptic building blocks in hippocampus circuits. Sci. Rep. 6, 1–14 (2016).
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).
Zeng, M. et al. Phase transition in postsynaptic densities underlies formation of synaptic complexes and synaptic plasticity. Cell 166, 1163–1175.e12 (2016).
Beutel, O., Maraspini, R., Pombo-García, K., Martin-Lemaitre, C. & Honigmann, A. Phase separation of zonula occludens proteins drives formation of tight junctions. Cell 179, 923–936.e11 (2019).
Schwayer, C. et al. Mechanosensation of tight junctions depends on ZO-1 phase separation and flow. Cell 179, 937–952.e18 (2019).
Kourtidis, A. et al. Cadherin complexes recruit mRNAs and RISC to regulate epithelial cell signaling. J. Cell Biol. 216, 3073–3085 (2017).
Benham-Pyle, B. W., Pruitt, B. L. & Nelson, W. J. Cell adhesion. Mechanical strain induces E-cadherin-dependent Yap1 and β-catenin activation to drive cell cycle entry. Science 348, 1024–1027 (2015).
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).
Yap. A. S. (Ed.), Cellular Adhesion in Development and Disease 1st edn Vol. 112 (Elsevier, 2015).
Alberts, B. et al. in Molecular Biology of the Cell 4th edn 567 (Garland Science, 2002).
Kim, J. H. & Chen, E. H. The fusogenic synapse at a glance. J. Cell Sci. 132, jcs213124 (2019).
Bertocchi, C. et al. Nanoscale architecture of cadherin-based cell adhesions. Nat. Cell Biol. 19, 28–37 (2017).
Mangeol, P., Massey-Harroche, D., Bivic, A. L. & Lenne, P.-F. Nectins rather than E-cadherin anchor the actin belts at cell–cell junctions of epithelia. Preprint at bioRxiv https://doi.org/10.1101/809343 (2019).
Parthasarathy, R. & Groves, J. T. Optical techniques for imaging membrane topography. Cell Biochem. Biophys. 41, 391–414 (2004).
Paszek, M. J. et al. Scanning angle interference microscopy reveals cell dynamics at the nanoscale. Nat. Methods 9, 825–827 (2012).
Son, S. et al. Molecular height measurement by cell surface optical profilometry (CSOP). Proc. Natl Acad. Sci. USA 117, 14209–14219 (2020).
Buckley, C. D. et al. Cell adhesion. The minimal cadherin–catenin complex binds to actin filaments under force. Science 346, 1254211 (2014).
Priest, A. V., Shafraz, O. & Sivasankar, S. Biophysical basis of cadherin mediated cell–cell adhesion. Exp. Cell Res. 358, 10–13 (2017).
Sivasankar, S., Brieher, W., Lavrik, N., Gumbiner, B. & Leckband, D. Direct molecular force measurements of multiple adhesive interactions between cadherin ectodomains. Proc. Natl Acad. Sci. USA 96, 11820–11824 (1999).
Freikamp, A., Cost, A.-L. & Grashoff, C. The picoNewton force awakens: quantifying mechanics in cells. Trends Cell Biol. 26, 838–847 (2016).
Ma, V. P.-Y. & Salaita, K. DNA nanotechnology as an emerging tool to study mechanotransduction in living systems. Small 15, 1900961 (2019).
Schmid, E. M., Richmond, D. L. & Fletcher, D. A. Reconstitution of proteins on electroformed giant unilamellar vesicles. Methods Cell Biol. 128, 319–338 (2015).
Biswas, K. H. & Groves, J. T. Hybrid live cell-supported membrane interfaces for signaling studies. Annu. Rev. Biophys. 48, 537–562 (2019).
Stachowiak, J. C. et al. Unilamellar vesicle formation and encapsulation by microfluidic jetting. Proc. Natl Acad. Sci. USA 105, 4697–4702 (2008).
Dürre, K. et al. Capping protein-controlled actin polymerization shapes lipid membranes. Nat. Commun. 9, 1–11 (2018).
Lemière, J., Carvalho, K. & Sykes, C. Cell-sized liposomes that mimic cell motility and the cell cortex. Methods Cell Biol. 128, 271–285 (2015).
Cohen, D. J., Gloerich, M. & Nelson, W. J. Epithelial self-healing is recapitulated by a 3D biomimetic E-cadherin junction. Proc. Natl Acad. Sci. USA 113, 14698–14703 (2016).
Smirnova, Y. G., Risselada, H. J. & Müller, M. Thermodynamically reversible paths of the first fusion intermediate reveal an important role for membrane anchors of fusion proteins. Proc. Natl Acad. Sci. USA 116, 2571–2576 (2019).
Ma, Z., Janmey, P. A. & Finkel, T. H. The receptor deformation model of TCR triggering. FASEB J. 22, 1002–1008 (2008).
Kaizuka, Y., Douglass, A. D., Vardhana, S., Dustin, M. L. & Vale, R. D. The coreceptor CD2 uses plasma membrane microdomains to transduce signals in T cells. J. Cell Biol. 185, 521–534 (2009).
Morsut, L. et al. Engineering customized cell sensing and response behaviors using synthetic notch receptors. Cell 164, 780–791 (2016).
Kang, T. H. & Jung, S. T. Boosting therapeutic potency of antibodies by taming Fc domain functions. Exp. Mol. Med. 51, 138 (2019).
Swaminathan, V. et al. Mechanical stiffness grades metastatic potential in patient tumor cells and in cancer cell lines. Cancer Res. 71, 5075–5080 (2011).
Al Absi, A. et al. Actin cytoskeleton remodeling drives breast cancer cell escape from natural killer-mediated cytotoxicity. Cancer Res. 78, 5631–5643 (2018).
Ito, S. et al. Induced cortical tension restores functional junctions in adhesion-defective carcinoma cells. Nat. Commun. 8, 1–16 (2017).
Suter, E. C., Schmid, E. M., Voets, E., Francica, B. & Fletcher, D. A. Antibody:CD47 ratio regulates macrophage phagocytosis through competitive receptor phosphorylation. Preprint at bioRxiv https://doi.org/10.1101/2020.07.31.231779 (2020).
Xiao, H., Woods, E. C., Vukojicic, P. & Bertozzi, C. R. Precision glycocalyx editing as a strategy for cancer immunotherapy. Proc. Natl Acad. Sci. USA 113, 10304–10309 (2016).
Acknowledgements
The authors thank E. Schmid for her research contributions and guidance on this Review, and the Fletcher and Dustin laboratories for helpful discussions. B.B. was supported by the National Institutes of Health (NIH) Ruth L. Kirschstein NRSA fellowship from the NIH (1F32GM115091). S.S. was supported by the Life Science Research Foundation. J.H.F. and M.L.D. were supported by the Wellcome Trust. This work was supported in part by the NIH R01 GM114671 (D.A.F.), the Immunotherapeutics and Vaccine Research Initiative at UC Berkeley (D.A.F.), the Miller Institute for Basic Research (D.A.F.), the NSF Center for Cellular Construction (DBI-1548297) and the Chan Zuckerberg Biohub (D.A.F.).
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Glossary
- Myoblast
-
A mononucleated, undifferentiated muscle cell precursor.
- Tight junctions
-
Cell–cell junctions that seal adjacent epithelial cells together, preventing the passage of most dissolved molecules from one side of the epithelial sheet to the other.
- Physical boundary conditions
-
A set of constraints that define a closed physical system.
- Desmosomes
-
Anchoring cell–cell junctions, usually formed between two epithelial cells, characterized by dense plaques of protein into which intermediate filaments in the two adjoining cells insert.
- Integrins
-
A large family of transmembrane proteins, consisting of α and β-subunits, involved in the adhesion of cells to the extracellular matrix and to each other.
- Glycocalyx
-
A carbohydrate-rich layer that forms the outer coat of a eukaryotic cell, composed of the oligosaccharides linked to plasma membrane glycoproteins and glycolipids, as well as glycoproteins and proteoglycans that have been secreted and reabsorbed onto the cell surface.
- Natural killer cells
-
Cytotoxic cells of the innate immune system that can kill virus-infected and cancer cells.
- Antigen-presenting cells
-
Highly specialized cells that can process antigens and display their peptide fragments on the cell surface together with other, co-stimulatory, proteins required for activating naive T cells.
- Adherens junction
-
A cell–cell interface that holds neighbouring cells together through transmembrane cadherin family proteins. The cytoplasmic face of the junction is attached to actin filaments. Examples include the adhesion belts linking adjacent epithelial cells.
- Ideal bond
-
A bond whose lifetime is not influenced by force.
- Slip bond
-
A bond whose lifetime is shortened under force.
- Catch bond
-
A bond that increases its lifetime under force.
- Kinetic trap
-
A system trapped in a local energy minimum of an energetic landscape due to high-energy transition barriers.
- Enthalpy
-
A thermodynamic quantity equivalent to the sum of the system’s internal energy and the product of its pressure and volume.
- Entropy
-
A thermodynamic quantity that measures the degree of disorder in a system.
- Cell cortex
-
A thin network of actin filaments and actin-binding proteins that underlies the plasma membrane in most eukaryotic cells.
- α-Catenin
-
An adaptor protein of the adherens junction that is part of the E-cadherin–catenin complex and can bind to the actin cytoskeleton.
- Vinculin
-
An adaptor protein of both the adherens junction and focal adhesions that reinforces connections to the actin cytoskeleton.
- Podosome
-
A dynamic, actin-rich cellular protrusion that degrades the extracellular matrix and is involved in cell invasion.
- Wiscott–Aldrich syndrome protein
-
(WASP). A nucleation-promoting factor of the actin cytoskeleton that acts on the Arp2/3 complex.
- Arp2/3 complex
-
A protein complex that nucleates the assembly of branched actin filament networks.
- Biomolecular condensates
-
Membraneless compartments in cells that concentrate specific collections of proteins and nucleic acids into a dynamic assembly.
- Liquid–liquid phase separation
-
The demixing of a fluid mixture into two distinct liquid phases.
- Kinetic proofreading
-
A specificity mechanism in biochemical reactions that achieves high fidelity beyond what is possible by free-energy differences. In the immunological synapse, kinetic proofreading allows small differences in ligand binding half-life to be amplified into larger differences in signalling through intermediate steps.
- Postsynaptic density
-
A protein-dense specialization in neurons attached to postsynaptic membranes.
- Antibody-dependent cellular cytotoxicity
-
The killing of antibody-coated target cells by cells with Fc receptors that recognize the constant region of the bound antibody. Most antibody-dependent cellular cytotoxicity is mediated by natural killer cells that have the Fc receptor FcγRIII on their surface.
- Chimeric antigen receptor
-
(CAR). Engineered fusion proteins composed of extracellular antigen-specific receptors (for example, single-chain antibody) and intracellular signalling domains that activate and co-stimulate, expressed in T cells for use in cancer immunotherapy.
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Belardi, B., Son, S., Felce, J.H. et al. Cell–cell interfaces as specialized compartments directing cell function. Nat Rev Mol Cell Biol 21, 750–764 (2020). https://doi.org/10.1038/s41580-020-00298-7
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DOI: https://doi.org/10.1038/s41580-020-00298-7
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