Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

Mechanical forces in the immune system

Key Points

  • Immune cells lead physically active 'lifestyles' that are characterized by shape change, migration and the formation of dynamic cell–cell interactions.

  • Leukocytes use mechanical cues to adjust their mode of migration to the prevailing environmental conditions.

  • Mechanically induced catch bonds have a crucial role in immune cell trafficking, lymphocyte activation and immunological synapse formation.

  • Lymphocytes use mechanical force to discriminate between high-affinity and low-affinity antigens.

  • Force exertion enables immune cells to control the strength and the specificity of their effector responses.

Abstract

Leukocytes can completely reorganize their cytoskeletal architecture within minutes. This structural plasticity, which facilitates their migration and communicative function, also enables them to exert a substantial amount of mechanical force against the extracellular matrix and the surfaces of interacting cells. In recent years, it has become increasingly clear that these forces have crucial roles in immune cell activation and subsequent effector responses. Here, I review our current understanding of how mechanical force regulates cell-surface receptor activation, cell migration, intracellular signalling and intercellular communication, highlighting the biological ramifications of these effects in various immune cell types.

This is a preview of subscription content, access via your institution

Access options

Buy this article

Prices may be subject to local taxes which are calculated during checkout

Figure 1: Force exertion and mechanotransduction.
Figure 2: Mechanical forces in cell migration.
Figure 3: Mechanical forces in immune cell–cell interactions.

Similar content being viewed by others

References

  1. Iskratsch, T., Wolfenson, H. & Sheetz, M. P. Appreciating force and shape — the rise of mechanotransduction in cell biology. Nat. Rev. Mol. Cell Biol. 15, 825–833 (2014).

    CAS  PubMed  Google Scholar 

  2. Vicente-Manzanares, M., Ma, X., Adelstein, R. S. & Horwitz, A. R. Non-muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol. Cell Biol. 10, 778–790 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  3. Smith, A. et al. The role of the integrin LFA-1 in T-lymphocyte migration. Immunol. Rev. 218, 135–146 (2007).

    CAS  PubMed  Google Scholar 

  4. Matsumura, F. Regulation of myosin II during cytokinesis in higher eukaryotes. Trends Cell Biol. 15, 371–377 (2005).

    CAS  PubMed  Google Scholar 

  5. Hui, K. L., Balagopalan, L., Samelson, L. E. & Upadhyaya, A. Cytoskeletal forces during signaling activation in Jurkat T-cells. Mol. Biol. Cell 26, 685–695 (2015).

    PubMed  PubMed Central  Google Scholar 

  6. Basu, R. et al. Cytotoxic T cells use mechanical force to potentiate target cell killing. Cell 165, 100–110 (2016). This study reveals that CTLs use mechanical force to increase the activity of secreted perforin, thus demonstrating that immune cells use a physical output to potentiate chemical responses.

    CAS  PubMed  PubMed Central  Google Scholar 

  7. Natkanski, E. et al. B cells use mechanical energy to discriminate antigen affinities. Science 340, 1587–1590 (2013). This study shows that B cells apply myosin II-dependent forces during antigen internalization that allow them to discriminate between high-affinity and low-affinity antigens.

    CAS  PubMed  PubMed Central  Google Scholar 

  8. Lammermann, T. et al. Rapid leukocyte migration by integrin-independent flowing and squeezing. Nature 453, 51–55 (2008).

    PubMed  Google Scholar 

  9. Jannat, R. A., Dembo, M. & Hammer, D. A. Traction forces of neutrophils migrating on compliant substrates. Biophys. J. 101, 575–584 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  10. Toyjanova, J., Flores-Cortez, E., Reichner, J. S. & Franck, C. Matrix confinement plays a pivotal role in regulating neutrophil-generated tractions, speed, and integrin utilization. J. Biol. Chem. 290, 3752–3763 (2015).

    CAS  PubMed  Google Scholar 

  11. Chesarone, M. A., DuPage, A. G. & Goode, B. L. Unleashing formins to remodel the actin and microtubule cytoskeletons. Nat. Rev. Mol. Cell Biol. 11, 62–74 (2010).

    CAS  PubMed  Google Scholar 

  12. Goley, E. D. & Welch, M. D. The ARP2/3 complex: an actin nucleator comes of age. Nat. Rev. Mol. Cell Biol. 7, 713–726 (2006).

    CAS  PubMed  Google Scholar 

  13. Yu, C. H., Law, J. B., Suryana, M., Low, H. Y. & Sheetz, M. P. Early integrin binding to Arg-Gly-Asp peptide activates actin polymerization and contractile movement that stimulates outward translocation. Proc. Natl Acad. Sci. USA 108, 20585–20590 (2011).

    CAS  PubMed  Google Scholar 

  14. Carman, C. V. et al. Transcellular diapedesis is initiated by invasive podosomes. Immunity 26, 784–797 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  15. Buccione, R., Orth, J. D. & McNiven, M. A. Foot and mouth: podosomes, invadopodia and circular dorsal ruffles. Nat. Rev. Mol. Cell Biol. 5, 647–657 (2004).

    CAS  PubMed  Google Scholar 

  16. Geiger, B., Spatz, J. P. & Bershadsky, A. D. Environmental sensing through focal adhesions. Nat. Rev. Mol. Cell Biol. 10, 21–33 (2009).

    CAS  PubMed  Google Scholar 

  17. Gardel, M. L., Schneider, I. C., Aratyn-Schaus, Y. & Waterman, C. M. Mechanical integration of actin and adhesion dynamics in cell migration. Annu. Rev. Cell Dev. Biol. 26, 315–333 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  18. Kanchanawong, P. et al. Nanoscale architecture of integrin-based cell adhesions. Nature 468, 580–584 (2010). The study applies super-resolution imaging to define the nanoarchitecture of focal adhesions.

    CAS  PubMed  PubMed Central  Google Scholar 

  19. Prager-Khoutorsky, M. et al. Fibroblast polarization is a matrix-rigidity-dependent process controlled by focal adhesion mechanosensing. Nat. Cell Biol. 13, 1457–1465 (2011).

    CAS  PubMed  Google Scholar 

  20. Goffin, J. M. et al. Focal adhesion size controls tension-dependent recruitment of α-smooth muscle actin to stress fibers. J. Cell Biol. 172, 259–268 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  21. Galbraith, C. G., Yamada, K. M. & Sheetz, M. P. The relationship between force and focal complex development. J. Cell Biol. 159, 695–705 (2002).

    CAS  PubMed  PubMed Central  Google Scholar 

  22. Riveline, D. et al. Focal contacts as mechanosensors: externally applied local mechanical force induces growth of focal contacts by an mDia1-dependent and ROCK-independent mechanism. J. Cell Biol. 153, 1175–1186 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  23. Balaban, N. Q. et al. Force and focal adhesion assembly: a close relationship studied using elastic micropatterned substrates. Nat. Cell Biol. 3, 466–472 (2001).

    CAS  PubMed  Google Scholar 

  24. Wolfenson, H., Lavelin, I. & Geiger, B. Dynamic regulation of the structure and functions of integrin adhesions. Dev. Cell 24, 447–458 (2013).

    CAS  PubMed  Google Scholar 

  25. Stanley, P. et al. Intermediate-affinity LFA-1 binds α-actinin-1 to control migration at the leading edge of the T cell. EMBO J. 27, 62–75 (2008).

    CAS  PubMed  Google Scholar 

  26. Monks, C. R., Freiberg, B. A., Kupfer, H., Sciaky, N. & Kupfer, A. Three-dimensional segregation of supramolecular activation clusters in T cells. Nature 395, 82–86 (1998).

    CAS  PubMed  Google Scholar 

  27. Santos, L. C. et al. Actin polymerization-dependent activation of Cas-L promotes immunological synapse stability. Immunol. Cell Biol. 94, 981–993 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  28. Ingber, D. E. Cellular mechanotransduction: putting all the pieces together again. FASEB J. 20, 811–827 (2006).

    CAS  PubMed  Google Scholar 

  29. Orr, A. W., Helmke, B. P., Blackman, B. R. & Schwartz, M. A. Mechanisms of mechanotransduction. Dev. Cell 10, 11–20 (2006).

    CAS  PubMed  Google Scholar 

  30. Pathak, M. M. et al. Stretch-activated ion channel Piezo1 directs lineage choice in human neural stem cells. Proc. Natl Acad. Sci. USA 111, 16148–16153 (2014).

    CAS  PubMed  Google Scholar 

  31. Lee, W. et al. Synergy between Piezo1 and Piezo2 channels confers high-strain mechanosensitivity to articular cartilage. Proc. Natl Acad. Sci. USA 111, E5114–E5122 (2014).

    CAS  PubMed  Google Scholar 

  32. Ranade, S. S. et al. Piezo1, a mechanically activated ion channel, is required for vascular development in mice. Proc. Natl Acad. Sci. USA 111, 10347–10352 (2014).

    CAS  PubMed  Google Scholar 

  33. del Rio, A. et al. Stretching single talin rod molecules activates vinculin binding. Science 323, 638–641 (2009). This study shows that the physical stretching of talin generates binding sites for vinculin, thus providing a mechanism for mechanotransduction in focal contacts.

    CAS  PubMed  Google Scholar 

  34. Elosegui-Artola, A. et al. Mechanical regulation of a molecular clutch defines force transmission and transduction in response to matrix rigidity. Nat. Cell Biol. 18, 540–548 (2016).

    CAS  PubMed  Google Scholar 

  35. Dembo, M., Torney, D. C., Saxman, K. & Hammer, D. The reaction-limited kinetics of membrane-to-surface adhesion and detachment. Proc. R. Soc. Lond. B Biol. Sci. 234, 55–83 (1988). This theoretical paper provides the conceptual basis for the existence of catch bonds.

    CAS  PubMed  Google Scholar 

  36. Thomas, W. E., Vogel, V. & Sokurenko, E. Biophysics of catch bonds. Annu. Rev. Biophys. 37, 399–416 (2008).

    CAS  PubMed  Google Scholar 

  37. Engler, A. J., Sen, S., Sweeney, H. L. & Discher, D. E. Matrix elasticity directs stem cell lineage specification. Cell 126, 677–689 (2006).

    CAS  PubMed  Google Scholar 

  38. Assoian, R. K. & Klein, E. A. Growth control by intracellular tension and extracellular stiffness. Trends Cell Biol. 18, 347–352 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  39. Trichet, L. et al. Evidence of a large-scale mechanosensing mechanism for cellular adaptation to substrate stiffness. Proc. Natl Acad. Sci. USA 109, 6933–6938 (2012).

    CAS  PubMed  Google Scholar 

  40. Judokusumo, E., Tabdanov, E., Kumari, S., Dustin, M. L. & Kam, L. C. Mechanosensing in T lymphocyte activation. Biophys. J. 102, L5–L7 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  41. Saez, A., Buguin, A., Silberzan, P. & Ladoux, B. Is the mechanical activity of epithelial cells controlled by deformations or forces? Biophys. J. 89, L52–L54 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  42. Califano, J. P. & Reinhart-King, C. A. Substrate stiffness and cell area predict cellular traction stresses in single cells and cells in contact. Cell. Mol. Bioeng. 3, 68–75 (2010).

    PubMed  PubMed Central  Google Scholar 

  43. Ghibaudo, M. et al. Traction forces and rigidity sensing regulate cell functions. Soft Matter 4, 1836–1843 (2008).

    CAS  Google Scholar 

  44. Renkawitz, J. & Sixt, M. Mechanisms of force generation and force transmission during interstitial leukocyte migration. EMBO Rep. 11, 744–750 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  45. Yakubenko, V. P., Lishko, V. K., Lam, S. C. & Ugarova, T. P. A molecular basis for integrin αMβ2 ligand binding promiscuity. J. Biol. Chem. 277, 48635–48642 (2002).

    CAS  PubMed  Google Scholar 

  46. Vorup-Jensen, T. et al. Exposure of acidic residues as a danger signal for recognition of fibrinogen and other macromolecules by integrin αXβ2. Proc. Natl Acad. Sci. USA 102, 1614–1619 (2005).

    CAS  PubMed  Google Scholar 

  47. Renkawitz, J. et al. Adaptive force transmission in amoeboid cell migration. Nat. Cell Biol. 11, 1438–1443 (2009).

    Article  CAS  PubMed  Google Scholar 

  48. Vicente-Manzanares, M., Choi, C. K. & Horwitz, A. R. Integrins in cell migration — the actin connection. J. Cell Sci. 122, 199–206 (2009). This study demonstrates that the engagement of the F-actin–talin–integrin clutch regulates the speed of retrograde F-actin flow, enabling cells to adjust their migratory strategy in response to substrate adhesiveness.

    CAS  PubMed  Google Scholar 

  49. Lin, C. H. & Forscher, P. Growth cone advance is inversely proportional to retrograde F-actin flow. Neuron 14, 763–771 (1995).

    CAS  PubMed  Google Scholar 

  50. Pollard, T. D. & Borisy, G. G. Cellular motility driven by assembly and disassembly of actin filaments. Cell 112, 453–465 (2003).

    CAS  PubMed  Google Scholar 

  51. Ponti, A., Machacek, M., Gupton, S. L., Waterman-Storer, C. M. & Danuser, G. Two distinct actin networks drive the protrusion of migrating cells. Science 305, 1782–1786 (2004).

    CAS  PubMed  Google Scholar 

  52. Medeiros, N. A., Burnette, D. T. & Forscher, P. Myosin II functions in actin-bundle turnover in neuronal growth cones. Nat. Cell Biol. 8, 215–226 (2006).

    CAS  PubMed  Google Scholar 

  53. Wilson, C. A. et al. Myosin II contributes to cell-scale actin network treadmilling through network disassembly. Nature 465, 373–377 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  54. Sanchez-Madrid, F. & Serrador, J. M. Bringing up the rear: defining the roles of the uropod. Nat. Rev. Mol. Cell Biol. 10, 353–359 (2009).

    CAS  PubMed  Google Scholar 

  55. Eddy, R. J., Pierini, L. M., Matsumura, F. & Maxfield, F. R. Ca2+-dependent myosin II activation is required for uropod retraction during neutrophil migration. J. Cell Sci. 113, 1287–1298 (2000).

    CAS  PubMed  Google Scholar 

  56. Luo, B. H., Carman, C. V. & Springer, T. A. Structural basis of integrin regulation and signaling. Annu. Rev. Immunol. 25, 619–647 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  57. Kim, C., Ye, F. & Ginsberg, M. H. Regulation of integrin activation. Annu. Rev. Cell Dev. Biol. 27, 321–345 (2011).

    CAS  PubMed  Google Scholar 

  58. Astrof, N. S., Salas, A., Shimaoka, M., Chen, J. & Springer, T. A. Importance of force linkage in mechanochemistry of adhesion receptors. Biochemistry 45, 15020–15028 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  59. Friedland, J. C., Lee, M. H. & Boettiger, D. Mechanically activated integrin switch controls α5β1 function. Science 323, 642–644 (2009).

    CAS  PubMed  Google Scholar 

  60. Woolf, E. et al. Lymph node chemokines promote sustained T lymphocyte motility without triggering stable integrin adhesiveness in the absence of shear forces. Nat. Immunol. 8, 1076–1085 (2007).

    CAS  PubMed  Google Scholar 

  61. Kong, F., Garcia, A. J., Mould, A. P., Humphries, M. J. & Zhu, C. Demonstration of catch bonds between an integrin and its ligand. J. Cell Biol. 185, 1275–1284 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  62. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  63. Rosetti, F. et al. A lupus-associated Mac-1 variant has defects in integrin allostery and interaction with ligands under force. Cell Rep. 10, 1655–1664 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  64. Case, L. B. & Waterman, C. M. Integration of actin dynamics and cell adhesion by a three-dimensional, mechanosensitive molecular clutch. Nat. Cell Biol. 17, 955–963 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  65. Chan, C. E. & Odde, D. J. Traction dynamics of filopodia on compliant substrates. Science 322, 1687–1691 (2008). This study combines computational modelling with biophysical measurements to investigate the applicability of the molecular clutch hypothesis to cell shape changes and migration.

    CAS  PubMed  Google Scholar 

  66. Smith, L. A., Aranda-Espinoza, H., Haun, J. B., Dembo, M. & Hammer, D. A. Neutrophil traction stresses are concentrated in the uropod during migration. Biophys. J. 92, L58–L60 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  67. Dixit, N., Yamayoshi, I., Nazarian, A. & Simon, S. I. Migrational guidance of neutrophils is mechanotransduced via high-affinity LFA-1 and calcium flux. J. Immunol. 187, 472–481 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  68. Green, C. E. et al. Dynamic shifts in LFA-1 affinity regulate neutrophil rolling, arrest, and transmigration on inflamed endothelium. Blood 107, 2101–2111 (2006).

    CAS  PubMed  PubMed Central  Google Scholar 

  69. Ricart, B. G., Yang, M. T., Hunter, C. A., Chen, C. S. & Hammer, D. A. Measuring traction forces of motile dendritic cells on micropost arrays. Biophys. J. 101, 2620–2628 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  70. Hind, L. E., Dembo, M. & Hammer, D. A. Macrophage motility is driven by frontal-towing with a force magnitude dependent on substrate stiffness. Integr. Biol. (Camb.) 7, 447–453 (2015).

    CAS  Google Scholar 

  71. Kozlov, M. M. & Mogilner, A. Model of polarization and bistability of cell fragments. Biophys. J. 93, 3811–3819 (2007).

    CAS  PubMed  PubMed Central  Google Scholar 

  72. Houk, A. R. et al. Membrane tension maintains cell polarity by confining signals to the leading edge during neutrophil migration. Cell 148, 175–188 (2012). This study reveals that leukocytes use membrane tension to maintain cell polarity during migration.

    CAS  PubMed  PubMed Central  Google Scholar 

  73. Keren, K. et al. Mechanism of shape determination in motile cells. Nature 453, 475–480 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  74. Diz-Munoz, A. et al. Membrane tension acts through PLD2 and mTORC2 to limit actin network assembly during neutrophil migration. PLoS Biol. 14, e1002474 (2016).

    PubMed  PubMed Central  Google Scholar 

  75. Ley, K. The role of selectins in inflammation and disease. Trends Mol. Med. 9, 263–268 (2003).

    CAS  PubMed  Google Scholar 

  76. Finger, E. B. et al. Adhesion through L-selectin requires a threshold hydrodynamic shear. Nature 379, 266–269 (1996).

    CAS  PubMed  Google Scholar 

  77. Lawrence, M. B., Kansas, G. S., Kunkel, E. J. & Ley, K. Threshold levels of fluid shear promote leukocyte adhesion through selectins (CD62L,P,E). J. Cell Biol. 136, 717–727 (1997).

    CAS  PubMed  PubMed Central  Google Scholar 

  78. Marshall, B. T. et al. Direct observation of catch bonds involving cell-adhesion molecules. Nature 423, 190–193 (2003). This study is the first to document the existence of catch bonds in biological systems.

    CAS  PubMed  Google Scholar 

  79. Yago, T. et al. Catch bonds govern adhesion through L-selectin at threshold shear. J. Cell Biol. 166, 913–923 (2004).

    CAS  PubMed  PubMed Central  Google Scholar 

  80. Thelen, M. & Stein, J. V. How chemokines invite leukocytes to dance. Nat. Immunol. 9, 953–959 (2008).

    CAS  PubMed  Google Scholar 

  81. Shamri, R. et al. Lymphocyte arrest requires instantaneous induction of an extended LFA-1 conformation mediated by endothelium-bound chemokines. Nat. Immunol. 6, 497–506 (2005).

    CAS  PubMed  Google Scholar 

  82. Alon, R. & Ley, K. Cells on the run: shear-regulated integrin activation in leukocyte rolling and arrest on endothelial cells. Curr. Opin. Cell Biol. 20, 525–532 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  83. Zhu, J. et al. Structure of a complete integrin ectodomain in a physiologic resting state and activation and deactivation by applied forces. Mol. Cell 32, 849–861 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  84. Nordenfelt, P., Elliott, H. L. & Springer, T. A. Coordinated integrin activation by actin-dependent force during T-cell migration. Nat. Commun. 7, 13119 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  85. Sarangapani, K. K. et al. Regulation of catch bonds by rate of force application. J. Biol. Chem. 286, 32749–32761 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  86. Klopocki, A. G. et al. Replacing a lectin domain residue in L-selectin enhances binding to P-selectin glycoprotein ligand-1 but not to 6-sulfo-sialyl Lewis x. J. Biol. Chem. 283, 11493–11500 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  87. Nourshargh, S. & Alon, R. Leukocyte migration into inflamed tissues. Immunity 41, 694–707 (2014).

    CAS  PubMed  Google Scholar 

  88. Sage, P. T. et al. Antigen recognition is facilitated by invadosome-like protrusions formed by memory/effector T cells. J. Immunol. 188, 3686–3699 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  89. Nourshargh, S., Hordijk, P. L. & Sixt, M. Breaching multiple barriers: leukocyte motility through venular walls and the interstitium. Nat. Rev. Mol. Cell Biol. 11, 366–378 (2010).

    CAS  PubMed  Google Scholar 

  90. Rabodzey, A., Alcaide, P., Luscinskas, F. W. & Ladoux, B. Mechanical forces induced by the transendothelial migration of human neutrophils. Biophys. J. 95, 1428–1438 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  91. Thiam, H. R. et al. Perinuclear Arp2/3-driven actin polymerization enables nuclear deformation to facilitate cell migration through complex environments. Nat. Commun. 7, 10997 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  92. Davidson, P. M., Denais, C., Bakshi, M. C. & Lammerding, J. Nuclear deformability constitutes a rate-limiting step during cell migration in 3D environments. Cell. Mol. Bioeng. 7, 293–306, (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  93. Raab, M. et al. ESCRT III repairs nuclear envelope ruptures during cell migration to limit DNA damage and cell death. Science 352, 359–362 (2016).

    CAS  PubMed  Google Scholar 

  94. Denais, C. M. et al. Nuclear envelope rupture and repair during cancer cell migration. Science 352, 353–358 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  95. Charras, G. & Paluch, E. Blebs lead the way: how to migrate without lamellipodia. Nat. Rev. Mol. Cell Biol. 9, 730–736 (2008).

    CAS  PubMed  Google Scholar 

  96. Chabaud, M. et al. Cell migration and antigen capture are antagonistic processes coupled by myosin II in dendritic cells. Nat. Commun. 6, 7526 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  97. Vargas, P. et al. Innate control of actin nucleation determines two distinct migration behaviours in dendritic cells. Nat. Cell Biol. 18, 43–53 (2016).

    CAS  PubMed  Google Scholar 

  98. Bergert, M. et al. Force transmission during adhesion-independent migration. Nat. Cell Biol. 17, 524–529 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  99. Jacobelli, J., Bennett, F. C., Pandurangi, P., Tooley, A. J. & Krummel, M. F. Myosin-IIA and ICAM-1 regulate the interchange between two distinct modes of T cell migration. J. Immunol. 182, 2041–2050 (2009).

    CAS  PubMed  Google Scholar 

  100. Franck, C., Maskarinec, S. A., Tirrell, D. A. & Ravichandran, G. Three-dimensional traction force microscopy: a new tool for quantifying cell–matrix interactions. PLoS ONE 6, e17833 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  101. Stout, D. A. et al. Mean deformation metrics for quantifying 3D cell–matrix interactions without requiring information about matrix material properties. Proc. Natl Acad. Sci. USA 113, 2898–2903 (2016).

    CAS  PubMed  Google Scholar 

  102. Dustin, M. L., Chakraborty, A. K. & Shaw, A. S. Understanding the structure and function of the immunological synapse. Cold Spring Harb. Perspect. Biol. 2, a002311 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  103. Harwood, N. E. & Batista, F. D. Early events in B cell activation. Annu. Rev. Immunol. 28, 185–210 (2010).

    CAS  PubMed  Google Scholar 

  104. O'Connor, R. S. et al. Substrate rigidity regulates human T cell activation and proliferation. J. Immunol. 189, 1330–1339 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  105. Wan, Z. et al. B cell activation is regulated by the stiffness properties of the substrate presenting the antigens. J. Immunol. 190, 4661–4675 (2013).

    CAS  PubMed  Google Scholar 

  106. Zeng, Y. et al. Substrate stiffness regulates B-cell activation, proliferation, class switch, and T-cell-independent antibody responses in vivo. Eur. J. Immunol. 45, 1621–1634 (2015).

    CAS  PubMed  Google Scholar 

  107. Kim, S. T. et al. The αβ T cell receptor is an anisotropic mechanosensor. J. Biol. Chem. 284, 31028–31037 (2009).

    CAS  PubMed  PubMed Central  Google Scholar 

  108. Das, D. K. et al. Force-dependent transition in the T-cell receptor β-subunit allosterically regulates peptide discrimination and pMHC bond lifetime. Proc. Natl Acad. Sci. USA 112, 1517–1522 (2015).

    CAS  PubMed  Google Scholar 

  109. 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 demonstrates that the TCR forms catch bonds with peptide–MHC complexes and that catch bonds are crucial for discrimination between high-affinity and low-affinity antigens.

    CAS  PubMed  PubMed Central  Google Scholar 

  110. Aivazian, D. & Stern, L. J. Phosphorylation of T cell receptor ζ is regulated by a lipid dependent folding transition. Nat. Struct. Biol. 7, 1023–1026 (2000).

    CAS  PubMed  Google Scholar 

  111. Xu, C. et al. Regulation of T cell receptor activation by dynamic membrane binding of the CD3ɛ cytoplasmic tyrosine-based motif. Cell 135, 702–713 (2008).

    CAS  PubMed  PubMed Central  Google Scholar 

  112. Lee, M. S. et al. A mechanical switch couples T cell receptor triggering to the cytoplasmic juxtamembrane regions of CD3ζζ. Immunity 43, 227–239 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  113. Swamy, M. et al. A cholesterol-based allostery model of T cell receptor phosphorylation. Immunity 44, 1091–1101 (2016).

    CAS  PubMed  Google Scholar 

  114. Wan, Z. et al. The activation of IgM- or isotype-switched IgG- and IgE-BCR exhibits distinct mechanical force sensitivity and threshold. eLife 4, e06925 (2015).

    PubMed Central  Google Scholar 

  115. 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).

    CAS  PubMed  PubMed Central  Google Scholar 

  116. Yi, J., Wu, X. S., Crites, T. & Hammer, J. A. 3rd. Actin retrograde flow and actomyosin II arc contraction drive receptor cluster dynamics at the immunological synapse in Jurkat T cells. Mol. Biol. Cell 23, 834–852 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  117. Cai, E. et al. Visualizing dynamic microvillar search and stabilization during ligand detection by T cells. Science 356, eaal3118 (2017).

    PubMed  PubMed Central  Google Scholar 

  118. 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).

    PubMed Central  Google Scholar 

  119. Pribila, J. T., Quale, A. C., Mueller, K. L. & Shimizu, Y. Integrins and T cell-mediated immunity. Annu. Rev. Immunol. 22, 157–180 (2004).

    CAS  PubMed  Google Scholar 

  120. Comrie, W. A., Babich, A. & Burkhardt, J. K. F-Actin flow drives affinity maturation and spatial organization of LFA-1 at the immunological synapse. J. Cell Biol. 208, 475–491 (2015).

    CAS  PubMed  PubMed Central  Google Scholar 

  121. Comrie, W. A., Li, S., Boyle, S. & Burkhardt, J. K. The dendritic cell cytoskeleton promotes T cell adhesion and activation by constraining ICAM-1 mobility. J. Cell Biol. 208, 457–473 (2015). This study shows that DCs immobilize cell-surface ICAM1 to promote integrin catch-bond formation within the immunological synapse, thus demonstrating that cells control the function of adhesion molecules by regulating their biophysical state.

    CAS  PubMed  PubMed Central  Google Scholar 

  122. Freeman, S. A. & Grinstein, S. Phagocytosis: receptors, signal integration, and the cytoskeleton. Immunol. Rev. 262, 193–215 (2014).

    CAS  PubMed  Google Scholar 

  123. Goodridge, H. S. et al. Activation of the innate immune receptor dectin-1 upon formation of a 'phagocytic synapse'. Nature 472, 471–475 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  124. Niedergang, F., Di Bartolo, V. & Alcover, A. Comparative anatomy of phagocytic and immunological synapses. Front. Immunol. 7, 18 (2016).

    PubMed  PubMed Central  Google Scholar 

  125. Freeman, S. A. et al. Integrins form an expanding diffusional barrier that coordinates phagocytosis. Cell 164, 128–140 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  126. Moller, J., Luhmann, T., Chabria, M., Hall, H. & Vogel, V. Macrophages lift off surface-bound bacteria using a filopodium-lamellipodium hook-and-shovel mechanism. Sci. Rep. 3, 2884 (2013).

    PubMed  PubMed Central  Google Scholar 

  127. Victora, G. D. & Nussenzweig, M. C. Germinal centers. Annu. Rev. Immunol. 30, 429–457 (2012).

    CAS  PubMed  Google Scholar 

  128. Nowosad, C. R., Spillane, K. M. & Tolar, P. Germinal center B cells recognize antigen through a specialized immune synapse architecture. Nat. Immunol. 17, 870–877 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  129. Olazabal, I. M. et al. Rho-kinase and myosin-II control phagocytic cup formation during CR, but not FcγR, phagocytosis. Curr. Biol. 12, 1413–1418 (2002).

    CAS  PubMed  Google Scholar 

  130. Araki, N., Hatae, T., Furukawa, A. & Swanson, J. A. Phosphoinositide-3-kinase-independent contractile activities associated with Fcγ-receptor-mediated phagocytosis and macropinocytosis in macrophages. J. Cell Sci. 116, 247–257 (2003).

    CAS  PubMed  Google Scholar 

  131. Dart, A. E., Tollis, S., Bright, M. D., Frankel, G. & Endres, R. G. The motor protein myosin 1G functions in FcγR-mediated phagocytosis. J. Cell Sci. 125, 6020–6029 (2012).

    CAS  PubMed  Google Scholar 

  132. Swanson, J. A. et al. A contractile activity that closes phagosomes in macrophages. J. Cell Sci. 112, 307–316 (1999).

    CAS  PubMed  Google Scholar 

  133. Cox, D. et al. Myosin X is a downstream effector of PI(3)K during phagocytosis. Nat. Cell Biol. 4, 469–477 (2002).

    CAS  PubMed  Google Scholar 

  134. Masters, T. A., Pontes, B., Viasnoff, V., Li, Y. & Gauthier, N. C. Plasma membrane tension orchestrates membrane trafficking, cytoskeletal remodeling, and biochemical signaling during phagocytosis. Proc. Natl Acad. Sci. USA 110, 11875–11880 (2013). This study identifies membrane tension as a key trigger for particle internalization during phagocytosis.

    CAS  PubMed  Google Scholar 

  135. Keefe, D. et al. Perforin triggers a plasma membrane-repair response that facilitates CTL induction of apoptosis. Immunity 23, 249–262 (2005).

    CAS  PubMed  Google Scholar 

  136. Thiery, J. et al. Perforin pores in the endosomal membrane trigger the release of endocytosed granzyme B into the cytosol of target cells. Nat. Immunol. 12, 770–777 (2011).

    CAS  PubMed  PubMed Central  Google Scholar 

  137. Stinchcombe, J. C. & Griffiths, G. M. Secretory mechanisms in cell-mediated cytotoxicity. Annu. Rev. Cell Dev. Biol. 23, 495–517 (2007).

    CAS  PubMed  Google Scholar 

  138. Bashour, K. T. et al. CD28 and CD3 have complementary roles in T-cell traction forces. Proc. Natl Acad. Sci. USA 111, 2241–2246 (2014).

    CAS  PubMed  Google Scholar 

  139. Le Floc'h, A. et al. Annular PIP3 accumulation controls actin architecture and modulates cytotoxicity at the immunological synapse. J. Exp. Med. 210, 2721–2737 (2013).

    PubMed  PubMed Central  Google Scholar 

  140. Huang, H. W., Chen, F. Y. & Lee, M. T. Molecular mechanism of peptide-induced pores in membranes. Phys. Rev. Lett. 92, 198304 (2004).

    PubMed  Google Scholar 

  141. Lee, M. T., Hung, W. C., Chen, F. Y. & Huang, H. W. Mechanism and kinetics of pore formation in membranes by water-soluble amphipathic peptides. Proc. Natl Acad. Sci. USA 105, 5087–5092 (2008).

    CAS  PubMed  Google Scholar 

  142. Polozov, I. V., Anantharamaiah, G. M., Segrest, J. P. & Epand, R. M. Osmotically induced membrane tension modulates membrane permeabilization by class L amphipathic helical peptides: nucleation model of defect formation. Biophys. J. 81, 949–959 (2001).

    CAS  PubMed  PubMed Central  Google Scholar 

  143. Guck, J. et al. Optical deformability as an inherent cell marker for testing malignant transformation and metastatic competence. Biophys. J. 88, 3689–3698 (2005).

    CAS  PubMed  PubMed Central  Google Scholar 

  144. Hou, H. W. et al. Deformability study of breast cancer cells using microfluidics. Biomed. Microdevices 11, 557–564 (2009).

    CAS  PubMed  Google Scholar 

  145. Xu, W. et al. Cell stiffness is a biomarker of the metastatic potential of ovarian cancer cells. PLoS ONE 7, e46609 (2012).

    CAS  PubMed  PubMed Central  Google Scholar 

  146. Liu, Y. et al. DNA-based nanoparticle tension sensors reveal that T-cell receptors transmit defined pN forces to their antigens for enhanced fidelity. Proc. Natl Acad. Sci. USA 113, 5610–5615 (2016).

    CAS  PubMed  Google Scholar 

  147. Zhang, Y., Ge, C., Zhu, C. & Salaita, K. DNA-based digital tension probes reveal integrin forces during early cell adhesion. Nat. Commun. 5, 5167 (2014).

    CAS  PubMed  PubMed Central  Google Scholar 

  148. Grashoff, C. et al. Measuring mechanical tension across vinculin reveals regulation of focal adhesion dynamics. Nature 466, 263–266 (2010).

    CAS  PubMed  PubMed Central  Google Scholar 

  149. 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).

    CAS  PubMed  Google Scholar 

  150. Conway, D. E. et al. Fluid shear stress on endothelial cells modulates mechanical tension across VE-cadherin and PECAM-1. Curr. Biol. 23, 1024–1030 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

  151. Polacheck, W. J. & Chen, C. S. Measuring cell-generated forces: a guide to the available tools. Nat. Methods 13, 415–423 (2016).

    CAS  PubMed  PubMed Central  Google Scholar 

  152. Liu, B., Chen, W. & Zhu, C. Molecular force spectroscopy on cells. Annu. Rev. Phys. Chem. 66, 427–451 (2015).

    CAS  PubMed  Google Scholar 

  153. Cost, A. L., Ringer, P., Chrostek-Grashoff, A. & Grashoff, C. How to measure molecular forces in cells: a guide to evaluating genetically-encoded FRET-based tension sensors. Cell. Mol. Bioeng. 8, 96–105 (2015).

    CAS  PubMed  Google Scholar 

  154. Evans, E., Ritchie, K. & Merkel, R. Sensitive force technique to probe molecular adhesion and structural linkages at biological interfaces. Biophys. J. 68, 2580–2587 (1995).

    CAS  PubMed  PubMed Central  Google Scholar 

  155. Dembo, M. & Wang, Y. L. Stresses at the cell-to-substrate interface during locomotion of fibroblasts. Biophys. J. 76, 2307–2316 (1999).

    CAS  PubMed  PubMed Central  Google Scholar 

  156. Tan, J. L. et al. Cells lying on a bed of microneedles: an approach to isolate mechanical force. Proc. Natl Acad. Sci. USA 100, 1484–1489 (2003).

    CAS  PubMed  Google Scholar 

  157. Wang, X. & Ha, T. Defining single molecular forces required to activate integrin and Notch signaling. Science 340, 991–994 (2013).

    CAS  PubMed  PubMed Central  Google Scholar 

Download references

Acknowledgements

The author thanks L. Kam and S. Grinstein for advice and discussions. This work was supported by the US National Institutes of Health National Institute of Allergy and Infectious Diseases (grant R01AI087644) and the US National Science Foundation (grant CMMI-1562905).

Author information

Authors and Affiliations

Authors

Corresponding author

Correspondence to Morgan Huse.

Ethics declarations

Competing interests

The author declares no competing financial interests.

PowerPoint slides

Glossary

Focal adhesions

Force-bearing multiprotein complexes (composed of integrins, cytoskeletal adaptors and signalling molecules) that anchor the cortical filamentous actin cytoskeleton of adherent cells to the extracellular matrix.

Mechanosensing

The capacity of cells or molecules to detect physical properties or perturbations.

Mechanotransduction

The process through which cells sense and respond to their mechanical environment, such as the extracellular matrix, adjacent cells or external stresses. During mechanotransduction, mechanical signals are sensed and activate intracellular biochemical signalling pathways.

Slip bonds

Bonds that are characterized by having a lifetime that decreases with applied force.

Catch bond

A bond that is characterized by having a lifetime that increases up to an optimal applied force, after which it decreases with increasing force.

Lamellipodium

A broad, flat protrusion at the leading edge of a moving cell that is enriched with a branched network of elongating actin filaments that generate the force to push the cell membrane forwards.

Uropod

A slender appendage that is formed at the trailing, rear edge of fast-migrating cells such as amoebae, neutrophils and lymphocytes.

Inside-out signalling

The process by which intracellular signalling mechanisms result in the activation of a cell-surface receptor, such as an integrin. By contrast, outside-in signalling is the process by which the ligation of a cell-surface receptor activates signalling pathways inside the cell.

WASP-family verprolin-homologous protein 2 complex

(WAVE2 complex). A complex that comprises five proteins and that regulates actin-related protein 2/3 (ARP2/3) complex-mediated actin assembly. The defining member of this complex, WAVE, is related to Wiskott–Aldrich syndrome protein and directly stimulates the ARP2/3 complex to nucleate actin assembly, while other components of the complex regulate WAVE.

Extravasate

To move out of the circulatory system, typically through vessel walls. Extravasation is also called diapedesis.

Immunological synapse

A large junctional structure that forms between a lymphocyte and its antigen-presenting cell (APC). The immunological synapse regulates lymphocyte activation, cell adhesion and the directional secretion of cytokines and cytolytic factors.

Rights and permissions

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Huse, M. Mechanical forces in the immune system. Nat Rev Immunol 17, 679–690 (2017). https://doi.org/10.1038/nri.2017.74

Download citation

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/nri.2017.74

This article is cited by

Search

Quick links

Nature Briefing

Sign up for the Nature Briefing newsletter — what matters in science, free to your inbox daily.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing