Synthetic mechanotransduction

Posted on
synthetic-mechanotransduction
Synthetic mechanotransduction

References

  1. Xie, M. & Fussenegger, M. Designing cell function: assembly of synthetic gene circuits for cell biology applications. Nat. Rev. Mol. Cell Biol. 19, 507–525 (2018).

    Article  Google Scholar 

  2. Lim, W. A. The emerging era of cell engineering: harnessing the modularity of cells to program complex biological function. Science 378, 848–852 (2022).

    Article  Google Scholar 

  3. Saxena, P. et al. A programmable synthetic lineage-control network that differentiates human IPSCs into glucose-sensitive insulin-secreting beta-like cells. Nat. Commun. 7, 11247 (2016).

    Article  Google Scholar 

  4. Courbet, A., Endy, D., Renard, E., Molina, F. & Bonnet, J. Detection of pathological biomarkers in human clinical samples via amplifying genetic switches and logic gates. Sci. Transl. Med. 7, 289ra83 (2015).

    Article  Google Scholar 

  5. Weber, W. et al. A synthetic mammalian gene circuit reveals antituberculosis compounds. Proc. Natl Acad. Sci. 105, 9994–9998 (2008).

    Article  Google Scholar 

  6. Nissim, L. et al. Synthetic RNA-based immunomodulatory gene circuits for cancer immunotherapy. Cell 171, 1138–1150.e15 (2017).

    Article  Google Scholar 

  7. Peng, L., Sferruzza, G., Yang, L., Zhou, L. & Chen, S. CAR-T and CAR-NK as cellular cancer immunotherapy for solid tumors. Cell. Mol. Immunol. 21, 1089–1108 (2024).

    Article  Google Scholar 

  8. Garreta, E. et al. Fine tuning the extracellular environment accelerates the derivation of kidney organoids from human pluripotent stem cells. Nat. Mater. 18, 397–405 (2019).

    Article  Google Scholar 

  9. Barriga, E. H., Franze, K., Charras, G. & Mayor, R. Tissue stiffening coordinates morphogenesis by triggering collective cell migration in vivo. Nature 554, 523–527 (2018).

    Article  Google Scholar 

  10. Barnes, J. M., Przybyla, L. & Weaver, V. M. Tissue mechanics regulate brain development, homeostasis and disease. J. Cell Sci. 130, 71–82 (2017).

    Article  Google Scholar 

  11. Tschumperlin, D. J., Ligresti, G., Hilscher, M. B. & Shah, V. H. Mechanosensing and fibrosis. J. Clin. Invest. 128, 74–84 (2018).

    Article  Google Scholar 

  12. Chatterjee, S. Endothelial mechanotransduction, redox signaling and the regulation of vascular inflammatory pathways. Front. Physiol. 9, 524 (2018).

    Article  Google Scholar 

  13. Nia, H. T., Munn, L. L. & Jain, R. K. Physical traits of cancer. Science 370, eaaz0868 (2020).

    Article  Google Scholar 

  14. Lampi, M. C. & Reinhart-King, C. A. Targeting extracellular matrix stiffness to attenuate disease: from molecular mechanisms to clinical trials. Sci. Transl. Med. 10, eaao0475 (2018).

    Article  Google Scholar 

  15. Paszek, M. J. et al. Tensional homeostasis and the malignant phenotype. Cancer Cell 8, 241–254 (2005).

    Article  Google Scholar 

  16. Gaggioli, C. et al. Fibroblast-led collective invasion of carcinoma cells with differing roles for RhoGTPases in leading and following cells. Nat. Cell Biol. 9, 1392–1400 (2007).

    Article  Google Scholar 

  17. Gomez, E. W., Chen, Q. K., Gjorevski, N. & Nelson, C. M. Tissue geometry patterns epithelial–mesenchymal transition via intercellular mechanotransduction. J. Cell Biochem. 110, 44–51 (2010).

    Article  Google Scholar 

  18. Kechagia, J. Z., Ivaska, J. & Roca-Cusachs, P. Integrins as biomechanical sensors of the microenvironment. Nat. Rev. Mol. Cell Biol. 20, 457–473 (2019).

    Article  Google Scholar 

  19. Kirby, T. J. & Lammerding, J. Emerging views of the nucleus as a cellular mechanosensor. Nat. Cell Biol. 20, 373–381 (2018).

    Article  Google Scholar 

  20. Kefauver, J. M., Ward, A. B. & Patapoutian, A. Discoveries in structure and physiology of mechanically activated ion channels. Nature 587, 567–576 (2020).

    Article  Google Scholar 

  21. Faure, L. M., Venturini, V. & Roca-Cusachs, P. Cell compression — relevance, mechanotransduction mechanisms and tools. J. Cell Sci. 138, jcs263704 (2025).

    Article  Google Scholar 

  22. Chen, W., Lou, J. & Zhu, C. Forcing switch from short- to intermediate- and long-lived states of the alphaA domain generates LFA-1/ICAM-1 catch bonds. J. Biol. Chem. 285, 35967–35978 (2010).

    Article  Google Scholar 

  23. Kong, F., García, 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).

    Article  Google Scholar 

  24. del Rio, A. et al. Stretching single talin rod molecules activates vinculin binding. Science 323, 638–641 (2009).

    Article  Google Scholar 

  25. Yao, M. et al. Mechanical activation of vinculin binding to talin locks talin in an unfolded conformation. Sci. Rep. 4, 4610 (2014).

    Article  Google Scholar 

  26. 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).

    Article  Google Scholar 

  27. Oria, R. et al. Force loading explains spatial sensing of ligands by cells. Nature 552, 219–224 (2017).

    Article  Google Scholar 

  28. 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).

    Article  Google Scholar 

  29. Desprat, N., Supatto, W., Pouille, P. A., Beaurepaire, E. & Farge, E. Tissue deformation modulates twist expression to determine anterior midgut differentiation in Drosophila embryos. Dev. Cell 15, 470–477 (2008).

    Article  Google Scholar 

  30. Röper, J.-C. et al. The major β-catenin/E-cadherin junctional binding site is a primary molecular mechano-transductor of differentiation in vivo. eLife 7, e33381 (2018).

    Article  Google Scholar 

  31. Zhou, B. et al. Notch signaling pathway: architecture, disease, and therapeutics. Signal. Transduct. Target. Ther. 7, 95 (2022).

    Article  Google Scholar 

  32. Gordon, W. R. et al. Mechanical allostery: evidence for a force requirement in the proteolytic activation of Notch. Dev. Cell 33, 729–736 (2015).

    Article  Google Scholar 

  33. Langridge, P. D. & Struhl, G. Epsin-dependent ligand endocytosis activates notch by force. Cell 171, 1383–1396.e12 (2017).

    Article  Google Scholar 

  34. Khamaisi, B., Luca, V. C., Blacklow, S. C. & Sprinzak, D. Functional comparison between endogenous and synthetic Notch systems. ACS Synth. Biol. 11, 3343–3353 (2022).

    Article  Google Scholar 

  35. Coste, B. et al. Piezo1 and Piezo2 are essential components of distinct mechanically activated cation channels. Science 330, 55–60 (2010).

    Article  Google Scholar 

  36. Servin-Vences, M. R., Moroni, M., Lewin, G. R. & Poole, K. Direct measurement of TRPV4 and Piezo1 activity reveals multiple mechanotransduction pathways in chondrocytes. eLife 6, e21074 (2017).

    Article  Google Scholar 

  37. O’Conor, C. J., Leddy, H. A., Benefield, H. C., Liedtke, W. B. & Guilak, F. TRPV4-mediated mechanotransduction regulates the metabolic response of chondrocytes to dynamic loading. Proc. Natl Acad. Sci. USA 111, 1316–1321 (2014).

    Article  Google Scholar 

  38. Huang, M. & Chalfie, M. Gene interactions affecting mechanosensory transduction in Caenorhabditis elegans. Nature 367, 467–470 (1994).

    Article  Google Scholar 

  39. Xiao, B. Mechanisms of mechanotransduction and physiological roles of PIEZO channels. Nat. Rev. Mol. Cell Biol. 25, 886–903 (2024).

    Article  Google Scholar 

  40. Sinha, B. et al. Cells respond to mechanical stress by rapid disassembly of caveolae. Cell 144, 402–413 (2011).

    Article  Google Scholar 

  41. Quiroga, X. et al. A mechanosensing mechanism controls plasma membrane shape homeostasis at the nanoscale. eLife 12, e72316 (2023).

    Article  Google Scholar 

  42. Diz-Muñoz, A. et al. Membrane tension acts through PLD2 and mTORC2 to limit actin network assembly during neutrophil migration. PLoS Biol. 14, 1–30 (2016).

    Article  Google Scholar 

  43. Roux, A.-L. L. e, Quiroga, X., Walani, N., Arroyo, M. & Roca-Cusachs, P. The plasma membrane as a mechanochemical transducer. Phil. Trans. R. Soc. B 374, 20180221 (2019).

    Article  Google Scholar 

  44. Chachisvilis, M., Zhang, Y. L. & Frangos, J. A. G protein-coupled receptors sense fluid shear stress in endothelial cells. Proc. Natl Acad. Sci. USA 103, 15463–15468 (2006).

    Article  Google Scholar 

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

    Article  Google Scholar 

  46. Elosegui-Artola, A. et al. Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171, 1397–1410.e14 (2017).

    Article  Google Scholar 

  47. Lombardi, M. L. et al. The interaction between nesprins and SUN proteins at the nuclear envelope is critical for force transmission between the nucleus and cytoskeleton. J. Biol. Chem. 286, 26743–26753 (2011).

    Article  Google Scholar 

  48. Niethammer, P. Components and mechanisms of nuclear mechanotransduction. Annu. Rev. Cell Dev. Biol. 37, 233–256 (2021).

    Article  Google Scholar 

  49. Enyedi, B., Jelcic, M. & Niethammer, P. The cell nucleus serves as a mechanotransducer of tissue damage-induced inflammation. Cell 165, 1160–1170 (2016).

    Article  Google Scholar 

  50. Venturini, V. et al. The nucleus measures shape changes for cellular proprioception to control dynamic cell behavior. Science 370, eaba2644 (2020).

    Article  Google Scholar 

  51. Lomakin, A. J. et al. The nucleus acts as a ruler tailoring cell responses to spatial constraints. Science 370, eaba2894 (2020).

    Article  Google Scholar 

  52. Tajik, A. et al. Transcription upregulation via force-induced direct stretching of chromatin. Nat. Mater. 15, 1287–1296 (2016).

    Article  Google Scholar 

  53. Zimmerli, C. E. et al. Nuclear pores dilate and constrict in cellulo. Science 374, eabd9776 (2021).

    Article  Google Scholar 

  54. Aureille, J. et al. Nuclear envelope deformation controls cell cycle progression in response to mechanical force. EMBO Rep. 20, e48084 (2019).

    Article  Google Scholar 

  55. Andreu, I. et al. Mechanical force application to the nucleus regulates nucleocytoplasmic transport. Nat. Cell Biol. 24, 896–905 (2022).

    Article  Google Scholar 

  56. Dupont, S. et al. Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183 (2011).

    Article  Google Scholar 

  57. Luciano, M. et al. Cell monolayers sense curvature by exploiting active mechanics and nuclear mechanoadaptation. Nat. Phys. 17, 1382–1390 (2021).

    Article  Google Scholar 

  58. Koushki, N., Ghagre, A., Srivastava, L. K., Molter, C. & Ehrlicher, A. J. Nuclear compression regulates YAP spatiotemporal fluctuations in living cells. Proc. Natl Acad. Sci. USA 120, e2301285120 (2023).

    Article  Google Scholar 

  59. Tharp, K. M. et al. Adhesion-mediated mechanosignaling forces mitohormesis. Cell Metab. 33, 1322–1341.e13 (2021).

    Article  Google Scholar 

  60. Phuyal, S. et al. Mechanical strain stimulates COPII-dependent secretory trafficking via Rac1. EMBO J. 41, e110596 (2022).

    Article  Google Scholar 

  61. Ucar, H. et al. Mechanical actions of dendritic-spine enlargement on presynaptic exocytosis. Nature 600, 686–689 (2021).

    Article  Google Scholar 

  62. Liu, L. et al. Mechanoresponsive stem cells to target cancer metastases through biophysical cues. Sci. Transl. Med. 9, eaan2966 (2017). This paper describes synthetic mechanotransduction triggered by tissue mechanical properties, based on the mechanosensor YAP.

    Article  Google Scholar 

  63. Pan, Y. et al. Mechanogenetics for the remote and noninvasive control of cancer immunotherapy. Proc. Natl Acad. Sci. USA 115, 992–997 (2018).

    Article  Google Scholar 

  64. Woo Yoon, C. et al. Tumor priming by ultrasound mechanogenetics for with SynNotch CAR T therapy. Preprint at bioRxiv https://doi.org/10.1101/2024.10.01.615989 (2024). This paper describes synthetic mechanotransduction triggered by ultrasound, providing both mechanical and chemical control, based on the mechanosensor Piezo1.

  65. Lee, J. U. et al. Non-contact long-range magnetic stimulation of mechanosensitive ion channels in freely moving animals. Nat. Mater. 20, 1029–1036 (2021).

    Article  Google Scholar 

  66. Shin, W. et al. Magnetogenetics with Piezo1 mechanosensitive ion channel for CRISPR gene editing. Nano Lett. 22, 7415–7422 (2022). This paper describes synthetic mechanotransduction triggered by magnetic fields resulting in gene editing, based on the mechanosensor Piezo1.

    Article  Google Scholar 

  67. Nims, R. J. et al. A synthetic mechanogenetic gene circuit for autonomous drug delivery in engineered tissues. Sci. Adv. 7, 9858–9885 (2021).

    Article  Google Scholar 

  68. Yang, X., Zeng, H., Wang, L., Luo, S. & Zhou, Y. Activation of Piezo1 downregulates renin in juxtaglomerular cells and contributes to blood pressure homeostasis. Cell Biosci. 12, 197 (2022).

    Article  Google Scholar 

  69. Kim, Y. S., Steward, N., Kim, A., Fehle, I. & Guilak, F. Tuning the response of synthetic mechanogenetic gene circuits using mutations in TRPV4. Tissue Eng. A 31, 174–183 (2025).

    Article  Google Scholar 

  70. Heureaux, J., Chen, D., Murray, V. L., Deng, C. X. & Liu, A. P. Activation of a bacterial mechanosensitive channel in mammalian cells by cytoskeletal stress. Cell Mol. Bioeng. 7, 307–319 (2014).

    Article  Google Scholar 

  71. Soloperto, A. et al. Mechano-sensitization of mammalian neuronal networks through expression of the bacterial large-conductance mechanosensitive ion channel. J. Cell Sci. 131, jcs210393 (2018).

    Article  Google Scholar 

  72. Yoshimura, K., Batiza, A., Schroeder, M., Blount, P. & Kung, C. Hydrophilicity of a single residue within MscL correlates with increased channel mechanosensitivity. Biophys. J. 77, 1960–1972 (1999).

    Article  Google Scholar 

  73. Iscla, I. & Blount, P. Sensing and responding to membrane tension: the bacterial MscL channel as a model system. Biophys. J. 103, 169–174 (2012).

    Article  Google Scholar 

  74. Zhao, H. et al. Tuning of cellular insulin release by music for real-time diabetes control. Lancet Diabetes Endocrinol. 11, 637–640 (2023).

    Article  Google Scholar 

  75. Liu, Y. et al. Robotic actuation-mediated quantitative mechanogenetics for noninvasive and on-demand cancer therapy. Adv. Sci. 11, 2401611 (2024).

    Article  Google Scholar 

  76. Pocaterra, A., Romani, P. & Dupont, S. YAP/TAZ functions and their regulation at a glance. J. Cell Sci. 133, jcs230425 (2020).

    Article  Google Scholar 

  77. Luan, S. & Wang, C. Calcium signaling mechanisms across kingdoms. Annu. Rev. Cell Dev. Biol. 37, 311–340 (2021).

    Article  Google Scholar 

  78. Kis, Z. et al. Development of a synthetic gene network to modulate gene expression by mechanical forces. Sci. Rep. 6, 29643 (2016).

    Article  Google Scholar 

  79. Barnea, G. et al. The genetic design of signaling cascades to record receptor activation. Proc. Natl Acad. Sci. USA 105, 64–69 (2008).

    Article  Google Scholar 

  80. Morsut, L. et al. Engineering customized cell sensing and response behaviors using synthetic notch receptors. Cell 164, 780–791 (2016).

    Article  Google Scholar 

  81. Meloty-Kapella, L., Shergill, B., Kuon, J., Botvinick, E. & Weinmaster, G. Notch ligand endocytosis generates mechanical pulling force dependent on dynamin, epsins, and actin. Dev. Cell 22, 1299–1312 (2012).

    Article  Google Scholar 

  82. McMillan, B. J. et al. A tail of two sites: a bipartite mechanism for recognition of Notch ligands by mind bomb E3 ligases. Mol. Cell 57, 912–924 (2015).

    Article  Google Scholar 

  83. Garibyan, M. et al. Engineering programmable material-to-cell pathways via synthetic Notch receptors to spatially control differentiation in multicellular constructs. Nat. Commun. 15, 5891 (2024).

    Article  Google Scholar 

  84. Handa, H., Idesako, N. & Itoh, M. Immobilized DLL4-induced Notch signaling is mediated by dynamics of the actin cytoskeleton. Biochem. Biophys. Res. Commun. 602, 179–185 (2022). This paper describes engineering of synthetic cell receptors is based on Notch, with customizable extra- and intracellular ligands.

    Article  Google Scholar 

  85. Sloas, D. C., Tran, J. C., Marzilli, A. M. & Ngo, J. T. Tension-tuned receptors for synthetic mechanotransduction and intercellular force detection. Nat. Biotechnol. 41, 1287–1295 (2023). Using the synthetic Notch developed by Handa et al., this paper describes engineering of synthetic Notch receptors sensitive to specific ranges of applied force.

    Article  Google Scholar 

  86. Yang, S. et al. DNA-functionalized artificial mechanoreceptor for de novo force-responsive signaling. Nat. Chem. Biol. 20, 1066–1077 (2024). This paper describes a DNA-based synthetic mechanosensor.

    Article  Google Scholar 

  87. Simmel, F. C., Yurke, B. & Singh, H. R. Principles and applications of nucleic acid strand displacement reactions. Chem. Rev. 119, 6326–6369 (2019).

    Article  Google Scholar 

  88. Walbrun, A. et al. Single-molecule force spectroscopy of toehold-mediated strand displacement. Nat. Commun. 15, 7564 (2024).

    Article  Google Scholar 

  89. Hsu, Y. Y., Resto Irizarry, A. M., Fu, J. & Liu, A. P. Mechanosensitive channel-based optical membrane tension reporter. ACS Sens. 8, 12–18 (2023).

    Article  Google Scholar 

  90. Granero-Moya, I. et al. Nucleocytoplasmic transport senses mechanical forces independently of cell density in cell monolayers. J. Cell Sci. 137, jcs262363 (2024).

    Article  Google Scholar 

  91. Oakes, P. W., Banerjee, S., Marchetti, M. C. & Gardel, M. L. Geometry regulates traction stresses in adherent cells. Biophys. J. 107, 825–833 (2014).

    Article  Google Scholar 

  92. Faure, L. M. et al. 3D micropatterned traction force microscopy: a technique to control 3D cell shape while measuring cell-substrate force transmission. Adv. Sci. 11, 2406932 (2024).

    Article  Google Scholar 

  93. Elosegui-Artola, A. et al. Rigidity sensing and adaptation through regulation of integrin types. Nat. Mater. 13, 631–637 (2014).

    Article  Google Scholar 

  94. Chan, C. E. & Odde, D. J. Traction dynamics of filopodia on compliant substrates. Science 322, 1687–1691 (2008).

    Article  Google Scholar 

  95. Elosegui-Artola, A., Trepat, X. & Roca-Cusachs, P. Control of mechanotransduction by molecular clutch dynamics. Trends Cell Biol. 28, 356–367 (2018).

    Article  Google Scholar 

  96. Krishnan, R. et al. Reinforcement versus fluidization in cytoskeletal mechanoresponsiveness. PLoS ONE 4, e5486 (2009).

    Article  Google Scholar 

  97. Andreu, I. et al. The force loading rate drives cell mechanosensing through both reinforcement and cytoskeletal softening. Nat. Commun. 12, 4229 (2021).

    Article  Google Scholar 

  98. Asano, T., Ishizua, T. & Yawo, H. Optically controlled contraction of photosensitive skeletal muscle cells. Biotechnol. Bioeng. 109, 199–204 (2012).

    Article  Google Scholar 

  99. Sakar, M. S. et al. Formation and optogenetic control of engineered 3D skeletal muscle bioactuators. Lab. Chip 12, 4976–4985 (2012).

    Article  Google Scholar 

  100. Raman, R. et al. Optogenetic skeletal muscle-powered adaptive biological machines. Proc. Natl Acad. Sci. USA 113, 3497–3502 (2016).

    Article  Google Scholar 

  101. Raman, R. Biofabrication of living actuators. Annu. Rev. Biomed. Eng. 26, 223–245 (2024).

    Article  Google Scholar 

  102. Lee, K. Y. et al. An autonomously swimming biohybrid fish designed with human cardiac biophysics. Science 375, 639–647 (2022).

    Article  Google Scholar 

  103. Izquierdo, E., Quinkler, T. & De Renzis, S. Guided morphogenesis through optogenetic activation of Rho signalling during early Drosophila embryogenesis. Nat. Commun. 9, 2366 (2018).

    Article  Google Scholar 

  104. Valon, L., Marín-Llauradó, A., Wyatt, T., Charras, G. & Trepat, X. Optogenetic control of cellular forces and mechanotransduction. Nat. Commun. 8, 14396 (2017). This paper describes a molecular system that triggers cell force generation with light.

    Article  Google Scholar 

  105. Wagner, E. & Glotzer, M. Local RhoA activation induces cytokinetic furrows independent of spindle position and cell cycle stage. J. Cell Biol. 213, 641–649 (2016).

    Article  Google Scholar 

  106. Oakes, P. W. et al. Optogenetic control of RhoA reveals zyxin-mediated elasticity of stress fibres. Nat. Commun. 8, 15817 (2017).

    Article  Google Scholar 

  107. Kennedy, M. J. et al. Rapid blue-light-mediated induction of protein interactions in living cells. Nat. Methods 7, 973–975 (2010).

    Article  Google Scholar 

  108. Ruppel, A. et al. Force propagation between epithelial cells depends on active coupling and mechano-structural polarization. eLife 12, e83588 (2023).

    Article  Google Scholar 

  109. Cavanaugh, K. E., Staddon, M. F., Munro, E., Banerjee, S. & Gardel, M. L. RhoA mediates epithelial cell shape changes via mechanosensitive endocytosis. Dev. Cell 52, 152–166.e5 (2020).

    Article  Google Scholar 

  110. Yamamoto, K. et al. Optogenetic relaxation of actomyosin contractility uncovers mechanistic roles of cortical tension during cytokinesis. Nat. Commun. 12, 7145 (2021).

    Article  Google Scholar 

  111. Herrera-Perez, R. M., Cupo, C., Allan, C., Lin, A. & Kasza, K. E. Using optogenetics to link myosin patterns to contractile cell behaviors during convergent extension. Biophys. J. 120, 4214–4229 (2021).

    Article  Google Scholar 

  112. MacHacek, M. et al. Coordination of Rho GTPase activities during cell protrusion. Nature 461, 99–103 (2009).

    Article  Google Scholar 

  113. Valon, L. et al. Predictive spatiotemporal manipulation of signaling perturbations using optogenetics. Biophys. J. 109, 1785–1797 (2015).

    Article  Google Scholar 

  114. O’Neill, P. R., Kalyanaraman, V. & Gautam, N. Subcellular optogenetic activation of Cdc42 controls local and distal signaling to drive immune cell migration. Mol. Biol. Cell 27, 1442–1450 (2016).

    Article  Google Scholar 

  115. Valon, L. et al. Quantitative subcellular control of Cdc42, Rac1 and RhoA GTPases using the Cry2/CIBN optogenetic dimerizer. Biophys. J. 106, 244a (2014).

    Article  Google Scholar 

  116. Rossetti, L. et al. Optogenetic generation of leader cells reveals a force–velocity relation for collective cell migration. Nat. Phys. 20, 1659–1669 (2024).

    Article  Google Scholar 

  117. Rao, M. V., Chu, P. H., Hahn, K. M. & Zaidel-Bar, R. An optogenetic tool for the activation of endogenous diaphanous-related formins induces thickening of stress fibers without an increase in contractility. Cytoskeleton 70, 394–407 (2013).

    Article  Google Scholar 

  118. Wu, Y. I. et al. A genetically encoded photoactivatable Rac controls the motility of living cells. Nature 461, 104–108 (2009).

    Article  Google Scholar 

  119. Wang, X., He, L., Wu, Y. I., Hahn, K. M. & Montell, D. J. Light-mediated activation reveals a key role for Rac in collective guidance of cell movement in vivo. Nat. Cell Biol. 12, 591–597 (2010).

    Article  Google Scholar 

  120. Martin, A. C. & Goldstein, B. Apical constriction: themes and variations on a cellular mechanism driving morphogenesis. Development 141, 1987–1998 (2014).

    Article  Google Scholar 

  121. Martínez-Ara, G. et al. Optogenetic control of apical constriction induces synthetic morphogenesis in mammalian tissues. Nat. Commun. 13, 5400 (2022). This paper describes a molecular system that induces differential cell forces on the apical and basal sides of cells, thus controlling tissue morphogenesis.

    Article  Google Scholar 

  122. Zhang, W. et al. Optogenetic control with a photocleavable protein, PhoCl. Nat. Methods 14, 391–394 (2017).

    Article  Google Scholar 

  123. Endo, M., Iwawaki, T., Yoshimura, H. & Ozawa, T. Photocleavable cadherin inhibits cell-to-cell mechanotransduction by light. ACS Chem. Biol. 14, 2206–2214 (2019).

    Google Scholar 

  124. Ollech, D. et al. An optochemical tool for light-induced dissociation of adherens junctions to control mechanical coupling between cells. Nat. Commun. 11, 472 (2020). This paper describes control of cell–cell adhesion with light.

    Article  Google Scholar 

  125. Nzigou Mombo, B. et al. Reversible photoregulation of cell-cell adhesions with opto-E-cadherin. Nat. Commun. 14, 6292 (2023).

    Article  Google Scholar 

  126. Liao, Z. & Shattil, S. J. Talin, a Rap1 effector for integrin activation at the plasma membrane, also promotes Rap1 activity by disrupting sequestration of Rap1 by SHANK3. J. Cell Sci. 138, JCS263595 (2025).

    Article  Google Scholar 

  127. Liao, Z., Kasirer-Friede, A. & Shattil, S. J. Optogenetic interrogation of integrin αVβ3 function in endothelial cells. J. Cell Sci. 130, 3532–3541 (2017).

    Article  Google Scholar 

  128. Baaske, J. et al. Optogenetic control of integrin–matrix interaction. Commun. Biol. 2, 15 (2019).

    Article  Google Scholar 

  129. Petersen, S. et al. Phototriggering of cell adhesion by caged cyclic RGD peptides. Angew. Chem. 120, 3236–3239 (2008).

    Article  Google Scholar 

  130. Lee, T. T. et al. Light-triggered in vivo activation of adhesive peptides regulates cell adhesion, inflammation and vascularization of biomaterials. Nat. Mater. 14, 352–360 (2014). This paper describes the control of cell–matrix adhesion with light in vivo.

    Article  Google Scholar 

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

    Article  Google Scholar 

  132. 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).

    Article  Google Scholar 

  133. Jo, M. H. et al. Determination of single-molecule loading rate during mechanotransduction in cell adhesion. Science 383, 1374–1379 (2024).

    Article  Google Scholar 

  134. Combs, J. D. et al. Measuring integrin force loading rates using a two-step DNA tension sensor. J. Am. Chem. Soc. 146, 23034–23043 (2024).

    Article  Google Scholar 

  135. Schoenit, A. et al. Tuning epithelial cell–cell adhesion and collective dynamics with functional DNA-E-cadherin hybrid linkers. Nano Lett. 22, 302–310 (2022).

    Article  Google Scholar 

  136. Stevens, A. J. et al. Programming multicellular assembly with synthetic cell adhesion molecules. Nature 614, 144–152 (2023).

    Article  Google Scholar 

  137. Baba, H., Fujita, T., Mizuno, K., Tambo, M. & Toda, S. Programming spatial cell sorting by engineering cadherin intracellular activity. ACS Synth. Biol. 13, 1705–1715 (2024).

    Article  Google Scholar 

  138. Bijonowski, B. M. et al. Intercellular adhesion boots collective cell migration through elevated membrane tension. Nat. Commun. 16, 1588 (2025).

    Article  Google Scholar 

  139. Heinisch, J. J. et al. Atomic force microscopy — looking at mechanosensors on the cell surface. J. Cell Sci. 125, 4189–4195 (2012).

    Google Scholar 

  140. Wang, C. & Yadavalli, V. K. Investigating biomolecular recognition at the cell surface using atomic force microscopy. Micron 60, 5–17 (2014).

    Article  Google Scholar 

  141. Liu, J. et al. Tension gauge tethers as tension threshold and duration sensors. ACS Sens. 8, 704–711 (2023).

    Article  Google Scholar 

  142. Chaudhuri, O., Cooper-White, J., Janmey, P. A., Mooney, D. J. & Shenoy, V. B. Effects of extracellular matrix viscoelasticity on cellular behaviour. Nature 584, 535–546 (2020).

    Article  Google Scholar 

  143. Joseph, J. G. & Liu, A. P. Mechanical regulation of endocytosis: new insights and recent advances. Adv. Biosyst. 4, 1900278 (2020).

    Article  Google Scholar 

  144. Boulant, S., Kural, C., Zeeh, J. C., Ubelmann, F. & Kirchhausen, T. Actin dynamics counteract membrane tension during clathrin-mediated endocytosis. Nat. Cell Biol. 13, 1124–1131 (2011).

    Article  Google Scholar 

  145. Baschieri, F. et al. Frustrated endocytosis controls contractility-independent mechanotransduction at clathrin-coated structures. Nat. Commun. 9, 3825 (2018).

    Article  Google Scholar 

  146. Zhao, W. et al. Nanoscale manipulation of membrane curvature for probing endocytosis in live cells. Nat. Nanotechnol. 12, 750–756 (2017).

    Article  Google Scholar 

  147. Kaksonen, M. & Roux, A. Mechanisms of clathrin-mediated endocytosis. Nat. Rev. Mol. Cell Biol. 19, 313–326 (2018).

    Article  Google Scholar 

  148. Lovett, D. B., Shekhar, N., Nickerson, J. A., Roux, K. J. & Lele, T. P. Modulation of nuclear shape by substrate rigidity. Cell Mol. Bioeng. 6, 230–238 (2013).

    Article  Google Scholar 

  149. Swift, J. et al. Nuclear lamin-A scales with tissue stiffness and enhances matrix-directed differentiation. Science 341, 1240104 (2013).

    Article  Google Scholar 

  150. Guilak, F. Compression-induced changes in the shape and volume of the chondrocyte nucleus. J. Biomech. 28, 1529–1541 (1995).

    Article  Google Scholar 

  151. Lemière, J., Real-Calderon, P., Holt, L. J., Fai, T. G. & Chang, F. Control of nuclear size by osmotic forces in Schizosaccharomyces pombe. eLife 11, e76075 (2022).

    Article  Google Scholar 

  152. Finan, J. D., Chalut, K. J., Wax, A. & Guilak, F. Nonlinear osmotic properties of the cell nucleus. Ann. Biomed. Eng. 37, 477–491 (2009).

    Article  Google Scholar 

  153. Versaevel, M., Grevesse, T. & Gabriele, S. Spatial coordination between cell and nuclear shape within micropatterned endothelial cells. Nat. Commun. 3, 671 (2012).

    Article  Google Scholar 

  154. Wu, J. et al. Actomyosin pulls to advance the nucleus in a migrating tissue cell. Biophys. J. 106, 7–15 (2014).

    Article  Google Scholar 

  155. Kalukula, Y., Stephens, A. D., Lammerding, J. & Gabriele, S. Mechanics and functional consequences of nuclear deformations. Nat. Rev. Mol. Cell Biol. 23, 583–602 (2022).

    Article  Google Scholar 

  156. Malashicheva, A. & Perepelina, K. Diversity of nuclear lamin A/C action as a key to tissue-specific regulation of cellular identity in health and disease. Front. Cell Dev. Biol. 9, 761469 (2021).

    Article  Google Scholar 

  157. Furusawa, T. et al. Chromatin decompaction by the nucleosomal binding protein HMGN5 impairs nuclear sturdiness. Nat. Commun. 6, 6138 (2015).

    Article  Google Scholar 

  158. Stephens, A. D. et al. Chromatin histone modifications and rigidity affect nuclear morphology independent of lamins. Mol. Biol. Cell 29, 220–233 (2018).

    Article  Google Scholar 

  159. Kosmalska, A. J. et al. Physical principles of membrane remodelling during cell mechanoadaptation. Nat. Commun. 6, 7292 (2015).

    Article  Google Scholar 

  160. Kechagia, Z. et al. The laminin–keratin link shields the nucleus from mechanical deformation and signalling. Nat. Mater. 22, 1409–1420 (2023).

    Article  Google Scholar 

  161. Beedle, A. E. M. et al. Fibrillar adhesion dynamics govern the timescales of nuclear mechano-response via the vimentin cytoskeleton. Preprint at bioRxiv https://doi.org/10.1101/2023.11.08.566191 (2023).

  162. Nava, M. M. et al. Heterochromatin-driven nuclear softening protects the genome against mechanical stress-induced damage. Cell 181, 800–817.e22 (2020).

    Article  Google Scholar 

  163. Santorelli, M. et al. Control of spatio-temporal patterning via cell growth in a multicellular synthetic gene circuit. Nat. Commun. 15, 9867 (2024).

    Article  Google Scholar 

  164. Roca-Cusachs, P., Conte, V. & Trepat, X. Quantifying forces in cell biology. Nat. Cell Biol. 19, 742–751 (2017).

    Article  Google Scholar 

  165. Villeneuve, C., McCreery, K. P. & Wickström, S. A. Measuring and manipulating mechanical forces during development. Nat. Cell Biol. 27, 575–590 (2025).

    Article  Google Scholar 

  166. Brophy, J. A. N. & Voigt, C. A. Principles of genetic circuit design. Nat. Methods 11, 508–520 (2014).

    Article  Google Scholar 

  167. Kohn, J. C. et al. Cooperative effects of matrix stiffness and fluid shear stress on endothelial cell behavior. Biophys. J. 108, 471–478 (2015).

    Article  Google Scholar 

  168. Zieman, S. J., Melenovsky, V. & Kass, D. A. Mechanisms, pathophysiology, and therapy of arterial stiffness. Arterioscler. Thromb. Vasc. Biol. 25, 932–943 (2005).

    Article  Google Scholar 

  169. O’Rourke, M. F. & Nichols, W. W. Aortic diameter, aortic stiffness, and wave reflection increase with age and isolated systolic hypertension. Hypertension 45, 652–658 (2005).

    Article  Google Scholar 

  170. Gossen, M. & Bujard, H. Tight control of gene expression in mammalian cells by tetracycline-responsive promoters. Proc. Natl Acad. Sci. USA 89, 5547–5551 (1992).

    Article  Google Scholar 

  171. Chassin, H. et al. A modular degron library for synthetic circuits in mammalian cells. Nat. Commun. 10, 2013 (2019).

    Article  Google Scholar 

  172. Simsek, H. & Klotzsch, E. The solid tumor microenvironment — breaking the barrier for T cells. BioEssays 44, 2100285 (2022).

    Article  Google Scholar 

  173. Chitty, J. L. et al. A first-in-class pan-lysyl oxidase inhibitor impairs stromal remodeling and enhances gemcitabine response and survival in pancreatic cancer. Nat. Cancer 4, 1326–1344 (2023).

    Article  Google Scholar 

  174. Nakanishi, H. & Kato, Y. Protein-based systems for translational regulation of synthetic mRNAs in mammalian cells. Life 11, 1192 (2021).

    Article  Google Scholar 

  175. Yang, X. et al. Engineering synthetic phosphorylation signaling networks in human cells. Science 387, 74–81 (2025).

    Article  Google Scholar 

  176. Nielsen, A. A. K. et al. Genetic circuit design automation. Science 352, aac7341 (2016).

    Article  Google Scholar 

  177. Jones, T. S., Oliveira, S. M. D., Myers, C. J., Voigt, C. A. & Densmore, D. Genetic circuit design automation with Cello 2.0. Nat. Protoc. 17, 1097–1113 (2022).

    Article  Google Scholar 

  178. Gala, M. & Žoldák, G. Classifying residues in mechanically stable and unstable substructures based on a protein sequence: the case study of the DnaK Hsp70 chaperone. Nanomaterials 11, 2198 (2021).

    Article  Google Scholar 

  179. Asgharzadeh, P. et al. A NanoFE simulation-based surrogate machine learning model to predict mechanical functionality of protein networks from live confocal imaging. Comput. Struct. Biotechnol. J. 18, 2774–2788 (2020).

    Article  Google Scholar 

  180. Kulkarni, J. A. et al. The current landscape of nucleic acid therapeutics. Nat. Nanotechnol. 16, 630–643 (2021).

    Article  Google Scholar 

  181. Procko, C. et al. Mutational analysis of mechanosensitive ion channels in the carnivorous Venus flytrap plant. Curr. Biol. 33, 3257–3264.e4 (2023).

    Article  Google Scholar 

  182. Garamella, J., Majumder, S., Liu, A. P. & Noireaux, V. An adaptive synthetic cell based on mechanosensing, biosensing, and inducible gene circuits. ACS Synth. Biol. 8, 1913–1920 (2019).

    Article  Google Scholar 

  183. Jahnke, K. et al. DNA origami signaling units transduce chemical and mechanical signals in synthetic cells. Adv. Funct. Mater. 34, 2301176 (2024).

    Article  Google Scholar 

Download references

Related Posts