Vincent, T. L. & Wann, A. K. Mechanoadaptation: articular cartilage through thick and thin. J. Physiol. 597, 1271–1281 (2019).
Google Scholar
Chang, S. H. et al. Excessive mechanical loading promotes osteoarthritis through the gremlin-1–NF-κB pathway. Nat. Commun. 10, 1442 (2019).
Google Scholar
Goldring, S. R. & Goldring, M. B. Changes in the osteochondral unit during osteoarthritis: structure, function and cartilage–bone crosstalk. Nat. Rev. Rheumatol. 12, 632–644 (2016).
Google Scholar
Pap, T. & Korb-Pap, A. Cartilage damage in osteoarthritis and rheumatoid arthritis — two unequal siblings. Nat. Rev. Rheumatol. 11, 606–615 (2015).
Google Scholar
Burr, D. B. & Gallant, M. A. Bone remodelling in osteoarthritis. Nat. Rev. Rheumatol. 8, 665–673 (2012).
Google Scholar
Collins, K. H. et al. Adipose tissue is a critical regulator of osteoarthritis. Proc. Natl Acad. Sci. USA 118, e2021096118 (2021).
Google Scholar
Watt, F. E. Posttraumatic osteoarthritis: what have we learned to advance osteoarthritis? Curr. Opin. Rheumatol. 33, 74–83 (2021).
Google Scholar
Greene, M. A. & Loeser, R. F. Aging-related inflammation in osteoarthritis. Osteoarthritis Cartilage 23, 1966–1971 (2015).
Google Scholar
McNulty, M. A. et al. Histopathology of naturally occurring and surgically induced osteoarthritis in mice. Osteoarthritis Cartilage 20, 949–956 (2012).
Google Scholar
Lotz, M. & Loeser, R. F. Effects of aging on articular cartilage homeostasis. Bone 51, 241–248 (2012).
Google Scholar
Loeser, R. F. et al. Microarray analysis reveals age‐related differences in gene expression during the development of osteoarthritis in mice. Arthritis Rheum. 64, 705–717 (2012).
Google Scholar
Thomas, A. C., Hubbard-Turner, T., Wikstrom, E. A. & Palmieri-Smith, R. M. Epidemiology of posttraumatic osteoarthritis. J. Athlet. Train. 52, 491–496 (2017).
Glasson, S. S. et al. Deletion of active ADAMTS5 prevents cartilage degradation in a murine model of osteoarthritis. Nature 434, 644–648 (2005).
Google Scholar
Burleigh, A. et al. Joint immobilization prevents murine osteoarthritis and reveals the highly mechanosensitive nature of protease expression in vivo. Arthritis Rheum. 64, 2278–2288 (2012).
Google Scholar
Ismail, H. M. et al. Interleukin‐1 acts via the JNK‐2 signaling pathway to induce aggrecan degradation by human chondrocytes. Arthritis Rheumatol. 67, 1826–1836 (2015).
Google Scholar
Zhang, M. et al. Induced superficial chondrocyte death reduces catabolic cartilage damage in murine posttraumatic osteoarthritis. J. Clin. Invest. 126, 2893–2902 (2016).
Google Scholar
Gilbert, S. J. & Blain, E. J. in Mechanobiology in Health and Disease (ed. Verbruggen, S. W.) 99–126 (Elsevier, 2018).
Guilak, F., Nims, R. J., Dicks, A., Wu, C.-L. & Meulenbelt, I. Osteoarthritis as a disease of the cartilage pericellular matrix. Matrix Biol. 71–72, 40–50 (2018).
Google Scholar
Vincent, T. L. Targeting mechanotransduction pathways in osteoarthritis: a focus on the pericellular matrix. Curr. Opin. Pharmacol. 13, 449–454 (2013).
Google Scholar
Agarwal, P. et al. A dysfunctional TRPV4–GSK3β pathway prevents osteoarthritic chondrocytes from sensing changes in extracellular matrix viscoelasticity. Nat. Biomed. Eng. 5, 1472–1484 (2021).
Google Scholar
Nims, R. J. et al. A synthetic mechanogenetic gene circuit for autonomous drug delivery in engineered tissues. Sci. Adv. 7, eabd9858 (2021).
Google Scholar
Deng, Y. et al. Reciprocal inhibition of YAP/TAZ and NF-κB regulates osteoarthritic cartilage degradation. Nat. Commun. 9, 4564 (2018).
Google Scholar
Eckstein, F. et al. Intra-articular sprifermin reduces cartilage loss in addition to increasing cartilage gain independent of location in the femorotibial joint: post-hoc analysis of a randomised, placebo-controlled phase II clinical trial. Ann. Rheum. Dis. 79, 525–528 (2020).
Google Scholar
Peredo, A. P. et al. Mechano-activated biomolecule release in regenerating load-bearing tissue microenvironments. Biomaterials 265, 120255 (2021).
Google Scholar
Nims, R. J., Pferdehirt, L. & Guilak, F. Mechanogenetics: harnessing mechanobiology for cellular engineering. Curr. Opin. Biotechnol. 73, 374–379 (2022).
Google Scholar
Poole, A. R. et al. Composition and structure of articular cartilage: a template for tissue repair. Clin. Orthop. Relat. Res. 391, S26–S33 (2001).
Mow, V. C., Ratcliffe, A. & Poole, A. R. Cartilage and diarthrodial joints as paradigms for hierarchical materials and structures. Biomaterials 13, 67–97 (1992).
Google Scholar
Schätti, O. R., Marková, M., Torzilli, P. A. & Gallo, L. M. Mechanical loading of cartilage explants with compression and sliding motion modulates gene expression of lubricin and catabolic enzymes. Cartilage 6, 185–193 (2015).
Google Scholar
Melrose, J., Hayes, A. J., Whitelock, J. M. & Little, C. B. Perlecan, the “jack of all trades” proteoglycan of cartilaginous weight‐bearing connective tissues. Bioessays 30, 457–469 (2008).
Google Scholar
Poole, C. A. Articular cartilage chondrons: form, function and failure. J. Anat. 191, 1–13 (1997).
Google Scholar
Schinagl, R. M., Gurskis, D., Chen, A. C. & Sah, R. L. Depth‐dependent confined compression modulus of full‐thickness bovine articular cartilage. J. Ortho Res. 15, 499–506 (1997).
Google Scholar
Xia, Y., Moody, J. B., Alhadlaq, H. & Hu, J. Imaging the physical and morphological properties of a multi‐zone young articular cartilage at microscopic resolution. J. Mag. Reson. Imaging 17, 365–374 (2003).
Ratcliffe, A., Fryer, P. R. & Hardingham, T. E. The distribution of aggregating proteoglycans in articular cartilage: comparison of quantitative immunoelectron microscopy with radioimmunoassay and biochemical analysis. J. Histochem. Cytochem. 32, 193–201 (1984).
Google Scholar
Maroudas, A., Muir, H. & Wingham, J. The correlation of fixed negative charge with glycosaminoglycan content of human articular cartilage. Biochim. Biophys. Acta 177, 492–500 (1969).
Google Scholar
Wilusz, R. E., Zauscher, S. & Guilak, F. Micromechanical mapping of early osteoarthritic changes in the pericellular matrix of human articular cartilage. Osteoarthritis Cartilage 21, 1895–1903 (2013).
Google Scholar
Chery, D. R. et al. Early changes in cartilage pericellular matrix micromechanobiology portend the onset of post-traumatic osteoarthritis. Acta Biomater. 111, 267–278 (2020).
Google Scholar
Simon, W. H. Scale effects in animal joints. I. Articular cartilage thickness and compressive stress. Arthritis Rheum. 13, 244–255 (1970).
Google Scholar
Loeser, R. F. Integrins and cell signaling in chondrocytes. Biorheology 39, 119–124 (2002).
Google Scholar
Millward-Sadler, S. J. & Salter, D. M. Integrin-dependent signal cascades in chondrocyte mechanotransduction. Ann. Biomed. Eng. 32, 435–446 (2004).
Google Scholar
Ross, T. D. et al. Integrins in mechanotransduction. Curr. Opin. Cell Biol. 25, 613–618 (2013).
Google Scholar
Blain, E. J. Involvement of the cytoskeletal elements in articular cartilage homeostasis and pathology. Int. J. Exp. Pathol. 90, 1–15 (2009).
Google Scholar
Barrett-Jolley, R., Lewis, R., Fallman, R. & Mobasheri, A. The emerging chondrocyte channelome. Front. Physiol. 1, 135 (2010).
Google Scholar
Matta, C., Zákány, R. & Mobasheri, A. Voltage-dependent calcium channels in chondrocytes: roles in health and disease. Curr. Rheumatol. Rep. 17, 43 (2015).
Google Scholar
Mobasheri, A. et al. The chondrocyte channelome: a narrative review. Jt. Bone Spine 86, 29–35 (2019).
Google Scholar
Ruhlen, R. & Marberry, K. The chondrocyte primary cilium. Osteoarthritis Cartilage 22, 1071–1076 (2014).
Google Scholar
Tao, F., Jiang, T., Tao, H., Cao, H. & Xiang, W. Primary cilia: versatile regulator in cartilage development. Cell Prolif. 53, e12765 (2020).
Google Scholar
Guilak, F. et al. The pericellular matrix as a transducer of biomechanical and biochemical signals in articular cartilage. Ann. N. Y. Acad. Sci. 1068, 498–512 (2006).
Google Scholar
Youn, I., Choi, J., Cao, L., Setton, L. & Guilak, F. Zonal variations in the three-dimensional morphology of the chondron measured in situ using confocal microscopy. Osteoarthritis Cartilage 14, 889–897 (2006).
Google Scholar
Martin, J., Miller, B., Scherb, M., Lembke, L. & Buckwalter, J. Co-localization of insulin-like growth factor binding protein 3 and fibronectin in human articular cartilage. Osteoarthritis Cartilage 10, 556–563 (2002).
Google Scholar
Vincent, T., Hermansson, M., Bolton, M., Wait, R. & Saklatvala, J. Basic FGF mediates an immediate response of articular cartilage to mechanical injury. Proc. Natl Acad. Sci. USA 99, 8259–8264 (2002).
Google Scholar
Vincent, T. L., Hermansson, M. A., Hansen, U. N., Amis, A. A. & Saklatvala, J. Basic fibroblast growth factor mediates transduction of mechanical signals when articular cartilage is loaded. Arthritis Rheum. 50, 526–533 (2004).
Google Scholar
Vincent, T. L., McLean, C. J., Full, L. E., Peston, D. & Saklatvala, J. FGF-2 is bound to perlecan in the pericellular matrix of articular cartilage, where it acts as a chondrocyte mechanotransducer. Osteoarthritis Cartilage 15, 752–763 (2007).
Google Scholar
Vincent, T. L. Fibroblast growth factor 2: good or bad guy in the joint? Arthritis Res. Ther. 13, 127 (2011).
Google Scholar
Xie, Y., Zinkle, A., Chen, L. & Mohammadi, M. Fibroblast growth factor signalling in osteoarthritis and cartilage repair. Nat. Rev. Rheumatol. 16, 547–564 (2020).
Google Scholar
Makarenkova, H. P. et al. Differential interactions of FGFs with heparan sulfate control gradient formation and branching morphogenesis. Sci. Signal. 2, ra55 (2009).
Google Scholar
Eckstein, F., Wirth, W., Guermazi, A., Maschek, S. & Aydemir, A. Brief report: intraarticular sprifermin not only increases cartilage thickness, but also reduces cartilage loss: location‐independent post hoc analysis using magnetic resonance imaging. Arthritis Rheumatol. 67, 2916–2922 (2015).
Google Scholar
Lohmander, L. S. et al. Intraarticular sprifermin (recombinant human fibroblast growth factor 18) in knee osteoarthritis: a randomized, double‐blind, placebo‐controlled trial. Arthritis Rheumatol. 66, 1820–1831 (2014).
Google Scholar
Hochberg, M. C. et al. Effect of intra-articular sprifermin vs placebo on femorotibial joint cartilage thickness in patients with osteoarthritis: the FORWARD randomized clinical trial. JAMA 322, 1360–1370 (2019).
Google Scholar
Zhen, G. et al. Mechanical stress determines the configuration of TGFβ activation in articular cartilage. Nat. Commun. 12, 1706 (2021).
Google Scholar
Kechagia, J. Z., Ivaska, J. & Roca-Cusachs, P. Integrins as biomechanical sensors of the microenvironment. Nat. Rev. Mol. Cell Biol. 20, 457–473 (2019).
Google Scholar
Seetharaman, S. & Etienne‐Manneville, S. Integrin diversity brings specificity in mechanotransduction. Biol. Cell 110, 49–64 (2018).
Google Scholar
Cantini, M., Donnelly, H., Dalby, M. J. & Salmeron‐Sanchez, M. The plot thickens: the emerging role of matrix viscosity in cell mechanotransduction. Adv. Healthc. Mater. 9, 1901259 (2020).
Google Scholar
Parsons, J. T., Horwitz, A. R. & Schwartz, M. A. Cell adhesion: integrating cytoskeletal dynamics and cellular tension. Nat. Rev. Mol. Cell Biol. 11, 633–643 (2010).
Google Scholar
Puklin-Faucher, E. & Sheetz, M. P. The mechanical integrin cycle. J. Cell Sci. 122, 179–186 (2009).
Google Scholar
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).
Google Scholar
Elosegui-Artola, A. et al. Rigidity sensing and adaptation through regulation of integrin types. Nat. Mater. 13, 631–637 (2014).
Google Scholar
Franz, F., Daday, C. & Gräter, F. Advances in molecular simulations of protein mechanical properties and function. Curr. Opin. Struct. Biol. 61, 132–138 (2020).
Google Scholar
Gouttenoire, J. et al. BMP-2 and TGF-β1 differentially control expression of type II procollagen and α10 and α11 integrins in mouse chondrocytes. Eur. J. Cell Biol. 89, 307–314 (2010).
Google Scholar
Salter, D., Hughes, D., Simpson, R. & Gardner, D. Integrin expression by human articular chondrocytes. Rheumatology 31, 231–234 (1992).
Google Scholar
Zhang, W.-M. et al. Analysis of the human integrin α11 gene (ITGA11) and its promoter. Matrix Biol. 21, 513–523 (2002).
Google Scholar
Loeser, R. F. Integrins and chondrocyte–matrix interactions in articular cartilage. Matrix Biol. 39, 11–16 (2014).
Google Scholar
Orazizadeh, M. et al. CD47 associates with alpha 5 integrin and regulates responses of human articular chondrocytes to mechanical stimulation in an in vitro model. Arthritis Res. Ther. 10, R4 (2008).
Google Scholar
Ostergaard, K. et al. Expression of α and β subunits of the integrin superfamily in articular cartilage from macroscopically normal and osteoarthritic human femoral heads. Ann. Rheum. Dis. 57, 303–308 (1998).
Google Scholar
Lucchinetti, E., Bhargava, M. M. & Torzilli, P. A. The effect of mechanical load on integrin subunits α5 and β1 in chondrocytes from mature and immature cartilage explants. Cell Tissue Res. 315, 385–391 (2004).
Google Scholar
Millward-Sadler, S. et al. Integrin-regulated secretion of interleukin 4: a novel pathway of mechanotransduction in human articular chondrocytes. J. Cell Biol. 145, 183–189 (1999).
Google Scholar
Millward‐Sadler, S., Wright, M., Davies, L., Nuki, G. & Salter, D. Mechanotransduction via integrins and interleukin‐4 results in altered aggrecan and matrix metalloproteinase 3 gene expression in normal, but not osteoarthritic, human articular chondrocytes. Arthritis Rheum. 43, 2091–2099 (2000).
Google Scholar
Steward, A. et al. Cell–matrix interactions regulate mesenchymal stem cell response to hydrostatic pressure. Acta Biomater. 8, 2153–2159 (2012).
Google Scholar
Jablonski, C. L., Ferguson, S., Pozzi, A. & Clark, A. L. Integrin α1β1 participates in chondrocyte transduction of osmotic stress. Biochem. Biophys. Res. Commun. 445, 184–190 (2014).
Google Scholar
Wright, M. et al. Hyperpolarisation of cultured human chondrocytes following cyclical pressure‐induced strain: evidence of a role for α5β1 integrin as a chondrocyte mechanoreceptor. J. Ortho Res. 15, 742–747 (1997).
Google Scholar
Camper, L., Hellman, U. & Lundgren-Åkerlund, E. Isolation, cloning, and sequence analysis of the integrin subunit α10, a β1-associated collagen binding integrin expressed on chondrocytes. J. Biol. Chem. 273, 20383–20389 (1998).
Google Scholar
Bengtsson, T., Camper, L., Schneller, M. & Lundgren-Åkerlund, E. Characterization of the mouse integrin subunit α10 gene and comparison with its human homologue: genomic structure, chromosomal localization and identification of splice variants. Matrix Biol. 20, 565–576 (2001).
Google Scholar
Lehnert, K. et al. Cloning, sequence analysis, and chromosomal localization of the novel human integrin α11 subunit (ITGA11). Genomics 60, 179–187 (1999).
Google Scholar
Varas, L. et al. α10 integrin expression is up-regulated on fibroblast growth factor-2-treated mesenchymal stem cells with improved chondrogenic differentiation potential. Stem Cell Dev. 16, 965–978 (2007).
Google Scholar
Delco, M. L. et al. Integrin α10β1-selected mesenchymal stem cells mitigate the progression of osteoarthritis in an equine talar impact model. Am. J. Sports Med. 48, 612–623 (2020).
Google Scholar
Hirose, N. et al. Protective effects of cilengitide on inflammation in chondrocytes under excessive mechanical stress. Cell Biol. Int. 44, 966–974 (2020).
Google Scholar
Chao, P. H., West, A. C. & Hung, C. T. Chondrocyte intracellular calcium, cytoskeletal organization, and gene expression responses to dynamic osmotic loading. Am. J. Physiol. Cell Physiol. 291, 718–725 (2006).
Erickson, G. R., Northrup, D. L. & Guilak, F. Hypo-osmotic stress induces calcium-dependent actin reorganization in articular chondrocytes. Osteoarthritis Cartilage 11, 187–197 (2003).
Google Scholar
Grodzinsky, A. J., Levenston, M. E., Jin, M. & Frank, E. H. Cartilage tissue remodeling in response to mechanical forces. Annu. Rev. Biomed. Eng. 2, 691–713 (2000).
Google Scholar
Guilak, F. Compression-induced changes in the shape and volume of the chondrocyte nucleus. J. Biomech. 28, 1529–1541 (1995).
Google Scholar
Blain, E. J., Mason, D. J. & Duance, V. C. The effect of thymosin β4 on articular cartilage chondrocyte matrix metalloproteinase expression. Biochem. Soc. Trans. 30, 879–882 (2002).
Google Scholar
Fioravanti, A., Nerucci, F., Annefeld, M., Collodel, G. & Marcolongo, R. Morphological and cytoskeletal aspects of cultivated normal and osteoarthritic human articular chondrocytes after cyclical pressure: a pilot study. Clin. Exp. Rheumatol. 21, 739–746 (2003).
Google Scholar
Fioravanti, A., Benetti, D., Coppola, G. & Collodel, G. Effect of continuous high hydrostatic pressure on the morphology and cytoskeleton of normal and osteoarthritic human chondrocytes cultivated in alginate gels. Clin. Exp. Rheumatol. 23, 847–853 (2005).
Google Scholar
Isermann, P. & Lammerding, J. Nuclear mechanics and mechanotransduction in health and disease. Curr. Biol. 23, R1113–R1121 (2013).
Google Scholar
Khilan, A. A., Al-Maslamani, N. A. & Horn, H. F. Cell stretchers and the LINC complex in mechanotransduction. Arch. Biochem. Biophys. 702, 108829 (2021).
Google Scholar
Lee, D. A. et al. Chondrocyte deformation within compressed agarose constructs at the cellular and sub-cellular levels. J. Biomech. 33, 81–95 (2000).
Google Scholar
Irianto, J. et al. Osmotic challenge drives rapid and reversible chromatin condensation in chondrocytes. Biophys. J. 104, 759–769 (2013).
Google Scholar
Hopewell, B. & Urban, J. P. Adaptation of articular chondrocytes to changes in osmolality. Biorheology 40, 73–77 (2003).
Google Scholar
Hung, C. T. et al. Disparate aggrecan gene expression in chondrocytes subjected to hypotonic and hypertonic loading in 2D and 3D culture. Biorheology 40, 61–72 (2003).
Google Scholar
Killaars, A. R., Walker, C. J. & Anseth, K. S. Nuclear mechanosensing controls MSC osteogenic potential through HDAC epigenetic remodeling. Proc. Natl Acad. Sci. USA 117, 21258 (2020).
Google Scholar
Elosegui-Artola, A. et al. Force triggers YAP nuclear entry by regulating transport across nuclear pores. Cell 171, 1397–1410 (2017).
Google Scholar
Delco, M. L. & Bonassar, L. J. Targeting calcium-related mechanotransduction in early OA. Nat. Rev. Rheumatol. 17, 445–446 (2021).
Google Scholar
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).
Google Scholar
Phan, M. N. et al. Functional characterization of TRPV4 as an osmotically sensitive ion channel in porcine articular chondrocytes. Arthritis Rheum. 60, 3028–3037 (2009).
Google Scholar
Clark, A. L., Votta, B. J., Kumar, S., Liedtke, W. & Guilak, F. Chondroprotective role of the osmotically sensitive ion channel transient receptor potential vanilloid 4: age- and sex-dependent progression of osteoarthritis in Trpv4-deficient mice. Arthritis Rheum. 62, 2973–2983 (2010).
Google Scholar
O’Conor, C. J. et al. Cartilage-specific knockout of the mechanosensory ion channel TRPV4 decreases age-related osteoarthritis. Sci. Rep. 6, 29053 (2016).
Google Scholar
Drexler, S. K., Wann, A. K. T. & Vincent, T. L. Are cellular mechanosensors potential therapeutic targets in osteoarthritis. Int. J. Clin. Rheumatol. 9, 155–167 (2014).
Google Scholar
Lee, W., Guilak, F. & Liedtke, W. Role of Piezo channels in joint health and injury. Curr. Top. Membr. 79, 263–273 (2017).
Google Scholar
Sun, Y. et al. Mechanism of abnormal chondrocyte proliferation induced by Piezo1-siRNA exposed to mechanical stretch. BioMed. Res. Int. 2020, 8538463 (2020).
Google Scholar
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).
Google Scholar
Gnanasambandam, R. et al. GsMTx4: mechanism of inhibiting mechanosensitive ion channels. Biophys. J. 112, 31–45 (2017).
Google Scholar
Suchyna, T. M. Piezo channels and GsMTx4: two milestones in our understanding of excitatory mechanosensitive channels and their role in pathology. Prog. Biophys. Mol. Biol. 130, 244–253 (2017).
Google Scholar
Xiao, W. F., Li, Y. S., Deng, A., Yang, Y. T. & He, M. Functional role of hedgehog pathway in osteoarthritis. Cell Biochem. Funct. 38, 122–129 (2020).
Google Scholar
McGlashan, S. R., Cluett, E. C., Jensen, C. G. & Poole, C. A. Primary cilia in osteoarthritic chondrocytes: from chondrons to clusters. Dev. Dyn. 237, 2013–2020 (2008).
Google Scholar
McGlashan, S. R., Jensen, C. G. & Poole, C. A. Localization of extracellular matrix receptors on the chondrocyte primary cilium. J. Histochem. Cytochem. 54, 1005–1014 (2006).
Google Scholar
Chang, C. F., Ramaswamy, G. & Serra, R. Depletion of primary cilia in articular chondrocytes results in reduced Gli3 repressor to activator ratio, increased Hedgehog signaling, and symptoms of early osteoarthritis. Osteoarthritis Cartilage 20, 152–161 (2012).
Google Scholar
Irianto, J., Ramaswamy, G., Serra, R. & Knight, M. M. Depletion of chondrocyte primary cilia reduces the compressive modulus of articular cartilage. J. Biomech. 47, 579–582 (2014).
Google Scholar
Wann, A. K. T. et al. Primary cilia mediate mechanotransduction through control of ATP-induced Ca2+ signaling in compressed chondrocytes. FASEB J. 26, 1663–1671 (2012).
Google Scholar
Shao, Y. Y., Wang, L., Welter, J. F. & Ballock, R. T. Primary cilia modulate Ihh signal transduction in response to hydrostatic loading of growth plate chondrocytes. Bone 50, 79–84 (2012).
Google Scholar
Pingguan‐Murphy, B., El‐Azzeh, M., Bader, D. & Knight, M. Cyclic compression of chondrocytes modulates a purinergic calcium signalling pathway in a strain rate‐and frequency‐dependent manner. J. Cell Physiol. 209, 389–397 (2006).
Google Scholar
Zhang, J. et al. Connexin43 hemichannels mediate small molecule exchange between chondrocytes and matrix in biomechanically-stimulated temporomandibular joint cartilage. Osteoarthritis Cartilage 22, 822–830 (2014).
Google Scholar
Garcia, M. & Knight, M. M. Cyclic loading opens hemichannels to release ATP as part of a chondrocyte mechanotransduction pathway. J. Orthop. Res. 28, 510–515 (2010).
Google Scholar
Chowdhury, T. & Knight, M. Purinergic pathway suppresses the release of NO and stimulates proteoglycan synthesis in chondrocyte/agarose constructs subjected to dynamic compression. J. Cell Physiol. 209, 845–853 (2006).
Google Scholar
Huang, C., Holfeld, J., Schaden, W., Orgill, D. & Ogawa, R. Mechanotherapy: revisiting physical therapy and recruiting mechanobiology for a new era in medicine. Trends Mol. Med. 19, 555–564 (2013).
Google Scholar
Thompson, W. R., Scott, A., Loghmani, M. T., Ward, S. R. & Warden, S. J. Understanding mechanobiology: physical therapists as a force in mechanotherapy and musculoskeletal regenerative rehabilitation. Phys. Ther. 96, 560–569 (2016).
Google Scholar
Dell’Accio, F., De Bari, C., Eltawil, N. M., Vanhummelen, P. & Pitzalis, C. Identification of the molecular response of articular cartilage to injury, by microarray screening: Wnt-16 expression and signaling after injury and in osteoarthritis. Arthritis Rheum. 58, 1410–1421 (2008).
Google Scholar
Loeser, R. F., Erickson, E. A. & Long, D. L. Mitogen-activated protein kinases as therapeutic targets in osteoarthritis. Curr. Opin. Rheumatol. 20, 581–586 (2008).
Google Scholar
Fanning, P. J. et al. Mechanical regulation of mitogen-activated protein kinase signaling in articular cartilage. J. Biol. Chem. 278, 50940–50948 (2003).
Google Scholar
Forsyth, C. B., Pulai, J. & Loeser, R. F. Fibronectin fragments and blocking antibodies to α2β1 and α5β1 integrins stimulate mitogen‐activated protein kinase signaling and increase collagenase 3 (matrix metalloproteinase 13) production by human articular chondrocytes. Arthritis Rheum. 46, 2368–2376 (2002).
Google Scholar
Im, H.-J. et al. Inhibitory effects of insulin-like growth factor-1 and osteogenic protein-1 on fibronectin fragment- and interleukin-1β-stimulated matrix metalloproteinase-13 expression in human chondrocytes. J. Biol. Chem. 278, 25386–25394 (2003).
Google Scholar
Loeser, R. F., Forsyth, C. B., Samarel, A. M. & Im, H.-J. Fibronectin fragment activation of proline-rich tyrosine kinase PYK2 mediates integrin signals regulating collagenase-3 expression by human chondrocytes through a protein kinase C-dependent pathway. J. Biol. Chem. 278, 24577–24585 (2003).
Google Scholar
Pulai, J. I. et al. NF-κB mediates the stimulation of cytokine and chemokine expression by human articular chondrocytes in response to fibronectin fragments. J. Immunol. 174, 5781–5788 (2005).
Google Scholar
Del Carlo, M., Schwartz, D., Erickson, E. A. & Loeser, R. F. Endogenous production of reactive oxygen species is required for stimulation of human articular chondrocyte matrix metalloproteinase production by fibronectin fragments. Free. Radic. Biol. Med. 42, 1350–1358 (2007).
Google Scholar
Long, D. L., Willey, J. S. & Loeser, R. F. Rac1 is required for matrix metalloproteinase 13 production by chondrocytes in response to fibronectin fragments. Arthritis Rheum. 65, 1561–1568 (2013).
Google Scholar
Gemba, T., Valbracht, J., Alsalameh, S. & Lotz, M. Focal adhesion kinase and mitogen-activated protein kinases are involved in chondrocyte activation by the 29-kDa amino-terminal fibronectin fragment. J. Biol. Chem. 277, 907–911 (2002).
Google Scholar
Ding, L., Guo, D. & Homandberg, G. The cartilage chondrolytic mechanism of fibronectin fragments involves MAP kinases: comparison of three fragments and native fibronectin. Osteoarthritis Cartilage 16, 1253–1262 (2008).
Google Scholar
Ding, L., Guo, D. & Homandberg, G. Fibronectin fragments mediate matrix metalloproteinase upregulation and cartilage damage through proline rich tyrosine kinase 2, c-src, NF-κB and protein kinase Cδ. Osteoarthritis Cartilage 17, 1385–1392 (2009).
Google Scholar
Fitzgerald, J. B. et al. Shear- and compression-induced chondrocyte transcription requires MAPK activation in cartilage explants. J. Biol. Chem. 283, 6735–6743 (2008).
Google Scholar
Zhang, J., Shen, B. & Lin, A. Novel strategies for inhibition of the p38 MAPK pathway. Trends Pharmacol. Sci. 28, 286–295 (2007).
Google Scholar
González-Vázquez, A. et al. Accelerating bone healing in vivo by harnessing the age-altered activation of c-Jun N-terminal kinase 3. Biomaterials 268, 120540 (2021).
Google Scholar
Agarwal, S. et al. A central role for the nuclear factor-κB pathway in anti-inflammatory and proinflammatory actions of mechanical strain. FASEB J. 17, 899–901 (2003).
Google Scholar
Yang, Y. et al. Mechanical stress protects against osteoarthritis via regulation of the AMPK/NF-κB signaling pathway. J. Cell Physiol. 234, 9156–9167 (2019).
Google Scholar
Vincent, T. L. Mechanoflammation in osteoarthritis pathogenesis. Semin. Arthritis Rheum. 49, S36–S38 (2019).
Google Scholar
Ismail, H. M., Didangelos, A., Vincent, T. L. & Saklatvala, J. Rapid activation of transforming growth factor β-activated kinase 1 in chondrocytes by phosphorylation and K(63)-linked polyubiquitination upon injury to animal articular cartilage. Arthritis Rheumatol. 69, 565–575 (2017).
Google Scholar
Lee, W. et al. Inflammatory signaling sensitizes Piezo1 mechanotransduction in articular chondrocytes as a pathogenic feed-forward mechanism in osteoarthritis. Proc. Natl Acad. Sci. USA 118, e2001611118 (2021).
Google Scholar
Nam, S. et al. Cell cycle progression in confining microenvironments is regulated by a growth-responsive TRPV4-PI3K/Akt-p27(Kip1) signaling axis. Sci. Adv. 5, eaaw6171 (2019).
Google Scholar
Lee, H. P., Gu, L., Mooney, D. J., Levenston, M. E. & Chaudhuri, O. Mechanical confinement regulates cartilage matrix formation by chondrocytes. Nat. Mater. 16, 1243–1251 (2017).
Google Scholar
Miller, J. R. The Wnts. Genome Biol. 3, reviews3001.1 (2001).
Blom, A. B. et al. Involvement of the Wnt signaling pathway in experimental and human osteoarthritis: prominent role of Wnt‐induced signaling protein 1. Arthritis Rheum. 60, 501–512 (2009).
Google Scholar
De Santis, M. et al. The role of Wnt pathway in the pathogenesis of OA and its potential therapeutic implications in the field of regenerative medicine. BioMed. Res. Int. 2018, 7402947 (2018).
Google Scholar
Dell’Accio, F. et al. Activation of WNT and BMP signaling in adult human articular cartilage following mechanical injury. Arthritis Res. Ther. 8, R139 (2006).
Google Scholar
Bougault, C. et al. Protective role of frizzled-related protein B on matrix metalloproteinase induction in mouse chondrocytes. Arthritis Res. Ther. 16, R137 (2014).
Google Scholar
Nalesso, G. et al. WNT16 antagonises excessive canonical WNT activation and protects cartilage in osteoarthritis. Ann. Rheum. Dis. 76, 218–226 (2017).
Google Scholar
Wang, Y., Fan, X., Xing, L. & Tian, F. Wnt signaling: a promising target for osteoarthritis therapy. Cell Commun. Signal. 17, 97 (2019).
Google Scholar
Lories, R. J. & Monteagudo, S. Review article: is Wnt signaling an attractive target for the treatment of osteoarthritis? Rheumatol. Ther. 7, 259–270 (2020).
Google Scholar
Monteagudo, S. & Lories, R. J. Cushioning the cartilage: a canonical Wnt restricting matter. Nat. Rev. Rheumatol. 13, 670–681 (2017).
Google Scholar
Deshmukh, V. et al. A small-molecule inhibitor of the Wnt pathway (SM04690) as a potential disease modifying agent for the treatment of osteoarthritis of the knee. Osteoarthritis Cartilage 26, 18–27 (2018).
Google Scholar
Yazici, Y. et al. A phase 2b randomized trial of lorecivivint, a novel intra-articular CLK2/DYRK1A inhibitor and Wnt pathway modulator for knee osteoarthritis. Osteoarthritis Cartilage 29, 654–666 (2021).
Google Scholar
Deshmukh, V. et al. Modulation of the Wnt pathway through inhibition of CLK2 and DYRK1A by lorecivivint as a novel, potentially disease-modifying approach for knee osteoarthritis treatment. Osteoarthritis Cartilage 27, 1347–1360 (2019).
Google Scholar
Monteagudo, S. et al. DOT1L safeguards cartilage homeostasis and protects against osteoarthritis. Nat. Commun. 8, 15889 (2017).
Google Scholar
Cornelis, F. M. F. et al. Increased susceptibility to develop spontaneous and post-traumatic osteoarthritis in Dot1l-deficient mice. Osteoarthritis Cartilage 27, 513–525 (2019).
Google Scholar
Castaño Betancourt, M. C. et al. Genome-wide association and functional studies identify the DOT1L gene to be involved in cartilage thickness and hip osteoarthritis. Proc. Natl Acad. Sci. USA 109, 8218–8223 (2012).
Google Scholar
Deng, Y. et al. Yap1 regulates multiple steps of chondrocyte differentiation during skeletal development and bone repair. Cell Rep. 14, 2224–2237 (2016).
Google Scholar
Gumbiner, B. M. & Kim, N.-G. The Hippo-YAP signaling pathway and contact inhibition of growth. J. Cell Sci. 127, 709–717 (2014).
Google Scholar
Dupont, S. et al. Role of YAP/TAZ in mechanotransduction. Nature 474, 179–183 (2011).
Google Scholar
Baker, B. M. & Chen, C. S. Deconstructing the third dimension–how 3D culture microenvironments alter cellular cues. J. Cell Sci. 125, 3015–3024 (2012).
Google Scholar
Caliari, S. R., Vega, S. L., Kwon, M., Soulas, E. M. & Burdick, J. A. Dimensionality and spreading influence MSC YAP/TAZ signaling in hydrogel environments. Biomaterials 103, 314–323 (2016).
Google Scholar
Karystinou, A. et al. Yes-associated protein (YAP) is a negative regulator of chondrogenesis in mesenchymal stem cells. Arthritis Res. Ther. 17, 147 (2015).
Google Scholar
Mobasheri, A. et al. The role of metabolism in the pathogenesis of osteoarthritis. Nat. Rev. Rheumatol. 13, 302–311 (2017).
Google Scholar
Salinas, D., Mumey, B. M. & June, R. K. Physiological dynamic compression regulates central energy metabolism in primary human chondrocytes. Biomech. Model. Mechanobiol. 18, 69–77 (2019).
Google Scholar
Lehtinen, M. K. et al. A conserved MST-FOXO signaling pathway mediates oxidative-stress responses and extends life span. Cell 125, 987–1001 (2006).
Google Scholar
Niehoff, A. et al. Dynamic and static mechanical compression affects Akt phosphorylation in porcine patellofemoral joint cartilage. J. Orthop. Res. 26, 616–623 (2008).
Google Scholar
Holledge, M. M., Millward-Sadler, S. J., Nuki, G. & Salter, D. M. Mechanical regulation of proteoglycan synthesis in normal and osteoarthritic human articular chondrocytes–roles for α5 and αVβ5 integrins. Biorheology 45, 275–288 (2008).
Google Scholar
Delco, M. L., Bonnevie, E. D., Bonassar, L. J. & Fortier, L. A. Mitochondrial dysfunction is an acute response of articular chondrocytes to mechanical injury. J. Orthop. Res. 36, 739–750 (2018).
Google Scholar
Waller, K. A., Zhang, L. X. & Jay, G. D. Friction-induced mitochondrial dysregulation contributes to joint deterioration in Prg4 knockout mice. Int. J. Mol. Sci. 18, 1252 (2017).
Google Scholar
Bartell, L. R. et al. Mitoprotective therapy prevents rapid, strain-dependent mitochondrial dysfunction after articular cartilage injury. J. Orthop. Res. 38, 1257–1267 (2020).
Google Scholar
Jutila, A. A. et al. Candidate mediators of chondrocyte mechanotransduction via targeted and untargeted metabolomic measurements. Arch. Biochem. Biophys. 545, 116–123 (2014).
Google Scholar
Zignego, D. L., Jutila, A. A., Gelbke, M. K., Gannon, D. M. & June, R. K. The mechanical microenvironment of high concentration agarose for applying deformation to primary chondrocytes. J. Biomech. 47, 2143–2148 (2014).
Google Scholar
Hodgkinson, T. et al. The use of nanovibration to discover specific and potent bioactive metabolites that stimulate osteogenic differentiation in mesenchymal stem cells. Sci. Adv. 7, eabb7921 (2021).
Google Scholar
Bonnevie, E. D. et al. Microscale frictional strains determine chondrocyte fate in loaded cartilage. J. Biomech. 74, 72–78 (2018).
Google Scholar
Irwin, R. M. et al. Distinct tribological endotypes of pathological human synovial fluid reveal characteristic biomarkers and variation in efficacy of viscosupplementation at reducing local strains in articular cartilage. Osteoarthritis Cartilage 28, 492–501 (2020).
Google Scholar
Xie, R. et al. Biomimetic cartilage-lubricating polymers regenerate cartilage in rats with early osteoarthritis. Nat. Biomed. Eng. 5, 1189–1201 (2021).
Google Scholar
Grither, W. R. & Longmore, G. D. Inhibition of tumor-microenvironment interaction and tumor invasion by small-molecule allosteric inhibitor of DDR2 extracellular domain. Proc. Natl Acad. Sci. USA 115, E7786–E7794 (2018).
Google Scholar
Kumar, A., Choudhury, M. D., Ghosh, P. & Palit, P. Discoidin domain receptor 2: an emerging pharmacological drug target for prospective therapy against osteoarthritis. Pharmacol. Rep. 71, 399–408 (2019).
Google Scholar
Occhetta, P. et al. Hyperphysiological compression of articular cartilage induces an osteoarthritic phenotype in a cartilage-on-a-chip model. Nat. Biomed. Eng. 3, 545–557 (2019).
Google Scholar
Lee, J. et al. Combinatorial screening of biochemical and physical signals for phenotypic regulation of stem cell-based cartilage tissue engineering. Sci. Adv. 6, eaaz5913 (2020).
Google Scholar
Wang, J., Lü, D., Mao, D. & Long, M. Mechanomics: an emerging field between biology and biomechanics. Protein Cell 5, 518–531 (2014).
Google Scholar
Gabriel, S. E., Crowson, C. S. & O’Fallon, W. M. Comorbidity in arthritis. J. Rheumatol. 26, 2475–2479 (1999).
Google Scholar
Shi, S., Man, Z., Li, W., Sun, S. & Zhang, W. Silencing of Wnt5a prevents interleukin-1β-induced collagen type II degradation in rat chondrocytes. Exp. Ther. Med. 12, 3161–3166 (2016).
Google Scholar
Yan, H. et al. Suppression of NF-κB activity via nanoparticle-based siRNA delivery alters early cartilage responses to injury. Proc. Natl Acad. Sci. USA 113, E6199–E6208 (2016).
Google Scholar
Rai, M. F. et al. Applications of RNA interference in the treatment of arthritis. Transl. Res. 214, 1–16 (2019).
Google Scholar
Cheleschi, S. et al. Hydrostatic pressure regulates microRNA expression levels in osteoarthritic chondrocyte cultures via the Wnt/β-catenin pathway. Int. J. Mol. Sci. 18, 133 (2017).
Google Scholar
De Palma, A. et al. Hydrostatic pressure as epigenetic modulator in chondrocyte cultures: a study on miRNA-155, miRNA-181a and miRNA-223 expression levels. J. Biomech. 66, 165–169 (2018).
Google Scholar
Yang, X. et al. Mechanical and IL-1β responsive miR-365 contributes to osteoarthritis development by targeting histone deacetylase 4. Int. J. Mol. Sci. 17, 436 (2016).
Google Scholar
Stadnik, P. S. et al. Regulation of microRNA-221, -222, -21 and -27 in articular cartilage subjected to abnormal compressive forces. J. Physiol. 599, 143–155 (2021).
Google Scholar
Dunn, W., DuRaine, G. & Reddi, A. H. Profiling microRNA expression in bovine articular cartilage and implications for mechanotransduction. Arthritis Rheum. 60, 2333–2339 (2009).
Google Scholar
Iliopoulos, D., Malizos, K. N., Oikonomou, P. & Tsezou, A. Integrative microRNA and proteomic approaches identify novel osteoarthritis genes and their collaborative metabolic and inflammatory networks. PLoS ONE 3, e3740 (2008).
Google Scholar
Song, J. et al. MicroRNA-222 regulates MMP-13 via targeting HDAC-4 during osteoarthritis pathogenesis. BBA Clin. 3, 79–89 (2014).
Google Scholar
Hecht, N., Johnstone, B., Angele, P., Walker, T. & Richter, W. Mechanosensitive MiRs regulated by anabolic and catabolic loading of human cartilage. Osteoarthritis Cartilage 27, 1208–1218 (2019).
Google Scholar
Lolli, A., Colella, F., De Bari, C. & van Osch, G. J. V. M. Targeting anti-chondrogenic factors for the stimulation of chondrogenesis: a new paradigm in cartilage repair. J. Orthop. Res. 37, 12–22 (2019).
Google Scholar
Mohanraj, B. et al. Mechanically activated microcapsules for “on-demand” drug delivery in dynamically loaded musculoskeletal tissues. Adv. Funct. Mater. 29, 1807909 (2019).
Google Scholar
Cambré, I. et al. Mechanical strain determines the site-specific localization of inflammation and tissue damage in arthritis. Nat. Commun. 9, 4613 (2018).
Google Scholar
Lin, X., Bai, Y., Zhou, H. & Yang, L. Mechano-active biomaterials for tissue repair and regeneration. J. Mater. Sci. Technol. 59, 227–233 (2020).
Zhang, Y., Yu, J., Bomba, H. N., Zhu, Y. & Gu, Z. Mechanical force-triggered drug delivery. Chem. Rev. 116, 12536–12563 (2016).
Google Scholar
Xiao, L. et al. Hyaluronic acid-based hydrogels containing covalently integrated drug depots: implication for controlling inflammation in mechanically stressed tissues. Biomacromolecules 14, 3808–3819 (2013).
Google Scholar

