Connective tissues within the synovial joints are characterized by their dense extracellular matrix and sparse cellularity. With injury or disease, however, tissues commonly experience an influx of cells owing to proliferation and migration of endogenous mesenchymal cell populations, as well as invasion of the tissue by other cell types, including immune cells. Although this process is critical for successful wound healing, aberrant immune-mediated cell infiltration can lead to pathological inflammation of the joint. Importantly, cells of mesenchymal or haematopoietic origin use distinct modes of migration and thus might respond differently to similar biological cues and microenvironments. Furthermore, cell migration in the physiological microenvironment of musculoskeletal tissues differs considerably from migration in vitro. This Review addresses the complexities of cell migration in fibrous connective tissues from three separate but interdependent perspectives: physiology (including the cellular and extracellular factors affecting 3D cell migration), pathophysiology (cell migration in the context of synovial joint autoimmune disease and injury) and tissue engineering (cell migration in engineered biomaterials). Improved understanding of the fundamental mechanisms governing interstitial cell migration might lead to interventions that stop invasion processes that culminate in deleterious outcomes and/or that expedite migration to direct endogenous cell-mediated repair and regeneration of joint tissues.
Interstitial cell migration in the fibrous microenvironments of intra-articular tissues is regulated by biophysical and biochemical factors.
Immune cells are recruited to and retained within the synovium by inflammatory cytokines and chemokines in rheumatic disorders.
High matrix density and stiffness of adult dense connective tissues restrict the mobility of endogenous cells, impeding wound healing after injury.
Early cell migration into biomaterial scaffolds is a critical but challenging step towards engineering functional musculoskeletal tissues.
Targeted strategies that limit inflammatory cell invasion while promoting the migration of endogenous reparative cells might enhance joint tissue formation and regeneration.
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Franz, C. M., Jones, G. E. & Ridley, A. J. Cell migration in development and disease. Dev. Cell 2, 153–158 (2002).
Cukierman, E., Pankov, R., Stevens, D. R. & Yamada, K. M. Taking cell-matrix adhesions to the third dimension. Science 294, 1708–1712 (2001).
Baker, B. M. & Chen, C. S. Deconstructing the third dimension: how 3D culture microenvironments alter cellular cues. J. Cell Sci. 125, 3015–3024 (2012).
Pathak, A. & Kumar, S. Biophysical regulation of tumor cell invasion: moving beyond matrix stiffness. Integr. Biol. 3, 267–278 (2011).
Friedl, P. & Wolf, K. Plasticity of cell migration: a multiscale tuning model. J. Cell Biol. 188, 11–19 (2010).
Lo, C. M., Wang, H. B., Dembo, M. & Wang, Y. L. Cell movement is guided by the rigidity of the substrate. Biophys. J. 79, 144–152 (2000).
Stoker, M. & Gherardi, E. Regulation of cell movement: the motogenic cytokines. Biochim. Biophys. Acta 1072, 81–102 (1991).
Walters, N. J. & Gentleman, E. Evolving insights in cell-matrix interactions: elucidating how non-soluble properties of the extracellular niche direct stem cell fate. Acta Biomater. 11, 3–16 (2015).
Petrie, R. J. & Yamada, K. M. At the leading edge of three-dimensional cell migration. J. Cell Sci. 125, 5917–5926 (2012).
Wolf, K. et al. Physical limits of cell migration: control by ECM space and nuclear deformation and tuning by proteolysis and traction force. J. Cell Biol. 201, 1069–1084 (2013).
Kurosaka, S. & Kashina, A. Cell biology of embryonic migration. Birth Defects Res. C Embryo Today 84, 102–122 (2008).
Friedl, P., Zanker, K. S. & Brocker, E. B. Cell migration strategies in 3D extracellular matrix: differences in morphology, cell matrix interactions, and integrin function. Microsc. Res. Tech. 43, 369–378 (1998).
Friedl, P. & Brocker, E. B. T cell migration in three-dimensional extracellular matrix: guidance by polarity and sensations. Dev. Immunol. 7, 249–266 (2000).
Liu, Y. J. et al. Confinement and low adhesion induce fast amoeboid migration of slow mesenchymal cells. Cell 160, 659–672 (2015).
Ruprecht, V. et al. Cortical contractility triggers a stochastic switch to fast amoeboid cell motility. Cell 160, 673–685 (2015).
Petrie, R. J., Koo, H. & Yamada, K. M. Generation of compartmentalized pressure by a nuclear piston governs cell motility in a 3D matrix. Science 345, 1062–1065 (2014).
Stroka, K. M. et al. Water permeation drives tumor cell migration in confined microenvironments. Cell 157, 611–623 (2014).
Mouw, J. K., Ou, G. & Weaver, V. M. Extracellular matrix assembly: a multiscale deconstruction. Nat. Rev. Mol. Cell. Biol. 15, 771–785 (2014).
Lu, P., Takai, K., Weaver, V. M. & Werb, Z. Extracellular matrix degradation and remodeling in development and disease. Cold Spring Harb. Perspect. Biol. 3, a005058 (2011).
Friedl, P. & Brocker, E. B. The biology of cell locomotion within three-dimensional extracellular matrix. Cell. Mol. Life Sci. 57, 41–64 (2000).
Friedl, P. & Wolf, K. Proteolytic interstitial cell migration: a five-step process. Cancer Metastasis Rev. 28, 129–135 (2009).
Guilak, F., Nims, R., Dicks, A., Wu, C.-L. & Meulenbelt, I. Osteoarthritis as a disease of the cartilage pericellular matrix. Matrix Biol. 71–72, 40–50 (2018).
Guilak, F., Tedrow, J. R. & Burgkart, R. Viscoelastic properties of the cell nucleus. Biochem. Biophys. Res. Commun. 269, 781–786 (2000).
Lautscham, L. A. et al. Migration in confined 3D environments is determined by a combination of adhesiveness, nuclear volume, contractility, and cell stiffness. Biophys. J. 109, 900–913 (2015).
Friedl, P., Wolf, K. & Lammerding, J. Nuclear mechanics during cell migration. Curr. Opin. Cell Biol. 23, 55–64 (2011).
Denais, C. M. et al. Nuclear envelope rupture and repair during cancer cell migration. Science 352, 353–358 (2016).
Harada, T. et al. Nuclear lamin stiffness is a barrier to 3D migration, but softness can limit survival. J. Cell Biol. 204, 669–682 (2014).
Rowat, A. C. et al. Nuclear envelope composition determines the ability of neutrophil-type cells to passage through micron-scale constrictions. J. Biol. Chem. 288, 8610–8618 (2013).
Pajerowski, J. D., Dahl, K. N., Zhong, F. L., Sammak, P. J. & Discher, D. E. Physical plasticity of the nucleus in stem cell differentiation. Proc. Natl Acad. Sci. USA 104, 15619–15624 (2007).
Lammerding, J. et al. Lamins A and C but not lamin B1 regulate nuclear mechanics. J. Biol. Chem. 281, 25768–25780 (2006).
Booth-Gauthier, E. A. et al. Hutchinson-Gilford progeria syndrome alters nuclear shape and reduces cell motility in three dimensional model substrates. Integr. Biol. 5, 569–577 (2013).
Greiner, A. M. et al. Multifunctional polymer scaffolds with adjustable pore size and chemoattractant gradients for studying cell matrix invasion. Biomaterials 35, 611–619 (2014).
Infante, E. et al. LINC complex-Lis1 interplay controls MT1-MMP matrix digest-on-demand response for confined tumor cell migration. Nat. Commun. 9, 2443 (2018).
Olins, A. L. et al. Nuclear envelope and chromatin compositional differences comparing undifferentiated and retinoic acid- and phorbol ester-treated HL-60 cells. Exp. Cell Res. 268, 115–127 (2001).
Ekpenyong, A. E. et al. Viscoelastic properties of differentiating blood cells are fate- and function-dependent. PLOS ONE 7, e45237 (2012).
Graham, D. M. et al. Enucleated cells reveal differential roles of the nucleus in cell migration, polarity, and mechanotransduction. J. Cell Biol. 217, 895–914 (2018).
Singh, S. P., Schwartz, M. P., Lee, J. Y., Fairbanks, B. D. & Anseth, K. S. A peptide functionalized poly(ethylene glycol) (PEG) hydrogel for investigating the influence of biochemical and biophysical matrix properties on tumor cell migration. Biomater. Sci. 2, 1024–1034 (2014).
Lowin, T. & Straub, R. H. Integrins and their ligands in rheumatoid arthritis. Arthritis Res. Ther. 13, 244 (2011).
Loeser, R. F. Integrins and chondrocyte-matrix interactions in articular cartilage. Matrix Biol. 39, 11–16 (2014).
Hinz, B. The extracellular matrix and transforming growth factor-β1: tale of a strained relationship. Matrix Biol. 47, 54–65 (2015).
Ni, G. X., Li, Z. & Zhou, Y. Z. The role of small leucine-rich proteoglycans in osteoarthritis pathogenesis. Osteoarthr. Cartil. 22, 896–903 (2014).
Merline, R., Schaefer, R. M. & Schaefer, L. The matricellular functions of small leucine-rich proteoglycans (SLRPs). J. Cell Commun. Signal 3, 323–335 (2009).
Tufvesson, E. & Westergren-Thorsson, G. Tumour necrosis factor-alpha interacts with biglycan and decorin. FEBS Lett. 530, 124–128 (2002).
Dormann, D. & Weijer, C. J. Chemotactic cell movement during development. Curr. Opin. Genet. Dev. 13, 358–364 (2003).
Wu, J. et al. Gradient biomaterials and their influences on cell migration. Interface Focus 2, 337–355 (2012).
Turner, M. D., Nedjai, B., Hurst, T. & Pennington, D. J. Cytokines and chemokines: at the crossroads of cell signalling and inflammatory disease. Biochim. Biophys. Acta 1843, 2563–2582 (2014).
Qu, F. et al. Maturation state and matrix microstructure regulate interstitial cell migration in dense connective tissues. Sci. Rep. 8, 3295 (2018).
Swift, J. et al. Nuclear lamin-A scales with tissue stiffness and enhances matrix-directed differentiation. Science 341, 1240104 (2013).
Vogel, V. & Sheetz, M. Local force and geometry sensing regulate cell functions. Nat. Rev. Mol. Cell. Biol. 7, 265–275 (2006).
Peyton, S. R. & Putnam, A. J. Extracellular matrix rigidity governs smooth muscle cell motility in a biphasic fashion. J. Cell. Physiol. 204, 198–209 (2005).
Zaman, M. H. et al. Migration of tumor cells in 3D matrices is governed by matrix stiffness along with cell-matrix adhesion and proteolysis. Proc. Natl Acad. Sci. USA 103, 10889–10894 (2006).
Pathak, A. & Kumar, S. Independent regulation of tumor cell migration by matrix stiffness and confinement. Proc. Natl Acad. Sci. USA 109, 10334–10339 (2012).
Peyton, S. R. et al. Marrow-derived stem cell motility in 3D synthetic scaffold is governed by geometry along with adhesivity and stiffness. Biotechnol. Bioeng. 108, 1181–1193 (2011).
Zaman, M. H., Matsudaira, P. & Lauffenburger, D. A. Understanding effects of matrix protease and matrix organization on directional persistence and translational speed in three-dimensional cell migration. Ann. Biomed. Eng. 35, 91–100 (2007).
Kim, M. C. et al. Integrating focal adhesion dynamics, cytoskeleton remodeling, and actin motor activity for predicting cell migration on 3D curved surfaces of the extracellular matrix. Integr. Biol. 4, 1386–1397 (2012).
Provenzano, P. P., Inman, D. R., Eliceiri, K. W., Trier, S. M. & Keely, P. J. Contact guidance mediated three-dimensional cell migration is regulated by Rho/ROCK-dependent matrix reorganization. Biophys. J. 95, 5374–5384 (2008).
Petrie, R. J., Doyle, A. D. & Yamada, K. M. Random versus directionally persistent cell migration. Nat. Rev. Mol. Cell. Biol. 10, 538–549 (2009).
Fraley, S. I. et al. Three-dimensional matrix fiber alignment modulates cell migration and MT1-MMP utility by spatially and temporally directing protrusions. Sci. Rep. 5, 14580 (2015).
Dickinson, R. B., Guido, S. & Tranquillo, R. T. Biased cell migration of fibroblasts exhibiting contact guidance in oriented collagen gels. Ann. Biomed. Eng. 22, 342–356 (1994).
Riching, K. M. et al. 3D collagen alignment limits protrusions to enhance breast cancer cell persistence. Biophys. J. 107, 2546–2558 (2014).
Smith, M. D. The normal synovium. Open Rheumatol. J. 5, 100–106 (2011).
Nevius, E., Gomes, A. C. & Pereira, J. P. Inflammatory cell migration in rheumatoid arthritis: a comprehensive review. Clin. Rev. Allergy Immunol. 51, 59–78 (2016).
Tak, P. P. et al. Analysis of the synovial cell infiltrate in early rheumatoid synovial tissue in relation to local disease activity. Arthritis. Rheum. 40, 217–225 (1997).
Mulherin, D., Fitzgerald, O. & Bresnihan, B. Synovial tissue macrophage populations and articular damage in rheumatoid arthritis. Arthritis Rheum. 39, 115–124 (1996).
Mellado, M. et al. T cell migration in rheumatoid arthritis. Front. Immunol. 6, 384 (2015).
Iwamoto, T., Okamoto, H., Toyama, Y. & Momohara, S. Molecular aspects of rheumatoid arthritis: chemokines in the joints of patients. FEBS J. 275, 4448–4455 (2008).
Koelink, P. J. et al. Targeting chemokine receptors in chronic inflammatory diseases: an extensive review. Pharmacol. Ther. 133, 1–18 (2012).
Buckley, C. D. Why does chronic inflammatory joint disease persist? Clin. Med. 3, 361–366 (2003).
Solari, R., Pease, J. E. & Begg, M. Chemokine receptors as therapeutic targets: why aren’t there more drugs? Eur. J. Pharmacol. 746, 363–367 (2015).
Hitchon, C. A. & El-Gabalawy, H. S. The synovium in rheumatoid arthritis. Open Rheumatol. J. 5, 107–114 (2011).
Amin, M. A. et al. Interleukin-18 induces angiogenic factors in rheumatoid arthritis synovial tissue fibroblasts via distinct signaling pathways. Arthritis Rheum. 56, 1787–1797 (2007).
Bruhl, H. et al. Functional expression of the chemokine receptor CCR7 on fibroblast-like synoviocytes. Rheumatology 47, 1771–1774 (2008).
Garcia-Vicuna, R. et al. CC and CXC chemokine receptors mediate migration, proliferation, and matrix metalloproteinase production by fibroblast-like synoviocytes from rheumatoid arthritis patients. Arthritis Rheum. 50, 3866–3877 (2004).
Nanki, T., Nagasaka, K., Hayashida, K., Saita, Y. & Miyasaka, N. Chemokines regulate IL-6 and IL-8 production by fibroblast-like synoviocytes from patients with rheumatoid arthritis. J. Immunol. 167, 5381–5385 (2001).
Lefevre, S. et al. Disease-specific effects of matrix and growth factors on adhesion and migration of rheumatoid synovial fibroblasts. J. Immunol. 198, 4588–4595 (2017).
Tolboom, T. C. et al. Invasive properties of fibroblast-like synoviocytes: correlation with growth characteristics and expression of MMP-1, MMP-3, and MMP-10. Ann. Rheum. Dis. 61, 975–980 (2002).
Li, D., Xiao, Z., Wang, G. & Song, X. Knockdown of ADAM10 inhibits migration and invasion of fibroblast-like synoviocytes in rheumatoid arthritis. Mol. Med. Rep. 12, 5517–5523 (2015).
Tolboom, T. C. et al. Invasiveness of fibroblast-like synoviocytes is an individual patient characteristic associated with the rate of joint destruction in patients with rheumatoid arthritis. Arthritis Rheum. 52, 1999–2002 (2005).
El-Zayadi, A. A. et al. Interleukin-22 drives the proliferation, migration and osteogenic differentiation of mesenchymal stem cells: a novel cytokine that could contribute to new bone formation in spondyloarthropathies. Rheumatology 56, 488–493 (2017).
Justa, S., Zhou, X. & Sarkar, S. Endogenous IL-22 plays a dual role in arthritis: regulation of established arthritis via IFN-gamma responses. PLOS ONE 9, e93279 (2014).
Qin, Y. et al. Increased CCL19 and CCL21 levels promote fibroblast ossification in ankylosing spondylitis hip ligament tissue. BMC Musculoskelet. Disord. 15, 316 (2014).
Lee, J. et al. Stimulation of osteoclast migration and bone resorption by C-C chemokine ligands 19 and 21. Exp. Mol. Med. 49, e358 (2017).
Meng, X. H. et al. Quantitative evaluation of knee cartilage and meniscus destruction in patients with rheumatoid arthritis using T1rho and T2 mapping. Eur. J. Radiol. 96, 91–97 (2017).
Fuhrmann, I. K., Steinhagen, J., Ruther, W. & Schumacher, U. Comparative immunohistochemical evaluation of the zonal distribution of extracellular matrix and inflammation markers in human meniscus in osteoarthritis and rheumatoid arthritis. Acta Histochem. 117, 243–254 (2015).
Lopez-Franco, M. et al. Meniscal degeneration in human knee osteoarthritis: in situ hybridization and immunohistochemistry study. Arch. Orthop. Trauma Surg. 136, 175–183 (2016).
van de Sande, M. A., de Groot, J. H. & Rozing, P. M. Clinical implications of rotator cuff degeneration in the rheumatic shoulder. Arthritis Rheum. 59, 317–324 (2008).
Meyer, C. et al. Rheumatoid arthritis affecting the upper cervical spine: biomechanical assessment of the stabilizing ligaments. Biomed. Res. Int. 2017, 6131703 (2017).
Puttlitz, C. M. et al. Biomechanical rationale for the pathology of rheumatoid arthritis in the craniovertebral junction. Spine 25, 1607–1616 (2000).
Yang, G., Rothrauff, B. B. & Tuan, R. S. Tendon and ligament regeneration and repair: clinical relevance and developmental paradigm. Birth Defects Res. C Embryo Today 99, 203–222 (2013).
McNulty, A. L., Moutos, F. T., Weinberg, J. B. & Guilak, F. Enhanced integrative repair of the porcine meniscus in vitro by inhibition of interleukin-1 or tumor necrosis factor alpha. Arthritis Rheum. 56, 3033–3042 (2007).
Mohanraj, B. et al. Chondrocyte and mesenchymal stem cell derived engineered cartilage exhibits differential sensitivity to pro-inflammatory cytokines. J. Orthop. Res. 36, 2901–2910 (2018).
Arnoczky, S. P. & Warren, R. F. The microvasculature of the meniscus and its response to injury. An experimental study in the dog. Am. J. Sports Med. 11, 131–141 (1983).
Hiraki, Y. & Shukunami, C. Angiogenesis inhibitors localized in hypovascular mesenchymal tissues: chondromodulin-I and tenomodulin. Connect. Tissue Res. 46, 3–11 (2005).
Manske, P. R., Lesker, P. A., Gelberman, R. H. & Rucinsky, T. E. Intrinsic restoration of the flexor tendon surface in the nonhuman primate. J. Hand Surg. Am. 10, 632–637 (1985).
Sharma, P. & Maffulli, N. Biology of tendon injury: healing, modeling and remodeling. J. Musculoskelet. Neuronal Interact. 6, 181–190 (2006).
Lo, I. K., Chi, S., Ivie, T., Frank, C. B. & Rattner, J. B. The cellular matrix: a feature of tensile bearing dense soft connective tissues. Histol. Histopathol. 17, 523–537 (2002).
Cadby, J. A., Buehler, E., Godbout, C., van Weeren, P. R. & Snedeker, J. G. Differences between the cell populations from the peritenon and the tendon core with regard to their potential implication in tendon repair. PLOS ONE 9, e92474 (2014).
Shen, W. et al. Intra-articular injection of human meniscus stem/progenitor cells promotes meniscus regeneration and ameliorates osteoarthritis through stromal cell-derived factor-1/CXCR4-mediated homing. Stem Cells Transl Med. 3, 387–394 (2014).
Mauck, R. L., Martinez-Diaz, G. J., Yuan, X. & Tuan, R. S. Regional multilineage differentiation potential of meniscal fibrochondrocytes: implications for meniscus repair. Anat. Rec. 290, 48–58 (2007).
Mesiha, M. et al. Pathologic characteristics of the torn human meniscus. Am. J. Sports Med. 35, 103–112 (2007).
de Albornoz, P. M. & Forriol, F. The meniscal healing process. Muscles Ligaments Tendons J. 2, 10–18 (2012).
Newman, A. P., Anderson, D. R., Daniels, A. U. & Dales, M. C. Mechanics of the healed meniscus in a canine model. Am. J. Sports Med. 17, 164–175 (1989).
Russo, V. et al. Cellular and molecular maturation in fetal and adult ovine calcaneal tendons. J. Anat. 226, 126–142 (2015).
Clark, C. R. & Ogden, J. A. Development of the menisci of the human knee joint. Morphological changes and their potential role in childhood meniscal injury. J. Bone Joint Surg. Am. 65, 538–547 (1983).
Marturano, J. E., Arena, J. D., Schiller, Z. A., Georgakoudi, I. & Kuo, C. K. Characterization of mechanical and biochemical properties of developing embryonic tendon. Proc. Natl Acad. Sci. USA 110, 6370–6375 (2013).
Li, Q. et al. Impacts of maturation on the micromechanics of the meniscus extracellular matrix. J. Biomech. 72, 252–257 (2018).
Ionescu, L. C. et al. Maturation state-dependent alterations in meniscus integration: implications for scaffold design and tissue engineering. Tissue Eng. Part A 17, 193–204 (2011).
Melrose, J., Smith, S., Cake, M., Read, R. & Whitelock, J. Comparative spatial and temporal localisation of perlecan, aggrecan and type I, II and IV collagen in the ovine meniscus: an ageing study. Histochem. Cell Biol. 124, 225–235 (2005).
Di Giancamillo, A., Deponti, D., Addis, A., Domeneghini, C. & Peretti, G. M. Meniscus maturation in the swine model: changes occurring along with anterior to posterior and medial to lateral aspect during growth. J. Cell. Mol. Med. 18, 1964–1974 (2014).
Morales, T. I. Chondrocyte moves: clever strategies? Osteoarthr. Cartil. 15, 861–871 (2007).
Qu, F., Holloway, J. L., Esterhai, J. L., Burdick, J. A. & Mauck, R. L. Programmed biomolecule delivery to enable and direct cell migration for connective tissue repair. Nat. Commun. 8, 1780 (2017).
Qu, F. et al. Repair of dense connective tissues via biomaterial-mediated matrix reprogramming of the wound interface. Biomaterials 39, 85–94 (2015).
Hunziker, E. B. & Kapfinger, E. Removal of proteoglycans from the surface of defects in articular cartilage transiently enhances coverage by repair cells. J. Bone Joint Surg. Br. 80, 144–150 (1998).
Kim, M., Farrell, M. J., Steinberg, D. R., Burdick, J. A. & Mauck, R. L. Enhanced nutrient transport improves the depth-dependent properties of tri-layered engineered cartilage constructs with zonal co-culture of chondrocytes and MSCs. Acta Biomater. 58, 1–11 (2017).
Cheng, N. C., Estes, B. T., Young, T. H. & Guilak, F. Genipin-crosslinked cartilage-derived matrix as a scaffold for human adipose-derived stem cell chondrogenesis. Tissue Eng. Part A 19, 484–496 (2013).
Baker, B. M. et al. The potential to improve cell infiltration in composite fiber-aligned electrospun scaffolds by the selective removal of sacrificial fibers. Biomaterials 29, 2348–2358 (2008).
Sundararaghavan, H. G. & Burdick, J. A. Gradients with depth in electrospun fibrous scaffolds for directed cell behavior. Biomacromolecules 12, 2344–2350 (2011).
Bhargava, M. M. et al. The effect of cytokines on the proliferation and migration of bovine meniscal cells. Am. J. Sports Med. 27, 636–643 (1999).
Caliari, S. R. & Harley, B. A. The effect of anisotropic collagen-GAG scaffolds and growth factor supplementation on tendon cell recruitment, alignment, and metabolic activity. Biomaterials 32, 5330–5340 (2011).
Hannafin, J. A., Attia, E. T., Warren, R. F. & Bhargava, M. M. Characterization of chemotatic migration and growth kinetics of canine knee ligament fibroblasts. J. Orthop. Res. 17, 398–404 (1999).
Kucia, M. et al. CXCR4–SDF-1 signalling, locomotion, chemotaxis and adhesion. J. Mol. Histol. 35, 233–245 (2004).
Moore, K., Macsween, M. & Shoichet, M. Immobilized concentration gradients of neurotrophic factors guide neurite outgrowth of primary neurons in macroporous scaffolds. Tissue Eng. 12, 267–278 (2006).
Wiles, K., Fishman, J. M., De Coppi, P. & Birchall, M. A. The host immune response to tissue-engineered organs: current problems and future directions. Tissue Eng. Part B Rev. 22, 208–219 (2016).
Swartz, M. A., Hirosue, S. & Hubbell, J. A. Engineering approaches to immunotherapy. Sci. Transl Med. 4, 148rv149 (2012).
Stano, A., Scott, E. A., Dane, K. Y., Swartz, M. A. & Hubbell, J. A. Tunable T cell immunity towards a protein antigen using polymersomes versus solid-core nanoparticles. Biomaterials 34, 4339–4346 (2013).
Nembrini, C. et al. Nanoparticle conjugation of antigen enhances cytotoxic T cell responses in pulmonary vaccination. Proc. Natl Acad. Sci. USA 108, E989–E997 (2011).
McCarthy, D. P. et al. An antigen-encapsulating nanoparticle platform for TH1/17 immune tolerance therapy. Nanomedicine 13, 191–200 (2017).
Spiller, K. L. et al. Sequential delivery of immunomodulatory cytokines to facilitate the M1-to-M2 transition of macrophages and enhance vascularization of bone scaffolds. Biomaterials 37, 194–207 (2015).
Boehler, R. M. et al. Lentivirus delivery of IL-10 to promote and sustain macrophage polarization towards an anti-inflammatory phenotype. Biotechnol. Bioeng. 111, 1210–1221 (2014).
Sridharan, R. et al. Biomaterial based modulation of macrophage polarization: a review and suggested design principles. Mater. Today 18, 313–325 (2015).
Ratanavaraporn, J., Furuya, H. & Tabata, Y. Local suppression of pro-inflammatory cytokines and the effects in BMP-2-induced bone regeneration. Biomaterials 33, 304–316 (2012).
Chen, W. C. et al. Controlled dual delivery of fibroblast growth factor-2 and Interleukin-10 by heparin-based coacervate synergistically enhances ischemic heart repair. Biomaterials 72, 138–151 (2015).
Geckil, H., Xu, F., Zhang, X., Moon, S. & Demirci, U. Engineering hydrogels as extracellular matrix mimics. Nanomedicine 5, 469–484 (2010).
Kim, D.-H., Provenzano, P. P., Smith, C. L. & Levchenko, A. Matrix nanotopography as a regulator of cell function. J. Cell Biol. 197, 351–360 (2012).
Wade, R. J., Bassin, E. J., Gramlich, W. M. & Burdick, J. A. Nanofibrous hydrogels with spatially patterned biochemical signals to control cell behavior. Adv. Mater. 27, 1356–1362 (2015).
Kim, I. L., Khetan, S., Baker, B. M., Chen, C. S. & Burdick, J. A. Fibrous hyaluronic acid hydrogels that direct MSC chondrogenesis through mechanical and adhesive cues. Biomaterials 34, 5571–5580 (2013).
Billiet, T., Vandenhaute, M., Schelfhout, J., Van Vlierberghe, S. & Dubruel, P. A review of trends and limitations in hydrogel-rapid prototyping for tissue engineering. Biomaterials 33, 6020–6041 (2012).
Bullough, P. G., Munuera, L., Murphy, J. & Weinstein, A. M. The strength of the menisci of the knee as it relates to their fine structure. J. Bone Joint Surg. Br. 52, 564–567 (1970).
Proctor, C. S., Schmidt, M. B., Whipple, R. R., Kelly, M. A. & Mow, V. C. Material properties of the normal medial bovine meniscus. J. Orthop. Res. 7, 771–782 (1989).
LeRoux, M. A. & Setton, L. A. Experimental and biphasic FEM determinations of the material properties and hydraulic permeability of the meniscus in tension. J. Biomech. Eng. 124, 315–321 (2002).
Mauck, R. L. et al. Engineering on the straight and narrow: the mechanics of nanofibrous assemblies for fiber-reinforced tissue regeneration. Tissue Eng. Part B Rev. 15, 171–193 (2009).
Lim, J. Y. & Donahue, H. J. Cell sensing and response to micro-and nanostructured surfaces produced by chemical and topographic patterning. Tissue Eng. 13, 1879–1891 (2007).
Gilchrist, C. L., Ruch, D. S., Little, D. & Guilak, F. Micro-scale and meso-scale architectural cues cooperate and compete to direct aligned tissue formation. Biomaterials 35, 10015–10024 (2014).
Baker, B. M. & Mauck, R. L. The effect of nanofiber alignment on the maturation of engineered meniscus constructs. Biomaterials 28, 1967–1977 (2007).
Orr, S. B. et al. Aligned multilayered electrospun scaffolds for rotator cuff tendon tissue engineering. Acta Biomater. 24, 117–126 (2015).
Ionescu, L. C. & Mauck, R. L. Porosity and cell preseeding influence electrospun scaffold maturation and meniscus integration in vitro. Tissue Eng. Part A 19, 538–547 (2013).
Nam, J., Huang, Y., Agarwal, S. & Lannutti, J. Improved cellular infiltration in electrospun fiber via engineered porosity. Tissue Eng. 13, 2249–2257 (2007).
Simonet, M., Schneider, O. D., Neuenschwander, P. & Stark, W. J. Ultraporous 3D polymer meshes by low-temperature electrospinning: use of ice crystals as a removable void template. Polym. Eng. Sci. 47, 2020–2026 (2007).
Lee, B. L.-P. et al. Synovial stem cells and their responses to the porosity of microfibrous scaffold. Acta Biomater. 9, 7264–7275 (2013).
Phipps, M. C., Clem, W. C., Grunda, J. M., Clines, G. A. & Bellis, S. L. Increasing the pore sizes of bone-mimetic electrospun scaffolds comprised of polycaprolactone, collagen I and hydroxyapatite to enhance cell infiltration. Biomaterials 33, 524–534 (2012).
Baker, B. M. et al. Sacrificial nanofibrous composites provide instruction without impediment and enable functional tissue formation. Proc. Natl Acad. Sci. USA 109, 14176–14181 (2012).
Cai, S., Xu, H., Jiang, Q. & Yang, Y. Novel 3D electrospun scaffolds with fibers oriented randomly and evenly in three dimensions to closely mimic the unique architectures of extracellular matrices in soft tissues: fabrication and mechanism study. Langmuir 29, 2311–2318 (2013).
Moutos, F. T., Freed, L. E. & Guilak, F. A biomimetic three-dimensional woven composite scaffold for functional tissue engineering of cartilage. Nat. Mater. 6, 162–167 (2007).
Lee, J., Jang, J., Oh, H., Jeong, Y. H. & Cho, D.-W. Fabrication of a three-dimensional nanofibrous scaffold with lattice pores using direct-write electrospinning. Mater. Lett. 93, 397–400 (2013).
Daly, A. C. et al. 3D bioprinting for cartilage and osteochondral tissue engineering. Adv. Healthc. Mater. 6, 1700298 (2017).
Miller, J. S. et al. Rapid casting of patterned vascular networks for perfusable engineered three-dimensional tissues. Nat. Mater. 11, 768–774 (2012).
Freeman, F. E. & Kelly, D. J. Tuning alginate bioink stiffness and composition for controlled growth factor delivery and to spatially direct MSC fate within bioprinted tissues. Sci. Rep. 7, 17042 (2017).
Skardal, A. et al. A hydrogel bioink toolkit for mimicking native tissue biochemical and mechanical properties in bioprinted tissue constructs. Acta Biomater. 25, 24–34 (2015).
Albritton, J. L. & Miller, J. S. 3D bioprinting: improving in vitro models of metastasis with heterogeneous tumor microenvironments. Dis. Model. Mech. 10, 3–14 (2017).
Miller, J. S. et al. Bioactive hydrogels made from step-growth derived PEG–peptide macromers. Biomaterials 31, 3736–3743 (2010).
Ehrbar, M. et al. Elucidating the role of matrix stiffness in 3D cell migration and remodeling. Biophys. J. 100, 284–293 (2011).
Bott, K. et al. The effect of matrix characteristics on fibroblast proliferation in 3D gels. Biomaterials 31, 8454–8464 (2010).
Wade, R. J., Bassin, E. J., Rodell, C. B. & Burdick, J. A. Protease-degradable electrospun fibrous hydrogels. Nat. Commun. 6, 6639 (2015).
Schultz, K. M., Kyburz, K. A. & Anseth, K. S. Measuring dynamic cell–material interactions and remodeling during 3D human mesenchymal stem cell migration in hydrogels. Proc. Natl Acad. Sci. USA 112, E3757–E3764 (2015).
Lee, S. H., Miller, J. S., Moon, J. J. & West, J. L. Proteolytically degradable hydrogels with a fluorogenic substrate for studies of cellular proteolytic activity and migration. Biotechnol. Prog. 21, 1736–1741 (2005).
Feng, Q., Zhu, M., Wei, K. & Bian, L. Cell-mediated degradation regulates human mesenchymal stem cell chondrogenesis and hypertrophy in MMP-sensitive hyaluronic acid hydrogels. PLOS ONE 9, e99587 (2014).
Salinas, C. N. & Anseth, K. S. The enhancement of chondrogenic differentiation of human mesenchymal stem cells by enzymatically regulated RGD functionalities. Biomaterials 29, 2370–2377 (2008).
Chaudhuri, O. et al. Substrate stress relaxation regulates cell spreading. Nat. Commun. 6, 6364 (2015).
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).
Brunger, J. M., Zutshi, A., Willard, V. P., Gersbach, C. A. & Guilak, F. CRISPR/Cas9 editing of murine induced pluripotent stem cells for engineering inflammation-resistant tissues. Arthritis Rheumatol. 69, 1111–1121 (2017).
Brunger, J. M., Zutshi, A., Willard, V. P., Gersbach, C. A. & Guilak, F. Genome engineering of stem cells for autonomously regulated, closed-loop delivery of biologic drugs. Stem Cell Rep. 8, 1202–1213 (2017).
Brunger, J. M. et al. Scaffold-mediated lentiviral transduction for functional tissue engineering of cartilage. Proc. Natl Acad. Sci. USA 111, E798–806 (2014).
Glass, K. A. et al. Tissue-engineered cartilage with inducible and tunable immunomodulatory properties. Biomaterials 35, 5921–5931 (2014).
The work of F.Q., F.G. and R.L.M. is supported by the US National Institutes of Health (AR060719, AR056624, EB008722, AR50245, AR48852, AG46927, AG15768, AR48182, AR067467 and AR057235), the US Department of Veterans’ Affairs (I01 RX000174), the Arthritis Foundation, the Nancy Taylor Foundation for Chronic Diseases and the Collaborative Research Center of the AO Foundation in Davos.
Nature Reviews Rheumatology thanks D. Grande and the other anonymous reviewer(s) for their contribution to the peer review of this work.
- Stress fibre
Contractile bundles in non-muscle cells composed of actin filaments and non-muscle myosin II; myosin motor activity results in contraction of the actomyosin bundles.
The mechanical properties of a material assessed at a local level (that is, at the micrometre scale). This approach can identify heterogeneities in materials or tissues that are indicative of the constituent materials and their properties at that location.
The microscopic structure of a material or tissue.
Directional cell movement along a soluble biochemical gradient.
Directional cell movement along a substrate-bound insoluble gradient.
- Collective migration
The process by which a group of cells move together while maintaining cell–cell contact.
- Tensile modulus
Young’s modulus of a material evaluated in tension (that is, a measurement of tensile strength, which is the ability of a material to withstand being stretched).
- Compressive modulus
Young’s modulus of a material evaluated in compression (that is, a measurement of compressive strength, which is the ability of a material to withstand being compressed).
- Shear modulus
Young’s modulus of a material evaluated in shear (that is, a measurement of the shear strength, which is the ability of a material to withstand forces that can cause the internal structure of the material to slide against itself).
- Young’s modulus
A mechanical property that defines the relationship between stress (force per unit area) and strain (proportional deformation) of a linearly elastic material during uniaxial deformation (also referred to as the elastic modulus; measured in MPa). Although commonly referred to as tissue stiffness or rigidity, these two terms are actually structural properties (that is, dependent on the size and shape of the tissue) and are not inherent material properties.
- Contact guidance
The response of cells to topographic cues; the direction of cell alignment and migration is affected by geometrical patterns such as grooves or fibres.
An abnormal layer of fibrovascular tissue, which can occur in rheumatoid arthritis.
- Heterotopic ossification
The presence of bone in soft tissue where bone does not normally exist.
- Scar tissue
Dense fibrous tissue that replaces original tissue during wound healing; the scar tissue is generally disordered and does not match the original tissue in terms of the biochemical content or mechanical properties.
Abnormal formation of scar tissue after injury that connects normally separated tissues and impedes joint motion.
A technique for generating microstructures using moulds with micrometre-scale features.
A technique that uses light to transfer geometric patterns from a photomask (an opaque plate that enables light to shine through in a defined pattern) to a light-sensitive chemical on a substrate.
A technique that produces nanofibres by charging and pulling a polymer solution through a spinneret under a high-voltage electrical field.
- Rapid prototyping
A group of techniques for constructing a 3D product using computer-aided design (for example, 3D printing).
Describes mechanical and physical properties that vary on the basis of the testing direction.
Space-filling particles used to create porous materials, which are later removed to generate voids.
Extrudable polymer solutions used in 3D bioprinting that can contain cells and/or other biologics that can solidify after printing.
- Stress relaxation
A time-dependent decrease in stress of a material in response to the same level of strain.