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Technical Forum

Harnessing Biomechanics to Develop Cartilage Regeneration Strategies

[+] Author and Article Information
Kyriacos A. Athanasiou

Distinguished Professor
of Biomedical Engineering
and Orthopaedic Surgery
Chair
Department of Biomedical Engineering,
College of Engineering,
University of California Davis,
One Shields Avenue,
Davis, CA 95616
e-mail: athanasiou@ucdavis.edu

Donald J. Responte

Zimmer,
1900 Aston Avenue,
Carlsbad, CA 92008

Wendy E. Brown, Jerry C. Hu

Department of Biomedical Engineering,
College of Engineering,
University of California Davis,
One Shields Avenue,
Davis, CA 95616

1Corresponding author.

Manuscript received August 10, 2014; final manuscript received October 6, 2014; published online January 26, 2015. Editor: Beth Winkelstein.

J Biomech Eng 137(2), 020901 (Feb 01, 2015) (14 pages) Paper No: BIO-14-1381; doi: 10.1115/1.4028825 History: Received August 10, 2014; Revised October 06, 2014; Online January 26, 2015

As this review was prepared specifically for the American Society of Mechanical Engineers H.R. Lissner Medal, it primarily discusses work toward cartilage regeneration performed in Dr. Kyriacos A. Athanasiou's laboratory over the past 25 years. The prevalence and severity of degeneration of articular cartilage, a tissue whose main function is largely biomechanical, have motivated the development of cartilage tissue engineering approaches informed by biomechanics. This article provides a review of important steps toward regeneration of articular cartilage with suitable biomechanical properties. As a first step, biomechanical and biochemical characterization studies at the tissue level were used to provide design criteria for engineering neotissues. Extending this work to the single cell and subcellular levels has helped to develop biochemical and mechanical stimuli for tissue engineering studies. This strong mechanobiological foundation guided studies on regenerating hyaline articular cartilage, the knee meniscus, and temporomandibular joint (TMJ) fibrocartilage. Initial tissue engineering efforts centered on developing biodegradable scaffolds for cartilage regeneration. After many years of studying scaffold-based cartilage engineering, scaffoldless approaches were developed to address deficiencies of scaffold-based systems, resulting in the self-assembling process. This process was further improved by employing exogenous stimuli, such as hydrostatic pressure, growth factors, and matrix-modifying and catabolic agents, both singly and in synergistic combination to enhance neocartilage functional properties. Due to the high cell needs for tissue engineering and the limited supply of native articular chondrocytes, costochondral cells are emerging as a suitable cell source. Looking forward, additional cell sources are investigated to render these technologies more translatable. For example, dermis isolated adult stem (DIAS) cells show potential as a source of chondrogenic cells. The challenging problem of enhanced integration of engineered cartilage with native cartilage is approached with both familiar and novel methods, such as lysyl oxidase (LOX). These diverse tissue engineering strategies all aim to build upon thorough biomechanical characterizations to produce functional neotissue that ultimately will help combat the pressing problem of cartilage degeneration. As our prior research is reviewed, we look to establish new pathways to comprehensively and effectively address the complex problems of musculoskeletal cartilage regeneration.

Copyright © 2015 by ASME
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Murphy, M. K., Masters, T. E., Hu, J. C., and Athanasiou, K. A., 2013, “Engineering a Fibrocartilage Spectrum Through Modulation of Aggregate Redifferentiation,” Cell Transplant., (preprint). [CrossRef]
Darling, E. M., and Athanasiou, K. A., 2005, “Rapid Phenotypic Changes in Passaged Articular Chondrocyte Subpopulations,” J. Orthop. Res., 23(2), pp. 425–432. [CrossRef] [PubMed]
Koay, E. J., and Athanasiou, K. A., 2008, “Hypoxic Chondrogenic Differentiation of Human Embryonic Stem Cells Enhances Cartilage Protein Synthesis and Biomechanical Functionality,” Osteoarthritis Cartilage, 16(12), pp. 1450–1456. [CrossRef] [PubMed]
Hoben, G. M., Willard, V. P., and Athanasiou, K. A., 2008, “Fibrochondrogenesis of hESCs: Growth Factor Combinations and Co-Cultures,” Stem Cells Dev., 18(2), pp. 283–292. [CrossRef]
Levenberg, S., Huang, N. F., Lavik, E., Rogers, A. B., Itskovitz-Eldor, J., and Langer, R., 2003, “Differentiation of Human Embryonic Stem Cells on Three-Dimensional Polymer Scaffolds,” Proc. Natl. Acad. Sci. U.S.A,100(22), pp. 12741–12746. [CrossRef]
Vats, A., Bielby, R. C., Tolley, N., Dickinson, S. C., Boccaccini, A. R., Hollander, A. P., Bishop, A. E., and Polak, J. M., 2006, “Chondrogenic Differentiation of Human Embryonic Stem Cells: The Effect of the Micro-Environment,” Tissue Eng., 12(6), pp. 1687–1697. [CrossRef] [PubMed]
Hoben, G. M., Koay, E. J., and Athanasiou, K. A., 2008, “Fibrochondrogenesis in Two Embryonic Stem Cell Lines: Effects of Differentiation Timelines,” Stem Cells, 26(2), pp. 422–430. [CrossRef] [PubMed]
Lee, J. K., Responte, D. J., Cissell, D. D., Hu, J. C., Nolta, J. A., and Athanasiou, K. A., 2014, “Clinical Translation of Stem Cells: Insight for Cartilage Therapies,” Crit. Rev. Biotechnol., 34(1), pp. 89–100. [CrossRef] [PubMed]
Sanchez-Adams, J., and Athanasiou, K. A., 2012, “Dermis Isolated Adult Stem Cells for Cartilage Tissue Engineering,” Biomaterials, 33(1), pp. 109–119. [CrossRef] [PubMed]
French, M. M., Rose, S., Canseco, J., and Athanasiou, K. A., 2004, “Chondrogenic Differentiation of Adult Dermal Fibroblasts,” Ann. Biomed. Eng., 32(1), pp. 50–56. [CrossRef] [PubMed]
Deng, Y., Hu, J. C., and Athanasiou, K. A., 2007, “Isolation and Chondroinduction of a Dermis-Isolated, Aggrecan-Sensitive Subpopulation With High Chondrogenic Potential,” Arthritis Rheum., 56(1), pp. 168–176. [CrossRef] [PubMed]
Kalpakci, K. N., Brown, W. E., Hu, J. C., and Athanasiou, K. A., 2014, “Cartilage Tissue Engineering Using Dermis Isolated Adult Stem Cells: The Use of Hypoxia During Expansion Versus Chondrogenic Differentiation,” PloS One, 9(5), p. e98570. [CrossRef] [PubMed]
Athens, A. A., Makris, E. A., and Hu, J. C., 2013, “Induced Collagen Cross-Links Enhance Cartilage Integration,” PloS One, 8(4), p. e60719. [CrossRef] [PubMed]
Huey, D. J., Hu, J. C., and Athanasiou, K. A., 2012, “Unlike Bone, Cartilage Regeneration Remains Elusive,” Science, 338(6109), pp. 917–921. [CrossRef] [PubMed]

Figures

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Fig. 1

The evolution of a comprehensive, multidisciplinary, and multiscale approach to elucidate cartilage physiology, pathology, and regeneration motivated by biomechanics

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Fig. 2

The single chondrocyte approach to elucidate mechanotransduction pathways and to select biomechanical forces as exogenous stimuli for tissue engineering strategies

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Fig. 3

Nucleus images of chondrocytes compressed at 0, 25, 50, and 100 nN (left to right). Transcriptional changes may be a direct result of conformational changes of chromatin. (Figure adapted from Leipzig and Athanasiou [19]).

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Fig. 4

Aggrecan gene expression levels for single chondrocytes subjected to various static compression conditions. There is a dose-dependent catabolic response of single cells to the applied static force. The growth factor, however, seems to offer a mechanoprotective effect. (Figure adapted from Leipzig and Athanasiou [19]).

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Fig. 5

The self-assembling process results in cartilage with clinically relevant dimensions. (Figure adapted from Hu and Athanasiou [62]).

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Fig. 6

In the first phase of the self-assembling process, chondrocytes were seeded at high density in a nonadherent agarose mold. In the second phase, cells began to aggregate following the differential adhesion hypothesis, which states that maximized intercellular adhesion occurs when the total free energy of the forming neocartilage is minimized. Tissue formation occurs during the self-assembling process via only cell–cell interactions, whereas in scaffold-based approaches it is achieved via cell-scaffold interactions. In the third phase, neotissue begins to form, and cells migrate apart and secrete ECM. In the fourth phase, neocartilage exhibits significant maturation, including distinct pericellular ECM formation and localization of collagen type VI. (Figure adapted from Athanasiou et al. [63]).

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Fig. 7

Tensile stiffness (A) and total collagen normalized to neocartilage WW (B). All three treatments resulted in an ∼95% increase in tensile stiffness compared with control, while groups treated with ouabain contained significantly more total collagen per wet weight than controls. Bars show the mean and SD. *= p < 0.05. (Figure adapted from Natoli et al. [69]).

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Fig. 8

C-ABC treatment increased both the tensile modulus and tensile strength of self-assembling neocartilage. (Figure adapted from Natoli et al. [73]).

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Fig. 9

Combination of hydrostatic pressure stimulation and growth factors increased compressive (a) and tensile (b) properties of tissue-engineered cartilage. (Figure adapted from Elder and Athanasiou [78]).

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Fig. 10

Collagen alignment, as confirmed by polarized microscopy, contributes to (a) a higher aggregate modulus (p  <  0.05) for articular neocartilage confined for 2 weeks; and (b) a threefold increase in circumferential, compared to radial, tensile modulus in meniscus neocartilage. (Figure adapted from Aufderheide and Athanasiou [83]).

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Fig. 11

Collagen type I, which is not normally expressed in cartilage, becomes more common as articular chondrocytes are passaged. In contrast, collagen type II and superficial SZP expression decreases precipitously after 1–2 passages, while aggrecan expression remains relatively constant. (Figure adapted from Darling and Athanasiou [89]).

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Fig. 12

The modular approach consists of chondrogenic differentiation of hESCs, followed by tissue engineering of the chondrocyte-like cells

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Fig. 13

Hypoxic conditions increased staining for collagen, indicating increased chondrogenesis. (Figure adapted from Koay and Athanasiou [90]).

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Fig. 14

Using coculture with a fibrochondrocyte feeder layer results in improved chondrogenic differentiation of hESCs. (Figure adapted from Hoben et al. [91]).

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Fig. 15

Cell line dramatically affects neocartilage tensile properties. (Figure adapted from Hoben et al. [94]).

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Fig. 16

Histological assessment of DIAS cell neocartilage treated with IGF-I, BMP-2, or TGF-β1, shows staining for collagen type I, II, and VI, indicated chondrogenic ECM content. (Figure adapted from Sanchez-Adams and Athanasiou [96]).

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Fig. 17

In Vivo integration interface histological and mechanical assessment shows (a) increased tissue integration and collagen staining, (b) increased tensile (Young's) modulus, and (c) increased interfacial tensile strength in LOX, C-ABC, and TGF-β1 treated groups, specifically the group that received a double LOX treatment. (Figure adapted from Makris et al. [82]).

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