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Depth-dependent biomechanical and biochemical properties of fetal, newborn, and tissue-engineered articular cartilage
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Journal of Biomechanics 40 (2007) 182–190 www.elsevier.com/locate/jbiomech www.JBiomech.com
Depth-dependent biomechanical and biochemical properties of fetal, newborn, and tissue-engineered articular cartilage Travis J. Kleina, Manu Chaudhrya, Won C. Baea, Robert L. Saha,b, a
Department of Bioengineering, 9500 Gilman Dr., Mail Code 0412, University of California, San Diego, La Jolla, CA 92093-0412, USA b Whitaker Institute of Biomedical Engineering, University of California, San Diego, La Jolla, CA, USA Accepted 26 October 2005
Abstract Adult articular cartilage has depth-dependent mechanical and biochemical properties which contribute to zone-specific functions. The compressive moduli of immature cartilage and tissue-engineered cartilage are known to be lower than those of adult cartilage. The objective of this study was to determine if such tissues exhibit depth-dependent compressive properties, and how these depthvarying properties were correlated with cell and matrix composition of the tissue. The compressive moduli of fetal and newborn bovine articular cartilage increased with depth (po0:05) by a factor of 4–5 from the top 0.1 mm (28713 kPa, 141710 kPa, respectively) to 1 mm deep into the tissue. Likewise, the glycosaminoglycan and collagen content increased with depth (both po0:001), and correlated with the modulus (both po0:01). In contrast, tissue-engineered cartilage formed by either layering or mixing cells from the superficial and middle zone of articular cartilage exhibited similarly soft regions at both construct surfaces, as exemplified by large equilibrium strains. The properties of immature cartilage may provide a template for developing tissueengineered cartilage which aims to repair cartilage defects by recapitulating the natural development and growth processes. These results suggest that while depth-dependent properties may be important to engineer into cartilage constructs, issues other than cell heterogeneity must be addressed to generate such tissues. r 2005 Elsevier Ltd. All rights reserved. Keywords: Cartilage; Tissue engineering; Compressive modulus; Depth-depedent properties; Proteoglycan; Collagen; Chondrocte; Growth
1. Introduction Adult articular cartilage is a highly organized tissue that covers the ends of long bones and functions as a low-friction, wear-resistant, load-bearing surface for efficient joint articulation. Chondrocytes vary in organization and phenotype with depth from the articular surface (Stockwell and Meachim, 1979), and are responsible for synthesizing and remodeling extracellular matrix that governs functional properties of the tissue. Near the surface, in the superficial zone, Corresponding author. Department of Bioengineering, 9500 Gilman Dr., Mail Code 0412, University of California, San Diego, La Jolla, CA 92093-0412, USA. Tel.: +1 858 534 0821; fax: +1 858 822 1614. E-mail address: [email protected] (R.L. Sah).
0021-9290/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.jbiomech.2005.11.002
chondrocytes are densely populated and have relatively low metabolic rates, but uniquely secrete molecules that may aid in boundary lubrication, including proteoglycan 4 (PRG4) (Jay et al., 2001; Schumacher et al., 1994). Deeper into the tissue, in the middle and deep zones, chondrocyte density decreases, while biosynthesis rates of matrix molecules, including type II collagen and aggrecan, increase (Poole et al., 1989; Wong et al., 1996). Associated with the zonal cellular variations are changes in the content, organization, and mechanical properties of the extracellular matrix (Maroudas, 1979). Proteoglycan content increases with depth and is primarily responsible for the depth-increasing compressive modulus (Maroudas, 1979; Schinagl et al., 1997). Collagen content remains fairly constant with depth, but fiber orientation varies from parallel to the surface in the
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superficial zone, to perpendicular in the deep zone (Hunziker et al., 1997). The collagen network is primarily responsible for tensile and shear properties, especially in the superficial zone (Bank et al., 1998; LeRoux et al., 2000), but also may contribute to compressive properties (Williamson et al., 2001). During development and growth, articular cartilage is subjected to increasing loads, and exhibits alteration in matrix content, organization, and mechanical properties. In utero, articular cartilage appears as a relatively homogeneous, very dense mixture of cells in a highly hydrated matrix of proteoglycan and collagen. The matrix is much softer than in mature tissue (Williamson et al., 2001), and the collagen network is organized differently (Bank et al., 1998; Keene et al., 1995). While bulk cartilage mechanical properties change markedly, depth-dependent compressive properties (Wang et al., 2003) have not been ascertained in developing articular cartilage at different stages of development and growth. Recapitulating development is one approach to engineering articular cartilage for the repair of cartilage defects (Sah et al., 2004). Focus has been to generate soft tissues with high cell density, with hopes that the tissue will be better able to integrate with surrounding cartilage (Obradovic et al., 2001) and mature in vivo. Little attention has been given to engineering tissues with depth-dependent properties, although such properties may be present in developing cartilage and may be functionally important. One method of fabricating tissues with depth-dependent properties involves layering chondrocytes isolated from different zones (Kim et al., 2003; Klein et al., 2003). When cells from the superficial layer were seeded atop cells from the middle layer, constructs exhibited localized expression of PRG4, and full-thickness compressive properties between those of constructs made from cells of the individual layers (Klein et al., 2003). However, the effects of creating stratified tissues on depth-dependent mechanical properties were not determined. Several methods have been used to determine depthdependent biochemical and biomechanical properties of cartilaginous tissues. Cartilage has been sectioned parallel to the articular surface, digested, and analyzed for cartilage components including cells, proteoglycan, and collagen, with resolution of microtome sections (0.1 mm) (Maroudas, 1979). Quantitative histological methods (Kiviranta et al., 1985) improve the resolution over the section-digest method, but lack the versatility to correlate multiple biochemical components to biomechanical properties. Biomechanical studies have been made on cartilage explants from different depths, but the resolution is limited to the thickness of the sample (typically, 40.25 mm (Chen et al., 2004)) and excessive cutting could lead to artifacts due to collagen network disruption or loss of molecules near the cut surface. These limitations have been overcome by imaging
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samples during compression tests, and determining strain using computer-aided tracking of chondrocytes (Schinagl et al., 1996, 1997), or digital image correlation (Wang et al., 2002). Image correlation methods may be applied to determine depth-varying compressive properties of both native and tissue-engineered cartilage. The objectives of the study were (1) to determine the depth-dependent compressive modulus of the articular surface region (top 1 mm) of cartilage from fetal and newborn bovines, (2) to determine the effects of layering subpopulations on the relative depth-dependent properties of tissue-engineered cartilage, and (3) to correlate the local cartilage mechanical properties with local cell and matrix content.
2. Materials and methods 2.1. Immature cartilage harvest and preparation Cartilage specimens from the surface region of fetal and newborn bovine knees were harvested and prepared for analysis of depth-varying mechanical and biochemical properties. Osteochondral cores (9 mm diameter) were obtained from the femoral condyles of fetal (n ¼ 4, gestational age 244713 days) and newborn (n ¼ 4, 2–3 week old) bovine knees using the Osteochondral Autograft Transfer System (Arthrex, Naples, FL). Punches (3 mm diameter) were made adjacent to the 9 mm cores using a dermal punch. Cores were incubated in phosphate buffered saline (PBS) with protease inhibitors (+PIs, 2 mM EDTA, 10 mM N-ethylmaleimide, 5 mM benzamidine-HCl, 1 mM PMSF) (Frank et al., 1987) at 4 1C for 1 h. Cores were then placed with the articular surface touching a flat optimal cutting temperature compound (OCT, Fisher Scientific, Tustin, CA) surface, covered with OCT, frozen, and cryosectioned until the remaining cartilage was 1.0 mm thick, with an intact articular surface. The cartilage disk was removed from OCT by immersing in an excess volume of PBS+PIs, and then frozen at 20 1C. One hour prior to biomechanical testing, the 9 mm disk was cut into a 6.4 mm half-disk (diameter secant to the 9 mm disk) and a 3.2 mm disk using custom punches and a razor blade. 2.2. Construct formation Tissue-engineered cartilage constructs were formed using cells from different cartilage layers, and prepared for biomechanical analysis. Cartilage constructs were formed by mixing (n ¼ 3) or layering (n ¼ 3) cells from the superficial (0–0.2 mm) and middle zones (0.8–1.6 mm) of newborn bovine articular cartilage, using a modified alginate-recovered chondrocyte (ARC) method (Masuda et al., 2003), essentially as described previously (Klein et al., 2003). Briefly, cells
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encapsulated in 1.2% alginate and cultured for 1 week in DMEM/F12 with additives (10% fetal bovine serum, 25 mg/ml ascorbic acid, 100 U/ml penicillin, 100 mg/ml streptomycin, 0.25 mg/ml amphotericin B, 0.1 mM MEM non-essential amino acids, 0.4 mM L-proline, 2 mM L-glutamine), with medium changed daily (1 ml/(million cells day)), to develop a pericellular matrix. Cells with their associated matrix were then released from alginate with 55 mM sodium citrate in 150 mM NaCl, and seeded in a 1:1 ratio (5 106 cells/cm2) onto 0.4 mm pore-size, 6.5 mm Transwells inserts (Corning Inc., Corning, NY). Constructs were cultured for 12 additional days with daily medium changes, removed from the membrane, halved, and analyzed for strain. 2.3. Biomechanical analyses Immature cartilage and tissue-engineered cartilage were analyzed for depth-dependent biomechanical properties by monitoring deformation on a microscope during compressive loading. To facilitate cell tracking, the cell nuclei of 6.4 mm half-disks were labeled fluorescently by immersing the tissue in PBS+PIs with 20 mM propidium iodide for 1 h at 4 1C. Samples were then rinsed in PBS+PIs and mounted into a custom confining chamber on an inverted microscope, between two porous stainless steel platens (Chen et al., 2001). The cartilage or construct was compressed 0%, 10%, and 20% of the initial thickness, to encompass in vivo compression levels (Eckstein et al., 2000) and similar in vitro tests (Wang et al., 2003), at a rate of 0.02%/s (Schinagl et al., 1997), while monitoring load. Fluorescence images were taken using a Nikon D70 camera and a 4X objective (2.3 1.5 mm field of view, 0.79 mm/pixel resolution) at each compression level after 30 min of stress-relaxation, a time sufficient to reach equilibrium (stress change of less than 0.001 MPa in 5 min). Image contrast was enhanced with the unsharp mask filter in Adobe Photoshop, and images were correlated using VIC2D (Correlated Solutions, Inc. West Columbia, SC) to determine the two-dimensional Lagrangian strain during compression (Sutton et al., 2000), using the 0% compression image as a reference. Regions that did not correlate (strain not determined) were excluded from further analysis, and intra-tissue axial strain data (Ezz) were averaged over 0.1 mm-thick regions (175 points) for each specimen to give a depth-profile of strain. To determine the compressive modulus of immature cartilage, 3.2 mm disks were placed in a radially confining chamber filled with PBS and compressed 0%, 10%, 20%, and 30% of the initial thickness, using the above stress-relaxation protocol on a mechanical spectrometer (Chen et al., 2001). Equilibrium stress– strain data were fit to a non-linear model (Kwan et al., 1990) to determine the confined compressive modulus for the whole tissue using platen–platen strain (including
30%, where non-linearities are apparent), and for 0.1 mm thick regions using intra-tissue strain. 2.4. Biochemical analyses The cell and matrix content of immature cartilage were determined on sequential sections from the articular surface. 3 mm punches were frozen in OCT and sectioned in 70 mm increments to 1 mm deep into the tissue. Slices were removed from OCT and digested overnight with 0.5 mg/ml Proteinase K at 60 1C. Tubes were inspected to ensure complete digestion, and digest solutions were analyzed for DNA as a measure of cellularity (7.7 pg DNA/cell) (McGowan et al., 2002), glycosaminoglycan (GAG) (Farndale et al., 1986), and hydroxyproline as a measure of collagen (7.25 g collagen/g hydroxyproline) (Pal et al., 1981), and normalized to the calculated volume of the slices (1.0 ml). 2.5. Statistical analysis Data are presented as mean7SEM. Biomechanical (Ezz, HA0) and biochemical (GAG, collagen, cells) data measured at several depths for each specimen were analyzed by ANOVA (a ¼ 0:05) for effects of developmental stage (fetal, newborn) or construct type (mixed, layered), and depth (repeated measure). When significant effects of depth were detected by ANOVA, posthoc tests with Bonferroni corrections were performed to determine which depths exhibited significantly higher strain than the other depths within a given tissue type. Additionally, fetal and newborn cartilage data were pooled and analyzed by univariate and multivariate regressions to determine if the local compressive modulus and local biochemical properties are associated in immature cartilage. All statistical analyses were performed with Systat version 10.2 (Systat Software, Inc., Richmond, CA).
3. Results 3.1. Immature cartilage mechanical properties Fetal and newborn articular cartilage equilibrium compressive moduli increased with depth from the articular surface. The top 1 mm of developing articular cartilage (Fig. 1) displayed non-linear stress–strain curve when analyzed as a whole (Fig. 2A). The aggregate modulus was significantly lower for fetal (89732 kPa) than newborn bovine cartilage (197721 kPa, po0:05). Intra-tissue strain of both fetal and newborn cartilage depended on depth from the articular surface (both po0:01, Fig. 2B), with the top 0.3 mm compressing significantly more than deeper layers (both po0:05).
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Fig. 1. Epifluourescent images of the top 1 mm of (A–C) 3rd trimester fetal bovine and (D–F) 2–3 week-old bovine calf articular cartilage in confined compression at (A, D) 0%, (B, E) 10%, and (C, F) 20% strain. The coordinate frame is indicated in panel A, and the articular surface is indicated in panels C and F.
Correspondingly, the confined compressive modulus was depth-dependent (po0:01, Fig. 2C), with the softest regions near the articular surface and the deepest regions 4–5-fold stiffer. 3.2. Tissue-engineered cartilage mechanical properties In contrast to the immature cartilage, the tissueengineered cartilage constructs displayed regions that were soft at both surfaces and stiffer in the center of the tissue. Constructs formed by mixing or layering chondrocyte subpopulations developed similar thickness (mixed 0.9970.04 mm, layered 1.0770.03, p ¼ 0:13) and strain profiles with depth (p ¼ 0:6). Unlike the native cartilage (Fig. 2B), the constructs showed regions of high strain at both surfaces, indicating relatively soft tissue, with regions in the top and bottom 0.2 mm straining more than the center of the tissue (po0:01, Fig. 3). 3.3. Cartilage biochemical properties and correlations Cartilage matrix constituents increased with depth from the articular surface of fetal and newborn cartilage, and correlated with mechanical properties. There was a general trend of increasing GAG (po0:001, Fig. 4A) and collagen (po0:001, Fig. 4B) content per volume from the superficial to the deeper layers. Both constituents were higher in the calf than the fetal cartilage (po0:001), and the increase with depth was greater for the calf. Cell density decreased similarly with depth for fetal and calf cartilage (both po0:001, Fig. 4C), falling from 180 million cells/cm3 in the top 0.1 mm to a plateau at 80 million cells/cm3 by 0.5 mm into the tissue.
The compressive modulus of immature bovine cartilage (fetal and newborn) was positively associated with GAG and collagen content in univariate regressions (both po0:01, Fig. 5A and B). The strongest positive correlation was with GAG content (r2 ¼ 0:45), with weaker correlation with collagen (r2 ¼ 0:17), and negative correlation with cell content (po0:01, r2 ¼ 0:22, Fig. 5C). Multivariate regressions on combined data from fetal and newborn cartilage suggest that the modulus depends on both the GAG and cell content (r2 ¼ 0:55, both po0:001), but not collagen content (p ¼ 0:3).
4. Discussion The results of this study suggest that there is a depthincreasing compressive modulus near the articular surface of immature articular cartilage, and that tissueengineered constructs differ from immature cartilage by exhibiting soft regions at both construct surfaces. Using image correlation methods with confined compression tests, the local compressive modulus was found to increase with depth in both fetal and newborn bovine cartilage (Fig. 2), similar to previous findings with mature articular cartilage (Schinagl et al., 1997). In contrast, constructs displayed symmetric soft surfaces (regions of high strain) regardless of how cells from the superficial or middle zone were seeded (Fig. 3). Biochemical analysis of cartilage sections indicated that matrix components, GAG and collagen, increase with depth and development, and that the cell content decreases from the articular surface, all with correlations to the local compressive properties (Figs. 3 and 4).
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Fig. 2. Mechanical properties of the top 1 mm of fetal (K) and newborn (m) bovine articular cartilage. (A) The stress is plotted versus the platen to platen (total) strain. (B) The local strain (Ezz) at 10% compression, and (C) confined compressive modulus (HA0) are plotted as a function of depth from the articular surface (n ¼ 4).
Several choices were made in the design of the experiments that may limit the interpretation of the results. First, while there were subtle differences between fetal and newborn tissues, such as greater absolute depth-variation in compressive modulus (Fig. 2C) and matrix components (Fig. 4A and B) in the newborn cartilage, the tissues were relatively close in age (1 month). Tissues from earlier stages of fetal development may not display such depth-dependent properties. Also, samples were from the surface 1 mm, and therefore these
Fig. 3. Epifluourescent images of constructs formed by (A–C) layering or (D–F) mixing chondrocytes from the superficial and middle zone of newborn bovine articular cartilage. Constructs are imaged at equilibrium in confined compression of (A, D) 0%, (B, E) 10%, and (C, F) 20% of the initial thickness. (G) Local strain (Ezz) at 20% compression is plotted for layered (E, n ¼ 3) and mixed (’, n ¼ 3) constructs. The coordinate frame is indicated in panel A and the membrane surface is indicated in panel F.
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Fig. 4. Depth-dependent biochemical properties of the top 1 mm of fetal (K) and newborn (m) bovine calf articular cartilage. The (A) glycosaminoglycan (GAG), (B) collagen (COL), and (C) cell number were normalized to the tissue volume of each cryosection bin and plotted as a function of depth from the articular surface (n ¼ 4).
data do not indicate how properties vary in the deep, vascularized regions. While full-thickness measurements may provide further insight into the development of depth-dependent mechanical properties, there is not a well-defined boundary for the deep zone in immature cartilage. The superficial 1 mm was chosen to include portions of the superficial and middle zones, as well as a
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region of tissue that is avascular, and of similar thickness to full-thickness adult bovine cartilage (Schinagl et al., 1997). Although the sample size was small, the variation in properties was large and demonstrated statistical significance. Further studies could examine different sites and tissue types, if more precise values were desired. However, depth-dependent compressive properties have been identified in cartilage from the knee (Schinagl et al., 1997), carpometacarpal (Wang et al., 2002), and shoulder (Wang et al., 2003) joints; thus, it appears depth-dependent properties are conserved across many joints. Finally, as in prior studies using image analysis to determine strain (Schinagl et al., 1996; Wang et al., 2003), measurements were taken at equilibrium, and thus do not provide information on the dynamic, viscoelastic response in the various regions relevant to joint kinematics, information that could be attained using ultrasound imaging methods (Zheng et al., 2005). Presence of depth-dependent mechanical properties in immature cartilage gives rise to questions of how these properties are developed. While articular cartilage appears relatively homogeneous at early developmental stages, differences between cells near the surface and those deeper are evident. For example, expression of PRG4 has been documented in the cells near the articular surface even in the first trimester (Schumacher et al., 1994), and may have importance in joint cavity formation (Flannery et al., 1999). Increases in matrix content with depth from the articular surface (Fig. 4A and B) may indicate a greater biosynthetic rate by cells further from the articular surface (Aydelotte et al., 1988). Additionally, changes in matrix organization, particularly collagen crosslinks, may affect the tissue’s mechanical properties (Williamson et al., 2003). Possible explanations for zonal cellular differences could include genetic factors, mechanical stresses (Carter et al., 2004; Wong et al., 1997), or transport of nutrients and growth factors. The top 1 mm of immature cartilage is essentially avascular, suggesting that transport of nutrients and waste products is primarily through the articular surface. Thus, gradients of oxygen, growth factors, etc. develop from the superficial to the deep zone, and are likely to influence chondrocyte function (Grimshaw and Mason, 2001; Kellner et al., 2002). Additionally, transport and binding of matrix molecules can lead to biochemical properties that increase with depth, even in absence of direct effects of nutrient gradients on cellular metabolism (DiMicco and Sah, 2003). The depth-varying properties of immature cartilage have implications for normal joint function as well as cartilage tissue engineering. Depth-dependent mechanical properties serve several functions in mature articular cartilage. The soft superficial zone functions to increase contact area and distribute load (Mankin et al., 1994),
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Fig. 5. Relationship between the local compressive modulus and GAG, collagen, and cell content, per volume, in the top 1 mm of bovine fetal (K) and newborn (m) articular cartilage.
and increase interstitial fluid pressure to lower frictional properties at the surface (Krishnan et al., 2003, 2004). The functions of depth-dependent mechanical properties, especially in the superficial zone, and their presence in immature cartilage suggest that they are necessary for normal joint function. In osteoarthritis, the absence or disruption of the superficial zone reduces functionality of the tissue (Guilak, 1994). Cartilage tissue engineering aims to restore normal joint function and thus depthdependent mechanical properties may be required for a properly functioning cartilage replacement. Cartilage constructs created with the aim of attaining one soft and one stiffer layer by sequentially seeding cells from different zones, developed relatively soft regions near both surfaces (absolute mechanical properties were not determined, but could be obtained by measuring stress), similar to constructs formed by mixing the cells (Fig. 3). Diffusive loss of macromolecules from construct surfaces may be responsible for development of such a strain profile. Applying membranes with small poresizes (100 kDa molecular weight cut-off) at the construct surfaces has resulted in excellent matrix retention in a scaffold-free system (Mainil-Varlet et al., 2001). Tissueengineered cartilage formed with hydrogels or polymer scaffolds may exhibit strain profiles drastically different from tissues formed here. In both immature cartilage and tissue-engineered cartilage formed without a mechanically functional scaffold, there may be an influence of the cell component on the mechanical properties of the tissue. In addition to positive correlations of compressive modulus with matrix components, there was a weak negative correlation with cell content for immature cartilage (Fig. 5C). While the cell component is generally thought not to affect the mechanical properties of mature cartilage, where there is a stiff matrix and low cell density, the extracellular matrix is less stiff and the cell volume fraction can be relatively high in immature cartilage
(diameter ¼ 10.4 mm, cell density ¼ 2 108 cells/ml, volume fraction ¼ 0.12) (Wong et al., 1997). Thus, the cell component may have a marked effect on mechanical properties, when considering that the cell has an elastic modulus three orders of magnitude smaller than the extracellular matrix (Jones et al., 1999; Wu et al., 1999). The cell shape and orientation may also influence mechanical properties, but appear to be more homogeneous with depth in immature than in mature cartilage (Wu and Herzog, 2002). While there was no observed change in cellularity between the two developmental stages, there were increases in GAG and collagen content, which may explain the increase in compressive modulus. Although the tissue-engineered cartilage was not analyzed for cell content as a function of depth, the cell component is expected to influence mechanical properties early in the culture process, as constructs consist largely of cells, and relatively small amounts of newly elaborated matrix. Immature articular cartilage is often used as a model for cartilage tissue engineering, due to its ability to grow, develop, and repair (Namba et al., 1998; Obradovic et al., 2001). The levels of organization and maturation necessary to make a functional cartilage replacement are not fully established, but the presence of biochemical and biomechanical properties that vary with depth from the surface in articular cartilage before birth (Fig. 2B and C) implicate depth-dependent properties as important details to consider. Engineering tissues with strain-profiles similar to native articular cartilage may have the benefit of enhancing host–implant integration by reducing strain discontinuities and relative motion at interfaces. Approaches which use cells from different zones, vary scaffold properties, or use bioreactor stimulation to engineer cartilage with depth-dependent properties may benefit from analytical methods such as image correlation to verify that these properties are attained (Ng et al., 2005), as there is not a
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simple correspondence between the properties of the cells (Fig. 3) or scaffold and the local mechanical properties developed after time in culture.
Acknowledgments This work was supported by grants from NASA, NIH, and NSF.
References Aydelotte, M.B., Greenhill, R.R., Kuettner, K.E., 1988. Differences between sub-populations of cultured bovine articular chondrocytes. II. Proteoglycan metabolism. Connective Tissue Research 18, 223–234. Bank, R.A., Bayliss, M.T., Lafeber, F.P., Maroudas, A., Tekoppele, J.M., 1998. Ageing and zonal variation in post-translational modification of collagen in normal human articular cartilage. The age-related increase in non-enzymatic glycation affects biomechanical properties of cartilage. Biochemical Journal 330, 345–351. Carter, D.R., Beaupre, G.S., Wong, M., Smith, R.L., Andriacchi, T.P., Schurman, D.J., 2004. The mechanobiology of articular cartilage development and degeneration. Clinical Orthopaedics and Related Research 427, S69–S77. Chen, A.C., Bae, W.C., Schinagl, R.M., Sah, R.L., 2001. Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression. Journal of Biomechanics 34, 1–12. Chen, A.C., Klisch, S.M., Bae, W.C., Temple, M.M., McGowan, K.B., Gratz, K.R., Schumacher, B.L., Sah, R.L., 2004. Mechanical characterization of native and tissue-engineered cartilage. In: Sabatini, M., DeCeuninck, F., Pastoureau, P. (Eds.), Cartilage and Osteoarthritis. Humana Press, Totowa, NJ, pp. 157–190. DiMicco, M.A., Sah, R.L., 2003. Dependence of cartilage matrix composition on biosynthesis, diffusion, and reaction. Transport in Porous Media 50, 57–73. Eckstein, F., Lemberger, B., Stammberger, T., Englmeier, K.H., Reiser, M., 2000. Patellar cartilage deformation in vivo after static versus dynamic loading. Journal of Biomechanics 33, 819–825. Farndale, R.W., Buttle, D.J., Barrett, A.J., 1986. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochimica et Biophysica Acta 883, 173–177. Flannery, C.R., Hughes, C.E., Schumacher, B.L., Tudor, D., Aydelotte, M.B., Kuettner, K.E., Caterson, B., 1999. Articular cartilage superficial zone protein (SZP) is homologous to megakaryocyte stimulating factor precursor and is a multifunctional proteoglycan with potential growth-promoting, cytoprotective, and lubricating properties in cartilage metabolism. Biochemical and Biophysical Research Communications 254, 535–541. Frank, E.H., Grodzinsky, A.J., Koob, T.J., Eyre, D.R., 1987. Streaming potentials: a sensitive index of enzymatic degradation in articular cartilage. Journal of Orthopaedic Research 5, 497–508. Grimshaw, M.J., Mason, R.M., 2001. Modulation of bovine articular chondrocyte gene expression in vitro by oxygen tension. Osteoarthritis and Cartilage 9, 357–364. Guilak, F., 1994. Volume and surface area measurement of viable chondrocytes in situ using geometric modelling of serial confocal sections. Journal of Microscopy 173, 245–256. Hunziker, E.B., Michel, M., Studer, D., 1997. Ultrastructure of adult human articular cartilage matrix after cryotechnical processing. Microscopy Research and Technique 37, 271–284.
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Jay, G.D., Tantravahi, U., Britt, D.E., Barrach, H.J., Cha, C.J., 2001. Homology of lubricin and superficial zone protein (SZP): products of megakaryocyte stimulating factor (MSF) gene expression by human synovial fibroblasts and articular chondrocytes localized to chromosome 1q25. Journal of Orthopaedic Research 19, 677–687. Jones, W.R., Ting-Beall, H.P., Lee, G.M., Kelley, S.S., Hochmuth, R.M., Guilak, F., 1999. Alterations in the Young’s modulus and volumetric properties of chondrocytes isolated from normal and osteoarthritic human cartilage. Journal of Biomechanics 32, 119–127. Keene, D.R., Oxford, J.T., Morris, N.P., 1995. Ultrastructural localization of collagen types II, IX, and XI in the growth plate of human rib and fetal bovine epiphyseal cartilage: type XI collagen is restricted to thin fibrils. Journal of Histochemistry and Cytochemistry 43, 967–979. Kellner, K., Liebsch, G., Klimant, I., Wolfbeis, O.S., Blunk, T., Schulz, M.B., Gopferich, A., 2002. Determination of oxygen gradients in engineered tissue using a fluorescent sensor. Biotechnology and Bioengineering 80, 73–83. Kim, T.K., Sharma, B., Williams, C.G., Ruffner, M.A., Malik, A., McFarland, E.G., Elisseeff, J.H., 2003. Experimental model for cartilage tissue engineering to regenerate the zonal organization of articular cartilage. Osteoarthritis and Cartilage 11, 653–664. Kiviranta, I., Jurvelin, J., Tammi, M., Saamanen, A.-M., Helminen, H.J., 1985. Microspectrophotometric quantitation of glycosaminoglycans in articular cartilage sections stained with Safranin O. Histochemistry 82, 249–255. Klein, T.J., Schumacher, B.L., Schmidt, T.A., Li, K.W., Voegtline, M.S., Masuda, K., Thonar, E.J.-M.A., Sah, R.L., 2003. Tissue engineering of articular cartilage with stratification using chondrocyte subpopulations. Osteoarthritis and Cartilage 11, 595–602. Krishnan, R., Park, S., Eckstein, F., Ateshian, G.A., 2003. Inhomogeneous cartilage properties enhance superficial interstitial fluid support and frictional properties, but do not provide a homogeneous state of stress. Journal of Biomechanical Engineering 125, 569–577. Krishnan, R., Kopacz, M., Ateshian, G.A., 2004. Experimental verification of the role of interstitial fluid pressurization in cartilage lubrication. Journal of Orthopaedic Research 22, 565–570. Kwan, M.K., Lai, W.M., Mow, V.C., 1990. A finite deformation theory for cartilage and other soft hydrated connective tissues—I. Equilibrium results. Journal of Biomechanics 23, 145–155. LeRoux, M.A., Arokoski, J., Vail, T.P., Guilak, F., Hyttinen, M.M., Kirivanta, K., Setton, L.A., 2000. Simultaneous changes in the mechanical properties, quantitative collagen organization, and proteoglycan concentration of articular cartilage following canine meniscectomy. Journal of Orthopaedic Research 18, 383–392. Mainil-Varlet, P., Rieser, F., Grogan, S., Mueller, W., Saager, C., Jakob, R.P., 2001. Articular cartilage repair using a tissueengineered cartilage-like implant: an animal study. Osteoarthritis and Cartilage 9 (Suppl A), S6–S15. Mankin, H.J., Mow, V.C., Buckwalter, J.A., Iannotti, J.P., Ratcliffe, A., 1994. Form and function of articular cartilage. In: Simon, S.R. (Ed.), Orthopaedic Basic Science. American Academy of Orthopaedic Surgeons, Rosemont, IL, pp. 1–44. Maroudas, A., 1979. Physico-chemical properties of articular cartilage. In: Freeman, M.A.R. (Ed.), Adult Articular Cartilage. Pitman Medical, Tunbridge Wells, England, pp. 215–290. Masuda, K., Sah, R.L., Hejna, M.J., Thonar, E.J., 2003. A novel twostep method for the formation of tissue-engineered cartilage by mature bovine chondrocytes: the alginate-recovered- chondrocyte (ARC) method. Journal of Orthopaedic Research 21, 139–148. McGowan, K.B., Kurtis, M.S., Lottman, L.M., Watson, D., Sah, R.L., 2002. Biochemical quantification of DNA in human articular and septal cartilage using PicoGreens and Hoechst 33258. Osteoarthritis and Cartilage 10, 580–587.
ARTICLE IN PRESS 190
T.J. Klein et al. / Journal of Biomechanics 40 (2007) 182–190
Namba, R.S., Meuli, M., Sullivan, K.M., Le, A., Adzick, N.S., 1998. Spontaneous repair of superficial defects in articular cartilage in a fetal lamb model. Journal of Bone and Joint Surgery American volume 80-A, 4–10. Ng, K.W., Wang, C.C., Mauck, R.L., Kelly, T.N., Chahine, N.O., Costa, K.D., Ateshian, G.A., Hung, C.T., 2005. A layered agarose approach to fabricate depth-dependent inhomogeneity in chondrocyte-seeded constructs. Journal of Orthopaedic Research 23, 134–141. Obradovic, B., Martin, I., Padera, R.F., Treppo, S., Freed, L.E., Vunjak-Novakovic, G., 2001. Integration of engineered cartilage. Journal of Orthopaedic Research 19, 1089–1097. Pal, S., Tang, L.-H., Choi, H., Habermann, E., Rosenberg, L., Roughley, P., Poole, A.R., 1981. Structural changes during development in bovine fetal epiphyseal cartilage. Collagen Related Research 1, 151–176. Poole, A.R., Matsui, Y., Hinek, A., Lee, E.R., 1989. Cartilage macromolecules and the calcification of cartilage matrix. The Anatomical Record 224, 167–179. Sah, R.L., Klein, T.J., Schmidt, T.A., Albrecht, D.R., Bae, W.C., Nugent, G.E., McGowan, K.B., Temple, M.M., Jadin, K.D., Schumacher, B.L., Chen, A.C., Sandy, J.D., 2004. Articular cartilage repair, regeneration, and replacement. In: Koopman, W.J. (Ed.), Arthritis and Allied Conditions: A Textbook of Rheumatology. Lippincott Williams & Wilkins, Philadelphia, pp. 2277–2301. Schinagl, R.M., Ting, M.K., Price, J.H., Sah, R.L., 1996. Video microscopy to quantitate the inhomogeneous equilibrium strain within articular cartilage during confined compression. Annals of Biomedical Engineering 24, 500–512. Schinagl, R.M., Gurskis, D., Chen, A.C., Sah, R.L., 1997. Depthdependent confined compression modulus of full-thickness bovine articular cartilage. Journal of Orthopaedic Research 15, 499–506. Schumacher, B.L., Block, J.A., Schmid, T.M., Aydelotte, M.B., Kuettner, K.E., 1994. A novel proteoglycan synthesized and secreted by chondrocytes of the superficial zone of articular cartilage. Archives of Biochemistry and Biophysics 311, 144–152. Stockwell, R.A., Meachim, G., 1979. The chondrocytes. In: Freeman, M.A.R. (Ed.), Adult Articular Cartilage. Pitman Medical, Tunbridge Wells, England, pp. 69–144.
Sutton, M.A., McNeill, S.R., Helm, J.D., Chao, Y.J., 2000. Advances in two-dimensional and three-dimensional computer vision. In: Rastogi, P.K. (Ed.), Photomechanics. Springer, New York, pp. 323–372. Wang, C.C., Deng, J.M., Ateshian, G.A., Hung, C.T., 2002. An automated approach for direct measurement of two-dimensional strain distributions within articular cartilage under unconfined compression. Journal of Biomechanical Engineering 124, 557–567. Wang, C.C., Chahine, N.O., Hung, C.T., Ateshian, G.A., 2003. Optical determination of anisotropic material properties of bovine articular cartilage in compression. Journal of Biomechanics 36, 339–353. Williamson, A.K., Chen, A.C., Sah, R.L., 2001. Compressive properties and function-composition relationships of developing bovine articular cartilage. Journal of Orthopaedic Research 19, 1113–1121. Williamson, A.K., Chen, A.C., Masuda, K., Thonar, E.J.-M.A., Sah, R.L., 2003. Tensile mechanical properties of bovine articular cartilage: variations with growth and relationships to collagen network components. Journal of Orthopaedic Research 21, 872–880. Wong, M., Wuethrich, P., Eggli, P., Hunziker, E., 1996. Zone-specific cell biosynthetic activity in mature bovine articular cartilage: a new method using confocal microscopic stereology and quantitative autoradiography. Journal of Orthopaedic Research 14, 424–432. Wong, M., Wuethrich, P., Buschmann, M.D., Eggli, P., Hunziker, E., 1997. Chondrocyte biosynthesis correlates with local tissue strain in statically compressed adult articular cartilage. Journal of Orthopaedic Research 15, 189–196. Wu, J.Z., Herzog, W., 2002. Elastic anisotropy of articular cartilage is associated with the microstructures of collagen fibers and chondrocytes. Journal of Biomechanics 35, 931–942. Wu, J.Z., Herzog, W., Epstein, M., 1999. Modelling of location- and time-dependent deformation of chondrocytes during cartilage loading. Journal of Biomechanics 32, 563–572. Zheng, Y.P., Niu, H.J., Arthur Mak, F.T., Huang, Y.P., 2005. Ultrasonic measurement of depth-dependent transient behaviors of articular cartilage under compression. Journal of Biomechanics 38, 1830–1837.
Journal of Biomechanics 40 (2007) 182–190 www.elsevier.com/locate/jbiomech www.JBiomech.com
Depth-dependent biomechanical and biochemical properties of fetal, newborn, and tissue-engineered articular cartilage Travis J. Kleina, Manu Chaudhrya, Won C. Baea, Robert L. Saha,b, a
Department of Bioengineering, 9500 Gilman Dr., Mail Code 0412, University of California, San Diego, La Jolla, CA 92093-0412, USA b Whitaker Institute of Biomedical Engineering, University of California, San Diego, La Jolla, CA, USA Accepted 26 October 2005
Abstract Adult articular cartilage has depth-dependent mechanical and biochemical properties which contribute to zone-specific functions. The compressive moduli of immature cartilage and tissue-engineered cartilage are known to be lower than those of adult cartilage. The objective of this study was to determine if such tissues exhibit depth-dependent compressive properties, and how these depthvarying properties were correlated with cell and matrix composition of the tissue. The compressive moduli of fetal and newborn bovine articular cartilage increased with depth (po0:05) by a factor of 4–5 from the top 0.1 mm (28713 kPa, 141710 kPa, respectively) to 1 mm deep into the tissue. Likewise, the glycosaminoglycan and collagen content increased with depth (both po0:001), and correlated with the modulus (both po0:01). In contrast, tissue-engineered cartilage formed by either layering or mixing cells from the superficial and middle zone of articular cartilage exhibited similarly soft regions at both construct surfaces, as exemplified by large equilibrium strains. The properties of immature cartilage may provide a template for developing tissueengineered cartilage which aims to repair cartilage defects by recapitulating the natural development and growth processes. These results suggest that while depth-dependent properties may be important to engineer into cartilage constructs, issues other than cell heterogeneity must be addressed to generate such tissues. r 2005 Elsevier Ltd. All rights reserved. Keywords: Cartilage; Tissue engineering; Compressive modulus; Depth-depedent properties; Proteoglycan; Collagen; Chondrocte; Growth
1. Introduction Adult articular cartilage is a highly organized tissue that covers the ends of long bones and functions as a low-friction, wear-resistant, load-bearing surface for efficient joint articulation. Chondrocytes vary in organization and phenotype with depth from the articular surface (Stockwell and Meachim, 1979), and are responsible for synthesizing and remodeling extracellular matrix that governs functional properties of the tissue. Near the surface, in the superficial zone, Corresponding author. Department of Bioengineering, 9500 Gilman Dr., Mail Code 0412, University of California, San Diego, La Jolla, CA 92093-0412, USA. Tel.: +1 858 534 0821; fax: +1 858 822 1614. E-mail address: [email protected] (R.L. Sah).
0021-9290/$ - see front matter r 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.jbiomech.2005.11.002
chondrocytes are densely populated and have relatively low metabolic rates, but uniquely secrete molecules that may aid in boundary lubrication, including proteoglycan 4 (PRG4) (Jay et al., 2001; Schumacher et al., 1994). Deeper into the tissue, in the middle and deep zones, chondrocyte density decreases, while biosynthesis rates of matrix molecules, including type II collagen and aggrecan, increase (Poole et al., 1989; Wong et al., 1996). Associated with the zonal cellular variations are changes in the content, organization, and mechanical properties of the extracellular matrix (Maroudas, 1979). Proteoglycan content increases with depth and is primarily responsible for the depth-increasing compressive modulus (Maroudas, 1979; Schinagl et al., 1997). Collagen content remains fairly constant with depth, but fiber orientation varies from parallel to the surface in the
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superficial zone, to perpendicular in the deep zone (Hunziker et al., 1997). The collagen network is primarily responsible for tensile and shear properties, especially in the superficial zone (Bank et al., 1998; LeRoux et al., 2000), but also may contribute to compressive properties (Williamson et al., 2001). During development and growth, articular cartilage is subjected to increasing loads, and exhibits alteration in matrix content, organization, and mechanical properties. In utero, articular cartilage appears as a relatively homogeneous, very dense mixture of cells in a highly hydrated matrix of proteoglycan and collagen. The matrix is much softer than in mature tissue (Williamson et al., 2001), and the collagen network is organized differently (Bank et al., 1998; Keene et al., 1995). While bulk cartilage mechanical properties change markedly, depth-dependent compressive properties (Wang et al., 2003) have not been ascertained in developing articular cartilage at different stages of development and growth. Recapitulating development is one approach to engineering articular cartilage for the repair of cartilage defects (Sah et al., 2004). Focus has been to generate soft tissues with high cell density, with hopes that the tissue will be better able to integrate with surrounding cartilage (Obradovic et al., 2001) and mature in vivo. Little attention has been given to engineering tissues with depth-dependent properties, although such properties may be present in developing cartilage and may be functionally important. One method of fabricating tissues with depth-dependent properties involves layering chondrocytes isolated from different zones (Kim et al., 2003; Klein et al., 2003). When cells from the superficial layer were seeded atop cells from the middle layer, constructs exhibited localized expression of PRG4, and full-thickness compressive properties between those of constructs made from cells of the individual layers (Klein et al., 2003). However, the effects of creating stratified tissues on depth-dependent mechanical properties were not determined. Several methods have been used to determine depthdependent biochemical and biomechanical properties of cartilaginous tissues. Cartilage has been sectioned parallel to the articular surface, digested, and analyzed for cartilage components including cells, proteoglycan, and collagen, with resolution of microtome sections (0.1 mm) (Maroudas, 1979). Quantitative histological methods (Kiviranta et al., 1985) improve the resolution over the section-digest method, but lack the versatility to correlate multiple biochemical components to biomechanical properties. Biomechanical studies have been made on cartilage explants from different depths, but the resolution is limited to the thickness of the sample (typically, 40.25 mm (Chen et al., 2004)) and excessive cutting could lead to artifacts due to collagen network disruption or loss of molecules near the cut surface. These limitations have been overcome by imaging
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samples during compression tests, and determining strain using computer-aided tracking of chondrocytes (Schinagl et al., 1996, 1997), or digital image correlation (Wang et al., 2002). Image correlation methods may be applied to determine depth-varying compressive properties of both native and tissue-engineered cartilage. The objectives of the study were (1) to determine the depth-dependent compressive modulus of the articular surface region (top 1 mm) of cartilage from fetal and newborn bovines, (2) to determine the effects of layering subpopulations on the relative depth-dependent properties of tissue-engineered cartilage, and (3) to correlate the local cartilage mechanical properties with local cell and matrix content.
2. Materials and methods 2.1. Immature cartilage harvest and preparation Cartilage specimens from the surface region of fetal and newborn bovine knees were harvested and prepared for analysis of depth-varying mechanical and biochemical properties. Osteochondral cores (9 mm diameter) were obtained from the femoral condyles of fetal (n ¼ 4, gestational age 244713 days) and newborn (n ¼ 4, 2–3 week old) bovine knees using the Osteochondral Autograft Transfer System (Arthrex, Naples, FL). Punches (3 mm diameter) were made adjacent to the 9 mm cores using a dermal punch. Cores were incubated in phosphate buffered saline (PBS) with protease inhibitors (+PIs, 2 mM EDTA, 10 mM N-ethylmaleimide, 5 mM benzamidine-HCl, 1 mM PMSF) (Frank et al., 1987) at 4 1C for 1 h. Cores were then placed with the articular surface touching a flat optimal cutting temperature compound (OCT, Fisher Scientific, Tustin, CA) surface, covered with OCT, frozen, and cryosectioned until the remaining cartilage was 1.0 mm thick, with an intact articular surface. The cartilage disk was removed from OCT by immersing in an excess volume of PBS+PIs, and then frozen at 20 1C. One hour prior to biomechanical testing, the 9 mm disk was cut into a 6.4 mm half-disk (diameter secant to the 9 mm disk) and a 3.2 mm disk using custom punches and a razor blade. 2.2. Construct formation Tissue-engineered cartilage constructs were formed using cells from different cartilage layers, and prepared for biomechanical analysis. Cartilage constructs were formed by mixing (n ¼ 3) or layering (n ¼ 3) cells from the superficial (0–0.2 mm) and middle zones (0.8–1.6 mm) of newborn bovine articular cartilage, using a modified alginate-recovered chondrocyte (ARC) method (Masuda et al., 2003), essentially as described previously (Klein et al., 2003). Briefly, cells
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encapsulated in 1.2% alginate and cultured for 1 week in DMEM/F12 with additives (10% fetal bovine serum, 25 mg/ml ascorbic acid, 100 U/ml penicillin, 100 mg/ml streptomycin, 0.25 mg/ml amphotericin B, 0.1 mM MEM non-essential amino acids, 0.4 mM L-proline, 2 mM L-glutamine), with medium changed daily (1 ml/(million cells day)), to develop a pericellular matrix. Cells with their associated matrix were then released from alginate with 55 mM sodium citrate in 150 mM NaCl, and seeded in a 1:1 ratio (5 106 cells/cm2) onto 0.4 mm pore-size, 6.5 mm Transwells inserts (Corning Inc., Corning, NY). Constructs were cultured for 12 additional days with daily medium changes, removed from the membrane, halved, and analyzed for strain. 2.3. Biomechanical analyses Immature cartilage and tissue-engineered cartilage were analyzed for depth-dependent biomechanical properties by monitoring deformation on a microscope during compressive loading. To facilitate cell tracking, the cell nuclei of 6.4 mm half-disks were labeled fluorescently by immersing the tissue in PBS+PIs with 20 mM propidium iodide for 1 h at 4 1C. Samples were then rinsed in PBS+PIs and mounted into a custom confining chamber on an inverted microscope, between two porous stainless steel platens (Chen et al., 2001). The cartilage or construct was compressed 0%, 10%, and 20% of the initial thickness, to encompass in vivo compression levels (Eckstein et al., 2000) and similar in vitro tests (Wang et al., 2003), at a rate of 0.02%/s (Schinagl et al., 1997), while monitoring load. Fluorescence images were taken using a Nikon D70 camera and a 4X objective (2.3 1.5 mm field of view, 0.79 mm/pixel resolution) at each compression level after 30 min of stress-relaxation, a time sufficient to reach equilibrium (stress change of less than 0.001 MPa in 5 min). Image contrast was enhanced with the unsharp mask filter in Adobe Photoshop, and images were correlated using VIC2D (Correlated Solutions, Inc. West Columbia, SC) to determine the two-dimensional Lagrangian strain during compression (Sutton et al., 2000), using the 0% compression image as a reference. Regions that did not correlate (strain not determined) were excluded from further analysis, and intra-tissue axial strain data (Ezz) were averaged over 0.1 mm-thick regions (175 points) for each specimen to give a depth-profile of strain. To determine the compressive modulus of immature cartilage, 3.2 mm disks were placed in a radially confining chamber filled with PBS and compressed 0%, 10%, 20%, and 30% of the initial thickness, using the above stress-relaxation protocol on a mechanical spectrometer (Chen et al., 2001). Equilibrium stress– strain data were fit to a non-linear model (Kwan et al., 1990) to determine the confined compressive modulus for the whole tissue using platen–platen strain (including
30%, where non-linearities are apparent), and for 0.1 mm thick regions using intra-tissue strain. 2.4. Biochemical analyses The cell and matrix content of immature cartilage were determined on sequential sections from the articular surface. 3 mm punches were frozen in OCT and sectioned in 70 mm increments to 1 mm deep into the tissue. Slices were removed from OCT and digested overnight with 0.5 mg/ml Proteinase K at 60 1C. Tubes were inspected to ensure complete digestion, and digest solutions were analyzed for DNA as a measure of cellularity (7.7 pg DNA/cell) (McGowan et al., 2002), glycosaminoglycan (GAG) (Farndale et al., 1986), and hydroxyproline as a measure of collagen (7.25 g collagen/g hydroxyproline) (Pal et al., 1981), and normalized to the calculated volume of the slices (1.0 ml). 2.5. Statistical analysis Data are presented as mean7SEM. Biomechanical (Ezz, HA0) and biochemical (GAG, collagen, cells) data measured at several depths for each specimen were analyzed by ANOVA (a ¼ 0:05) for effects of developmental stage (fetal, newborn) or construct type (mixed, layered), and depth (repeated measure). When significant effects of depth were detected by ANOVA, posthoc tests with Bonferroni corrections were performed to determine which depths exhibited significantly higher strain than the other depths within a given tissue type. Additionally, fetal and newborn cartilage data were pooled and analyzed by univariate and multivariate regressions to determine if the local compressive modulus and local biochemical properties are associated in immature cartilage. All statistical analyses were performed with Systat version 10.2 (Systat Software, Inc., Richmond, CA).
3. Results 3.1. Immature cartilage mechanical properties Fetal and newborn articular cartilage equilibrium compressive moduli increased with depth from the articular surface. The top 1 mm of developing articular cartilage (Fig. 1) displayed non-linear stress–strain curve when analyzed as a whole (Fig. 2A). The aggregate modulus was significantly lower for fetal (89732 kPa) than newborn bovine cartilage (197721 kPa, po0:05). Intra-tissue strain of both fetal and newborn cartilage depended on depth from the articular surface (both po0:01, Fig. 2B), with the top 0.3 mm compressing significantly more than deeper layers (both po0:05).
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Fig. 1. Epifluourescent images of the top 1 mm of (A–C) 3rd trimester fetal bovine and (D–F) 2–3 week-old bovine calf articular cartilage in confined compression at (A, D) 0%, (B, E) 10%, and (C, F) 20% strain. The coordinate frame is indicated in panel A, and the articular surface is indicated in panels C and F.
Correspondingly, the confined compressive modulus was depth-dependent (po0:01, Fig. 2C), with the softest regions near the articular surface and the deepest regions 4–5-fold stiffer. 3.2. Tissue-engineered cartilage mechanical properties In contrast to the immature cartilage, the tissueengineered cartilage constructs displayed regions that were soft at both surfaces and stiffer in the center of the tissue. Constructs formed by mixing or layering chondrocyte subpopulations developed similar thickness (mixed 0.9970.04 mm, layered 1.0770.03, p ¼ 0:13) and strain profiles with depth (p ¼ 0:6). Unlike the native cartilage (Fig. 2B), the constructs showed regions of high strain at both surfaces, indicating relatively soft tissue, with regions in the top and bottom 0.2 mm straining more than the center of the tissue (po0:01, Fig. 3). 3.3. Cartilage biochemical properties and correlations Cartilage matrix constituents increased with depth from the articular surface of fetal and newborn cartilage, and correlated with mechanical properties. There was a general trend of increasing GAG (po0:001, Fig. 4A) and collagen (po0:001, Fig. 4B) content per volume from the superficial to the deeper layers. Both constituents were higher in the calf than the fetal cartilage (po0:001), and the increase with depth was greater for the calf. Cell density decreased similarly with depth for fetal and calf cartilage (both po0:001, Fig. 4C), falling from 180 million cells/cm3 in the top 0.1 mm to a plateau at 80 million cells/cm3 by 0.5 mm into the tissue.
The compressive modulus of immature bovine cartilage (fetal and newborn) was positively associated with GAG and collagen content in univariate regressions (both po0:01, Fig. 5A and B). The strongest positive correlation was with GAG content (r2 ¼ 0:45), with weaker correlation with collagen (r2 ¼ 0:17), and negative correlation with cell content (po0:01, r2 ¼ 0:22, Fig. 5C). Multivariate regressions on combined data from fetal and newborn cartilage suggest that the modulus depends on both the GAG and cell content (r2 ¼ 0:55, both po0:001), but not collagen content (p ¼ 0:3).
4. Discussion The results of this study suggest that there is a depthincreasing compressive modulus near the articular surface of immature articular cartilage, and that tissueengineered constructs differ from immature cartilage by exhibiting soft regions at both construct surfaces. Using image correlation methods with confined compression tests, the local compressive modulus was found to increase with depth in both fetal and newborn bovine cartilage (Fig. 2), similar to previous findings with mature articular cartilage (Schinagl et al., 1997). In contrast, constructs displayed symmetric soft surfaces (regions of high strain) regardless of how cells from the superficial or middle zone were seeded (Fig. 3). Biochemical analysis of cartilage sections indicated that matrix components, GAG and collagen, increase with depth and development, and that the cell content decreases from the articular surface, all with correlations to the local compressive properties (Figs. 3 and 4).
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Fig. 2. Mechanical properties of the top 1 mm of fetal (K) and newborn (m) bovine articular cartilage. (A) The stress is plotted versus the platen to platen (total) strain. (B) The local strain (Ezz) at 10% compression, and (C) confined compressive modulus (HA0) are plotted as a function of depth from the articular surface (n ¼ 4).
Several choices were made in the design of the experiments that may limit the interpretation of the results. First, while there were subtle differences between fetal and newborn tissues, such as greater absolute depth-variation in compressive modulus (Fig. 2C) and matrix components (Fig. 4A and B) in the newborn cartilage, the tissues were relatively close in age (1 month). Tissues from earlier stages of fetal development may not display such depth-dependent properties. Also, samples were from the surface 1 mm, and therefore these
Fig. 3. Epifluourescent images of constructs formed by (A–C) layering or (D–F) mixing chondrocytes from the superficial and middle zone of newborn bovine articular cartilage. Constructs are imaged at equilibrium in confined compression of (A, D) 0%, (B, E) 10%, and (C, F) 20% of the initial thickness. (G) Local strain (Ezz) at 20% compression is plotted for layered (E, n ¼ 3) and mixed (’, n ¼ 3) constructs. The coordinate frame is indicated in panel A and the membrane surface is indicated in panel F.
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Fig. 4. Depth-dependent biochemical properties of the top 1 mm of fetal (K) and newborn (m) bovine calf articular cartilage. The (A) glycosaminoglycan (GAG), (B) collagen (COL), and (C) cell number were normalized to the tissue volume of each cryosection bin and plotted as a function of depth from the articular surface (n ¼ 4).
data do not indicate how properties vary in the deep, vascularized regions. While full-thickness measurements may provide further insight into the development of depth-dependent mechanical properties, there is not a well-defined boundary for the deep zone in immature cartilage. The superficial 1 mm was chosen to include portions of the superficial and middle zones, as well as a
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region of tissue that is avascular, and of similar thickness to full-thickness adult bovine cartilage (Schinagl et al., 1997). Although the sample size was small, the variation in properties was large and demonstrated statistical significance. Further studies could examine different sites and tissue types, if more precise values were desired. However, depth-dependent compressive properties have been identified in cartilage from the knee (Schinagl et al., 1997), carpometacarpal (Wang et al., 2002), and shoulder (Wang et al., 2003) joints; thus, it appears depth-dependent properties are conserved across many joints. Finally, as in prior studies using image analysis to determine strain (Schinagl et al., 1996; Wang et al., 2003), measurements were taken at equilibrium, and thus do not provide information on the dynamic, viscoelastic response in the various regions relevant to joint kinematics, information that could be attained using ultrasound imaging methods (Zheng et al., 2005). Presence of depth-dependent mechanical properties in immature cartilage gives rise to questions of how these properties are developed. While articular cartilage appears relatively homogeneous at early developmental stages, differences between cells near the surface and those deeper are evident. For example, expression of PRG4 has been documented in the cells near the articular surface even in the first trimester (Schumacher et al., 1994), and may have importance in joint cavity formation (Flannery et al., 1999). Increases in matrix content with depth from the articular surface (Fig. 4A and B) may indicate a greater biosynthetic rate by cells further from the articular surface (Aydelotte et al., 1988). Additionally, changes in matrix organization, particularly collagen crosslinks, may affect the tissue’s mechanical properties (Williamson et al., 2003). Possible explanations for zonal cellular differences could include genetic factors, mechanical stresses (Carter et al., 2004; Wong et al., 1997), or transport of nutrients and growth factors. The top 1 mm of immature cartilage is essentially avascular, suggesting that transport of nutrients and waste products is primarily through the articular surface. Thus, gradients of oxygen, growth factors, etc. develop from the superficial to the deep zone, and are likely to influence chondrocyte function (Grimshaw and Mason, 2001; Kellner et al., 2002). Additionally, transport and binding of matrix molecules can lead to biochemical properties that increase with depth, even in absence of direct effects of nutrient gradients on cellular metabolism (DiMicco and Sah, 2003). The depth-varying properties of immature cartilage have implications for normal joint function as well as cartilage tissue engineering. Depth-dependent mechanical properties serve several functions in mature articular cartilage. The soft superficial zone functions to increase contact area and distribute load (Mankin et al., 1994),
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Fig. 5. Relationship between the local compressive modulus and GAG, collagen, and cell content, per volume, in the top 1 mm of bovine fetal (K) and newborn (m) articular cartilage.
and increase interstitial fluid pressure to lower frictional properties at the surface (Krishnan et al., 2003, 2004). The functions of depth-dependent mechanical properties, especially in the superficial zone, and their presence in immature cartilage suggest that they are necessary for normal joint function. In osteoarthritis, the absence or disruption of the superficial zone reduces functionality of the tissue (Guilak, 1994). Cartilage tissue engineering aims to restore normal joint function and thus depthdependent mechanical properties may be required for a properly functioning cartilage replacement. Cartilage constructs created with the aim of attaining one soft and one stiffer layer by sequentially seeding cells from different zones, developed relatively soft regions near both surfaces (absolute mechanical properties were not determined, but could be obtained by measuring stress), similar to constructs formed by mixing the cells (Fig. 3). Diffusive loss of macromolecules from construct surfaces may be responsible for development of such a strain profile. Applying membranes with small poresizes (100 kDa molecular weight cut-off) at the construct surfaces has resulted in excellent matrix retention in a scaffold-free system (Mainil-Varlet et al., 2001). Tissueengineered cartilage formed with hydrogels or polymer scaffolds may exhibit strain profiles drastically different from tissues formed here. In both immature cartilage and tissue-engineered cartilage formed without a mechanically functional scaffold, there may be an influence of the cell component on the mechanical properties of the tissue. In addition to positive correlations of compressive modulus with matrix components, there was a weak negative correlation with cell content for immature cartilage (Fig. 5C). While the cell component is generally thought not to affect the mechanical properties of mature cartilage, where there is a stiff matrix and low cell density, the extracellular matrix is less stiff and the cell volume fraction can be relatively high in immature cartilage
(diameter ¼ 10.4 mm, cell density ¼ 2 108 cells/ml, volume fraction ¼ 0.12) (Wong et al., 1997). Thus, the cell component may have a marked effect on mechanical properties, when considering that the cell has an elastic modulus three orders of magnitude smaller than the extracellular matrix (Jones et al., 1999; Wu et al., 1999). The cell shape and orientation may also influence mechanical properties, but appear to be more homogeneous with depth in immature than in mature cartilage (Wu and Herzog, 2002). While there was no observed change in cellularity between the two developmental stages, there were increases in GAG and collagen content, which may explain the increase in compressive modulus. Although the tissue-engineered cartilage was not analyzed for cell content as a function of depth, the cell component is expected to influence mechanical properties early in the culture process, as constructs consist largely of cells, and relatively small amounts of newly elaborated matrix. Immature articular cartilage is often used as a model for cartilage tissue engineering, due to its ability to grow, develop, and repair (Namba et al., 1998; Obradovic et al., 2001). The levels of organization and maturation necessary to make a functional cartilage replacement are not fully established, but the presence of biochemical and biomechanical properties that vary with depth from the surface in articular cartilage before birth (Fig. 2B and C) implicate depth-dependent properties as important details to consider. Engineering tissues with strain-profiles similar to native articular cartilage may have the benefit of enhancing host–implant integration by reducing strain discontinuities and relative motion at interfaces. Approaches which use cells from different zones, vary scaffold properties, or use bioreactor stimulation to engineer cartilage with depth-dependent properties may benefit from analytical methods such as image correlation to verify that these properties are attained (Ng et al., 2005), as there is not a
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simple correspondence between the properties of the cells (Fig. 3) or scaffold and the local mechanical properties developed after time in culture.
Acknowledgments This work was supported by grants from NASA, NIH, and NSF.
References Aydelotte, M.B., Greenhill, R.R., Kuettner, K.E., 1988. Differences between sub-populations of cultured bovine articular chondrocytes. II. Proteoglycan metabolism. Connective Tissue Research 18, 223–234. Bank, R.A., Bayliss, M.T., Lafeber, F.P., Maroudas, A., Tekoppele, J.M., 1998. Ageing and zonal variation in post-translational modification of collagen in normal human articular cartilage. The age-related increase in non-enzymatic glycation affects biomechanical properties of cartilage. Biochemical Journal 330, 345–351. Carter, D.R., Beaupre, G.S., Wong, M., Smith, R.L., Andriacchi, T.P., Schurman, D.J., 2004. The mechanobiology of articular cartilage development and degeneration. Clinical Orthopaedics and Related Research 427, S69–S77. Chen, A.C., Bae, W.C., Schinagl, R.M., Sah, R.L., 2001. Depth- and strain-dependent mechanical and electromechanical properties of full-thickness bovine articular cartilage in confined compression. Journal of Biomechanics 34, 1–12. Chen, A.C., Klisch, S.M., Bae, W.C., Temple, M.M., McGowan, K.B., Gratz, K.R., Schumacher, B.L., Sah, R.L., 2004. Mechanical characterization of native and tissue-engineered cartilage. In: Sabatini, M., DeCeuninck, F., Pastoureau, P. (Eds.), Cartilage and Osteoarthritis. Humana Press, Totowa, NJ, pp. 157–190. DiMicco, M.A., Sah, R.L., 2003. Dependence of cartilage matrix composition on biosynthesis, diffusion, and reaction. Transport in Porous Media 50, 57–73. Eckstein, F., Lemberger, B., Stammberger, T., Englmeier, K.H., Reiser, M., 2000. Patellar cartilage deformation in vivo after static versus dynamic loading. Journal of Biomechanics 33, 819–825. Farndale, R.W., Buttle, D.J., Barrett, A.J., 1986. Improved quantitation and discrimination of sulphated glycosaminoglycans by use of dimethylmethylene blue. Biochimica et Biophysica Acta 883, 173–177. Flannery, C.R., Hughes, C.E., Schumacher, B.L., Tudor, D., Aydelotte, M.B., Kuettner, K.E., Caterson, B., 1999. Articular cartilage superficial zone protein (SZP) is homologous to megakaryocyte stimulating factor precursor and is a multifunctional proteoglycan with potential growth-promoting, cytoprotective, and lubricating properties in cartilage metabolism. Biochemical and Biophysical Research Communications 254, 535–541. Frank, E.H., Grodzinsky, A.J., Koob, T.J., Eyre, D.R., 1987. Streaming potentials: a sensitive index of enzymatic degradation in articular cartilage. Journal of Orthopaedic Research 5, 497–508. Grimshaw, M.J., Mason, R.M., 2001. Modulation of bovine articular chondrocyte gene expression in vitro by oxygen tension. Osteoarthritis and Cartilage 9, 357–364. Guilak, F., 1994. Volume and surface area measurement of viable chondrocytes in situ using geometric modelling of serial confocal sections. Journal of Microscopy 173, 245–256. Hunziker, E.B., Michel, M., Studer, D., 1997. Ultrastructure of adult human articular cartilage matrix after cryotechnical processing. Microscopy Research and Technique 37, 271–284.
189
Jay, G.D., Tantravahi, U., Britt, D.E., Barrach, H.J., Cha, C.J., 2001. Homology of lubricin and superficial zone protein (SZP): products of megakaryocyte stimulating factor (MSF) gene expression by human synovial fibroblasts and articular chondrocytes localized to chromosome 1q25. Journal of Orthopaedic Research 19, 677–687. Jones, W.R., Ting-Beall, H.P., Lee, G.M., Kelley, S.S., Hochmuth, R.M., Guilak, F., 1999. Alterations in the Young’s modulus and volumetric properties of chondrocytes isolated from normal and osteoarthritic human cartilage. Journal of Biomechanics 32, 119–127. Keene, D.R., Oxford, J.T., Morris, N.P., 1995. Ultrastructural localization of collagen types II, IX, and XI in the growth plate of human rib and fetal bovine epiphyseal cartilage: type XI collagen is restricted to thin fibrils. Journal of Histochemistry and Cytochemistry 43, 967–979. Kellner, K., Liebsch, G., Klimant, I., Wolfbeis, O.S., Blunk, T., Schulz, M.B., Gopferich, A., 2002. Determination of oxygen gradients in engineered tissue using a fluorescent sensor. Biotechnology and Bioengineering 80, 73–83. Kim, T.K., Sharma, B., Williams, C.G., Ruffner, M.A., Malik, A., McFarland, E.G., Elisseeff, J.H., 2003. Experimental model for cartilage tissue engineering to regenerate the zonal organization of articular cartilage. Osteoarthritis and Cartilage 11, 653–664. Kiviranta, I., Jurvelin, J., Tammi, M., Saamanen, A.-M., Helminen, H.J., 1985. Microspectrophotometric quantitation of glycosaminoglycans in articular cartilage sections stained with Safranin O. Histochemistry 82, 249–255. Klein, T.J., Schumacher, B.L., Schmidt, T.A., Li, K.W., Voegtline, M.S., Masuda, K., Thonar, E.J.-M.A., Sah, R.L., 2003. Tissue engineering of articular cartilage with stratification using chondrocyte subpopulations. Osteoarthritis and Cartilage 11, 595–602. Krishnan, R., Park, S., Eckstein, F., Ateshian, G.A., 2003. Inhomogeneous cartilage properties enhance superficial interstitial fluid support and frictional properties, but do not provide a homogeneous state of stress. Journal of Biomechanical Engineering 125, 569–577. Krishnan, R., Kopacz, M., Ateshian, G.A., 2004. Experimental verification of the role of interstitial fluid pressurization in cartilage lubrication. Journal of Orthopaedic Research 22, 565–570. Kwan, M.K., Lai, W.M., Mow, V.C., 1990. A finite deformation theory for cartilage and other soft hydrated connective tissues—I. Equilibrium results. Journal of Biomechanics 23, 145–155. LeRoux, M.A., Arokoski, J., Vail, T.P., Guilak, F., Hyttinen, M.M., Kirivanta, K., Setton, L.A., 2000. Simultaneous changes in the mechanical properties, quantitative collagen organization, and proteoglycan concentration of articular cartilage following canine meniscectomy. Journal of Orthopaedic Research 18, 383–392. Mainil-Varlet, P., Rieser, F., Grogan, S., Mueller, W., Saager, C., Jakob, R.P., 2001. Articular cartilage repair using a tissueengineered cartilage-like implant: an animal study. Osteoarthritis and Cartilage 9 (Suppl A), S6–S15. Mankin, H.J., Mow, V.C., Buckwalter, J.A., Iannotti, J.P., Ratcliffe, A., 1994. Form and function of articular cartilage. In: Simon, S.R. (Ed.), Orthopaedic Basic Science. American Academy of Orthopaedic Surgeons, Rosemont, IL, pp. 1–44. Maroudas, A., 1979. Physico-chemical properties of articular cartilage. In: Freeman, M.A.R. (Ed.), Adult Articular Cartilage. Pitman Medical, Tunbridge Wells, England, pp. 215–290. Masuda, K., Sah, R.L., Hejna, M.J., Thonar, E.J., 2003. A novel twostep method for the formation of tissue-engineered cartilage by mature bovine chondrocytes: the alginate-recovered- chondrocyte (ARC) method. Journal of Orthopaedic Research 21, 139–148. McGowan, K.B., Kurtis, M.S., Lottman, L.M., Watson, D., Sah, R.L., 2002. Biochemical quantification of DNA in human articular and septal cartilage using PicoGreens and Hoechst 33258. Osteoarthritis and Cartilage 10, 580–587.
ARTICLE IN PRESS 190
T.J. Klein et al. / Journal of Biomechanics 40 (2007) 182–190
Namba, R.S., Meuli, M., Sullivan, K.M., Le, A., Adzick, N.S., 1998. Spontaneous repair of superficial defects in articular cartilage in a fetal lamb model. Journal of Bone and Joint Surgery American volume 80-A, 4–10. Ng, K.W., Wang, C.C., Mauck, R.L., Kelly, T.N., Chahine, N.O., Costa, K.D., Ateshian, G.A., Hung, C.T., 2005. A layered agarose approach to fabricate depth-dependent inhomogeneity in chondrocyte-seeded constructs. Journal of Orthopaedic Research 23, 134–141. Obradovic, B., Martin, I., Padera, R.F., Treppo, S., Freed, L.E., Vunjak-Novakovic, G., 2001. Integration of engineered cartilage. Journal of Orthopaedic Research 19, 1089–1097. Pal, S., Tang, L.-H., Choi, H., Habermann, E., Rosenberg, L., Roughley, P., Poole, A.R., 1981. Structural changes during development in bovine fetal epiphyseal cartilage. Collagen Related Research 1, 151–176. Poole, A.R., Matsui, Y., Hinek, A., Lee, E.R., 1989. Cartilage macromolecules and the calcification of cartilage matrix. The Anatomical Record 224, 167–179. Sah, R.L., Klein, T.J., Schmidt, T.A., Albrecht, D.R., Bae, W.C., Nugent, G.E., McGowan, K.B., Temple, M.M., Jadin, K.D., Schumacher, B.L., Chen, A.C., Sandy, J.D., 2004. Articular cartilage repair, regeneration, and replacement. In: Koopman, W.J. (Ed.), Arthritis and Allied Conditions: A Textbook of Rheumatology. Lippincott Williams & Wilkins, Philadelphia, pp. 2277–2301. Schinagl, R.M., Ting, M.K., Price, J.H., Sah, R.L., 1996. Video microscopy to quantitate the inhomogeneous equilibrium strain within articular cartilage during confined compression. Annals of Biomedical Engineering 24, 500–512. Schinagl, R.M., Gurskis, D., Chen, A.C., Sah, R.L., 1997. Depthdependent confined compression modulus of full-thickness bovine articular cartilage. Journal of Orthopaedic Research 15, 499–506. Schumacher, B.L., Block, J.A., Schmid, T.M., Aydelotte, M.B., Kuettner, K.E., 1994. A novel proteoglycan synthesized and secreted by chondrocytes of the superficial zone of articular cartilage. Archives of Biochemistry and Biophysics 311, 144–152. Stockwell, R.A., Meachim, G., 1979. The chondrocytes. In: Freeman, M.A.R. (Ed.), Adult Articular Cartilage. Pitman Medical, Tunbridge Wells, England, pp. 69–144.
Sutton, M.A., McNeill, S.R., Helm, J.D., Chao, Y.J., 2000. Advances in two-dimensional and three-dimensional computer vision. In: Rastogi, P.K. (Ed.), Photomechanics. Springer, New York, pp. 323–372. Wang, C.C., Deng, J.M., Ateshian, G.A., Hung, C.T., 2002. An automated approach for direct measurement of two-dimensional strain distributions within articular cartilage under unconfined compression. Journal of Biomechanical Engineering 124, 557–567. Wang, C.C., Chahine, N.O., Hung, C.T., Ateshian, G.A., 2003. Optical determination of anisotropic material properties of bovine articular cartilage in compression. Journal of Biomechanics 36, 339–353. Williamson, A.K., Chen, A.C., Sah, R.L., 2001. Compressive properties and function-composition relationships of developing bovine articular cartilage. Journal of Orthopaedic Research 19, 1113–1121. Williamson, A.K., Chen, A.C., Masuda, K., Thonar, E.J.-M.A., Sah, R.L., 2003. Tensile mechanical properties of bovine articular cartilage: variations with growth and relationships to collagen network components. Journal of Orthopaedic Research 21, 872–880. Wong, M., Wuethrich, P., Eggli, P., Hunziker, E., 1996. Zone-specific cell biosynthetic activity in mature bovine articular cartilage: a new method using confocal microscopic stereology and quantitative autoradiography. Journal of Orthopaedic Research 14, 424–432. Wong, M., Wuethrich, P., Buschmann, M.D., Eggli, P., Hunziker, E., 1997. Chondrocyte biosynthesis correlates with local tissue strain in statically compressed adult articular cartilage. Journal of Orthopaedic Research 15, 189–196. Wu, J.Z., Herzog, W., 2002. Elastic anisotropy of articular cartilage is associated with the microstructures of collagen fibers and chondrocytes. Journal of Biomechanics 35, 931–942. Wu, J.Z., Herzog, W., Epstein, M., 1999. Modelling of location- and time-dependent deformation of chondrocytes during cartilage loading. Journal of Biomechanics 32, 563–572. Zheng, Y.P., Niu, H.J., Arthur Mak, F.T., Huang, Y.P., 2005. Ultrasonic measurement of depth-dependent transient behaviors of articular cartilage under compression. Journal of Biomechanics 38, 1830–1837.