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In situ chondrocyte viscoelasticity
Journal of Biomechanics 45 (2012) 2450–2456
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2011 American Society of Biomechanics Journal of Biomechanics Award
In situ chondrocyte viscoelasticity Sang-Kuy Han a,b,1, Ryan Madden a,c, Ziad Abusara a, Walter Herzog a,b,c,n a b c
Human Performance Laboratory, University of Calgary, Canada Department of Mechanical and Manufacturing Engineering, University of Calgary, Canada Department of Biomedical Engineering, University of Calgary, Canada
a r t i c l e i n f o
a b s t r a c t
Article history: Accepted 24 June 2012
It has been proposed, based on theoretical considerations, that the strain rate-dependent viscoelastic response of cartilage reduces local tissue and cell deformations during cyclic compressions. However, experimental studies have not addressed the in situ viscoelastic response of chondrocytes under static and dynamic loading conditions. In particular, results obtained from experimental studies using isolated chondrocytes embedded in gel constructs cannot be used to predict the intrinsic viscoelastic responses of chondrocytes in situ or in vivo. Therefore, the purpose of this study was to investigate the viscoelastic response of chondrocytes in their native environment under static and cyclic mechanical compression using a novel in situ experimental approach. Cartilage matrix and chondrocyte recovery in situ following mechanical compressions was highly viscoelastic. The observed in situ behavior was consistent with a previous study on in vivo chondrocyte mechanics which showed that it took 5–7 min for chondrocytes to recover shape and volume following virtually instantaneous cell deformations during muscular loading of the knee in live mice. We conclude from these results that the viscoelastic properties of cartilage minimize chondrocyte deformations during cyclic dynamic loading as occurs, for example, in the lower limb joints during locomotion, thereby allowing the cells to reach mechanical and metabolic homeostasis even under highly dynamic loading conditions. & 2012 Elsevier Ltd. All rights reserved.
Keywords: Articular cartilage Chondrocyte Viscoelasticity Stress relaxation Cyclic compression
1. Introduction Articular cartilage is the thin layer of connective tissue covering the bony surfaces in synovial joints. It plays an essential role in joint health by providing surface lubrication and load distribution across articular surfaces. Articular cartilage experiences millions of cycles of mechanical loading during its lifetime. The health and integrity of the cartilage’s extracellular matrix (ECM) is maintained by the activity of chondrocytes, the cells within articular cartilage, which are responsible for synthesizing structural macromolecules (Stockwell, 1979). The biosynthetic activity of chondrocytes is thought to be regulated by multiple factors including genetic and environmental influences, the composition of the extracellular matrix, and mechanical stimuli (Stockwell, 1979; Guerne et al., 1990; Palmoski and Brandt, 1984). Although the detailed regulatory mechanisms remain unknown, there is convincing evidence that mechanical stimuli, such as
n Corresponding author at: Human Performance Laboratory, Faculty of Kinesiology, University of Calgary, 2500 University DR. N.W., Calgary, Canada T2N 1N4. Tel.: þ 1 403 220 8525; fax: þ1 403 220 2070. E-mail address: [email protected] (W. Herzog). 1 Current address: Fischell Department of Bioengineering, University of Maryland, MD, USA.
0021-9290/$ - see front matter & 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.jbiomech.2012.06.028
compression, shear stress, hydrostatic pressure, and osmotic stress, influence the biosynthetic activity of chondrocytes (Kim et al., 1994; Smith et al., 1995; Hall et al., 1996; Mizuno et al., 2002). Therefore, the health and integrity of articular cartilage and the synthesizing activity of chondrocytes are thought to depend largely on the mechanical stimuli experienced by the tissue. Excessive loads are thought to cause damage to the cartilage matrix, thereby inducing negative biosynthetic responses from chondrocytes (Wollheim and Lohmander, 2007). The material behavior of articular cartilage is characterized by viscoelastic properties such as creep, stress relaxation, and hysteresis (Hayes and Mockros, 1971; Armstrong et al., 1984, Mow et al., 1980; Holmes, 1986). The material properties of chondrocytes have been investigated using a variety of methods including compression of isolated cells within a gel construct, micropipette aspiration and cellular indentation of single cells using atomic force microscopy (Knight et al., 1998; Trickey et al., 2000; Darling et al., 2006). These works found that single chondrocytes are inherently viscoelastic. A possible functional role for articular cartilage viscoelasticity is a mechanical damping of external loads. It has been shown theoretically that the strain rate-dependent viscoelastic response of cartilage reduces periodic alterations of tissue strains (Suh et al., 1995) and cell deformations (Kim et al., 2008) during cyclic
S.-K. Han et al. / Journal of Biomechanics 45 (2012) 2450–2456
compression. However, neither theoretical nor experimental studies have fully addressed the viscoelastic responses of chondrocyte deformations in situ under static and dynamic mechanical loading conditions (Freeman et al., 1994; Knight et al., 1998; Wu and Herzog, 2006; Kim et al., 2008). Pilot results obtained in a recent study on in vivo chondrocyte deformations in the intact knee loaded by muscular contractions suggest that the intrinsic viscoelastic response of chondrocytes substantially differs from that observed in isolated chondrocytes embedded in gel constructs (Abusara et al., 2011; Knight et al., 1998). Since the material behavior of in situ chondrocytes is essential for understanding the relationship between cartilage loading and the adaptive/degenerative responses of chondrocytes, the purpose of this study was to investigate the viscoelastic responses of chondrocytes in their native environment using a novel in situ experimental approach. We hypothesized that the viscoelastic properties of the extracellular matrix and chondrocytes limit cell deformations during cyclic loading.
2. Materials and methods 2.1. Sample preparation Articular cartilage and its subchondral bone were taken from the femoral groove of skeletally mature cows on the day of slaughtering. Cylindrical cores (diameter¼ 6.4 mm) were extracted from the harvested osteochondral blocks (N ¼4, Fig. 1). Cartilage thickness was measured using a digital caliper at opposing circumferential locations to obtain an estimate of the average sample thickness. Fluorescein conjugated dextran (Alexa, excitation: 488 nm, emission: 517 nm; Molecular Probes, OR, USA) was suspended in DMEM (Dulbecco’s Modified Eagle’s Medium, Gibco, OR, USA) at a concentration of 6.7 mg/ml (2.2 mM). The osteochondral cores were incubated in the dextran solution for 4–10 h at 4 1C prior to mechanical testing and confocal imaging. Samples were rinsed in dye-free phosphate-buffered saline (PBS) for 20 min following staining. Cylindrical samples were rigidly fixed in a specimen holder using dental cement. During testing, samples were fully immersed in the PBS solution to prevent dehydration of the cartilage surface.
2.2. Mechanical testing All mechanical tests and imaging were conducted using a custom-designed indentation system that allows for imaging of cells in the intact cartilage attached to its native bone (Fig. 1A and B). Briefly, the system consists of a light-transmissible glass indenter mounted to the stage of an upright laser scanning microscope and a piezoactuator which pushes the tissue sample towards the indenter. These primary components are complemented by a load cell and displacement transducer which,
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coupled with custom-designed software, allow for controlled loading of the cartilage/ bone specimen. The microscope-based indentation system has been described in detail previously (Han et al., 2009). Three different mechanical tests, described in detail below, were conducted to characterize the viscoelastic properties of chondrocytes in their native environment. 2.2.1. Single stress relaxation test A single static compressive load of 10% nominal tissue strain was applied to the surface of the osteochondral blocks using a glass indenter (diameter¼2.00 mm) at an average strain rate of 3.1%/s (Fig. 2A). The target load was held for 20 min to allow the tissue to reach a steady-state. Confocal image sections were recorded through a 40 magnification, 0.8 numerical aperture, 0.17 mm coverglass-corrected objective: (i) before loading; (ii) at 2 and 17 min of relaxation; and (iii) at 1/2, 1, 3, 5, 7, and 10 min after removal of the load. Confocal scans were recorded from the cartilage surface to a depth of 60 mm, thereby capturing surface zone chondrocytes. Optical slice thickness (i.e. perpendicular distance between image planes) was set at 0.5 mm for all scans except for 30 s after load removal, where the optical slice thickness was set at 3 mm only for tissue recovery measurement to reduce scan time and capture the initial recovery of the tissue. Upon completion of the 10 min recovery phase, tissue samples were allowed to fully relax prior to the next test. Tissue samples were removed from the indentation system and rinsed with fresh PBS for 10 min before conducting the next test. 2.2.2. Three cycle compression test Next, a series of three 10% compressive strain loads were applied to the articular surface and held for 2 min each (Fig. 2B). Confocal scans were taken before loading, during each of the three loading and unloading phases; and at 1/2, 1, 3, 5, 7, and 10 min after removal of the load (Fig. 2B). As in the previous test, confocal scans were taken from the cartilage surface to a depth of 60 mm. Optical slice thickness was 0.5 mm for all scans except for 30 s after removal of the final load (3 mm). Tissue samples were allowed to fully relax prior to the next test, as described above. 2.2.3. Dynamic cyclic compression test The final test consisted of a half-sinusoidal compressive load (13% strain) applied for 100 cycles at 0.35 Hz (Fig. 2C). Confocal scans were taken before loading and after loading as outlined in the previous tests. Total tissue strains, local tissue strains (denoted as ECM strain), and compressive cell strains were quantified as described below. 2.3. Confocal image analysis Four cells from each femoral groove core (total of 16 cells) were used for cell morphology analysis. All chondrocytes were located in the superficial zone of the bovine cartilage between the cartilage surface and a depth of 60 mm. Cell height was quantified using a confocal image viewer (LSM Image Browser, Zeiss Inc.). Local compressive ECM strain was calculated by measuring the distance between paired cells (n¼ 3 pairs per sample) in the pre-test condition and at each experimental measurement point. Overall tissue recovery was quantified by measuring the distance from the cartilage surface to the bottom of the glass indenter observed in the images. Tissue strains during the recovery phase were calculated as engineering strains (eE ¼ (h h0)/h0, where h is the deformed tissue thickness and h0 is the original tissue thickness), and local ECM and cell strains were calculated as
Fig. 1. Sample preparation and indentation testing. (A) Photo of the actual indentation system on the x–y stage of a microscope. (B) Schematic illustration of the area marked with the dashed square in (A).
Nominal strain [-%]
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100 cycles A : Single stress relaxation test B : 3 cycle test C : 100 cycle test s : tissue measuring point : ECM measuring point : cell measuring point
13 10 0.35 [Hz] 5 0 0
1 3 time [m]
5
7
9 10
Fig. 2. Experimental loading protocol and measurement time points for confocal imaging: (A) single static compression test; (B) three-cycle compression test; and (C) dynamic cyclic compression test.
during loading
at 3 min recovery phase
at 7 min recovery phase
Cell
ECM
before loading
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N=3
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ECM, cell compressive strain [%]
Tissue compressive strain [%]
40
9 10
recovery phase
time [m] Fig. 3. Stress relaxation test. ECM: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (Po 0.05: ntissue, #ECM, þcell).
finite deformation strains for the large deformation theory (eF ¼ (l 1)/2, where l ¼ h/h0, h is the deformed distance between paired cells for ECM or height of cell, and h0 is the original distance of paired cells for ECM or height of cell).
2.4. Statistical analysis Tissue, ECM, and cell strains at different experimental time points were compared using one-way repeated measures ANOVA (SPSS 19, IBM) to determine if there was a significant time-effect during recovery. If indicated, Bonferroni posthoc testing was used to compare loaded and recovery time results with the initial unloaded condition. The level of significance was set at a ¼ 0.05. Results are presented as means7 1 standard deviation.
3. Results 3.1. Single stress relaxation test After 2 min of static loading at 10% compressive tissue strain, local compressive ECM strains in the superficial zone were 3971% and average compressive cell strains were 2976%. Local ECM compressive strains and average compressive cell strains increased further to 41 72% and 3175%, respectively, after 17 min of static loading. The cartilage recovered to its original shape 5 min after load removal and the chondrocytes to their original height 5 min after load removal (P40.05 for both,
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Figs. 3 and 4). The local tissue (ECM) strain did not recover completely within 10 min after load removal (P¼0.002, Fig. 3).
3.2. Three cycle compression test Average compressive ECM and chondrocyte strains in the superficial zone were 3672% and 2877% respectively, during each of the three loading cycles of 10% compressive tissue strain. The cartilage tissue did not fully recover during the 2 min unloading phase, and did not recover completely within 3 min after load removal (Po0.05, Fig. 5). Local tissue (ECM) strain and compressive chondrocyte strain were fully recovered to the initial unloaded values within 10 and 5 min, respectively (P40.05 for both, Fig. 5).
before
during load
3 min after
7 min after
surface y x z
Fig. 4. Three-dimensional reconstruction of local matrix (ECM) and cell deformations before loading, during loading, 3 min after load removal, and 7 min after load removal.
during loading phase
3.3. Dynamic cyclic compression test Cartilage tissue fully recovered to its original shape within 3 min following the completion of 100 compressive loading cycles at a frequency of 0.35 Hz (P4 0.05, Fig. 6). The local tissue (ECM) and cellular strains exhibited a similar recovery, returning to their original configurations within 7 min after load removal (P 40.05 for both, Fig. 6).
4. Discussion This study demonstrated the viscoelasticity of articular cartilage and chondrocytes in situ under static and dynamic compression loads. Our results of cellular recovery upon unloading are consistent with a previous study in the intact mouse tibia–femoral joint (Abusara et al., 2011), where it was found that chondrocytes deform rapidly upon loading but take minutes to recover shape and volume upon unloading. However, the results of our study are in contrast with findings from isolated chondrocytes embedded in agarose gel constructs, which recovered immediately and nearly elastically following mechanical unloading (Knight et al., 1998). This difference in chondrocyte recovery between in situ conditions and cells embedded in agarose gel constructs is likely related to the mechanical environment of the cells, as chondrocyte deformation is linked to the deformation of the extracellular matrix (Guilak, 2000). We conclude from these results that chondrocyte mechanics differ substantially between in situ and in vitro conditions, and that the mechanical properties of cells must be considered within the context of the experimental conditions and environment. The extracellular matrix also behaved highly viscoelastically, similar to the cells following compression testing (Fig. 3). It is well recognized that chondrocytes and surrounding ECM are physically connected by adhesion molecules through the pericellular matrix (Guilak et al., 2006), therefore it seems plausible that the
at 3 min at 7 min recovery phase recovery phase
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before loading
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20
10 N=3
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3 5 7 recovery phase
ECM, cell compressive strain [%]
30
N=2
Tissue compressive strain [%]
40
9 10
time [m] Fig. 5. Three cycle compression test: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼ 5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (P o0.05: ntissue, #ECM, þ cell).
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at 3 min recovery phase
at 7 min recovery phase
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ECM
before loading
13
Tissue compressive strain [%]
10
16
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8
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+ #
4 0 -2
0 0
1
3
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7
ECM, cell compressive strain [%]
18
9 10
recovery phase time [m] Fig. 6. Dynamic compression test: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼ 5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (Po 0.05: ntissue, #ECM, þcell).
Cyclic external load [cm]
[mm]
Time
[μm]
Time
elastic prediction viscoelastic
Time
Fig. 7. Schematic diagram for cyclic movement, load on the knee joint and possible cell deformation response to the cyclic loading of the knee joint.
shape and volume recovery of the cells is intimately linked to the recovery of the ECM, as has been suggested for articular cartilage when exposed to swelling following osmotic loading (Korhonen et al., 2010). Furthermore, given the three orders of magnitude difference in Young’s modulus between chondrocytes and surrounding ECM (Jones et al., 1999; Schinagl et al., 1997), one would expect that chondrocyte deformations are dominated by the ECM deformations in the vicinity of cells rather than the mechanical properties of the chondrocytes. Therefore, we propose that the viscoelastic response of chondrocytes in situ is largely governed by the viscoelastic behavior of the extracellular matrix. It has been well documented that mechanical loading of cartilage enhances chondrocyte biosynthesis (Sah et al., 1989; Kim et al., 1994; Little and Ghosh, 1997; Buschmann et al., 1999). However, excessive loads may damage the cartilage matrix and produce negative cellular effects (Wollheim and Lohmander, 2007). Therefore, it might be of advantage if cell deformations are minimized under conditions of cyclic mechanical loading.
Here we found, in agreement with pilot results in intact joints (Abusara et al., 2011), that cells and ECM deform quickly ( o1 s) upon mechanical loading, while cell shape and cell volume recovery take minutes to complete (Fig. 5). Therefore, for cyclic dynamic loading, cell deformations and volume changes are greatest in the first cycle of loading, but then become negligibly small in subsequent loading cycles, suggesting that one of the in situ properties of the ECM/cell construct is to minimize cell shape and volume changes, thereby allowing cells to reach homeostasis during cyclic mechanical loading. Physiological loading of joints is often cyclic, for example for the lower limb joints during locomotion, thus protecting cells from excessive strains in these situations and minimizing costly, cyclic cell deformations would be advantageous (Fig. 7). The viscoelasticity of the cartilage matrix and chondrocytes is associated with the fluid phase movement and the interaction between fluid and solid matrix within the tissue and the cells (Mow et al., 1980). When articular cartilage degenerates in
S.-K. Han et al. / Journal of Biomechanics 45 (2012) 2450–2456
Table 1 Steady-state compressive strains in the tissue, matrix and cells 17 min after applying a 10% nominal compressive tissue strain.
Compressive strain (%)
Tissue
Superficial cartilage
Superficial cell
10
417 2
31 7 5
disease, such as osteoarthritis (OA), material properties of the cartilage matrix and the chondrocytes change substantially (Alexopoulos et al., 2003, 2005; Hasler and Herzog, 1998; Seedholm et al., 1979). For instance, in early OA, the cartilage matrix becomes softer and permeability increases (Hasler and Herzog, 1998; Seedholm et al., 1979). Additionally, the collagen network in the superficial zone of early OA cartilage has been shown to undergo a disruption and remodeling process (Stockwell, 1991; Guilak et al., 1994; Buckwalter and Mankin, 1997), which may lead to changes in the local matrix environment of the chondrocytes. It has been suggested that these altered material properties of osteoarthritic cartilage affect cell deformation (Kim et al., 2008), and cell deformation-based signaling pathways (Kim et al., 1994). In order to understand the link between OA onset and progression, it would be important to investigate how disease-related changes in articular cartilage viscoelasticity affect chondrocyte deformations, signaling, and macromolecule synthesis. Cell deformations were affected by the viscoelastic properties of the local cartilage matrix in this study. Furthermore, the mechanical properties of the pericellular matrix likely also affected cell mechanics (Poole et al., 1987; Knight et al., 2001). It has been reported that the pericellular matrix provides a unique mechanical environment for chondrocytes and may protect cells from damaging strains in conjunction with the ECM (Youn et al., 2006). For instance, the pericellular matrix acts as a non-linear mechanical filter for controlling cell deformations under mechanical compression (Choi et al., 2007) and modulating chondrocyte biosynthesis (Guilak et al., 2006). The pericellular matrix has viscoelastic material properties, similar to the chondrocytes and extracellular matrix, and these properties are known to be altered when cartilage degenerates (Alexopoulos et al., 2003). Therefore, in order to completely understand the viscoelastic behavior of chondrocytes in situ, the detailed viscoelastic properties of the PCM and their potential effect on chondrocyte mechanics need to be studied further in the future. Consistent with previous studies (Guilak et al., 1995; Han et al., 2010; Schinagl et al., 1997), we found that the superficial zone of cartilage exhibited much greater compressive strains than the average strain imposed on the tissue (Table 1). It is well accepted that extracellular matrix (ECM) and chondrocyte deformations are depth dependent under static compression (Guilak et al., 1995; Schinagl et al., 1997). The superficial matrix and chondrocytes are expected to deform more than the matrix and cells in the deeper layers of the cartilage. With the indentation system and the scanning laser confocal microscopy approach used in this study, we are limited to studying the extracellular matrix and cellular deformations up to about 60 mm below the articular surface. Attempts at numerical modeling of chondrocytes subjected to cyclic loading also showed depth dependent cell deformation (Wu and Herzog, 2006). However, they did not capture the highly viscoelastic cell responses we observed here experimentally in situ, but rather showed cell deformations and recovery that were almost identical in scale, thereby predicting large cell shape and volume changes during each loading cycle. Biosynthetic responses of chondrocytes are also known to be depth dependent (Wong et al., 1997). For example, aggrecan synthesis is thought to occur primarily in deep zone chondrocytes
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(Wong et al., 1997), while proteoglycan-4 synthesis is localized in superficial zone cells (Nugent-Derfus et al., 2007). Furthermore, cartilage compression at different rates affects the biosynthetic response of chondrocytes (Buschmann et al., 1999). Together, these results suggest that cyclic mechanical loading produces depth dependent mechanical and biological responses. Therefore, studies in which chondrocytes at all depths can be visualized need to be performed to fully understand the functional roles of chondrocytes in the different structural layers of articular cartilage. In this study, we focused on the recovery phase of the tissue, the local extracellular matrix and chondrocytes following mechanical compression. Although we aimed at investigating the tissue, local ECM, and cellular deformations during cyclic loading, it was technically difficult to capture three-dimensional ECM and cell morphologies during the dynamic phases when deformations occurred rapidly. Furthermore, it was also technically difficult to capture the tissue recovery right after load removal. Therefore, the initial measurement of tissue recovery, right after load removal, was conducted using 3 mm thick slices, while all other measurements were made using 0.5 mm slices. This allowed for capturing the fast tissue recovery immediately after unloading, while providing excellent accuracy for cell shapes and volumes for measurements in near steady-state conditions. Future studies need to consider how the temporal resolution of confocal imaging can be improved substantially without losing spatial resolution, so that cell deformation and signaling can be studied during highly dynamic conditions. It seems to us that these dynamic conditions, that elude quantification with present day technology, may hold the key to our understanding of cartilage mechanobiology and to the mechanisms of cartilage adaptation and degeneration.
5. Conclusion The results of this study led to the conclusion (i) that chondrocyte mechanics are greatly affected by the local environment, thus cells in agarose gel and in the native cartilage matrix behave very differently, and (ii) that chondrocytes in their native environment have properties that minimize cell deformations during cyclic loading. We suggest that this latter property allows chondrocytes to achieve homeostasis during dynamic loading in a short period of time.
Conflict of interest statement The authors have no conflicts of interest.
Acknowledgments The authors would like to thank A. Jinha from the Human Performance Laboratory at the University of Calgary for assistance with LabView programming, the Canada Research Chair Programme, the Killam Foundation, the Canadian Institutes for Health Research, and the Alberta Innovates—Health Solutions Team Grant on Osteoarthritis. References Abusara, Z., Seerattan, R., Leumann, A., Thompson, R., Herzog, W., 2011. A novel method for determining articular cartilage chondrocyte mechanics in vivo. Journal of Biomechanics 44, 930–934. Alexopoulos, L., Haider, M.A., Vail, T.P., Guilak, F., 2003. Alterations in the mechanical properties of the human chondrocyte pericellular matrix with osteoarthritis. Journal of Biomechanical Engineering 125, 323–333.
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Knight, M.M., Ghori, S.A., Lee, D.A., Bader, D.L., 1998. Measurement of the deformation of isolated chondrocytes in agarose subjected to cyclic compression. Medical Engineering and Physics 20, 684–688. Knight, M.M., Ross, J.M., Sherwin, A.F., Lee, D.A., Bader, D.L., Poole, C.A., 2001. Chondrocyte deformation within mechanically and enzymatically extracted chondrons compressed in agarose. Biochimmica et Biophysica Acta 1526 (2), 141–146. Korhonen, R.K., Han, S.-K., Herzog, W., 2010. Osmotic loading of in situ chondrocytes in their native environment. Molecular and Cellular Biomechanics 7 (3), 125–134. Little, C.B., Ghosh, P.G., 1997. Variation in proteoglycan metabolism by articular chondrocytes in different joint regions is determined by post-natal mechanical loading. Osteoarthritis and Cartilage 5, 49–62. Mizuno, S., Tateishi, T., Ushida, T., Glowacki, J., 2002. Hydrostatic fluid pressure enhances matrix synthesis and accumulation by bovine chondrocytes in threedimensional culture. Journal of Cellular Physiology 193, 319–327. Mow, V.C., Kuei, S.C., Lai, W.M., Armstrong, C.G., 1980. Biphasic creep and stress relaxation of articular cartilage: theory and experiments. Journal of Biomechanical Engineering 102, 73–84. Nugent-Derfus, G.E., Chan, A.H., Schumacher, B.L., Sah, R.L., 2007. PRG4 exchange between the articular cartilage surface and synovial fluid. Journal of Orthopaedic Research 25, 1269–1276. Palmoski, M.J., Brandt, K.D., 1984. Effect of static and cyclic compressive loading on articular cartilage plugs in vitro. Arthritis and Rheumatism 27, 675–681. Poole, C.A., Flint, M.H., Beaumont, B.W., 1987. Chondrons in cartilage: ultrastructural analysis of the pericellular microenvironment in adult human articular cartilages. Journal of Orthopaedic Research 5 (4), 509–522. Sah, R.L., Kim, Y.-J., Doong, J.-Y.H., Grodzinsky, A.J., Plaas, A.H.K., Sandy, J.D., 1989. Biosynthetic response of cartilage explants to dynamic compression. Journal of Orthopaedic Research 7, 619–636. Schinagl, R.M., Gurskis, D., Chen, A.C., Sah, R.L., 1997. Depth-dependent confined compression modulus of full-thickness bovine articular cartilage. Journal of Orthopaedic Research 15, 499–506. Seedholm, B.B., Takeda, T., Tsubuku, M., Wright, V., 1979. Mechanical factors and patellofemoral osteoarthritis. Annals of the Rheumatic Diseases 38, 307–316. Smith, R.L., Donlon, B.S., Gupta, M.K., Mohtai, M., Das, P., Carter, D.R., Cooke, J., Gibbons, G., Hutchinson, N., Schurman, D.J., 1995. Effects of fluid-induced shear on articular chondrocyte morphology and metabolism in vitro. Journal of Orthopaedic Research 13, 824–831. Stockwell, R.A., 1979. Biology of Cartilage Cells. Cambridge University Press, Cambridge. Stockwell, R.A., 1991. Cartilage failure in osteoarthritis: relevance of normal structure and function. A review. Clinical Anatomy 4, 161–191. Suh, J.K., Li, Z., Woo, S.L., 1995. Dynamic behavior of a biphasic cartilage model under cyclic compressive loading. Journal of Biomechanics 28, 357–364. Trickey, W.R., Lee, G.M., Guilak, F., 2000. Viscoelastic properties of chondrocytes from normal and osteoarthritic human cartilage. Journal of Orthopaedic Research 18, 891–898. 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. Wollheim, F.A., Lohmander, S.L., 2007. Pathology and animal models of osteoarthritis. In: Sharma, L. (Ed.), Osteoarthritis: A Companion to Rheumatology. Mosby Elsevier, Philadelphia, pp. 104–112. Wu, J.Z., Herzog, W., 2006. Analysis of the mechanical behavior of chondrocytes in unconfined compression tests for cyclic loading. Journal of Biomechanics 39, 603–616. Youn, I., Choi, J.B., Cao, L., Setton, L.A., Guilak, F., 2006. Zonal variations in the three-dimensional morphology of the chondron measured in situ using confocal microscopy. Osteoarthritis and Cartilage 14, 889–897.
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Journal of Biomechanics journal homepage: www.elsevier.com/locate/jbiomech www.JBiomech.com
2011 American Society of Biomechanics Journal of Biomechanics Award
In situ chondrocyte viscoelasticity Sang-Kuy Han a,b,1, Ryan Madden a,c, Ziad Abusara a, Walter Herzog a,b,c,n a b c
Human Performance Laboratory, University of Calgary, Canada Department of Mechanical and Manufacturing Engineering, University of Calgary, Canada Department of Biomedical Engineering, University of Calgary, Canada
a r t i c l e i n f o
a b s t r a c t
Article history: Accepted 24 June 2012
It has been proposed, based on theoretical considerations, that the strain rate-dependent viscoelastic response of cartilage reduces local tissue and cell deformations during cyclic compressions. However, experimental studies have not addressed the in situ viscoelastic response of chondrocytes under static and dynamic loading conditions. In particular, results obtained from experimental studies using isolated chondrocytes embedded in gel constructs cannot be used to predict the intrinsic viscoelastic responses of chondrocytes in situ or in vivo. Therefore, the purpose of this study was to investigate the viscoelastic response of chondrocytes in their native environment under static and cyclic mechanical compression using a novel in situ experimental approach. Cartilage matrix and chondrocyte recovery in situ following mechanical compressions was highly viscoelastic. The observed in situ behavior was consistent with a previous study on in vivo chondrocyte mechanics which showed that it took 5–7 min for chondrocytes to recover shape and volume following virtually instantaneous cell deformations during muscular loading of the knee in live mice. We conclude from these results that the viscoelastic properties of cartilage minimize chondrocyte deformations during cyclic dynamic loading as occurs, for example, in the lower limb joints during locomotion, thereby allowing the cells to reach mechanical and metabolic homeostasis even under highly dynamic loading conditions. & 2012 Elsevier Ltd. All rights reserved.
Keywords: Articular cartilage Chondrocyte Viscoelasticity Stress relaxation Cyclic compression
1. Introduction Articular cartilage is the thin layer of connective tissue covering the bony surfaces in synovial joints. It plays an essential role in joint health by providing surface lubrication and load distribution across articular surfaces. Articular cartilage experiences millions of cycles of mechanical loading during its lifetime. The health and integrity of the cartilage’s extracellular matrix (ECM) is maintained by the activity of chondrocytes, the cells within articular cartilage, which are responsible for synthesizing structural macromolecules (Stockwell, 1979). The biosynthetic activity of chondrocytes is thought to be regulated by multiple factors including genetic and environmental influences, the composition of the extracellular matrix, and mechanical stimuli (Stockwell, 1979; Guerne et al., 1990; Palmoski and Brandt, 1984). Although the detailed regulatory mechanisms remain unknown, there is convincing evidence that mechanical stimuli, such as
n Corresponding author at: Human Performance Laboratory, Faculty of Kinesiology, University of Calgary, 2500 University DR. N.W., Calgary, Canada T2N 1N4. Tel.: þ 1 403 220 8525; fax: þ1 403 220 2070. E-mail address: [email protected] (W. Herzog). 1 Current address: Fischell Department of Bioengineering, University of Maryland, MD, USA.
0021-9290/$ - see front matter & 2012 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.jbiomech.2012.06.028
compression, shear stress, hydrostatic pressure, and osmotic stress, influence the biosynthetic activity of chondrocytes (Kim et al., 1994; Smith et al., 1995; Hall et al., 1996; Mizuno et al., 2002). Therefore, the health and integrity of articular cartilage and the synthesizing activity of chondrocytes are thought to depend largely on the mechanical stimuli experienced by the tissue. Excessive loads are thought to cause damage to the cartilage matrix, thereby inducing negative biosynthetic responses from chondrocytes (Wollheim and Lohmander, 2007). The material behavior of articular cartilage is characterized by viscoelastic properties such as creep, stress relaxation, and hysteresis (Hayes and Mockros, 1971; Armstrong et al., 1984, Mow et al., 1980; Holmes, 1986). The material properties of chondrocytes have been investigated using a variety of methods including compression of isolated cells within a gel construct, micropipette aspiration and cellular indentation of single cells using atomic force microscopy (Knight et al., 1998; Trickey et al., 2000; Darling et al., 2006). These works found that single chondrocytes are inherently viscoelastic. A possible functional role for articular cartilage viscoelasticity is a mechanical damping of external loads. It has been shown theoretically that the strain rate-dependent viscoelastic response of cartilage reduces periodic alterations of tissue strains (Suh et al., 1995) and cell deformations (Kim et al., 2008) during cyclic
S.-K. Han et al. / Journal of Biomechanics 45 (2012) 2450–2456
compression. However, neither theoretical nor experimental studies have fully addressed the viscoelastic responses of chondrocyte deformations in situ under static and dynamic mechanical loading conditions (Freeman et al., 1994; Knight et al., 1998; Wu and Herzog, 2006; Kim et al., 2008). Pilot results obtained in a recent study on in vivo chondrocyte deformations in the intact knee loaded by muscular contractions suggest that the intrinsic viscoelastic response of chondrocytes substantially differs from that observed in isolated chondrocytes embedded in gel constructs (Abusara et al., 2011; Knight et al., 1998). Since the material behavior of in situ chondrocytes is essential for understanding the relationship between cartilage loading and the adaptive/degenerative responses of chondrocytes, the purpose of this study was to investigate the viscoelastic responses of chondrocytes in their native environment using a novel in situ experimental approach. We hypothesized that the viscoelastic properties of the extracellular matrix and chondrocytes limit cell deformations during cyclic loading.
2. Materials and methods 2.1. Sample preparation Articular cartilage and its subchondral bone were taken from the femoral groove of skeletally mature cows on the day of slaughtering. Cylindrical cores (diameter¼ 6.4 mm) were extracted from the harvested osteochondral blocks (N ¼4, Fig. 1). Cartilage thickness was measured using a digital caliper at opposing circumferential locations to obtain an estimate of the average sample thickness. Fluorescein conjugated dextran (Alexa, excitation: 488 nm, emission: 517 nm; Molecular Probes, OR, USA) was suspended in DMEM (Dulbecco’s Modified Eagle’s Medium, Gibco, OR, USA) at a concentration of 6.7 mg/ml (2.2 mM). The osteochondral cores were incubated in the dextran solution for 4–10 h at 4 1C prior to mechanical testing and confocal imaging. Samples were rinsed in dye-free phosphate-buffered saline (PBS) for 20 min following staining. Cylindrical samples were rigidly fixed in a specimen holder using dental cement. During testing, samples were fully immersed in the PBS solution to prevent dehydration of the cartilage surface.
2.2. Mechanical testing All mechanical tests and imaging were conducted using a custom-designed indentation system that allows for imaging of cells in the intact cartilage attached to its native bone (Fig. 1A and B). Briefly, the system consists of a light-transmissible glass indenter mounted to the stage of an upright laser scanning microscope and a piezoactuator which pushes the tissue sample towards the indenter. These primary components are complemented by a load cell and displacement transducer which,
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coupled with custom-designed software, allow for controlled loading of the cartilage/ bone specimen. The microscope-based indentation system has been described in detail previously (Han et al., 2009). Three different mechanical tests, described in detail below, were conducted to characterize the viscoelastic properties of chondrocytes in their native environment. 2.2.1. Single stress relaxation test A single static compressive load of 10% nominal tissue strain was applied to the surface of the osteochondral blocks using a glass indenter (diameter¼2.00 mm) at an average strain rate of 3.1%/s (Fig. 2A). The target load was held for 20 min to allow the tissue to reach a steady-state. Confocal image sections were recorded through a 40 magnification, 0.8 numerical aperture, 0.17 mm coverglass-corrected objective: (i) before loading; (ii) at 2 and 17 min of relaxation; and (iii) at 1/2, 1, 3, 5, 7, and 10 min after removal of the load. Confocal scans were recorded from the cartilage surface to a depth of 60 mm, thereby capturing surface zone chondrocytes. Optical slice thickness (i.e. perpendicular distance between image planes) was set at 0.5 mm for all scans except for 30 s after load removal, where the optical slice thickness was set at 3 mm only for tissue recovery measurement to reduce scan time and capture the initial recovery of the tissue. Upon completion of the 10 min recovery phase, tissue samples were allowed to fully relax prior to the next test. Tissue samples were removed from the indentation system and rinsed with fresh PBS for 10 min before conducting the next test. 2.2.2. Three cycle compression test Next, a series of three 10% compressive strain loads were applied to the articular surface and held for 2 min each (Fig. 2B). Confocal scans were taken before loading, during each of the three loading and unloading phases; and at 1/2, 1, 3, 5, 7, and 10 min after removal of the load (Fig. 2B). As in the previous test, confocal scans were taken from the cartilage surface to a depth of 60 mm. Optical slice thickness was 0.5 mm for all scans except for 30 s after removal of the final load (3 mm). Tissue samples were allowed to fully relax prior to the next test, as described above. 2.2.3. Dynamic cyclic compression test The final test consisted of a half-sinusoidal compressive load (13% strain) applied for 100 cycles at 0.35 Hz (Fig. 2C). Confocal scans were taken before loading and after loading as outlined in the previous tests. Total tissue strains, local tissue strains (denoted as ECM strain), and compressive cell strains were quantified as described below. 2.3. Confocal image analysis Four cells from each femoral groove core (total of 16 cells) were used for cell morphology analysis. All chondrocytes were located in the superficial zone of the bovine cartilage between the cartilage surface and a depth of 60 mm. Cell height was quantified using a confocal image viewer (LSM Image Browser, Zeiss Inc.). Local compressive ECM strain was calculated by measuring the distance between paired cells (n¼ 3 pairs per sample) in the pre-test condition and at each experimental measurement point. Overall tissue recovery was quantified by measuring the distance from the cartilage surface to the bottom of the glass indenter observed in the images. Tissue strains during the recovery phase were calculated as engineering strains (eE ¼ (h h0)/h0, where h is the deformed tissue thickness and h0 is the original tissue thickness), and local ECM and cell strains were calculated as
Fig. 1. Sample preparation and indentation testing. (A) Photo of the actual indentation system on the x–y stage of a microscope. (B) Schematic illustration of the area marked with the dashed square in (A).
Nominal strain [-%]
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13 10 0.35 [Hz] 5 0 0
1 3 time [m]
5
7
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Fig. 2. Experimental loading protocol and measurement time points for confocal imaging: (A) single static compression test; (B) three-cycle compression test; and (C) dynamic cyclic compression test.
during loading
at 3 min recovery phase
at 7 min recovery phase
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ECM
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Tissue compressive strain [%]
40
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time [m] Fig. 3. Stress relaxation test. ECM: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (Po 0.05: ntissue, #ECM, þcell).
finite deformation strains for the large deformation theory (eF ¼ (l 1)/2, where l ¼ h/h0, h is the deformed distance between paired cells for ECM or height of cell, and h0 is the original distance of paired cells for ECM or height of cell).
2.4. Statistical analysis Tissue, ECM, and cell strains at different experimental time points were compared using one-way repeated measures ANOVA (SPSS 19, IBM) to determine if there was a significant time-effect during recovery. If indicated, Bonferroni posthoc testing was used to compare loaded and recovery time results with the initial unloaded condition. The level of significance was set at a ¼ 0.05. Results are presented as means7 1 standard deviation.
3. Results 3.1. Single stress relaxation test After 2 min of static loading at 10% compressive tissue strain, local compressive ECM strains in the superficial zone were 3971% and average compressive cell strains were 2976%. Local ECM compressive strains and average compressive cell strains increased further to 41 72% and 3175%, respectively, after 17 min of static loading. The cartilage recovered to its original shape 5 min after load removal and the chondrocytes to their original height 5 min after load removal (P40.05 for both,
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Figs. 3 and 4). The local tissue (ECM) strain did not recover completely within 10 min after load removal (P¼0.002, Fig. 3).
3.2. Three cycle compression test Average compressive ECM and chondrocyte strains in the superficial zone were 3672% and 2877% respectively, during each of the three loading cycles of 10% compressive tissue strain. The cartilage tissue did not fully recover during the 2 min unloading phase, and did not recover completely within 3 min after load removal (Po0.05, Fig. 5). Local tissue (ECM) strain and compressive chondrocyte strain were fully recovered to the initial unloaded values within 10 and 5 min, respectively (P40.05 for both, Fig. 5).
before
during load
3 min after
7 min after
surface y x z
Fig. 4. Three-dimensional reconstruction of local matrix (ECM) and cell deformations before loading, during loading, 3 min after load removal, and 7 min after load removal.
during loading phase
3.3. Dynamic cyclic compression test Cartilage tissue fully recovered to its original shape within 3 min following the completion of 100 compressive loading cycles at a frequency of 0.35 Hz (P4 0.05, Fig. 6). The local tissue (ECM) and cellular strains exhibited a similar recovery, returning to their original configurations within 7 min after load removal (P 40.05 for both, Fig. 6).
4. Discussion This study demonstrated the viscoelasticity of articular cartilage and chondrocytes in situ under static and dynamic compression loads. Our results of cellular recovery upon unloading are consistent with a previous study in the intact mouse tibia–femoral joint (Abusara et al., 2011), where it was found that chondrocytes deform rapidly upon loading but take minutes to recover shape and volume upon unloading. However, the results of our study are in contrast with findings from isolated chondrocytes embedded in agarose gel constructs, which recovered immediately and nearly elastically following mechanical unloading (Knight et al., 1998). This difference in chondrocyte recovery between in situ conditions and cells embedded in agarose gel constructs is likely related to the mechanical environment of the cells, as chondrocyte deformation is linked to the deformation of the extracellular matrix (Guilak, 2000). We conclude from these results that chondrocyte mechanics differ substantially between in situ and in vitro conditions, and that the mechanical properties of cells must be considered within the context of the experimental conditions and environment. The extracellular matrix also behaved highly viscoelastically, similar to the cells following compression testing (Fig. 3). It is well recognized that chondrocytes and surrounding ECM are physically connected by adhesion molecules through the pericellular matrix (Guilak et al., 2006), therefore it seems plausible that the
at 3 min at 7 min recovery phase recovery phase
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30
N=2
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40
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time [m] Fig. 5. Three cycle compression test: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼ 5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (P o0.05: ntissue, #ECM, þ cell).
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at 3 min recovery phase
at 7 min recovery phase
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Tissue compressive strain [%]
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16
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8
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0 0
1
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7
ECM, cell compressive strain [%]
18
9 10
recovery phase time [m] Fig. 6. Dynamic compression test: The distance between paired cells was used to determine local tissue deformations and recovery. Scale bar ¼ 5 mm. Cell: Three dimensional reconstruction of cell. Compressive strain graph: quantitative analysis of tissue, ECM and cell height (Po 0.05: ntissue, #ECM, þcell).
Cyclic external load [cm]
[mm]
Time
[μm]
Time
elastic prediction viscoelastic
Time
Fig. 7. Schematic diagram for cyclic movement, load on the knee joint and possible cell deformation response to the cyclic loading of the knee joint.
shape and volume recovery of the cells is intimately linked to the recovery of the ECM, as has been suggested for articular cartilage when exposed to swelling following osmotic loading (Korhonen et al., 2010). Furthermore, given the three orders of magnitude difference in Young’s modulus between chondrocytes and surrounding ECM (Jones et al., 1999; Schinagl et al., 1997), one would expect that chondrocyte deformations are dominated by the ECM deformations in the vicinity of cells rather than the mechanical properties of the chondrocytes. Therefore, we propose that the viscoelastic response of chondrocytes in situ is largely governed by the viscoelastic behavior of the extracellular matrix. It has been well documented that mechanical loading of cartilage enhances chondrocyte biosynthesis (Sah et al., 1989; Kim et al., 1994; Little and Ghosh, 1997; Buschmann et al., 1999). However, excessive loads may damage the cartilage matrix and produce negative cellular effects (Wollheim and Lohmander, 2007). Therefore, it might be of advantage if cell deformations are minimized under conditions of cyclic mechanical loading.
Here we found, in agreement with pilot results in intact joints (Abusara et al., 2011), that cells and ECM deform quickly ( o1 s) upon mechanical loading, while cell shape and cell volume recovery take minutes to complete (Fig. 5). Therefore, for cyclic dynamic loading, cell deformations and volume changes are greatest in the first cycle of loading, but then become negligibly small in subsequent loading cycles, suggesting that one of the in situ properties of the ECM/cell construct is to minimize cell shape and volume changes, thereby allowing cells to reach homeostasis during cyclic mechanical loading. Physiological loading of joints is often cyclic, for example for the lower limb joints during locomotion, thus protecting cells from excessive strains in these situations and minimizing costly, cyclic cell deformations would be advantageous (Fig. 7). The viscoelasticity of the cartilage matrix and chondrocytes is associated with the fluid phase movement and the interaction between fluid and solid matrix within the tissue and the cells (Mow et al., 1980). When articular cartilage degenerates in
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Table 1 Steady-state compressive strains in the tissue, matrix and cells 17 min after applying a 10% nominal compressive tissue strain.
Compressive strain (%)
Tissue
Superficial cartilage
Superficial cell
10
417 2
31 7 5
disease, such as osteoarthritis (OA), material properties of the cartilage matrix and the chondrocytes change substantially (Alexopoulos et al., 2003, 2005; Hasler and Herzog, 1998; Seedholm et al., 1979). For instance, in early OA, the cartilage matrix becomes softer and permeability increases (Hasler and Herzog, 1998; Seedholm et al., 1979). Additionally, the collagen network in the superficial zone of early OA cartilage has been shown to undergo a disruption and remodeling process (Stockwell, 1991; Guilak et al., 1994; Buckwalter and Mankin, 1997), which may lead to changes in the local matrix environment of the chondrocytes. It has been suggested that these altered material properties of osteoarthritic cartilage affect cell deformation (Kim et al., 2008), and cell deformation-based signaling pathways (Kim et al., 1994). In order to understand the link between OA onset and progression, it would be important to investigate how disease-related changes in articular cartilage viscoelasticity affect chondrocyte deformations, signaling, and macromolecule synthesis. Cell deformations were affected by the viscoelastic properties of the local cartilage matrix in this study. Furthermore, the mechanical properties of the pericellular matrix likely also affected cell mechanics (Poole et al., 1987; Knight et al., 2001). It has been reported that the pericellular matrix provides a unique mechanical environment for chondrocytes and may protect cells from damaging strains in conjunction with the ECM (Youn et al., 2006). For instance, the pericellular matrix acts as a non-linear mechanical filter for controlling cell deformations under mechanical compression (Choi et al., 2007) and modulating chondrocyte biosynthesis (Guilak et al., 2006). The pericellular matrix has viscoelastic material properties, similar to the chondrocytes and extracellular matrix, and these properties are known to be altered when cartilage degenerates (Alexopoulos et al., 2003). Therefore, in order to completely understand the viscoelastic behavior of chondrocytes in situ, the detailed viscoelastic properties of the PCM and their potential effect on chondrocyte mechanics need to be studied further in the future. Consistent with previous studies (Guilak et al., 1995; Han et al., 2010; Schinagl et al., 1997), we found that the superficial zone of cartilage exhibited much greater compressive strains than the average strain imposed on the tissue (Table 1). It is well accepted that extracellular matrix (ECM) and chondrocyte deformations are depth dependent under static compression (Guilak et al., 1995; Schinagl et al., 1997). The superficial matrix and chondrocytes are expected to deform more than the matrix and cells in the deeper layers of the cartilage. With the indentation system and the scanning laser confocal microscopy approach used in this study, we are limited to studying the extracellular matrix and cellular deformations up to about 60 mm below the articular surface. Attempts at numerical modeling of chondrocytes subjected to cyclic loading also showed depth dependent cell deformation (Wu and Herzog, 2006). However, they did not capture the highly viscoelastic cell responses we observed here experimentally in situ, but rather showed cell deformations and recovery that were almost identical in scale, thereby predicting large cell shape and volume changes during each loading cycle. Biosynthetic responses of chondrocytes are also known to be depth dependent (Wong et al., 1997). For example, aggrecan synthesis is thought to occur primarily in deep zone chondrocytes
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(Wong et al., 1997), while proteoglycan-4 synthesis is localized in superficial zone cells (Nugent-Derfus et al., 2007). Furthermore, cartilage compression at different rates affects the biosynthetic response of chondrocytes (Buschmann et al., 1999). Together, these results suggest that cyclic mechanical loading produces depth dependent mechanical and biological responses. Therefore, studies in which chondrocytes at all depths can be visualized need to be performed to fully understand the functional roles of chondrocytes in the different structural layers of articular cartilage. In this study, we focused on the recovery phase of the tissue, the local extracellular matrix and chondrocytes following mechanical compression. Although we aimed at investigating the tissue, local ECM, and cellular deformations during cyclic loading, it was technically difficult to capture three-dimensional ECM and cell morphologies during the dynamic phases when deformations occurred rapidly. Furthermore, it was also technically difficult to capture the tissue recovery right after load removal. Therefore, the initial measurement of tissue recovery, right after load removal, was conducted using 3 mm thick slices, while all other measurements were made using 0.5 mm slices. This allowed for capturing the fast tissue recovery immediately after unloading, while providing excellent accuracy for cell shapes and volumes for measurements in near steady-state conditions. Future studies need to consider how the temporal resolution of confocal imaging can be improved substantially without losing spatial resolution, so that cell deformation and signaling can be studied during highly dynamic conditions. It seems to us that these dynamic conditions, that elude quantification with present day technology, may hold the key to our understanding of cartilage mechanobiology and to the mechanisms of cartilage adaptation and degeneration.
5. Conclusion The results of this study led to the conclusion (i) that chondrocyte mechanics are greatly affected by the local environment, thus cells in agarose gel and in the native cartilage matrix behave very differently, and (ii) that chondrocytes in their native environment have properties that minimize cell deformations during cyclic loading. We suggest that this latter property allows chondrocytes to achieve homeostasis during dynamic loading in a short period of time.
Conflict of interest statement The authors have no conflicts of interest.
Acknowledgments The authors would like to thank A. Jinha from the Human Performance Laboratory at the University of Calgary for assistance with LabView programming, the Canada Research Chair Programme, the Killam Foundation, the Canadian Institutes for Health Research, and the Alberta Innovates—Health Solutions Team Grant on Osteoarthritis. References Abusara, Z., Seerattan, R., Leumann, A., Thompson, R., Herzog, W., 2011. A novel method for determining articular cartilage chondrocyte mechanics in vivo. Journal of Biomechanics 44, 930–934. Alexopoulos, L., Haider, M.A., Vail, T.P., Guilak, F., 2003. Alterations in the mechanical properties of the human chondrocyte pericellular matrix with osteoarthritis. Journal of Biomechanical Engineering 125, 323–333.
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