Department of Orthopedic Surgery, Brigham and Womens Hospital, Harvard Medical School, Boston, Massachusetts
Submitted 8 March 2004 ; accepted in final form 8 October 2004
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ABSTRACT |
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real-time optical measurement; intracellular calcium; mechanosignal transduction
We attempted to characterize the change in [Ca2+]i in response to the application of pure HP alone to chondrocytes in vitro. To elucidate real-time changes in [Ca2+]i due to HP at the single-cell level, we developed a pressure-proof optical chamber and HP application apparatus to conduct imaging of individual cells immediately following the application of pure hydrostatic fluid pressure (HFP).
The development of this method is a major breakthrough because it allows measurement of real-time changes in cellular composition in response to HFP as single-cell measurements. To evaluate motion-free fluid condition of adjacent cells, we measured the real-time motion of microbeads and diffusion profile of fluorescent dye with the same protocol developed for [Ca2+]i measurement. To mimic the extracellular environment, isolated chondrocytes should be allowed to accumulate some pericellular and extracellular matrix (ECM) and to maintain phenotype. Furthermore, in native articular cartilage, middle-zone (MZ) chondrocytes are surrounded by highly sulfated ECM and sandwiched between distinct surface (SZ) and deep zones (DZ). These depth-dependent zones of tissue organization have distinctive organization because cell shape, matrix components, pathophysiology, and stress distribution differ among the zones (8, 17, 24, 29, 35). However, little is known about zone-dependent characteristics due to mechanical stimuli with pure HP alone: HFP. Therefore, in this series of experiments, we used chondrocytes derived from distinct zones preincubated up to 5 days to allow ECM accumulation.
This study tests the hypothesis that cells in different depth zones from articular cartilage have unique mechanosensitivity due to pure HP. If our hypothesis proves correct, the mechanism of calcium signal cascade would be of importance. To elucidate the mechanism of dynamic mechanosignal transduction due to HFP, we measured [Ca2+]i after application of HFP in cultured confluent chondrocytes by using a fluorescent indicator (X-rhod-1 AM), a custom-made novel pressure-proof optical chamber and culture system, and laser confocal microscopy. To examine the calcium signal cascade, we used a calcium channel inhibitor (verapamil), a stretch-activated channel blocker (gadolinium), a calcium storage inhibitor (dantrolene), and a calcium-free buffer with EGTA.
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MATERIALS AND METHODS |
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Validation of fluid temperature during HFP application experiments. The temperature of water in the pressure-proof chamber before, during, and after HFP application was monitored using a water-proof thermosensor (E3M-42D; Sibauradenshi, Saitama, Japan) placed in the chamber and connected to a data logger (8421-51; Hioki USA, Cranbury, NJ). The chamber was filled and injected with HPLC-grade water (Sigma, St. Louis, MO) to insulate and minimize electrical noise. The electrical connection between the thermosensor and data logger was sealed with an epoxy adhesive to prevent water invasion. A reference thermosensor placed outside the chamber automatically recorded the temperature every 5 s.
Measurement of fluid flow between the coverslip and sapphire glass window. Fluid flow between the coverslip, where cells adhered, and the sapphire glass window was measured with microbeads as an indicator of fluid motion. One million rhodamine-labeled polystyrene microbeads (2-µm diameter; Sigma) were placed inside the pressure-proof chamber, at the center of its sapphire glass window, and a glass coverslip was placed over two suspension rails (Fig. 1D). The chamber was gently filled with balanced salt solution (BSS: 130 mM NaCl, 5.4 mM KCl, 20 mM HEPES, 2.5 mM CaCl2, 1 mM MgCl2, and 0.1% glucose, pH 7.4) to minimize dispersion of the beads. Fluorescent images of the beads were acquired using a x40 objective lens (Nikon, Melville, NY) attached to an inverted microscope (Eclipse TE-300; Nikon) and a video system (Sony, Tokyo, Japan). Real-time images were recorded as a digital movie, which was converted to individual frames at 1-s intervals for 50 s. Images of the beads were recorded under three experimental conditions: no flow, flow at 0.1 ml/min, and HFP at 0.5 MPa, 0.1 ml/min. Five beads in the frame were chosen, including a bead that did not move, three freely moving beads with no surrounding obstruction, and one bead in close proximity to another bead. The motion of these beads was representative of all other beads in the chamber. The x-y position of each bead was measured on each frame.
Measurement of fluorescent dye diffusion between the coverslip and sapphire glass window. Fluid flow between the coverslip and sapphire glass window was also measured using fluorescent dye as an indicator of diffusion and, consequently, of fluid motion. Fluorescein isothiocyanate (FITC)-dextran (mol/wt 70,000; Sigma) was dissolved in 1.5% sodium alginate solution (Inotec Biosystems International, Rockville, MD). One microliter of the FITC-dextran-alginate solution was dropped into a solution of 100 mM CaCl2 and 10 mM MOPS to form a FITC microcapsule. One FITC microcapsule (<100 µm in diameter) was placed in the chamber at the center of the sapphire glass window (Fig. 1E). A cover glass was placed on suspension rails, and then the chamber was filled with calcium-magnesium free BSS with EDTA (130 mM NaCl, 5.4 mM KCl, 20 mM HEPES, 0.1% glucose, and 10 mM EDTA, pH 7.4). The pressure-proof chamber was placed on an inverted microscope (Eclipse TE-300; Nikon) attached to a 2x objective lens (Nikon) and a digital camera (SPOT; Diagnostic Instruments, Sterling Heights, MI). Fluorescent images of the FITC-dextran were taken every 1 min at constant HFP at 0.5 MPa, 0.1 ml/min, for the first 5 min and then at no flow and no HFP for 5 min by using the same protocol as for [Ca2+]i measurement. Calcium alginate capsule-encapsulating FITC-dextran was gradually degraded with calcium-free BSS-EDTA buffer, allowing the FITC-dextran to diffuse out. Positions (x- and y-axis) of the fluorescent periphery of the microcapsule were measured with NIH Image software. The distance between the periphery of fluorescent dye and initial periphery of the bead was used to calculate diffusion profile.
Cell isolation and culture.
Bovine shoulders from 10 calves 23 wk old were obtained from a local abattoir. Pieces of cartilage (3 x 3 x 3 mm) with subchondral bone were harvested from the weight-bearing region of the humeral articular cartilage. Under a stereomicroscope, a no. 15 surgical blade and a 100-µm guide scale were used to divide the pieces into three zones: 100150 µm from the surface (SZ), 200400 µm from the subchondral bone (DZ), and the remaining section as the middle layer (MZ). The pieces were minced, rinsed with phosphate-buffered saline (PBS) three times, and digested with a solution of 0.15% collagenase (CLS 1; Worthington Biochemical, Freehold, NJ) in Hams F-12 medium (Invitrogen, Grand Island, NY) with gentle shaking overnight at 37°C. The isolated cells were rinsed in PBS three times.
Round glass coverslips (15-mm diameter; Fisher Scientific, Pittsburgh, PA) were coated with a 0.1% collagen type I solution (Vitrogen; Cohesion, Palo Alto, CA) and air-dried. A 150-µl aliquot containing 2 x 105 cells was deposited onto the coverslips. They were incubated in Hams F-12, 10% fetal bovine serum, 100 U/ml penicillin, and 100 µg/ml streptomycin (Invitrogen) for 2 h in a humidified incubator with 5% CO2 in air at 37°C, during which time the cells adhered to the coverslip. The coverslips were transferred to 12-well plates (Falcon; Becton Dickinson, Franklin Lakes, NJ), and 2 ml of medium were added. In preliminary experiments, chondrocytes became unstable and detached from the coverslip after 7 days in culture. Therefore, we incubated cells for 5 days in this experiment.
Colorimetric assay for sulfated glycosaminoglycan and DNA. The cells attached to each coverslip were digested with 1 ml of 125 µg/ml papain (Sigma) in 5 mM cysteine-HCl, 0.05 M EDTA, and 0.1 M sodium phosphate for 16 h at 60°C (19). Two milligrams of sodium formate (Sigma) were dissolved in 2 ml of formic acid (Sigma) and mixed with 900 ml of deionized water; 16 mg of 1,9-dimethylmethylene blue (DMB; Sigma-Aldrich, Milwaukee, WI) were dissolved in 5 ml of absolute ethanol and added to the formate solution, which was brought to a final volume of 1,000 ml with deionized water (9). A 30-µl volume of each papain-digested sample was added to 150 µl of the DMB solution in a 96-well titer plate (Falcon, Becton Dickinson). The optical density of the sample was measured at 540 and 595 nm with a microtiter plate reader (model 550; Bio-Rad, Cambridge, MA). The sulfated glycosaminoglycan (S-GAG) concentration was estimated from a standard curve for shark chondroitin sulfate (Sigma). The papain-digested samples (10 µl) were added to a solution of 2 ml of Hoechst 33258 (4.0 µg/ml; Polysciences, Warrington, PA), 0.1 M NaCl, 10 mM Tris·HCl, and 1 mM EDTA, pH 7.4. The sample was measured with a fluorometer (TKO 100; Hoefer, San Francisco, CA) with calf thymus DNA as the standard (Clontech, Palo Alto, CA).
Measurement of cell viability after application of HFP. Cell viability was evaluated after treatment with constant HFP at 0.5 MPa for 5 min in BSS. After the treatment, the cells (on the coverslip) were double-stained with 2 µM calcein-AM and 4 µM ethidium homodimer-1 (Live/Dead viability/cytotoxicity kit; Molecular Probes, Eugene, OR) for 20 min and rinsed with PBS three times. The live cells were identified at an excitation wavelength of 484 nm and an emission wavelength of 530 nm, and dead cells were identified at an excitation wavelength of 568 nm and an emission wavelength >590 nm by using a fluorescence microscope.
Image acquisition of calcium concentration in [Ca2+]i in bovine articular chondrocytes from different zones after application of HFP. The cells were rinsed with PBS twice, incubated with a calcium indicator (5 µM X-rhod-1 dissolved in Hams F-12; Molecular Probes) for 4060 min at room temperature, and rinsed and incubated in BSS for 1020 min. The coverslip was inverted and placed onto stainless steel guide rails (0.5 mm thick) inside the chamber over the sapphire glass window so that the cells were on the other side of the coverslip from the window, which was at the bottom of the chamber (Fig. 1B). Thus the cells were protected from deformation by the sapphire glass during application of HFP while suspended in buffer fluid. A water drop was placed on the water-immersion objective lens (long working distance, 3.3 mm; numerical aperture, 0.80; Olympus America), and the pressure-proof chamber was positioned on an inverted microscope stage (Diaphot 200; Nikon). This sample preparation for imaging acquisition required especially gentle handling to avoid inducing artifacts.
Constant HFP at 0.5 MPa for 5 min was applied by pumping BSS at 0.1 ml/min into the chamber (Fig. 1, B and C). The magnitude and duration of HFP and the pumping rate of BSS were optimized from preliminary experiments (data not shown). After HFP application, real-time fluorescent imaging of [Ca2+]i in a single focal plane was acquired with a confocal inverted microscope system (Noran Instruments, Middleton, WI). For these images, we used a confocal slit of 100 µm and a scan speed of 800 ns at 10-s intervals for 290 s at 568 nm (excitation) and 590+ nm (emission) (Fig. 1C). From preliminary experiments, the slit width and the scan speed were chosen to optimize resolution, fluorescent signal intensity, and total exposure time of the laser beam. Although a substantial optical signal was acquired from all cells without the use of the slit mode, spatial resolution was reduced. At the chosen setting, the baseline fluorescence of the nontreated control was maintained at the initial level. The peak fluorescence intensity was expressed as the peak increase in fluorescence (%increase over baseline). Experimental conditions included no flow and flow at 0.1 ml/min as no HFP controls, or calcium ion-free BSS with EGTA (130 mM NaCl, 5.4 mM KCl, 20 mM HEPES, 1 mM MgCl2, 0.1% glucose, and 10 mM EGTA, pH 7.4).
Image acquisition of [Ca2+]i in MZ-derived bovine articular chondrocytes treated with inhibitors after application of HFP. The mobility of calcium ions participating in the increase in [Ca2+]i was elucidated by adding the membrane stretch-activated channel blocker gadolinium (5 µM), the cytosolic calcium storage blocker dantrolene (5 µM), or the membrane calcium channel blocker verapamil (20 µM) to X-rhod-1 in Hams F-12 medium and then BSS. Constant HFP at 0.5 MPa, 5 min was applied to the cells with each calcium inhibitor, and [Ca2+]i was measured as described above. The samples were placed onto rails in the chamber by using the same methods as described above. A water drop was placed on the water-immersion objective lens (long working distance, 3.3 mm; numerical aperture, 0.80; Olympus America), and the pressure-proof chamber was positioned on a custom-made stage of an inverted microscope (Axiovert; Zeiss, Thornwood, NY) attached to a laser confocal system (MRC 1024; Bio-Rad, Hercules, CA). The fluorescent images of cells stained with X-rhod-1 were imaged with a resolution of 516 x 516 pixels in a full frame. With 10% laser power at 568 nm (excitation) and 590+ nm (emission), the images were acquired at normal scan speed every 10 s for 290 s at atmospheric pressure after 5-min HFP application at 0.5 MPa and BSS injection at 0.1 ml/min. The images were converted to TIFF files and analyzed for total fluorescence intensity with NIH Image software. At the chosen setting, the baseline fluorescence of the control was maintained at the initial level. The peak fluorescence intensity was expressed as the peak percentage increase in fluorescence over baseline.
Data analysis. For the zone-dependent experiment, eight cells from one coverslip culture were randomly selected in one frame of the display and isolated as a region of interest, and fluorescence intensity was measured. In the calcium inhibitor experiment, a frame of each sample was randomly selected and the fluorescence intensity was measured. A total of eight to twelve coverslips (samples) were measured at each condition over at least three separate isolations of chondrocytes to minimize sample-to-sample variation. The peak value of each sample represented [Ca2+]i, and comparisons were made using ANOVA for statistical significance.
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RESULTS |
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Fluid flow between the coverslip and sapphire glass window. The x-y position of each bead was traced and plotted at 1-s intervals for 50 s (Fig. 2). At each test condition (no flow with no HFP, flow with no HFP, and HFP with flow), the beads moved around within 36 µm of their original locations. Occasionally, a few beads stayed in the same position on the surface of the sapphire glass window, as determined by a lack of motion in the x-y axis, and yet motion was evident in the z-axis in focus due to deformation of the sapphire glass window.
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After the application of HFP at constant 0.5 MPa for 5 min, peak fluorescence in the MZ-derived cells increased twofold compared with the initial level (Fig. 7, a and b). The peak fluorescence value was detected 40120 s (73 ± 27 s) after HFP application. Fluorescence in SZ-derived cells increased 1.5-fold compared with the initial level but was 50% lower than in MZ-derived cells (P < 0.05; Fig. 8). The peak fluorescence value was detected 20140 s (72 ± 34 s) after HFP application. Fluorescence in DZ-derived cells increased 1.5-fold compared with the initial level but was 50% lower than in MZ-derived cells (P < 0.05; Fig. 8). As in the untreated control groups, a slight increase in the peak fluorescence in MZ-derived cells was detected with no-flow conditions (without HFP application or pumping) and with flow alone at 0.1 ml/min by pumping compared with baseline (Fig. 8). In the presence of a calcium ion-free buffer, the effect of HFP was reduced to 13% (P < 0.01) of the control with no calcium inhibitor (Fig. 8).
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DISCUSSION |
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Fluid flow between the coverslip and sapphire glass window was measured using the same protocol as for [Ca2+]i measurement in bovine articular chondrocytes. Movement of the beads in all three conditions seemed to be within a margin of Brownian motion. Because the movement was similar in all three conditions, there is minimal fluid flow near the cells, and, consequently, the cells are experiencing pure HFP. Thus our experimental model measuring dynamic [Ca2+]i due to pure HFP application by using a laser confocal microscope system is feasible.
During weight bearing and joint loading, the magnitude of HP placed on articular cartilage is approximately <10 MPa. Cartilage tissue is an incompressible material at the physiological HP (1). Nevertheless, HP itself is transmitted to all points intracellularly and extracellularly because the cytosolic membrane of the cultured chondrocytes is the only partition separating extracellular and cytosolic fluid. To elucidate the mechanism of dynamic mechanosignal transduction in single cells, we developed a novel, pressure-proof optical chamber that allowed measurement of real-time changes in [Ca2+]i with a fluorescent indicator and a confocal imaging system.
This study focused on the mechanism of [Ca2+]i increase due to HFP. MZ-, SZ-, and DZ-derived cells cultured for 5 days showed a significant increase in [Ca2+]i due to the application of HFP at 0.5 MPa for 5 min. However, after 2 days of culture, the peak fluorescence of MZ cells did not increase upon application of HFP at 0.5 MPa (data not shown). The degree of response in [Ca2+]i due to application of HFP depended on the zonal origin of the cell fractions. We examined the most highly responsive cell fraction (MZ-derived cells) under conditions of optimal duration and magnitude of HFP.
It is reasonable that mechanosignal transduction in MZ-derived cells themselves and/or their extracellular environment could differ from that in SZ- and DZ-derived cells. A key factor in mechanosignal transduction in MZ-derived cells was that cells required an extended culture period to respond to HFP. In addition to morphological differences between MZ-derived cells and SZ-derived cells, S-GAG accumulation was significantly greater for MZ-derived cells than for SZ-derived cells. Consistent with our findings are those indicating that the MZ of native tissue contains more sulfated ECM than do cells from other zones (10, 27, 31). After 5 days of culture, tissue depth-based cellular characteristics were reflected in morphological differences among cells, ECM, and free Ca2+ (20) and were subsequently influenced differently by HFP.
Another difference among cell fractions is that of stress distribution in native cartilage tissue with depth. MZ cells in vivo are subjected to more HP than distortional stress during weight bearing and joint motion (8). We formulated the working hypothesis that mechanosignal transduction of MZ-derived cells is stimulated predominantly by HP rather than by shear stress and that SZ-derived cells are stimulated predominantly by shear stress. Our results are consistent with this hypothesis, because the increase of [Ca2+]i in SZ-derived cells was less than that in MZ-derived cells.
It is unlikely that interstitial fluid flow accounts for our data. The [Ca2+]i in cells responding to fluid flow (shear stress) has been shown to increase with relatively large increases in flow rate [934 ml/min: 1037 dyn/cm2 (37); 16 kPa (16)] and to be inhibited by gadolinium (37), although the magnitudes of these increases were not equivalent to physiological changes in interstitial fluid flow [velocity: 100 µm/min (23); 12 µm/s (11)]. Our control experiment used continuous injection of BSS alone at 0.1 ml/min, which sustained peak fluorescence at baseline levels. Thus these results indicate that the change in magnitude of flow used in our study was negligible.
Our results do not address the effects of deformation (distortional stress involved) on [Ca2+]i. Compressive strain has been shown to induce a delayed increase in [Ca2+]i after the application of deformation in an agarose/cell construct (28). However, our data indicate that peak fluorescence increased 50150 s after HFP application, which was earlier than the change in peak calcium that occurred 200 s after deformation in the study by Roberts et al. (28). They speculated that upstream mechanosignal transduction might be caused by strain. Their data were acquired after application of compression. With sustained compressive strain on an agarose construct, both HFP and strain increased, then HFP gradually decreased (equilibrated), and strain was maintained during [Ca2+]i measurement. The ensuing time lag in peak [Ca2+]i between Robertss and our data may be due to the difference between the immediate HFP release in our experimental setting and a slower HP release in Robertss experiment as well as to the existence of a separate signal cascade, as per their hypothesis.
We sought to characterize the mechanism of the effects of HFP through the use of inhibitors of calcium mobility. Our data show inhibition of an HFP-stimulated increase in peak fluorescence in the presence of a stretch-activated channel blocker (gadolinium) or a calcium-free medium (EGTA). These same inhibitions were also found with other means of applying mechanical stress, i.e., poking (13) and fluid shear stress (36). In other studies, gadolinium was found to delay peak responses to mechanical forces (15). In studies with bone cells, calcium was increased during 3-min fluid flow (15) and after 25 s in response to 10-s cyclic HP at 10 lb/in2, 1 Hz (2). Increased [Ca2+]i at an early time point strongly suggests the involvement of inositol 1,4,5-trisphosphate (IP3)-mediated calcium increase due to HP in bone cells (2). In our experiments with chondrocytes, it is possible that early IP3-mediated calcium release from intracellular calcium storage could occur during the 5-min application of HFP. This speculation is supported by a parallel study. Wilkins et al. (32) showed that shorter (30 s) bursts of HFP stimulated [Ca2+]i via an IP3-dependent cascade. In addition, when calcium ions from cytosolic storage were blocked with dantrolene, the application of HFP inhibited an increase in their peak fluorescence. Dantrolene inhibits calcium release from storage, as in "calcium-induced calcium release" (5, 25). The current data were not sufficient for us to distinguish the order of events from the apparent magnitude. Calcium ion influx can follow release from stores (26), so if store depletion has been inhibited by dantrolene, an enhancement of influx will still be seen. It may be acceptable at present to offer two options: calcium influx triggers release, or IP3 triggers calcium release due to HFP and then flux. We propose that calcium flux was initiated through a stretch-activated channel, because verapamil did not inhibit the increase in [Ca2+]i.
Our novel, pressure-proof apparatus has made it possible to measure real-time [Ca2+]i attributable to HFP. In future studies, we need to elucidate the mechanosignal cascade due to HP application by using other pharmacological inhibitors and varied algorithms and magnitudes of HFP. Our present study indicates that an increase in peak fluorescence intensity due to HFP was related to the zonal origin of chondrocytes and the presence of ECM accumulation. Maturity of chondrocytes and pathological conditions may be encountered and reflected by sensitivity to HFP. The stimulation of the calcium cascade in these studies supports the positive effects of HFP on chondrogenesis in three-dimensional culture (22). This novel methodology will allow the responses of other signal cascades and cell types to be examined in response to a wide range of magnitudes of pressure (21).
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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