A novel method for assessing effects of hydrostatic fluid pressure on intracellular calcium: a study with bovine articular chondrocytes

Shuichi Mizuno

Department of Orthopedic Surgery, Brigham and Women’s Hospital, Harvard Medical School, Boston, Massachusetts

Submitted 8 March 2004 ; accepted in final form 8 October 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Chondrocytes in articular cartilage are exposed to hydrostatic pressure and distortional stress during weight bearing and joint loading. Because these stresses occur simultaneously in articular cartilage, the mechanism of mechanosignal transduction due to hydrostatic pressure alone in chondrocytes is not clear. In this study, we attempted to characterize the change in intracellular calcium concentration ([Ca2+]i) in response to the application of hydrostatic fluid pressure (HFP) to cultured bovine articular chondrocytes isolated from defined surface (SZ) and middle zones (MZ) by using a fluorescent indicator (X-rhod-1 AM), a novel custom-made pressure-proof optical chamber, and laser confocal microscopy. Critical methodology implemented in this experiment involved application of high levels of HFP to the cells and the use of a novel imaging apparatus to measure the peak [Ca2+]i in individual cells. The peak [Ca2+]i in MZ cells cultured for 5 days showed a significant twofold increase after the application of HFP at constant 0.5 MPa for 5 min. The peak [Ca2+]i in SZ cells was lower (43%) than that of MZ cells. The peak was suppressed with an inhibitor of dantrolene, gadolinium, or a calcium ion-free buffer, but not with verapamil. This study indicated that the increase in [Ca2+]i in chondrocytes to HFP is dependent on the zonal origin. HFP stimulates calcium mobilization and stretch-activated channels.

real-time optical measurement; intracellular calcium; mechanosignal transduction


THE HYDROSTATIC STRESS AND DISTORTIONAL STRESS that occur during weight bearing and joint loading in articular cartilage are integrated and interacting mechanobiological forces (4). Technically, it is difficult to distinguish between pure hydrostatic stress (pressure and tension) and pure distortional stress (shear stress) because compressive loading on a tissue or reconstituted cell construct (11) (e.g., agarose gel) involves gross tissue deformation. At the cellular level, both direct mechanical stress induced by poking cells (involving deformation) with a glass needle (6, 7, 12, 13) and indirect deformation induced by compressing cells in either an agarose construct (18, 28, 34) or a monolayer culture (30, 33) immediately increased intracellular calcium concentration ([Ca2+]i) in chondrocytes. The types of stress examined in previous studies included cellular deformation but not pure hydrostatic pressure (HP) alone in individual cells. [Ca2+]i and other ion transport in suspended chondrocytes have been examined at extremely high HP (3, 14) with a pressure vessel. It was necessary to examine changes in [Ca2+]i due to HP in an in vivo-like environment and at physiological HP.

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.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
HFP application apparatus and pressure-proof chamber. The pressure-proof imaging system was composed of a buffer container, a high-performance liquid chromatography (HPLC) isocratic pump (Acuflow series I; SSI, State College, PA), a pressure sensor attached to a custom-made pulse damper, a custom-made pressure-proof chamber, and a computer-assisted back-pressure controller (Takagi Industrial, Shizuoka, Japan) (Fig. 1A).



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Fig. 1. A: image acquisition system. Balanced salt solution (BSS) is injected into a pressure-proof chamber with an HPLC isocratic pump. The hydrostatic fluid pressure (HFP) in the chamber is regulated with a needle valve attached to a spring and a computer-driven actuator. B: the chamber has sapphire glass windows (2 mm thick and 30 mm in diameter, 20-mm-diameter available optical field) and holds up to 10 MPa pressure. The coverslip was inverted and placed onto stainless steel suspension rails (0.5 mm thick) over the sapphire glass window in the chamber so that the cells opposed the sapphire glass window and faced the lens. Thus the cells were isolated from deformation of the sapphire glass during application of HFP. C: HFP profile and a protocol of imaging acquisition. The chamber was positioned on the microscope stage, and constant HFP at 0.5 MPa for 5 min at 0.1 ml/min was applied by pumping BSS into the chamber. D: experimental setting used to measure fluid flow between the coverslip and the sapphire glass window, using rhodamine-labeled microbeads. E: experimental setting used to measure fluorescent dye diffusion between the coverslip and the sapphire glass window, using FITC-labeled alginate microcapsule.

 
The pressure-proof chamber, designed for application of HFP up to 10 MPa (Fig. 1B), was made of stainless steel with a sapphire glass window (30-mm diameter, 2-mm thickness) at the bottom and top of a cylinder body (20-mm diameter, 30-mm depth, 9-ml volume; Taiatsu Glass, Tokyo, Japan). The back-pressure controller was composed of a valve, a reverse spring (to open the valve completely), and a spring-loaded actuator. A set maximum HFP was regulated using a compressing spring. Reduction of HFP was controlled using a relaxing spring driven by an actuator. The medium reservoir, pressure-proof chamber, and outlet for drain tubing were placed at the same horizontal level to maintain atmospheric pressure. The buffer solution was injected into the chamber at a constant rate of 0.1 ml/min. Because of the pump stroke, the set constant pressure fluctuated ±10%. Other system components, such as tubing, ferrules, and connectors, were made of chemically inert polyetheretherketone and Tefzel (Upchurch, Oak Harbor, WA). All chamber units were maintained at room temperature.

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 2–3 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: 100–150 µm from the surface (SZ), 200–400 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 Ham’s 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 Ham’s 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 Ham’s F-12; Molecular Probes) for 40–60 min at room temperature, and rinsed and incubated in BSS for 10–20 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 Ham’s 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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Temperature changes due to fluid flow and HFP. The temperature of water in the pressure-proof chamber drifted ±0.03°C during HFP application and ±0.03°C during image acquisition (no HFP and no flow). Room temperature during the experiment was ±0.06°C.

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 3–6 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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Fig. 2. Real-time microbead movement with and without HFP treatment. A: 1 frame of the real-time tracking. The numbers 1 to 5 indicate the tracing of a single bead. The x-y position of each bead is shown at 1-s intervals for 50 s under conditions of no HFP and no BSS injection (B), BSS injection at 0.1 ml/min (C), and HFP at 0.5 MPa and BSS injection at 0.1 ml/min (D).

 
Diffusion of fluorescent dye between the coverslip and sapphire glass window. The distance between the periphery of fluorescence and an initial radius of the fluorescent bead in four directions (y± and x±) were within 100 µm during HFP application at 0.5 MPa, 0.1 ml/min, for the first 5 min (Fig. 3). After the HFP was released, the distance in each direction increased linearly with time to 350–450 µm. Because the objective lens (x2) has a long focus range, transitional changes from HFP to no flow and HFP (time-releasing HFP) were monitored for fluid flow detection without optical interruption changing the focus. Irregular diffusion of the fluorescence in each direction was not detected.



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Fig. 3. Photomicrograph of fluorescent dye diffusing from alginate microcapsule at time 0 (A) and 10 min (B). C: diagram of distance measured between the initial position of fluorescence and peripheral fluorescence with time. Dy±, distance on the ±y-axis; Dx±, distance on the ±x-axis. D: distance of diffusing fluorescence periphery from initial position taken at 1-min intervals.

 
Cell morphology, viability, and ECM accumulation. Two days after seeding, cells from the different zones had different morphologies: SZ-derived cells were stellate or round, MZ-derived cells were homogeneously polygonal or round, and DZ-derived cells were relatively larger and round. For all cell fractions, cell density increased, intercellular spaces decreased with time, and cells were confluent 5 days after seeding, but cell morphologies were similar to those seen at day 2 (Fig. 4).



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Fig. 4. Phase-contrast photomicrographs of bovine articular chondrocytes derived from the surface zone (SZ) (A), middle zone (MZ) (B), and deep zone (DZ) (C) after 5 days of culture (x50). SZ-derived cells were stellate or round, MZ-derived cells were homogeneously polygonal or round, and DZ-derived cells were relatively larger and round. The intercellular spaces decreased in size with time, and the cells were confluent 5 days after seeding.

 
After 5 min of no-flow conditions followed by 5 min of 0.5-MPa constant HFP with or without inhibitors, there was no evidence of decreased viability compared with preloaded cultures, because very few dead cells were seen (Fig. 5).



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Fig. 5. Double staining of live and dead MZ-derived chondrocytes. The cells were incubated with calcein-AM and ethidium homodimer-1 fluorescent indicators after treatment with constant HFP at 0.5 MPa for 5 min. The live cells (A) and dead cells (B) were identified using a fluorescence microscope (x20). No dead cells were detected in the same frame as any live cells.

 
The S-GAG accumulation in MZ-derived cells increased 2.7-fold from day 2 (5.1 µg/µg DNA) to day 5 (13.7 µg/µg DNA, P < 0.01) and was 4.3 times and 2.1 times greater than the accumulation in SZ-derived (3.2 µg/µg DNA, P < 0.01) and DZ-derived cells (6.4 µg/µg DNA, P < 0.01), respectively, on day 5 (Fig. 6).



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Fig. 6. Accumulation of sulfated glycosaminoglycan (S-GAG) based on the original location of cells. The accumulation of S-GAG in MZ-derived cells increased 2.7-fold from day 2 to day 5 (P < 0.01) and was 4.3 times and 2.1 times greater than the accumulation in SZ-derived cells (P < 0.01) and DZ-derived cells (P < 0.01), respectively.

 
Effects of HFP on intracellular calcium concentration. After 5 min of exposure to constant HFP at 0.5 MPa and after 290 s of image acquisition, either with or without calcium inhibitors or calcium-free buffer, chondrocytes remained attached to the coverslips.

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 40–120 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 20–140 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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Fig. 7. A: representative imaging of MZ-derived cells at 0 (I), 60 (II), and 290 s (III) after HFP application at constant 0.5 MPa, 0.1 ml/min. Pseudocolor indicates fluorescence intensity of 5 µM X-rhod-1. B: representative peak fluorescence of MZ-derived cells after constant HFP at 0.5 MPa, 0.1 ml/min. The cells were loaded with a calcium indicator, 5 µM X-rhod-1-AM for 40–60 min at room temperature. HFP was applied with BSS for 5 min before image acquisition. Dynamic fluorescent imaging of intracellular calcium concentration ([Ca2+]i) at a single focal plane was acquired with a confocal microscope system at an 800-ns scan speed at 10-s intervals for 5 min at 568 nm (excitation) and 590+ nm (emission).

 


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Fig. 8. Effects of original depth zone of cells on change in intracellular calcium stimulated at 0.5 MPa for 5 min. Peak fluorescence intensity of cells was determined using cells from each zone. No flow, no flow of BSS; flow, BSS flow rate of 0.1 ml/min; Ca-free, calcium ion-free BSS with EGTA. The number above each column represents the number of samples (coverslips). Eight cells from one coverslip culture were randomly selected in one frame of the display, isolated as a region of interest, and converted to fluorescence intensity. Eight to twelve coverslips (samples) over at least three separate isolations of chondrocytes were measured at each condition.

 
MZ-derived cells were used for determining the mechanism of the effect of HFP on calcium mobilization (Fig. 9). In the presence of a cell membrane stretch-activated channel blocker (gadolinium), the effect of HFP on peak fluorescence was reduced to 55% (P < 0.01) of the control with no calcium inhibitor. In the presence of an intracellular storage blocker (dantrolene), the effect of HFP was reduced to 36% (P < 0.01) of the control with no calcium inhibitor. In the presence of a calcium channel blocker (verapamil), the effect of HFP on peak fluorescence was maintained at the same level as the untreated control.



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Fig. 9. Effects of inhibitors of calcium mobilization on change in intracellular calcium stimulated with HFP at 0.5 MPa for 5 min. Peak fluorescence intensity of MZ-derived cells with the following inhibitors: 20 µM verapamil (Vera), a cell membrane channel blocker; 5 µM gadolinium (Gado), a stretch-activated channel blocker; or 5 µM dantrolene (Dant), a cytosolic calcium storage inhibitor. HFP was applied to the cells with each calcium inhibitor. The number above each column represents the number of samples (coverslips). Data represent the means (±SD) of 8–12 coverslips. *P < 0.01 (ANOVA).

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Before conducting experiments aimed at elucidating the effects of HFP, we verified that the temperature of water in the chamber was at room temperature. Changes in temperature of water during fluid flow at 0.1 ml/min and an HFP application at 0.5 MPa, 0.1 ml/min, was determined to be negligible. We determined that the temperature changes were negligible because the temperature fluctuations were within the guaranteed sensitivity range of the thermometer.

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 [9–34 ml/min: 10–37 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); 1–2 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 50–150 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 Roberts’s and our data may be due to the difference between the immediate HFP release in our experimental setting and a slower HP release in Roberts’s 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).


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This study was supported by National Institute of Arthritis and Musculoskeletal and Skin Diseases Grant R03 AR-45793.


    ACKNOWLEDGMENTS
 
We especially thank Ryosuke Fujimori, Kara Johnson, and Rachael Fiske for technical assistance; Scott Pomeroy and Jon Kornhauser at the Image Core Children’s Hospital, Boston, MA, for image acquisition consultation; Stephen Bolsover at the University of London for consultation on fluorescent dyes; Jill P. G. Urban at Oxford University for consultation on cartilage zone physiology; and Julie Glowacki at Harvard Medical School and Robert Wilkins at Oxford University for both critical review and supportive comments. Image acquisition was conducted at the Image Core Children’s Hospital, Boston; Adult Oncology Pathology Laboratory, Dana-Faber Cancer Institute; and Image Core Brigham and Women’s Hospital, Boston.


    FOOTNOTES
 

Address for reprint requests and other correspondence: S. Mizuno, Orthopedic Research, Brigham and Women’s Hospital, 75 Francis St., Boston, MA 02115 (E-mail: smizuno{at}rics.bwh.harvard.edu)

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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