1Life Sciences Division, National Aeronautics and Space Administration Ames Research Center, Moffett Field; 2Department of Mechanical Engineering, Stanford University, Stanford; and 3Department of Stomatology, University of California, San Francisco, San Francisco, California
Submitted 22 November 2003 ; accepted in final form 16 February 2005
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ABSTRACT |
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mechanotransduction; differentiation; bone
Because the weight of a cell depends on the amount of gravity acting on its mass, changes in gravity may serve as mechanical stimuli to adherent cells. By applying increased gravity to cells (hypergravity), the direction and magnitude of gravity can be varied to provide insight into its effects on cellular physiology. Coordinated shape and cytoskeletal changes may result from shifts in the position of organelles of different densities within the cell. In a gravity field, the denser nucleus (26, 48, 49) may shift downward, pulling the interconnected cytoskeleton and plasma membrane with it, resulting in a reduced cell height that would be expected to continue to decrease as the gravity field increased. Microtubules are thought to resist compressive loads (21, 22) and may themselves be compressed as a result of the hydrostatic pressure generated by extracellular fluid. Studies of osteoblasts and endothelial cells have shown that reorganization of the cytoskeleton correlates with increased release of prostaglandin E2 (PGE2), which is important for paracrine and autocrine signaling (42, 50). Whether gravity loading results in microtubule rearrangements related to PGE2 release is unclear.
Bone-forming osteoblasts play a mechanosensing role in vivo (9); thus we anticipated that they would be responsive to changes in the gravity vector. In vivo PGE2 stimulates bone formation and resorption and may mediate the response to mechanical loading (38). Osteoblasts respond to substrate deformation, fluid-induced shear stress, and hydrostatic pressure with changes in cell shape, cytoskeletal organization, and PGE2 production (13, 7, 10, 12, 19, 27, 32, 34, 37), but less is known about the influence of gravity. When gravity is decreased, such as in the microgravity environment of spaceflight, MC3T3-E1 osteoblasts adopt a more rounded morphology; yet, when corrected for cell number, PGE2 release is unchanged (20). Rat osteosarcoma (ROS 17/2.8) osteoblastic cells subjected to alternating gravity loading between microgravity and 2 g via parabolic aircraft flights exhibit increased cell shape irregularity, decreased cell area, and increased PGE2 (17). Neither of these two previous studies addressed possible changes in three-dimensional shape. Hypergravity increases PGE2 production from MC3T3-E1 osteoblast-like cells, but the relationship to cell shape or cytoskeletal changes has not been studied (14, 28, 31). In spaceflight, microtubule polymerization is impaired in intact leukocytes (36) and microtubules do not self-organize in in vitro assays (35), suggesting that altered microtubule polymerization or organization may contribute to observed changes in cell shape. Primary osteoblast cultures at progressive stages of differentiation undergo well-defined changes in cell shape (16, 33), and the influence of gravity on cytoskeleton and PGE2 production may differ as a consequence.
Primary osteoblasts offer the advantage that their function and regulation more closely mimic osteoblasts in vivo, which is not always the case with cell lines. Treatment of confluent primary osteoblasts with ascorbic acid (AA) and -glycerophosphate (
-GP) leads to a progression of events, including proliferation and multilayering, synthesis of an extracellular matrix, and mineralization of that matrix with associated changes in cell shape (5, 33). Spaceflight impairs differentiation of primary embryonic chick osteoblasts (25), suggesting that differentiation is sensitive to microgravity. This result and the cell shape changes associated with differentiation suggest that gravity loading may act differently depending on the stage of cell differentiation.
In this study, we tested the hypothesis that hypergravity loading of primary osteoblasts reduces the microtubule network and nuclear height in a dose-dependent manner, and this change in cell shape accompanies increased PGE2 release. Furthermore, we hypothesized that the stage of differentiation of the osteoblast culture influences these responses. To test these hypotheses, we developed and characterized a cell culture centrifuge that reproduces a standard tissue culture environment and therefore is suitable for both short- and long-term experiments. We showed that immature, confluent primary osteoblasts responded to hypergravity with increased PGE2 release, decreased microtubule network height but no measurable change in nuclear height, and no major morphological changes. Observed changes depended on the dose and duration of the hypergravity stimulus and were associated with progressive differentiation.
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MATERIALS AND METHODS |
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Cell number. Primary osteoblasts were counted in the differentiating cultures using the methods described by Komarova et al. (24). To release the osteoblasts from their matrix at the termination of the experiment, cultures at each time point were treated sequentially with phosphate-buffered saline (PBS) containing 10 mM EGTA and 20 mM HEPES, pH 7.4, for 20 min and then for 60 min at 37°C in 572 U/ml collagenase in 115 mM NaCl, 5.3 mM KCl, 3 mM K2HPO4, 1 mM CaCl2, 30 mM mannitol, 10 mM glucose, 2 g/l BSA, and 24 mM HEPES, pH 7.4. An equal volume of 0.25% trypsin in EDTA (1 mM) in Hanks' balanced salt solution without Ca2+ and Mg2+ (GIBCO-BRL) was added to the collagenase, and cells were incubated for another 30 min. Dispersed cells were counted using a hemocytometer.
Measurement of individual cell density.
Experiments to measure the density of an osteoblast cell (to calculate gravity-induced shear stress in cells placed parallel to the gravity vector) were performed with ROS 17/2.8 cells to represent a homogeneous pool of differentiated osteoblasts. ROS cells were cultured in Ham's F-12 medium (GIBCO-BRL) containing 10% fetal calf serum, L-glutamine (GIBCO-BRL), HEPES, and antibiotics in tissue culture dishes and were passaged once they were 80% confluent. Cells were trypsinized (0.25% trypsin; GIBCO-BRL), pelleted by centrifugation, and resuspended in culture medium to determine cell density. The cell density assay was based on the assumption that particles falling through a column of liquid reach a terminal velocity that can be calculated using Stokes flow theory. Terminal velocity is calculated as vt = (1 f/
p)gdp2/(18
f/
p), where
f is the fluid density,
p is the particle density, dp is the particle diameter, g is gravity, and
is the kinematic viscosity. For each step described below, a QuickScan automated vertical scan dispersion analyzer was used (Beckman Coulter, Miami, FL). The mixed sample of liquid and particles was placed in the sample tube, and the QuickScan detecting head moved vertically along the tube and measured transmitted or backscattered light from the tube. Periodic measurements at room temperature captured the settling behavior, and settling rates were calculated from the slope of the transmission curves. To ensure the experimental results correlated with the Stokes settling calculations, 15-µm polystyrene microspheres of known density (1.05 g/ml; Duke Scientific, Palo Alto, CA) were tested with water because all parameters (particle density, particle diameter, water density, and water viscosity) were known. The density of the ROS medium was determined by weighing a known medium volume and calculating the density (mass/vol). To determine the ROS medium kinematic viscosity at room temperature, the settling rate of microspheres suspended in medium was measured using the QuickScan analyzer. The diameter of suspended ROS cells was measured using a microscope with an ocular mounted reticle. Finally, the density of a ROS cell was determined by measuring the settling rate of cells suspended in medium using the QuickScan analyzer. Each experiment was repeated a minimum of three times.
Once cell density was known, the cell volume was calculated from the measured cell diameter and the cell mass was calculated from density and volume. With the use of these data, the force due to a 10-g hypergravity acceleration was calculated as force = mass x acceleration. The shear stress was then calculated by dividing the force by the average area of attached cells measured near the growth substrate using confocal microscopy.
Cell hypergravity stimulation. Hypergravity was applied using the 1-foot diameter centrifuge (1-FDC) at the NASA Ames Center for Gravitational Biology Research (see http://lifesci.arc.nasa.gov/CGBR/1_ft.html). To minimize evaporation of cell culture medium, the eight-well chamber slides were placed in a 10-cm culture dish with an open 35-mm dish containing 2 ml of water. Alternatively, cell orientation experiments used sealed eight-well chamber slides oriented either flat on the platform or normal to the platform and secured to flasks using adhesive tape. The cells were then placed at the center of the swinging bucket to reduce inertial shear and on a rubber pad to reduce vibration.
Immature confluent osteoblasts were exposed to 238 rpm (10 g) for 3 h to establish a baseline response to hypergravity. Dose response was determined at rotation rates ranging from 112 rpm (2.5 g) to 518 rpm (50 g). The applied gravity level was calculated as the resultant of Earth's 1 g and the centrifugal acceleration. The duration of a 10-g hypergravity stimulus was then varied from 10 min to 6 h, with acceleration to 10 g in 2 min and return to 1 g as a control for the stimulus of acceleration up to constant speed. For each time point <3 h, medium was replaced so that all cells conditioned the medium for 3 h to ensure comparable PGE2 values, e.g., medium was replaced 2 h, 50 min before the 10-min spin. To assess the influence of orientation, cells were placed on the centrifuge with the gravity vector both perpendicular and parallel to the cell growth substrate. Cells at various times in culture (associated with different stages of differentiation; Refs. 5, 24, and 29) were stimulated with 3-h exposure to 10 or 50 g. Immediately after loading, medium was collected for PGE2 analysis and cells were fixed for immunocytochemistry.
1-Foot-diameter centrifuge: characterization of apparatus. The 1-FDC consists of a tabletop centrifuge (model 6S-6RHT; Beckman Coulter), modified to provide lower rotation rates (451,000 rpm yielding 1.4180 g) and environmental monitoring for cultured cells and small organisms. The centrifuge was integrated with a tissue culture incubator (model 3851; Forma-Scientific) to control temperature, humidity, and CO2. Swinging platforms sized for a standard multiwell plate (8.9 cm in the radial direction and 13.3 cm in the circumferential direction) maintained the resultant gravity vector perpendicular to the cell layer. Fans circulated the air between the incubator and centrifuge through insulated ducts, and water traps collected condensation. An identical incubator adjacent to the 1-FDC was used for stationary 1-g controls. Environmental data from the integrated centrifuge-incubator system and the 1-g control incubator were displayed on analog data displays and recorded using a data acquisition system.
The 1-FDC was tested to ensure that the environmental conditions within the centrifuge (other than hypergravity) were similar to those within the adjacent control incubator. Temperature and CO2 were measured inside the centrifuge volume adjacent to the centrifuge lid and inside the control incubator at the back of the unit. The temperature for the 1-FDC supply incubator was set 1°C higher than the control incubator to achieve 37°C within the centrifuge chamber.
To characterize the mechanical loading environment within the centrifuge, vibrations were measured on the platform during rotation. A single-axis, high-sensitivity Bruel & Kjaer type 8318 accelerometer (Naerum, Denmark) was placed on one platform, and an equivalent weight was placed on the opposite platform. Data were transmitted from the rotating platform to the stationary data acquisition system via slip rings temporarily mounted on the centrifuge. Acceleration was measured in the direction of the resultant gravity vector when the centrifuge was spinning at 238 rpm (10 g). Data were collected from the control incubator for comparison. Data were collected at a sampling rate of 256 samples/s for 80 s, processed, and expressed as acceleration in units of gravity, root mean squared (rms).
PGE2 production. The amount of PGE2 released by the cells into the medium was measured using a commercial enzyme immunoassay kit (Amersham Pharmacia, Little Chalfont, UK) according to the manufacturer's protocol. Samples were read at 630-nm wavelength using a SpectraMax 250 microplate reader (Molecular Devices, Sunnyvale, CA). The data were analyzed using SoftMax Pro software version 1.1 (Molecular Devices) on an IBM-compatible personal computer. Concentrations measured were corrected for the amount of medium in the culture well and were expressed as picograms per milliliter. To control for additional proliferation as cultures matured, PGE2 values for d6, d9, and d19 cultures were normalized to cell number. For experiments measuring the effects of hypergravity duration <3 h, cells were grown and maintained in medium for 3 h and centrifuged at the end of the 3-h period, because short exposures were not sufficient to generate measurable changes in PGE2. For example, for the 10-min stimulus, cells were exposed to the medium for 2 h, 50 min before the 10-min spin. The rate of PGE2 release during the hypergravity stimulation was calculated as the difference between the total amount produced during 3 h and the amount produced during the 1-g precentrifugation period. We assumed that the rate of PGE2 release during the 1-g precentrifugation period was the same as that produced by the 1-g controls. The detailed calculation was made as follows. First, the amount of PGE2 produced at 1 g before hypergravity stimulation was calculated as PGE2 (amount during prehypergravity period in pg/ml) = PGE2 rate1-g controls (pg·ml1·min1) x [180 min thypergravity (min)], where PGE2 rate1-g controls is the rate of PGE2 produced in the corresponding 1-g controls during a 3-h period and thypergravity is the duration of hypergravity exposure. Next, the rate of PGE2 production during hypergravity stimulation was calculated as PGE2 ratehypergravity (pg·ml1·min1) = [PGE2 (amount in 3 h in pg/ml) PGE2 (amount during prehypergravity period in pg/ml)]/thypergravity (min).
Alkaline phosphatase and osteocalcin content. To measure alkaline phosphatase activity, cells were extracted in 1% Triton X-100 in HEPES buffer, pH 7.4, and then sonicated and centrifuged at 14,000 g for 4 min. Supernatants were stored at 80°C until analysis. A reaction buffer, pH 7.4, composed of 100 mM glycine buffer, 1 mM MgCl, and 1 mM ZnCl in distilled water was added with 60 mM p-nitrophenyl phosphate (Sigma) to the cell supernatants, and then alkaline phosphatase was measured spectrophotometrically (SpectraMax 250 microplate reader) at 405-nm wavelength. Osteocalcin levels in medium were measured after 24-h serum starvation using a commercial enzyme immunoassay kit (Biomedical Technologies, Stoughton, MA) according to the manufacturer's protocol. Samples were read at 450-nm wavelength using a SpectraMax 250 microplate reader.
Staining and imaging of differentiating cells. Mineralized nodules in d9 and d19 cultures were demonstrated using alizarin red staining of calcium salts. Cells were fixed in ethanol for 15 min, stained for 60 min in 1% alizarin red solution (Hartman-Leddon, Philadelphia, PA) in distilled water, pH 6.4, and then washed in distilled water. Images of alizarin red-stained cultures were acquired using phase-contrast microscopy. Images of separate osteoblast cultures at different stages of differentiation were acquired using an inverted scanning confocal microscope (Zeiss LSM 510) with differential interference contrast (DIC) microscopy to illustrate the formation of nodules with commensurate height and morphological changes.
Immunocytochemistry.
Within 5 min of stopping the centrifuge, cells for microtubule and nuclear staining were washed with Dulbecco's PBS (GIBCO-BRL/Invitrogen, Grand Island, NY) at 37°C, fixed in ice-cold methanol for 5 min, washed with PBS, and incubated for 30 min in a blocking solution containing 5% bovine serum albumin (Sigma), 0.1% Tween 20 (Fisher Scientific, Fair Lawn, NJ), 2% goat serum (Jackson ImmunoResearch, West Grove, PA), and PBS. Cells were incubated in primary antibody (1:200 dilution mouse monoclonal anti-chicken -tubulin, clone DM 1A; Sigma), diluted in the blocking solution for 1 h at room temperature, then washed with a solution containing 0.1% Tween 20 (Fisher Scientific), 5% bovine serum albumin (Sigma), and PBS, and then incubated for 20 min at room temperature with Texas Red goat anti-mouse secondary antibody (Jackson ImmunoResearch) diluted 1:200 in blocking solution with 0.5 mM Sytox Green nuclear stain (Molecular Probes, Eugene, OR). Finally, slides were washed with blocking solution and rinsed with distilled water. Coverslips were placed on slides with Aqua Polymount (Polysciences, Warrington, PA) and sealed with nail polish.
Microtubule and nuclear morphology. Serial optical images of the microtubule cytoskeleton and cell nuclei were obtained using a confocal laser scanning microscope equipped with DIC optics, a x63 magnification, 1.25 numerical aperture oil-immersion lens objective, and a 30-mW argon/krypton laser (LSM 510; Carl Zeiss, Thornwood, NY). Pinhole sizes and photomultipliers were set to produce the clearest possible image without saturating the signal. After the conditions of image acquisition were optimized for the 1-g control cells, images of hypergravity-stimulated and 1-g control cells were collected using identical settings. Several techniques were used to identify morphological changes, including characterization of optical slices through the midplane of the cell, construction of image galleries containing each serial slice in an image stack, and projection of all slices into one view. To analyze microtubule network height, image z-stacks were acquired and post-image analysis software was used to draw a horizontal line in the x-y plane across the center of the image field. The profile function was used to provide signal intensity for each pixel along the line, yielding a graphed plot of intensity vs. distance. The image stacks were viewed from the top slice to the bottom slice, the graphical intensity plot changes were observed, and all images with an intensity value >50 anywhere along the line were included in the calculation of height. The intensity threshold of 50 was determined in tests using fluorescent beads of known size to best represent actual signal and not background noise. The resulting number of slices was multiplied by the slice thickness (0.5 µm) to yield the microtubule network and nuclear height. Microtubule network height of the multicell layer in differentiating cultures was measured from the top to the bottom image in the confocal image stack to provide an indication of the overall height of the nodular and internodular regions. The number of nuclei in a z-axis orthogonal view was counted to provide an indication of the number of cell layers.
Statistics. Data are representative of three to six separate experiments performed with four individual culture wells per condition. PGE2 analyses were performed in duplicate. Cell number and alkaline phosphatase were performed on four samples and osteocalcin on six samples. Eight microscopic fields per condition were evaluated for microtubule morphology, network height, and alizarin red staining. Values are expressed as means ± SE. Statistical evaluation was performed using StatView version 5.0.1 software (SAS Institute, Cary, NC). Differences were compared using ANOVA with a significance level of 0.05. P < 0.05 was accepted as significant, with P values corrected by applying the Bonferroni adjustment to Fisher's protected least-significant difference post hoc analysis.
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RESULTS |
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Hypergravity increased PGE2 release and decreased microtubule network height in a dose-dependent manner. We initially assessed the influence of a 10-g hypergravity stimulus on PGE2 release; overall cell, microtubule cytoskeleton, and nuclear morphologies; microtubule network height; and nuclear height. We then tested stimuli <10 g and >10 g. Confluent osteoblasts (d3) subjected to 10 g for 3 h released 490 pg/ml PGE2 compared with 190 pg/ml released by 1-g control cells, representing a 2.5-fold increase (Fig. 2). A 2.5-g stimulus did not result in detectable PGE2 release compared with 1-g controls (data not shown). A 5-g stimulus resulted in a 2.5-fold increase in PGE2 release compared with controls; this change was not significantly different from the effects of a 10-g stimulus. Hypergravity stimuli >10 g applied for 3 h further increased PGE2 release. A 15-g stimulus resulted in a 3.4-fold increase, and a 50-g stimulus resulted in a 5.3-fold increase, compared with controls. The PGE2 release due to 50 g was significantly different from that at 5, 10, and 15 g. The difference between 10 and 15 g was significant, but the difference between 5 and 15 g was not.
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Time course of response. As a control for possible transient effects of acceleration and deceleration, cells were accelerated to 10 g and then immediately decelerated to 1 g and compared with the 1-g controls. No differences were noted (Fig. 5). Increasing the duration of the 10-g stimulus to 10 min resulted in a 1.5-fold increase in the amount of PGE2 released into the medium. Further increases were observed as the 10-g stimulus was lengthened to 1 h (1.5-fold), 3 h (2.8-fold), and 6 h (4.8-fold); all increases were significant. The rate of PGE2 release was calculated to determine whether the rate depended on the duration of the stimulus (see MATERIALS AND METHODS). Exposure to hypergravity for 10 min caused a 10-fold increase in the calculated rate of PGE2 release compared with the 1-g control (Table 1). The 1-, 3-, and 6-h exposures resulted in 2.6-, 2.9-, and 4.3-fold increases in the rate of PGE2 release, respectively, compared with the 1-g control. Changing the duration of 10-g hypergravity exposure from 10 min to 6 h did not affect the overall appearance of the cells or nuclei. Only the 3-h exposure resulted in a significant reduction in microtubule network height, while shorter durations slightly but not significantly reduced network height (data not shown). Nuclear height was not affected by hypergravity exposure at any duration tested (data not shown).
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To calculate the amount of shear stress applied to the osteoblasts when they were oriented on their side, the density of ROS 17/2.8 osteoblast-like cells was measured as described in MATERIALS AND METHODS. The ROS medium density was determined to be 0.99 g/ml, and medium kinematic viscosity was 1.4 x 106 m2/s. The average cell diameter was 19 µm, the cell density was 1.04 g/cm3, and the area near the growth substrate was 730 µm3. From these measured values, cell volume and mass were calculated as 3.6 x 1015 m3 and 3.73 x 109 g, respectively. On the basis of these values, 10-g stimulation resulted in shear stress of 0.5 Pa (5 dyn/cm2).
Characterization of osteoblast differentiation.
Osteoblast differentiation in a 1-g environment was characterized by evaluating the overall osteoblast culture morphology as shown using DIC imaging (Fig. 6, AD) and alizarin red staining of bone nodules (Fig. 6, E and F), alkaline phosphatase activity, osteocalcin content in the media, microtubule network height, and PGE2 release. Cells were grown for 34 days to confluence (Fig. 6A), and the medium was supplemented with AA and -GP to induce differentiation. Cells formed a uniform layer, and nuclear and microtubule network morphologies were as shown in Fig. 3. By d6, discrete regions of cuboidal cells (prenodules) appeared within the confluent layer (Fig. 6B, arrow) and multilayered nuclei and microtubule networks were observed (data not shown). By d9, cells had multilayered further and produced abundant extracellular matrix (Fig. 6C; arrow) and small alizarin red-stained mineralized nodules (Fig. 6E; arrow). By d19, mature mineralized nodules formed as shown using DIC (Fig. 6D, arrows) and alizarin red staining (Fig. 6F, arrows) and were surrounded by unmineralized internodular regions. Cell number, alkaline phosphatase activity, and osteocalcin increased as the cells differentiated (Table 2).
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In the latter stages of differentiation (d9 and d19), microtubule network height in nodular and internodular regions did not change consistently as a result of either 10 g or 50 g applied for 3 h (data not shown). In d6 cultures, 10 g resulted in a 10% decrease in internodular and nodular height, but this difference was not statistically significant (data not shown). Nuclear height was insensitive to hypergravity at all days in culture, and there was no evidence of apoptosis or other indications of poor cell health, regardless of the gravity level examined (data not shown).
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DISCUSSION |
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Validation of hypergravity as a model for studying the influence of the gravity vector. To develop a valid tool to study hypergravity, the physical environment of the centrifuge must replicate the environment of the control cultures. Furthermore, the impact of various physical factors that may influence cellular responses to centrifugation other than hypergravity per se should be considered. To characterize the physical environment, we ensured that the temperature, CO2, and humidity conditions in the centrifuge (1-FDC) and control incubator were similar. Next, we measured vibration in the centrifuge because low-frequency vibrations are known to regulate osteoblast activities (40). The 1-FDC displayed a low level of broad frequency vibration (106 to 105 g, rms; peak 104 g, rms at 4 Hz) in the direction of the resultant gravity vector; these values are well below the magnitude of a 10-g hypergravity stimulus (1% x 105 to 1% x 103). Furthermore, there were no changes in PGE2 release or microtubule network height in cultures maintained in the 1-FDC at 1 g (without rotation) compared with cultures maintained in the control incubator. On the basis of these results, we conclude that the 1-FDC and control incubators provide comparable physical environments for cell growth.
To evaluate the various physical factors contributing to the centrifuge environment, we calculated the gravity gradient, coriolis force, and inertial shear contributions to cell cultures for the 1-FDC in producing acceleration of 10 g (238 rpm). Objects on a centrifuge are exposed to a gravity gradient. Given the measured cell height of 4 µm, the difference in acceleration between the two opposite cell surfaces is 2.6% x 104 of 10 g. A centrifuge also causes motile cells to experience a coriolis force. Coriolis acceleration is defined as ac = 2 v
, where v is the radial velocity of a motile cell and
is the angular velocity of the centrifuge. Assuming an average osteoblast motility of 10 µm/h (13) on the 1-FDC at 10 g, the coriolis acceleration is only 1.41 x 108 g, or 1.41% x 107 of 10 g. Finally, because the culture surface is flat, the gravity vector is not uniform. This results in a net acceleration, termed inertial shear, toward the edges of the platform (46). From the center of the platform to the edge, the inertial shear varies from 0 to 3% of 10 g (given a platform width of 9 cm). By using the central four wells of the eight-well chamber slide (2 cm total width) and placing the slide in the middle of the platform, the inertial shear for these experiments was limited to 0.67% of 10 g. These gravity variations due to centrifugation in this apparatus are summarized in Table 3. While the inertial shear is the largest artifact resulting from using the 1-FDC to simulate hypergravity, it still represents only a small contribution to the gravity levels used in these studies (2.550 g).
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Hypergravity may cause increased strain due to cell deformation, in addition to increased hydrostatic pressure. To estimate the strain due to gravity acting on an osteoblast, Hatton et al. (18) modeled an osteoblast as a homogeneous elastic disk with the material properties of a chondrocyte. They concluded that hypergravity levels of 430 g resulted in 40300 microstrains, respectively, similar to those observed in human tibia during light exercise (8). To assess the combined effects of hydrostatic pressure and strain on an osteoblast with a heavier nucleus and a discrete cytoskeleton, we developed a cell model that included a plasma membrane, a nucleus 40% heavier than the surrounding cytoplasm, and actin and microtubule cytoskeletal networks with material properties derived from the literature (43, 44). Results indicated that hypergravity levels of 10 g resulted in a 5% reduction in cell height, and strains varied by several orders of magnitude, depending on location within the cell (e.g., outer plasma membrane vs. microtubules). On the basis of these analyses, we concluded that exposing osteoblasts grown on a substrate oriented perpendicular to the gravity vector to centrifugation from 2.5 to 50 g resulted in mechanical strains within the physiological range.
Cells oriented with the growth substrate parallel to the gravity vector are subject to shear stress due to gravity acting on the cell mass. With the use of measured values of ROS 17/2.8 osteoblast-like cell mass and area near the growth substrate, a 10-g stimulus resulted in a shear stress of 0.5 Pa (5 dyn/cm2). This value of shear stress is similar to shear induced by fluid flow, which has been shown to act as a mechanical stimulus to osteoblasts in vitro (3, 23, 39, 40), and is predicted to be applied to osteocytes in vivo (51). Therefore, cells in this orientation experienced a shear stress in the physiological range.
Response of immature osteoblasts to hypergravity. With the hypergravity model established, we investigated the characteristics of hypergravity-induced changes in microtubules and release of the paracrine signaling factor PGE2, which is a critical component of the anabolic response of bone to mechanical loads (11). Exposure of confluent, immature osteoblasts (d3) to hypergravity ranging from 5 to 50 g for 3 h resulted in a 2.5- to 5.3-fold increase in PGE2 release compared with 1-g controls. These results are consistent with the finding that a short, intensive pulse (5 min, 187 g) of centrifugation triggers PGE2 release from MC3T3-E1 osteoblasts (14). In our study, changes in PGE2 release were not observed after a 2.5-g stimulus, suggesting either insensitivity to small gravity changes or that PGE2 release is not a sensitive cellular response. The dose-response curve from 5 to 50 g indicates that 5 g was the minimum effective dose for PGE2 release, and this release increased to 50 g. Because the differences in PGE2 release between 5-, 10-, and 15-g stimuli were small, it may be that the response to gravity levels up to 15 g are the first phase of a response and that the response to 50 g represents a second phase. This hypothesis is consistent with osteoblast responses to mechanical deformation of the growth substrate; studies have shown biphasic responses to increasing levels of strain (30).
Centrifugation at increasing gravity levels also caused a gradual decline in microtubule network height to a 26% decrease in 50-g cultures relative to 1-g controls. These results show that there is a correlation between PGE2 release and microtubule network height as the magnitude of the hypergravity stimulus is raised. When ROS 17/2.8 osteoblastic cells were subjected to alternating hypergravity and microgravity in parabolic flight, a positive correlation between cell area and intracellular PGE2 levels was demonstrated when all gravity levels were considered (17), suggesting that rapid changes in the direction of the gravity vector may affect cell shape. However, we found that acceleration followed by immediate deceleration failed to exert the same effects on microtubule network height and PGE2 release as exposure to a continuous hypergravity stimulus.
Although the microtubule network height decreased with increasing gravity levels, the nuclear height did not appear to change. This suggests that the nucleus did not displace downward toward the substrate in the increased gravity field, although repositioning of the nucleus may have been below the limits of detection.
PGE2 release and microtubule network height demonstrated similar dose dependency, but the time courses of these responses appeared to differ. PGE2 release increased as the duration of the hypergravity stimulus was lengthened from 10 min (1.5-fold) to 6 h (4.8-fold); yet only a 3-h duration resulted in a significant reduction in microtubule network height. By 3 h, the microtubule network may have adapted to form a more stable configuration.
Hypergravity stimulated PGE2 release whether the cultures were oriented parallel (side) or perpendicular (flat) relative to the gravity vector. Because hydrostatic pressure acts in all directions, the cultures in the side orientation still experienced pressure. The strains applied to cultures in the side orientation due to the shear stress acting on the cell's mass were different from the strains applied to cultures in the flat orientation, owing to the compressive load in the flat orientation. The PGE2 release could be attributed to the influence of hydrostatic pressure in both cases.
Response of differentiating osteoblasts to hypergravity.
The hypergravity-induced release of PGE2 and reduction in microtubule network height differed depending on the duration of cell culture. We confirmed that differentiation in vitro recapitulates the major features of osteoblast differentiation in vivo as shown in other studies (5, 16, 24, 29, 33). Treatment with AA and -GP in the continuous presence of 10% serum caused progressive changes in characteristic features of the mature osteoblast, including acquisition of a cuboidal morphology, increased alkaline phosphatase activity, production of a collagenous extracellular matrix that mineralized, and osteocalcin production.
The PGE2 released in control cultures at 1 g was relatively constant from confluence (d3) through early nodular mineralization (d9) but fell to undetectable levels at latter stages of culture when nodules were mineralized (d19). These results are consistent with the decline in PGE2 during differentiation of adult rat calvarial osteoblasts reported by Fujieda et al. (15).
We found that hypergravity increased PGE2 release and reduced microtubule height in confluent (d3) and early nodule-forming (d6) cultures (3 h at 10 or 50 g), demonstrating sensitivity to hypergravity through the initiation of nodule formation. In contrast, hypergravity stimuli failed to induce PGE2 release or microtubule network height changes in more mature cultures (d9d19), demonstrating a possible decline in sensitivity during later stages of nodule maturation and mineralization.
Given the conditions of cell growth used in this study, variables other than differentiation per se also may contribute to the reduced gravity sensitivity observed at the later time points in culture (45). Continuous growth in relatively high concentrations of fetal calf serum (10%), together with supplementation with AA and -GP is currently the standard condition used for growth and differentiation of rat primary osteoblasts (5, 16, 24, 29, 33). However, other potentially important factors that may contribute to the changes observed over time in this study include cellular aging; sustained exposure to high concentrations of growth factors, hormones, and other ill-defined serum factors; and/or altered cell-cell interactions resulting from high cell density. To control for additional proliferation in maturing cultures, PGE2 release was corrected to cell number. Another possible explanation for the reduced sensitivity to hypergravity that we observed in mature cultures is that the abundant extracellular matrix and cell multilayering, which are present only in mature cultures, blunted transmission of gravity loads to the osteoblasts. Alternatively, the cyclooxygenase responsible for PGE2 production in response to the hypergravity stimulus may be present in lower levels in our mature cultures.
In any event, our results show that the sensitivity to hypergravity appears highest at less mature stages of osteoblast differentiation, when cells are confluent but are not yet producing a matrix that is mineralized. Miwa et al. (28) suggested that hypergravity stimulates proliferation in early cultures and that PGE2 mediates this response. Consistent with our findings, human fetal osteoblasts lose their sensitivity to mechanical stretch at late stages of differentiation (45).
In conclusion, we have shown that a continuous hypergravity stimulus induced PGE2 release and reduced the height of the microtubule network in primary fetal rat osteoblasts. These responses depended on the magnitude and duration of the stimulus. Immature osteoblasts appeared most sensitive to changes in gravity loading. Our results demonstrate the utility of centrifugation as an experimental tool to study the influence of changes in the gravity vector on cell structure and function.
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ACKNOWLEDGMENTS |
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