Influence of increased mechanical loading by hypergravity on the microtubule cytoskeleton and prostaglandin E2 release in primary osteoblasts

Nancy D. Searby,1,2 Charles R. Steele,2 and Ruth K. Globus1,3

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


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cells respond to a wide range of mechanical stimuli such as fluid shear and strain, although the contribution of gravity to cell structure and function is not understood. We hypothesized that bone-forming osteoblasts are sensitive to increased mechanical loading by hypergravity. A centrifuge suitable for cell culture was developed and validated, and then primary cultures of fetal rat calvarial osteoblasts at various stages of differentiation were mechanically loaded using hypergravity. We measured microtubule network morphology as well as release of the paracrine factor prostaglandin E2 (PGE2). In immature osteoblasts, a stimulus of 10x gravity (10 g) for 3 h increased PGE2 2.5-fold and decreased microtubule network height 1.12-fold without affecting cell viability. Hypergravity (3 h) caused dose-dependent (5–50 g) increases in PGE2 (5.3-fold at 50 g) and decreases (1.26-fold at 50 g) in microtubule network height. PGE2 release depended on duration but not orientation of the hypergravity load. As osteoblasts differentiated, sensitivity to hypergravity declined. We conclude that primary osteoblasts demonstrate dose- and duration-dependent sensitivity to gravitational loading, which appears to be blunted in mature osteoblasts.

mechanotransduction; differentiation; bone


LIFE AS WE KNOW IT HAS EVOLVED under the influence of Earth's gravity. Changes in other physical conditions in the environment (e.g., temperature) have profound effects on cells, and gravity may act at a fundamental level as well. Gravity is ubiquitous and influences the mechanical environment within tissues by affecting cell weight, extracellular hydrostatic pressure, and fluid convection. Despite the central importance of gravity, little is known about how it influences cellular physiology.

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 {beta}-glycerophosphate ({beta}-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.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Cell culture. Primary osteoblast cells were obtained by performing sequential collagenase digestion with embryonic day 21 fetal rat calvaria as previously described (5), with minor revisions. All animal procedures were reviewed and approved by the Institutional Animal Care and Use Committee at the National Aeronautics and Space Administration (NASA) Ames Research Center. Briefly, cells were subcultured the day after isolation and plated at a density of 36,000 cells/cm2 onto 0.2% gelatin-cross-linked (gelatin type A from porcine skin; Sigma, St. Louis, MO) eight-well chamber slides (0.81 cm2/well, Permanox; Nalge Nunc International, Naperville, IL) and grown in {alpha}-minimal essential medium containing 10% fetal calf serum (medium and serum; GIBCO-BRL/Life Technologies, Grand Island, NY) and antibiotics (penicillin-streptomycin and Fungizone; GIBCO-BRL) in 5% CO2 at 37°C. Cells were induced to differentiate according to standard methods (5) as described by Moursi et al. (29) and further characterized by Komarova et al. (24). Briefly, when cells reached confluence (day 3 or 4, referred to hereinafter as d3), they were induced to differentiate by supplementing the media with freshly prepared AA (50 µg/ml; GIBCO-BRL) to induce matrix formation and with {beta}-GP (3 mM; Sigma) to induce mineralization. Media were changed every 2–3 days. The medium pH was controlled in one of two ways. In the gravity orientation experiments, media were supplemented with HEPES buffer (10 mM, pH 7.4; GIBCO-BRL) before centrifugation and the wells were filled to the top with 700 µl of medium and sealed with liquid-tight gaskets. In other experiments, the wells were filled with 650 µl of medium (10 mm medium height) and the lids were vented for gas exchange. No differences were noted to be a result of the method of pH control in these short-term (<6 h) experiments (data not shown). To determine whether this volume or height of medium (larger than normal medium-to-cell ratio) affected the results, experiments were performed with 325 µl of medium (5 mm medium height), with no differences observed (data not shown).

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 – {rho}f/{rho}p)gdp2/(18{nu}{rho}f/{rho}p), where {rho}f is the fluid density, {rho}p is the particle density, dp is the particle diameter, g is gravity, and {nu} 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 (45–1,000 rpm yielding 1.4–180 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·ml–1·min–1) 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·ml–1·min–1) = [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 {alpha}-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.


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1-FDC provides a thermal and vibration environment comparable to that of a control incubator. Thermal control tests of 1-FDC (Fig. 1) showed that centrifugation for 3 h before the experiment stabilized the centrifuge temperature at 37 ± 0.2°C. By placing samples on prewarmed platforms, closing the centrifuge lid within 30 s, and then resetting the rotation rate, temperature within the centrifuge decreased transiently by 0.5°C and then returned to 37°C within 12 min. This transient was similar to that which occurred when opening and closing the door of the control incubator. The CO2 levels remained steady at 5 ± 0.1%, with a 1% transient decrease corresponding to opening and then closing the centrifuge; CO2 levels recovered to 5% within 12 min. Evaporation from the culture wells was assessed to validate humidity control. After a 3-h spin at 238 rpm (10 g), the amount of medium was reduced by <5%.



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Fig. 1. 1-foot diameter centrifuge (1-FDC). Osteoblasts were exposed to hypergravity using the 1-FDC at the National Aeronautics and Space Administration Ames Research Center for Gravitational Biology Research (see http://lifesci.arc.nasa.gov/CGBR/1_ft.html). A tabletop centrifuge with four swinging buckets was modified to provide lower rotation rates (45–1,000 rpm yielding 1.4–180 g) and environmental monitoring. The centrifuge was integrated with an incubator to control temperature, humidity, and CO2. Fans circulated air through insulated ducts, and water traps collected condensation. An identical incubator was used for stationary controls. Environmental data from the control incubator and the integrated centrifuge-incubator system were collected using a data acquisition system.

 
Tests to characterize the centrifuge vibration environment in the resultant gravity vector direction were performed. When the centrifuge was spinning at 238 rpm (10 g), a peak value of 10–4 g, rms was measured at a frequency of 4 Hz, with additional peaks of 2–6 x 10–5 g rms from ~4 to 85 Hz. The majority of the vibrations were measured in the range of 10–6 to 10–5 g, rms from 0 to 100 Hz, which was 1% x 10–5 to 1% x 10–4 that of a 10-g hypergravity stimulus. In comparison, the control incubator generated vibrations of 10–6 to 10–5 g, rms from 0 to 100 Hz, which was similar to the centrifuge, with a resonant peak of 10–2 g, rms at 17 Hz. Thus the centrifuge and control incubator provided comparable vibration environments.

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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Fig. 2. Influence of hypergravity on prostaglandin E2 (PGE2) release. Confluent osteoblasts (day 3, d3) were subjected to hypergravity from 5 to 50 g for 3 h. PGE2 release increased from 2.5-fold (5 g) to 5.3-fold (50 g). *P < 0.0001, significant difference compared with 1-g controls. +P < 0.05, significant difference between 10 and 15 g. {wedge}P < 0.0001, significant difference between 50 g and all other hypergravity levels.

 
The morphology of osteoblasts subjected to 10-g stimulation for 3 h was similar to that of controls (Fig. 3, AD). A healthy, densely confluent cell layer was observed in both 1- and 10-g cultures. Nuclear morphology in an image slice through the midplane of the cell (Fig. 3C) indicated no evidence of mechanical damage in 10-g cultures, and the overall cell morphology was indistinguishable from controls (Fig. 3A). Microtubules in the same confocal slice exhibited interconnected, radially emanating network structures in 1 g (Fig. 3B) and 10 g (Fig. 3D) conditions with similar architecture. Hypergravity caused a small increase in the intensity of the labeled microtubules, but the difference in fluorescence intensity was not significant (data not shown). Exposure to 10-g hypergravity decreased the microtubule network height by 12% (Fig. 4), but nuclear height was not different (data not shown). No differences were observed between cultures in the control incubator and cultures in the 1-FDC maintained without rotation at 1 g, showing that rotation was responsible for the changes induced by centrifugation.



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Fig. 3. Influence of hypergravity on cell and cytoskeletal morphologies. Confluent osteoblasts (d3) were subjected to 10-g hypergravity for 3 h. Hypergravity (10 g, 3 h; A and B) did not cause significant changes in overall morphologies of the cell, nucleus, or microtubule network of confluent osteoblasts compared with 1-g controls (C and D). Differential interference contrast (DIC) images revealed a healthy, confluent cell layer in both 1- and 10-g cultures (not shown). Nuclear morphology of a confocal slice mid-cell height above the substrate indicates no evidence of mechanical damage (A and C). Microtubules in the same confocal slice exhibit interconnected radially emanating network structures (B and D). Small increases were noted in the intensity of the labeled microtubules of centrifuged cells, but these differences were not quantifiable.

 


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Fig. 4. Influence of hypergravity on the height of the microtubule network. Confluent osteoblasts (d3) were subjected to hypergravity from 5 to 50 g for 3 h. Microtubule network height decreased as hypergravity increased. *P < 0.05, significant difference compared with 1-g controls. +P < 0.05, significant difference compared with 50 g.

 
Increasing levels of hypergravity exerted no detectable effects on cellular, microtubule network, or nuclear morphologies. In all cases, the cells appeared healthy, nuclei were intact, and radially emanating microtubule networks were observed. Microtubule network height did not change at 2.5-g stimulation (data not shown), but decreased from 5 g (7% decrease) to 50 g (26% decrease) (Fig. 4). The 50-g stimulation resulted in a significantly reduced microtubule network height compared with the 1-g control and the 5-g stimulation. Nuclear height was unaffected at any hypergravity dose tested (data not shown).

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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Fig. 5. Influence of hypergravity duration on PGE2 release. Confluent osteoblasts (d3) were subjected to 10-g hypergravity, and the PGE2 release was measured for durations varying from a control for possible transient effects of acceleration and deceleration (accel/decel) up to 6-h hypergravity exposure. PGE2 release increased significantly as the duration of hypergravity exposure increased. *P < 0.05, significant difference compared with corresponding 1-g controls. **P < 0.001, significant difference compared with 1-g controls.

 

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Table 1. Dependence of rate of hypergravity-induced PGE2 release on hypergravity duration

 
Influence of gravity vector direction. D3 cells were centrifuged for 3 h at 10 g in a flat or side orientation, either perpendicular or parallel to the gravity vector, respectively. Cells perpendicular to the gravity vector released 2.3-fold more PGE2 compared with the 1-g control, whereas cells parallel to the gravity vector released 2.9-fold more PGE2 compared with the 1-g control. While the amount of PGE2 released due to hypergravity was significantly different from that of the 1-g controls, the orientation of the gravity vector did not result in a significant difference in PGE2 release between the two hypergravity conditions. Hypergravity (10 g) applied to cells parallel to the gravity vector did not alter microtubule network and nuclear morphologies or nuclear height. Microtubule network height appeared reduced in cultures oriented parallel to the gravity vector, although this was not consistently observed (data not shown).

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 10–6 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 10–15 m3 and 3.73 x 10–9 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 3–4 days to confluence (Fig. 6A), and the medium was supplemented with AA and {beta}-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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Fig. 6. Osteoblast differentiation. Osteoblast cultures undergo characteristic morphological changes as they differentiate as shown in DIC images (AD) and images of alizarin red staining of mineralization (E and F) in separate cultures. Cells were grown for 3–4 days to confluence (A), and the medium was supplemented with ascorbic acid (AA) and {beta}-glycerophosphate ({beta}-GP) to induce differentiation. By d6, discrete regions of cuboidal cells (prenodules) appeared within the confluent layer (B, arrows). By d9, cells multilayered and began to produce abundant extracellular matrix (C, arrow) and small mineralized regions as shown by alizarin red staining (E, arrow). By d19, mature mineralized nodules formed (DIC image, D, arrows; alizarin red staining, F, arrows), surrounded by unmineralized internodular regions. Scale, 50 µm.

 

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Table 2. Characterization of differentiation of osteoblasts in culture

 
Quantification of the multicell microtubule network height in a mature, multilayered culture provides an indication of the overall height of nodular and internodular regions. Confocal images were analyzed with the same method used for confluent cell layers whereby the height was measured from the top to the bottom image in the confocal image stack. The number of nuclei in a z-axis orthogonal view were counted to provide an estimate of the number of cell layers. The number of cell layers within nodules appeared to increase from one layer on d3, one to two layers on d6, two layers on d9, and three to four layers on d19. The nodule height increased as the culture progressively differentiated (Fig. 7) (P < 0.001), due partially to increased numbers of cell layers but also to increased spacing between nuclei by d19. By d9, the height of the internodular regions was significantly less (7 µm) than that of the nodules (17 µm) (P < 0.001), but by d19, the height of both the nodular and internodular regions had increased to 25 µm. In the internodular regions, cell layer number increased from one layer on d3 to three layers on d19.



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Fig. 7. Influence of osteoblast culture differentiation on microtubule network height. The overall multicell microtubule network height of the multilayered, maturing osteoblast cultures was measured in the nodular and internodular regions at various times in culture as cells progressively differentiated. By d9, the multilayered nodular region was taller than the internodular region. By d19, both nodular and internodular regions were taller than the 1-g controls. *P < 0.001, significant difference compared with 1-g controls.

 
In contrast to nodular and internodular height, PGE2 released per 106 cells in a 1-g environment did not appear to change significantly with differentiation through d9 (Fig. 8). By d19 in culture, PGE2 release was not detectable (data not shown).



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Fig. 8. Influence of cell differentiation on hypergravity-induced PGE2 release. PGE2 release was measured at various times in culture as cells progressively differentiated and was normalized to cell number. Hypergravity-induced PGE2 release was maximal at d6 in culture but decreased by d9 in culture, with no statistical difference due to 10 g vs. 50 g. Fully mature cultures (d19) did not release detectable amounts of PGE2. *P < 0.001, significant difference compared with 1-g controls.

 
Effects of osteoblast differentiation on responses to hypergravity. We asked whether the response to a 10- or 50-g hypergravity stimulus was associated with the stage of differentiation. A 10-g stimulus for 3 h caused a fivefold increase and a 50-g stimulus for 3 h caused a sixfold increase in PGE2 released per 106 cells from d6 cultures compared with the 1-g control (P < 0.001), and the difference in PGE2 release between 10 g and 50 g was not significant. Neither 10 g nor 50 g resulted in increased PGE2 released per 106 cells from d9 cultures (Fig. 8). Hypergravity did not induce d19 cultures to release PGE2 (data not shown).

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).


    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 
As a first part of this study, we characterized the properties of the 1-FDC and found that the centrifuge provides a thermal and vibration environment comparable to that of a control incubator. The results revealed that both PGE2 release and microtubule network height in confluent primary osteoblasts were sensitive to hypergravity in a dose- and time-dependent manner and that hypergravity-induced PGE2 release appeared blunted in mature osteoblasts. Hypergravity applied to confluent (d3) cultures at 10 g for 3 h resulted in increased PGE2 release, decreased microtubule network height, and no major changes in overall morphology or viability. When the magnitude of the hypergravity stimulus was varied from 5 to 50 g, PGE2 release rose without reaching a plateau and microtubule network height gradually declined. PGE2 release in response to hypergravity was time dependent but was not influenced by the orientation of the gravity vector. When primary osteoblasts at different stages of differentiation were exposed to hypergravity, PGE2 release increased through the initial stages of matrix deposition when nodules first formed (d6). In contrast to immature cultures, cultures with mineralized matrix failed to show sensitivity to hypergravity stimulation. Taken together, these results indicate that changes in the magnitude of the gravity vector regulate cytoskeletal structure as indicated by microtubule network height, as well as function as indicated by release of the paracrine signaling factor PGE2. Furthermore, our results indicate that immature osteoblasts are more sensitive than more mature cells to a hypergravity stimulus.

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 (10–6 to 10–5 g, rms; peak 10–4 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 10–5 to 1% x 10–3). 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 10–4 of 10 g. A centrifuge also causes motile cells to experience a coriolis force. Coriolis acceleration is defined as ac = 2 v{omega}, where v is the radial velocity of a motile cell and {omega} 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 10–8 g, or 1.41% x 10–7 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 (2.5–50 g).


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Table 3. Mechanical components of loading by hypergravity

 
The final question we asked was what levels of hypergravity stimulation are likely to be physiologically relevant for cells that reside in bone. Gravity generates hydrostatic pressure due to the column of medium above the cultures when oriented with the gravity vector orthogonal to the cell growth substrate. Hydrostatic pressure is calculated as P = {rho}gh, where {rho} is the density of the medium, g is the acceleration of gravity, and h is the height of the column of medium. For our cell culture system, the hydrostatic pressure at the substrate was calculated to be 1.2 kPa at 10 g and 6 kPa at 50 g. These pressure levels are comparable to the intraosseous and intramedullary pressures applied to osteoblasts growing on trabeculae in vivo, which are 1–5 kPa in dogs (4) and 2 kPa in rats (6). Thus the hydrostatic pressure levels produced by centrifugation from 10 to 50 g are in the physiological range for the osseous environment.

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 4–30 g resulted in 40–300 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 {beta}-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 {beta}-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.


    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by NASA Grants NAGS5-6374 and ARC DDF-96-01.


    ACKNOWLEDGMENTS
 
We thank Soha Motlagh and Indroneal Banerjee for technical assistance. We thank the National Aeronautics and Space Administration (NASA) Ames Research Center for Gravitational Biology Research support team, including Tianna Shaw, Duncan Atchison, Anthony Purcell, and Ed Houston, for their help with 1-FDC modifications and testing and Marty Hasha for assistance with vibration testing. We thank Robert Majeska for the gracious gift of ROS 17/2.8 cells and Beckman Coulter for the loan of the QuickScan dispersion analyzer. We also thank Sigrid Reinsch and Eduardo Almeida for helpful advice during the course of the study and Wenonah Vercoutere and Emily Holton for critical reading of the manuscript.


    FOOTNOTES
 

Address for reprint requests and other correspondence: N. D. Searby, National Aeronautics and Space Administration Ames Research Center, MS/236-7, Moffett Field, CA 94035 (e-mail: nancy.d.searby{at}nasa.gov)

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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
1. Ajubi NE, Klein-Nulend J, Alblas MJ, Burger EH, and Nijweide PJ. Signal transduction pathways involved in fluid flow-induced PGE2 production by cultured osteocytes. Am J Physiol Endocrinol Metab 276: E171–E178, 1999.[Abstract/Free Full Text]

2. Ajubi NE, Klein-Nulend J, Nijweide PJ, Vrijheid-Lammers T, Alblas MJ, and Burger EH. Pulsating fluid flow increases prostaglandin production by cultured chicken osteocytes: a cytoskeleton-dependent process. Biochem Biophys Res Commun 225: 62–68, 1996.[CrossRef][ISI][Medline]

3. Bakker AD, Soejima K, Klein-Nulend J, and Burger EH. The production of nitric oxide and prostaglandin E2 by primary bone cells is shear stress dependent. J Biomech 34: 671–677, 2001.[CrossRef][ISI][Medline]

4. Bauer MS and Walker TL. Intramedullary pressure in canine long bones. Am J Vet Res 49: 425–427, 1988.[ISI][Medline]

5. Bellows C, Aubin J, Heersche J, and Antosz M. Mineralized bone nodules formed in vitro from enzymatically released rat calvaria cell populations. Calcif Tissue Int 38: 143–154, 1986.[ISI][Medline]

6. Bergula AP, Huang W, and Frangos JA. Femoral vein ligation increases bone mass in the hindlimb suspended rat. Bone 24: 171–177, 1999.[CrossRef][ISI][Medline]

7. Brighton CT, Strafford B, Gross SB, Leatherwood DF, Williams JL, and Pollack SR. The proliferative and synthetic response of isolated calvarial bone cells of rats to cyclic biaxial mechanical strain. J Bone Joint Surg Am 73: 320–331, 1991.[Abstract]

8. Burr DB, Milgrom C, Fyhrie D, Forwood M, Nyska M, Finestone A, Hoshaw S, Saiag E, Simkin A, Rubin C, Turner AS, Mallinckrodt C, Jerome C, McLeod K, and Bain S. In vivo measurement of human tibial strains during vigorous activity. Bone 18: 405–410, 1996.[CrossRef][ISI][Medline]

9. Carter DR and Beaupre GS. Skeletal Function and Form. Cambridge, UK: Cambridge University Press, 2001.

10. Carvalho RS, Scott JE, Suga EM, and Yen EHK. Stimulation of signal transduction pathways in osteoblasts by mechanical strain potentiated by parathyroid hormone. J Bone Miner Res 9: 999–1011, 1994.[ISI][Medline]

11. Chow JW and Chambers TJ. Indomethacin has distinct early and late actions on bone formation induced by mechanical stimulation. Am J Physiol Endocrinol Metab 267: E287–E292, 1994.[Abstract/Free Full Text]

12. Davies PF, Robotewskyj A, and Griem ML. Quantitative studies of endothelial cell adhesion: directional remodeling of focal adhesion sites in response to flow forces. J Clin Invest 93: 2031–2038, 1994.[ISI][Medline]

13. Dee KC, Anderson TT, and Bizios R. Osteoblast population migration characteristics on substrates modified with immobilized adhesive peptides. Biomaterials 20: 221–227, 1999.[CrossRef][ISI][Medline]

14. Fitzgerald J and Hughes-Fulford M. Mechanically induced c-fos expression is mediated by cAMP in MC3T3-E1 osteoblasts. FASEB J 13: 553–557, 1999.[Abstract/Free Full Text]

15. Fujieda M, Kiriu M, Mizuochi S, Hagiya K, Kaneki H, and Ide H. Formation of mineralized bone nodules by rat calvarial osteoblasts decreases with donor age due to a reduction in signaling through EP1 subtype of prostaglandin E2 receptor. J Cell Biochem 75: 215–225, 1999.[CrossRef][ISI][Medline]

16. Globus R, Doty S, Lull J, Holmuhamedov E, Humphries M, and Damsky C. Fibronectin is a survival factor for differentiated osteoblasts. J Cell Sci 111: 1385–1393, 1998.[Abstract/Free Full Text]

17. Guignandon A, Vico L, Alexandre C, and Lafage-Proust MH. Shape changes of osteoblastic cells under gravitational variations during parabolic flight: relationship with PGE2 synthesis. Cell Struct Funct 20: 369–375, 1995.[ISI][Medline]

18. Hatton JP, Pooran M, Li CF, Luzzio C, and Hughes-Fulford M. A short pulse of mechanical force induces gene expression and growth in MC3T3-E1 osteoblasts via an ERK 1/2 pathway. J Bone Miner Res 18: 58–66, 2003.[ISI][Medline]

19. Horikawa A, Okada K, Sato K, and Sato M. Morphological changes in osteoblastic cells (MC3T3-E1) due to fluid shear stress: cellular damage by prolonged application of fluid shear stress. Tohoku J Exp Med 191: 127–137, 2000.[CrossRef][ISI][Medline]

20. Hughes-Fulford M and Lewis M. Effects of microgravity on osteoblast growth activation. Exp Cell Res 224: 103–109, 1996.[CrossRef][ISI][Medline]

21. Ingber D. Integrins as mechanochemical transducers. Curr Biol 3: 841–848, 1991.[CrossRef]

22. Ingber DE. Cellular tensegrity: defining new rules of biological design that govern the cytoskeleton. J Cell Sci 104: 613–627, 1993.[Free Full Text]

23. Klein-Nulend J, Semeins CM, Ajubi NE, Nijweide PJ, and Burger EH. Pulsating fluid flow increases nitric oxide (NO) synthesis by osteocytes but not periosteal fibroblasts: correlation with prostaglandin upregulation. Biochem Biophys Res Commun 217: 640–648, 1995.[CrossRef][ISI][Medline]

24. Komarova SV, Ataullakhanov FI, and Globus RK. Bioenergetics and mitochondrial transmembrane potential during differentiation of cultured osteoblasts. Am J Physiol Cell Physiol 279: C1220–C1229, 2000.[Abstract/Free Full Text]

25. Landis WJ, Hodgens KJ, Block D, Toma CD, and Gerstenfeld LC. Spaceflight effects on cultured embryonic chick bone cells. J Bone Miner Res 15: 1099–1112, 2000.[ISI][Medline]

26. Marczinovits I, Szabo G, Komaromy L, Bajszar G, and Molnar J. Isolation and characterization of nuclear hnRNP complexes from Drosophila melanogaster tissue culture cells. Acta Biochim Biophys Acad Sci Hung 18: 185–198, 1983.[Medline]

27. Meazzini MC, Toma CD, Schaffer JL, Gray ML, and Gerstenfeld LC. Osteoblast cytoskeletal modulation in response to mechanical strain in vitro. J Orthop Res 16: 170–180, 1998.[CrossRef][ISI][Medline]

28. Miwa M, Kosawa O, Tokuda H, Kawakubo A, Yoneda M, Oiso Y, and Takatsuki K. Effects of hypergravity on proliferation and differentiation of osteoblast-like cells. Bone Miner 14: 15–25, 1991.[CrossRef][ISI][Medline]

29. Moursi AM, Damsky CH, Lull J, Zimmerman D, Doty SB, Aota S, and Globus RK. Fibronectin regulates calvarial osteoblast differentiation. J Cell Sci 109: 1369–1380, 1996.[Abstract/Free Full Text]

30. Murray DW and Rushton N. The effect of strain on bone cell prostaglandin E2 release: a new experimental method. Calcif Tissue Int 47: 35–39, 1990.[ISI][Medline]

31. Nakajima T. Effects of hypergravity on migration, proliferation and function of mouse osteoblastic cell line MC3T3-E1. Kokubyo Gakkai Zasshi 58: 529–544, 1991.[Medline]

32. Norvell SM, Ponik SM, Bowen DK, Gerard R, and Pavalko FM. Fluid shear stress induction of COX-2 protein and prostaglandin release in cultured MC3T3-E1 osteoblasts does not require intact microfilaments or microtubules. J Appl Physiol 96: 957–966, 2004.[Abstract/Free Full Text]

33. Owen TA, Aronow M, Shalhoub V, Barone LM, Wilming L, Tassinari MS, Kennedy MB, Pockwinse S, Lian JB, and Stein GS. Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol 143: 420–430, 1990.[CrossRef][ISI][Medline]

34. Ozawa H, Imamura K, Abe E, Takahashi N, Hiraide T, Shibasaki Y, Fukuhara T, and Suda T. Effect of a continuously applied compressive pressure on mouse osteoblast-like cells (MC3T3-E1) in vitro. J Cell Physiol 142: 177–185, 1990.[CrossRef][ISI][Medline]

35. Papaseit C, Pochon N, and Tabony J. Microtubule self-organization is gravity-dependent. Proc Natl Acad Sci USA 97: 8364–8368, 2000.[Abstract/Free Full Text]

36. Piepmeier EH, Kalns JE, McIntyre KM, and Lewis ML. Prolonged weightlessness affects promyelocytic multidrug resistance. Exp Cell Res 237: 410–418, 1997.[CrossRef][ISI][Medline]

37. Putnam AJ, Schultz K, and Mooney DJ. Control of microtubule assembly by extracellular matrix and externally applied strain. Am J Physiol Cell Physiol 280: C556–C564, 2001.[Abstract/Free Full Text]

38. Raisz LG. Prostaglandins and bone: physiology and pathophysiology. Osteoarthritis Cartilage 7: 419–421, 1999.[CrossRef][ISI][Medline]

39. Reich KM and Frangos JA. Effect of flow on prostaglandin E2 and inositol triphosphate levels in osteoblasts. Am J Physiol Cell Physiol 261: C428–C432, 1991.[Abstract/Free Full Text]

40. Reich KM, McAllister TN, Gudi S, and Frangos JA. Activation of G proteins mediates flow-induced prostaglandin E2 production in osteoblasts. Endocrinology 138: 1014–1018, 1997.[Abstract/Free Full Text]

41. Rubin C, Turner AS, Mallinckrodt C, Jerome C, McLeod K, and Bain S. Mechanical strain, induced noninvasively in the high-frequency domain, is anabolic to cancellous bone, but not cortical bone. Bone 30: 445–452, 2002.[CrossRef][ISI][Medline]

42. Sawyer SJ, Norvell SM, Ponik SM, and Pavalko FM. Regulation of PGE2 and PGI2 release from human umbilical vein endothelial cells by actin cytoskeleton. Am J Physiol Cell Physiol 281: C1038–C1045, 2001.[Abstract/Free Full Text]

43. Searby ND. Morphological responses of osteoblasts to mechanical loading: experimental and modeling studies. Doctoral thesis in mechanical engineering, Stanford University, 2002, p. 166.

44. Searby ND, Globus RK, and Steele CR. Structural modeling of an osteoblast subjected to hypergravity loading. Proceedings of the 2001 ASME Bioengineering Conference. Snowbird, UT: ASME BED, 2001.

45. Stein GS, Lian JB, Stein JL, Van Wijnen AJ, and Montecino M. Transcriptional control of osteoblast growth and differentiation. Physiol Rev 76: 593–629, 1996.[Abstract/Free Full Text]

46. Van Loon JJ, Folgering EH, Bouten CV, Veldhuijzen JP, and Smit TH. Inertial shear forces and the use of centrifuges in gravity research: what is the proper control? J Biomech Eng 125: 342–346, 2003.[CrossRef][ISI][Medline]

47. Weyts FA, Bosmans B, Niesing R, Leeuwen JP, and Weinans H. Mechanical control of human osteoblast apoptosis and proliferation in relation to differentiation. Calcif Tissue Int 72: 505–512, 2003.[CrossRef][ISI][Medline]

48. Worofka R and Sauermann G. Nuclear columns, kinetics of RNA synthesis and release in isolated rat liver nuclei. Biochim Biophys Acta 518: 61–80, 1978.[ISI][Medline]

49. Wyrobek AJ, Meistrich ML, Furrer R, and Bruce WR. Physical characteristics of mouse sperm nuclei. Biophys J 16: 811–825, 1976.[Abstract]

50. Yeh CK and Rodan GA. Microtubule disruption enhances prostaglandin E2 production in osteoblastic cells. Biochim Biophys Acta 927: 315–323, 1987.[CrossRef][ISI][Medline]

51. Zeng Y, Cowin SC, and Weinbaum S. A fiber matrix model for fluid flow and streaming potentials in the canaliculi of an osteon. Ann Biomed Eng 22: 280–292, 1994.[ISI][Medline]





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