• ATP release;
  • mechanotransduction;
  • Ca2+ signaling;
  • osteoblasts;
  • fluid shear


  1. Top of page
  2. Abstract
  7. Acknowledgements

ATP is rapidly released from osteoblasts in response to mechanical load. We examined the mechanisms involved in this release and established that shear-induced ATP release was mediated through vesicular fusion and was dependent on Ca2+ entry into the cell through L-type voltage-sensitive Ca2+ channels. Degradation of secreted ATP by apyrase prevented shear-induced PGE2 release.

Introduction Fluid shear induces a rapid rise in intracellular calcium ([Ca2+]i) in osteoblasts that mediates many of the cellular responses associated with mechanotransduction in bone. A potential mechanism for this increase in [Ca2+]i is the activation of purinergic (P2) receptors resulting from shear-induced extracellular release of ATP. This study was designed to determine the effects of fluid shear on ATP release and the possible mechanisms associated with this release.

Materials and Methods MC3T3-E1 preosteoblasts were plated on type I collagen, allowed to proliferate to 90% confluency, and subjected to 12 dynes/cm2 laminar fluid flow using a parallel plate flow chamber. ATP release into the flow media was measured using a luciferin/luciferase assay. Inhibitors of channels, gap junctional intercellular communication (GJIC), and vesicular formation were added before shear and maintained in the flow medium for the duration of the experiment.

Results and Conclusions Fluid shear produced a transient increase in ATP release compared with static MC3T3-E1 cells (59.8 ± 15.7 versus 6.2 ± 1.8 nM, respectively), peaking within 1 minute of onset. Inhibition of calcium entry through the L-type voltage-sensitive Ca2+ channel (L-VSCC) with nifedipine or verapamil significantly attenuated shear-induced ATP release. Channel inhibition had no effect on basal ATP release in static cells. Ca2+-dependent ATP release in response to shear seemed to result from vesicular release and not through gap hemichannels. Vesicle disruption with N-ethylmaleimide, brefeldin A, or monensin prevented increases in flow-induced ATP release, whereas inhibition of gap hemichannels with either 18α-glycyrrhetinic acid or 18β-glycyrrhetinic acid did not. Degradation of extracellular ATP with apyrase prevented shear-induced increases in prostaglandin E2 (PGE2) release. These data suggest a time line of mechanotransduction wherein fluid shear activates L-VSCCs to promote Ca2+ entry that, in turn, stimulates vesicular ATP release. Furthermore, these data suggest that P2 receptor activation by secreted ATP mediates flow-induced prostaglandin release.


  1. Top of page
  2. Abstract
  7. Acknowledgements

BONE IS A dynamic organ, with its architecture constantly changing in accordance with the mechanical usage required of it. As external forces placed on bone decrease, which occurs in prolonged bed rest, immobilization, or microgravity, the skeleton undergoes net resorption, resulting in significant bone loss.(1) Conversely, increased external forces on the skeleton can produce net bone accumulation.(2) Various in vitro loading techniques, including hypotonic swelling, substrate strain, and fluid shear stress (FSS), have been developed to study the cellular responses and mechanisms involved in the perception of mechanical stimuli by bone cells. Whereas none of these models completely replicate the stresses endured by bone, most produce osteoblastic responses that are considered anabolic in vivo. These responses include transients in intracellular calcium levels ([Ca2+]i),(3) changes in gene expression,(4-6) and prostaglandin release.(7,8) We have focused this study on the effects of fluid shear because we have shown that fluid shear, and not physiologic levels of mechanical strain, increases the expression of osteopontin, c-fos, cyclooxygenase 2, and TGFβ(4,9)

Osteoblasts respond to FSS with a rapid increase in intracellular Ca2+ that is dependent on both extracellular Ca2+ entry and intracellular Ca2+ release.(3) Whereas we have shown that FSS-induced increases in gene expression are mediated by intracellular Ca2+ release in osteoblasts,(10) Ca2+ entry in response to shear has been shown to be required for release of prostaglandins,(11) NO,(12) and TGFβ.(13) Rapid Ca2+ entry must occur through ion channels, and two potential candidates for mediation of FSS-induced Ca2+ entry in osteoblasts are the mechanosensitive, cation-selective channel (MSCC) and the L-type, voltage-sensitive Ca2+ channel (L-VSCC).(14) The MSCC has been shown to be important in the release of prostaglandins(11) and TGFβ,(13) whereas inhibition of the L-type VSCC has been shown to inhibit NO release in bone organ cultures(15) and reduce loading-induced bone formation in vivo.(16)

There is a significant body of evidence showing that ATP in the extracellular milieu induces a host of physiologic responses on activation of ATP-binding purinergic (P2) receptors. These receptors are found in a wide variety of cell types and tissues and have been shown to alter Ca2+ signaling in numerous cell types. P2 receptors can be divided into two families of receptors: metabotropic P2Y receptors that induce intracellular Ca2+ release through activation of G-proteins and ionotropic P2X receptors that are ligand-gated channels. Osteoblasts express a variety of P2Y and P2X receptors,(17) and activation of these receptors have been shown to increase [Ca2+]i, propagate calcium waves,(18) induce c-fos,(17) and increase proliferation.(19,20) Release of ATP from the cytosol to the pericellular environment is a regulated process, and its extracellular availability for P2 receptor binding is limited by the presence of membrane-bound nucleotidases.(21) The mechanism(s) of ATP release are unclear, yet chloride-conducting channels,(22,23) gap junctional hemichannels,(24,25) and vesicular mechanisms(26,27) have been implicated in the controlled release of ATP.

In this study, we examined the effects of fluid shear stress on ATP release in MC3T3-E1 osteoblasts. We showed that shear transiently increases ATP and that this release is Ca2+-dependent. We further show that the shear-induced release of ATP is blocked by inhibition of the L-VSCC mediated by vesicular fusion. Most significant is the observation that ATP activation of P2 receptors is important for shear-induced prostaglandin E2 (PGE2) release.


  1. Top of page
  2. Abstract
  7. Acknowledgements

Cell culture

MC3T3-E1 cells, a murine osteoblast-like cell line (a gift from Dr Mary C Farach-Carson, University of Delaware), were grown in α-MEM containing 10% FBS (Gibco, New York, NY, USA), 100 U/ml penicillin G, and 100 μg/ml streptomycin. Cells were maintained in a humidified incubator at 37°C with 5% CO2/95% air and subcultured every 72 h. For shear studies, 80,000 cells were seeded onto rat-tail type I collagen-coated (100 μg/ml; BD, Franklin Lakes, NJ, USA) glass slides. Fluid shear experiments were performed 2 days later, when the cells were 80-85% confluent. Flow media consisted of α-MEM containing 0.5% FBS, 100 U/ml penicillin G, 100 μg/ml streptomycin, and 20 mM HEPES (pH 7.4).

Fluid flow experiments

Fluid flow was applied to cells in a parallel plate flow chamber using a closed flow loop, as described previously(28) (Cytodyne, San Diego, CA, USA). This system uses a constant hydrostatic pressure head to drive media through the channel of the flow chamber to subject the cell monolayer to steady laminar flow resulting in a well-defined fluid shear stress of 12 dynes/cm2. The apparatus was maintained at 37°C throughout the duration of experimentation. The correlation between shear and flow rate was calculated using the equation

  • equation image(1)

where Q is the flow rate (cm3/s); μ is the viscosity of the flow media (0.01 dynes/cm2); h is the height of the channel (0.022cm); b is the slit width (3.2cm); and τ is the wall shear stress (dyne/cm2). For time course studies of ATP release, a programmable Harvard Syringe Pump (PHD programmable; Harvard Apparatus, Hollison, MA, USA) was used to perfuse the flow chamber with fresh media at the aforementioned shear rate of 12 dynes/cm2.

Luciferin/luciferase-dependent detection of ATP

An ATP bioluminescence assay containing luciferin/luciferase reagent was used to detect ATP (ATP Bioluminescence Assay kit HS II; Roche, Indianapolis, IN, USA). This assay uses the conversion of d-luciferin by luciferase into oxyluciferin and light that requires ATP as a cofactor. The resultant luminescence, measured using a Monolight 3010 (BD Biosciences Pharmingen, San Diego, CA, USA), reflects ATP concentration. Conditioned media samples were acquired in two separate protocols. To determine the time course of ATP release, media samples were taken at each time-point and immediately frozen at −80°C for further analysis. For studies using the closed flow loop, a known volume of media was added to the flow loop before fluid shear exposure for 5 minutes. After 5 minutes of fluid shear, 1 ml of media was removed and stored as above. Controls were performed with each drug solution to ensure that the added drugs had no effect on luciferase activity. Results were normalized to cellular protein concentration, as determined by the amido black method.

Pharmacologic agents

All drugs tested were purchased from Sigma Chemical (St Louis, MO, USA) and dissolved in distilled water unless otherwise specified. Drugs were added 45 minutes before the onset of flow and remained present for the duration of the experiment. The following concentrations were used: 5 μM nifedipine (from 5 mM stock in ethanol), 10 μM verapamil, 5 μM 18α-glycyrrhetinic acid (AGA; from 15 mM stock in DMSO), 5 μM 18β-glycyrrhetinic acid (BGA; from 15 mM stock in DMSO), 100 μM monensin (from 100 mM stock in MeOH), 10 μM brefeldin A (BFA; 35 mM stock in EtOH), and 100 μM N-ethylmaleimide (NEM; from 100 mM stock).

Assessment of plasma membrane integrity

Cell damage was assessed quantitatively by measuring the samples of recovered media for lactate dehydrogenase (LDH). Analysis of LDH levels was performed using the CytoTox96 Nonradioactive Cytotoxicity Assay (Promega, Madison, WI, USA). This assay uses NADH, generated by oxidation of lactate into pyruvate, with the conversion of iodonitrotetrazolium into a red formazan product in the presence of diaphorase. The absorbance at 490 nm is proportional to the amount of LDH in the media sample. For a positive control, cells were disrupted by lysing in 0.1% Triton X-100. Serial dilutions of this positive control were compared with LDH levels from media samples.

Gap junctional intercellular communication assays

Gap junctional intercellular communication (GJIC) assays were performed using double labeling and immunofluorescence as described previously.(29,30) In these experiments, cells were loaded with the dyes calcein-AM (Molecular Probes, Eugene, OR, USA) and 1,19-dioctadecyl-3,3,39,39-tetramethylindocarbocyanine perchlorate (DiI; Molecular Probes). Because of its small molecular weight (<1 kDa), calcein is gap junction-permeable and able to transfer to neighboring cells in the presence of functional gap junctions, whereas DiI, a larger, lipophilic dye, incorporates into the membrane and is unable to pass through functional gap junctions. These double-labeled (donor) cells were dropped onto nonlabeled (acceptor) cells in monolayer. If functional GJIC existed, calcein would be transferred to the acceptor cells, whereas DiI would stay in the donor cell. Donor cells can be distinguished from acceptor cells through double exposure with fluorescein and rhodamine filters: donor cells fluoresce yellow because of the presence of both calcein (green) and DiI (red), whereas acceptor cells only fluoresce green.

Two days before experimentation, MC3T3-E1 cells were seeded onto 35-mm glass coverslips in 6-well plates at a density of 60,000 cells/well (acceptor cells). On the day of the experiments, the preconfluent donor cells were removed from the incubator and washed with PBS followed by aspiration. The donor cells were labeled in a solution composed of 2 ml HBSS, 2% BSA, 7 μl DiI, 20 μl calcein-AM, and 20 μl pluronic acid (Molecular Probes) and incubated at 37°C for 30 minutes. After 30 minutes, the dye was aspirated, and the donor cells were detached by trypsinization and resuspended in fresh growth medium. A cell pellet was obtained by 5 minutes of centrifugation at 200g. Five hundred double-labeled donor cells were dropped onto the acceptor cells and incubated for 2 h at 37°C. After incubation, the coverslips were removed from the dishes, washed twice in HBSS, and mounted onto a glass slide using Fluoromount-G (Fischer Scientific). The slides were placed on a Nikon fluorescent microscope (Nikon EFD-3; Optical Apparatus, Ardmore, PA, USA) and visualized using fluorescein and rhodamine to locate the calcein- and DiI-loaded cells, respectively. For studies using AGA or BGA to inhibit GJIC, 5 μM AGA or BGA was added to acceptor cells 45 minutes before the addition of donor cells and maintained in the incubation media until the coverslips were mounted.

Quinacrine staining of intracellular ATP

Osteoblasts were seeded onto type I collagen-coated glass slides as described above. Two days later, when the slides were ∼80-85% confluent, the cells were incubated in 25 μM quinacrine for 30 minutes, rinsed twice in HBSS, and mounted with Fluoromount G. The slides were immediately examined using the same Nikon fluorescent microscope as for GJIC assays.

Prostaglandin measurement

For measurement of PGE2 in sheared cells, experiments were conducted as described above but exposed to fluid shear for 60 minutes instead of 5 minutes. After the 60-minute shear treatment, slides of cells were overlaid with 1 ml of fresh flow media (with or without drug, as appropriate) and incubated for 30 additional minutes at 37°C with 5% CO2. The media were collected, and PGE2 was measured using commercially available ELISA kits (Amersham Biosciences, Piscataway, NJ, USA) according to the manufacturer's instructions. Results were normalized to cellular protein levels. The effect of exogenous ATP addition to static cells was also addressed. Experiments were performed as above for PGE2 release, but overlaid with 1 ml of ATP (at concentrations of 100 nM-1 mM) in flow media for 30 minutes, after which time the media were collected and analyzed by ELISA.

Statistical analysis

A minimum of two slides per treatment was examined each day on at least 3 different days. Two-way ANOVA analyses were used to compare ATP release from MC3T3-E1 cells treated with pharmacological agents. When a significant difference was found between samples, a Fisher's PLSD was performed to localize the significant difference. Statistical significance was considered at p < 0.05, and samples are presented as mean ± SE.


  1. Top of page
  2. Abstract
  7. Acknowledgements

Fluid shear stress induces ATP release

MC3T3-E1 cells exhibited a basal release of ATP (6.2 ± 1.8 nM) that was significantly increased 10-fold (59.8 ± 15.7 nM; p < 0.001) when cells were subjected to 12 dynes/cm2 FSS (Fig. 1A). Because cytosolic ATP concentrations are in the millimolar range,(31) it was necessary to determine whether the changes in extracellular ATP levels resulted from an active release rather than shear-induced cell lysis. To ensure that membrane damage did not contribute to shear-mediated ATP release, we analyzed the conditioned media from sheared cells for the presence of the cytosolic enzyme, lactate dehydrogenase (LDH): if ATP release resulted from cellular damage, LDH would be found in the conditioned media. We consistently found that fluid shear-induced ATP release occurred in the absence of significant plasmalemmal damage compared with static controls (data not shown).

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Figure FIG. 1. (A) Effects of fluid shear stress at 12 dynes/cm2 on ATP release in MC3T3-E1 osteoblasts. After 5 minutes of flow, the flow media was removed for analysis of ATP content using an ATP-dependent luciferin-luciferase reaction. Fluid shear increased ATP release ∼5-fold compared with static controls (ap < 0.01 vs. static cells). Each bar represents the mean ± SE of 10 experiments. (B) Time course of ATP release in response to shear. ATP was released rapidly from MC3T3-E1 cells, peaking within 1 minute of shear application. ATP release returned to near, but elevated, baseline levels with sustained shear (ap < 0.01 vs. static cells at same time-point).20

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ATP release in response to a variety of stimuli has been shown to be transient, occurring rapidly after the stimulus and decreasing to basal levels over a period of 30-60 minutes.(32) To determine if a similar pattern occurs in osteoblastic MC3T3-E1 cells in response to FSS, we examined ATP release at time-points before and after 5 minutes using a programmable Harvard Syringe Pump to produce 12 dynes/cm2 shear. We found that FSS induced a rapid release of ATP within one minute of the onset of FSS that returned to preflow levels with prolonged fluid shear (Fig. 1B).

Shear-induced ATP release requires calcium entry

To determine whether ATP release was Ca2+-dependent, we exposed static MC3T3-E1 cells to the Ca2+ ionophore, ionomycin (1 μM), for 10 minutes, and removed the bathing medium for ATP analysis. Addition of ionomycin produced a 3-fold (p < 0.05) increase in ATP release compared with untreated controls (Fig. 2A). L-VSCC and MSCC channels have been implicated in the intracellular Ca2+ response to mechanical load.(3, 4, 15, 33) To test whether these channels are involved in shear-induced ATP release, we blocked the MSCC with GdCl3 (10 μM) and the L-VSCC with nifedipine (5 μM) and verapamil (10 μM). Nifedipine significantly attenuated the shear-induced ATP release (Fig. 2B) in a manner similar to verapamil inhibition (data not shown). Neither inhibitor altered basal ATP release. Block of the MSCC with GdCl3 did not alter either basal or shear-induced ATP release.

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Figure FIG. 2. ATP release is dependent on extracellular Ca2+ entry. (A) MC3T3-E1 cells were treated with ionomycin (1μM), a calcium ionophore, for 10 minutes. The media were collected for ATP analysis. The addition of ionomycin significantly increased ATP release compared with control cells (ap < 0.02 vs. vehicle control). (B) Effects of channel blockers on ATP release. MSCC inhibition with GdCl3 (10 μM) had no effect on either basal or FSS-induced ATP release, whereas L-type VSCC inhibition with nifedipine (5 μM) or verapamil (10 μM) attenuated FSS-induced ATP release but not basal release (ap < 0.05 vs. static cells in same conditions;bp > 0.05 vs. static cells in same conditions).20

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Shear-induced ATP release does not require GJIC

Gap junctions and GJIC have been implicated in the mechanism through which other cell types release ATP in response to a mechanical signals.(24,25) Because serum proteins can bind to AGA and titrate the effective AGA concentration,(34) it was first necessary to show that GJIC was inhibited under the flow conditions used. GJIC was evaluated using the double-labeling technique as described previously. In Fig. 3A, the green (calcein) fluorescence indicates the coupled cells in the monolayer, whereas the yellow (calcein and DiI) fluorescence indicates the donor cells. We found that 5 μM AGA pretreatment in flow media (containing 0.5% FBS) effectively inhibited GJIC in MC3T3-E1 osteoblasts. Under these conditions, we found that osteoblasts released ATP in the presence of 5 μM AGA when exposed to fluid shear stress (Fig. 3B). Similar results were found when cells were sheared in the presence of another GJ inhibitor, BGA (5μM). These results suggest that shear-induced ATP release in murine osteoblasts does not require GJIC or hemichannels.

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Figure FIG. 3. The role of GJIC in ATP release. (A) GJIC was assessed in MC3T3-E1 cells by dual-label dye transfer. Donor cells double labeled with the fluorescent dyes calcein and 1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI) were placed in contact with unloaded cells in the monolayer. Dye transfer was evaluated after 2 h. In the dual-exposure photographs, calcein has transferred to acceptor cells, which are green, showing functional GJIC; cells fluorescing yellow contain both calcein and DiI and are the dual-labeled donor cells. Preincubation with 5 μM 18α-glycyrrhetinic acid inhibited GJIC. (B) MC3T3-E1 cells pretreated with vehicle control (DMSO) or GJ inhibitors, 18α-glycyrrhetinic acid or 18β-glycyrrhetinic acid, showed that inhibition of GJIC had no significant effect on FSS-induced ATP release (ap < 0.01 vs. static cells in same conditions).20

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Localization of intracellular ATP stores

We examined the localization of intracellular ATP stores in MC3T3-E1 osteoblasts using quinacrine, a cell-permeant fluorophore that binds to ATP. After a 30-minute incubation with quinacrine, a high level of punctuated fluorescence was seen, localized primarily within the cytosol of the cells (Fig. 4A). To assess the role of vesicular exocytosis in ATP release, we used three pharmacologic agents: BFA, which causes disruption of the Golgi apparatus(26); monensin, which prevents vesicle formation from the Golgi apparatus(26,27); and NEM, which prevents vesicle fusion to the plasma membrane by interfering with vesicle-associated NSF proteins.(26,27) Fluid shear in the presence of each of these antagonists significantly attenuated ATP release compared with untreated controls (Figs. 4B4D). These data, combined with the highly granular and punctate intracellular localization of ATP as visualized by quinacrine staining, suggest that ATP is released from murine osteoblasts in a vesicular manner.

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Figure FIG. 4. (A) Quinacrine staining of MC3T3-E1 cells shows grainy, punctate localization of ATP, suggesting that ATP is contained in vesicles. (B) Brefeldin A, an agent that disrupts the Golgi and thereby prevents vesicle formation, attenuated FSS-induced increases in ATP release but had no effect on basal ATP release. (C) Monensin, which maintains the Golgi structure but prevents vesicle budding from the Golgi, similarly attenuated FSS-induced ATP release without affecting static ATP release. (D) N-ethylmaleimide prevents vesicular exocytosis by inhibiting NSF proteins. Addition of NEM also inhibited FSS-induced ATP release. (ap < 0.01 vs. static cells in same conditions).20

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Extracellular ATP is required for flow-induced increases in PGE2

Prostaglandins are rapidly released in response to shear in osteoblasts,(8,11) and their formation is required for load-induced bone formation.(35) To determine whether shear-induced PGE2 release was mediated by extracellular ATP, we added apyrase (5U/ml), a nucleotidase that degrades nucleotide triphosphates into nucleotide monophosphates, to the flow medium. Apyrase attenuated flow-induced increases in PGE2 release (Fig. 5A). Similar results were obtained from experiments performed with the nonspecific P2 antagonist PPADS, and additional experiments with heat-inactivated apyrase confirmed that the enzymatic activity of apyrase, and not some secondary, nonspecific effect, was responsible for attenuating PGE2 release (data not shown); Furthermore, when exogenous ATP was added to static cells, we observed significant increases in PGE2 release (Fig. 5B) without induction of Cox-2 (data not shown). These data suggest that shear-induced ATP secretion mediates the release of prostaglandins, paracrine factors that have been implicated in the anabolic response to exogenous mechanical load.

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Figure FIG. 5. (A) Addition of apyrase (5 U/ml), which hydrolyzes extracellular ATP, significantly decreased PGE2 release from sheared osteoblasts, suggesting that FSS-induced ATP secretion mediates PGE2 release. (B) Addition of exogenous ATP to static, nonsheared cells dose-dependently increased PGE2 release (ap < 0.01 to static control;bp < 0.01 to shear in the absence of apyrase).20

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  1. Top of page
  2. Abstract
  7. Acknowledgements

Several mechanisms have been proposed for stimuli-induced ATP release from various cell types. These proposed mechanisms include release through a chloride conducting pathway,(36) through gap junctional hemichannels,(24,37) and through Ca2+-dependent vesicular exocytosis.(26) Our data indicate that shear-induced ATP release results from Ca2+ dependent vesicular release. However, we did observe that the general chloride channel blocker, 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS), inhibited both basal and shear-induced ATP release (data not shown). While this is a focus of another study, previous reports suggest that DIDS does not block ATP release through inhibition of a chloride conductance, but rather, by binding to a sulfonylurea receptor on the vesicle to prevent fusion of the vesicle to the membrane.(38) Another potential pathway, ATP conductance through gap junctional hemichannels seems to be involved in other cell types,(24,39) but not in the MC3T3-E1 cell model. We showed that gap junction inhibition with either AGA or BGA prevented dye movement through the junctional complex, but had no effect on FSS-induced ATP release. These data support a previous report that found no difference in mechanically stimulated ATP release in human osteoblast-like cells overexpressing the Cx43 connexin compared with wildtype controls.(25) Quinacrine, a ATP-binding fluorophore, showed punctate, granular staining, suggesting that ATP can be localized in vesicles in MC3T3-E1 osteoblasts; further experimentation, such as density gradient fractionation, however, is required to unambiguously show vesicular ATP localization.

Similar to a previous report in epithelial cells,(26) we show that FSS-induced ATP release from osteoblastic cells is the result of Ca2+-dependent vesicular binding to the membrane. Three pharmacologic agents, which cause disruption of the Golgi apparatus,(26) prevent vesicle formation from the Golgi apparatus(26,40) or prevent vesicle fusion to the plasma membrane by interfering with vesicle-associated NSF proteins(26,40) all significantly decreased FSS-induced ATP release compared with untreated controls. Interestingly, none of these agents completely blocked either basal or FSS-induced ATP release. This observation would suggest that either each of these agents does not totally block vesicular fusion to the membrane or that a secondary pathway for ATP release exists.

Whereas numerous studies have shown that ATP binding to P2 receptors results in an increase in [Ca2+]i that is dependent on both extracellular Ca2+ entry and intracellular Ca2+release, few have examined the role of [Ca2+]i in ATP release. Osteoblasts and osteocytes respond to fluid shear with a rapid increase in intracellular Ca2+(3) that is essential for shear-induced changes in actin cytoskeletal organization and gene expression.(10) Whereas this [Ca2+]i response has been shown to be dependent on both extracellular Ca2+ entry and intracellular Ca2+ release, we have shown that only IP3-mediated Ca2+ release is required for the subsequent changes in cell architecture and protein production.(10) However, others have reported that Ca2+ entry through ion channels is important to shear-induced secretion of prostaglandins,(11) TGFβ,(13) and NO(15) from bone cells, leading us to postulate that Ca2+ entry mediates signal amplification, whereas intracellular Ca2+ release results in changes in gene expression in osteoblasts. Katsuragi et al.(41) have shown that inhibition of phospholipase C/IP3-mediated intracellular Ca2+ release blocks angiotensin II-stimulated ATP release in smooth muscle cells. However, in this study, we found no consistent effect of inhibition of phospholipase C with U73122 on either basal or shear-induced ATP release. This lack of consistency may be caused by the subsequent inhibition by U73122 of protein kinase C, which mediates a number of cellular responses, including phosphorylation of Ca2+ channels.

Calcium entry through membrane ion channels could also mediate ATP release in response to shear. This postulate is strengthened by the observation that, when extracellular Ca2+ is removed, ATP release in response to mechanical load is attenuated.(26) Osteoblasts express a number of ion channels capable of conducting Ca2+,(14) but to date, only two have been shown to play a role in the [Ca2+]i response to shear: the mechanosensitive, cation-selective channel (MSCC)(3,33) and the dihydropyridine- and voltage-sensitive L-type Ca2+ channel (VSCC).(33) Activation of the MSCC in response to loading has been associated with prostaglandin, TGFβ, and NO release.(11, 13, 15) Activation of the L-type VSCC has been linked to cellular responses of osteoblasts or osteocytes to shear(15) and hormonal stimulation.(33) We have recently shown that inhibition of this channel with either nifedipine or verapamil significantly reduces bone formation in mechanically loaded rat tibias and ulna, in vivo,(16) indicating the importance of this channel in mechanotransduction in bone.

How L-type VSCCs are activated by mechanical perturbation is unclear. We have postulated that shear-induced activation of the MSCC results in a membrane depolarization that, in turn, activates the VSCC current. However, the data reported here fail to support this premise. Inhibition of the MSCC with 10 μM GdCl3 did not significantly block ATP release, although a reduced ATP release level in response to shear was observed. Thus, either FSS depolarizes the membrane through a separate mechanism to activate the L-VSCC, or this stimulus can directly activate the L-VSCC.

One mechanism through which the L-VSCC could be activated directly is by the autocrine/paracrine action of ATP. ATP released from the cell can bind to P2 receptors that modulate a number of second messenger pathways, including [Ca2+]i. There are two subtypes of P2 receptors: ionotropic (P2X) receptors that allow entry of ions through receptor-mediated channels, and metabotropic (P2Y) receptors that induce G-protein-mediated intracellular Ca2+ release.(42) Whereas the P2Y receptor has been linked with IP3-mediated intracellular Ca2+ release, P2X receptors have been shown to activate both K+ channel and Ca2+ channels, including the L-type VSCC.(43,44) A number of isoforms for each of these subtypes have been described, based on the nucleotide binding selectivity of the receptor and inhibition. Osteoblasts exhibit many of these isoforms for both P2X and P2Y,(17) and ATP binding to these receptors has also been associated with fast, gap junction-independent Ca2+ waves(45) and potentiation of the [Ca2+]i response to PTH in osteoblasts.(46) Here, we showed that MC3T3-E1 osteoblasts respond to a defined fluid shear with release of ATP within 1 minute of shear onset. Because the [Ca2+]i response to shear exhibits a time lag from onset to intracellular response of 30-60 s,(3,33) it is possible that ATP initiates this event to enhance its own release.

Prostaglandin synthesis and release have been shown to occur rapidly in osteoblasts in response to shear(7, 8, 11) and are essential for the anabolic response of bone to mechanical loading.(35) PGE2 release from osteoblasts and osteocytes has been shown to be released in two stages. On application of shear, a brief burst of PGE2 is observed, peaking at 5-10 minutes and returning to levels near baseline.(7,11) This is followed at 45-60 minutes by a large, continuous increase in release that corresponds to an increase in Cox-2 production.(7,8) The importance of Cox-2 function in bone formation was shown when the Cox-2 specific inhibitor, NS398, was given to rats before mechanical loading: the presence of NS398 completely abolished tibial bone formation in response to four-point bending compared with loaded controls.(35) Whereas these results implicate Cox-2 in bone formation in response to exogenous load, it should be noted that prostaglandin synthesis and release does not directly correlate with bone formation, because prostaglandins can similarly promote bone resorption.(47) The addition of exogenous ATP to static MC3T3-E1 cells induced the release of PGE2, and the hydrolysis of ATP released in response to shear blocked flow-induced increase in PGE2 release, suggesting that FSS-induced ATP secretion induces PGE2 release through activation of a P2 receptor. Because Reich and Frangos(48,49) showed that flow-induced PGE2 release was mediated through a Gq-protein, we hypothesize that a metabotropic P2Y receptor is involved in this response.

Whereas the measured amount of secreted ATP in response to shear was on the order of 100 nM (i.e., 1000-fold less than the amount required to induce PGE2 release in static conditions), it should be noted that the data presented in Figs. 1, 2, 3, and 4 represents ATP found in solution. This does not accurately show the local ATP concentration at the cell surface immediately on its release from the cytosol or the local concentration at P2 receptors. Furthermore, whereas their expression has not yet been shown in osteoblasts, other ATP-releasing cells, such as chondrocytes and endothelial cells, express membrane-bound NTP-degrading enzymes (ecto-5′-nucleotidases and exonucleotidases) that regulate the extracellular availability of ATP.

Figure 6 shows our working model for the action of ATP release in osteoblasts. We show that osteoblasts exhibit a basal release of ATP in static cells and respond to a well-defined fluid shear regimen with a significant increase in ATP release. This release is mediated by Ca2+-dependent vesicular fusion to the membrane, although only the shear-induced release is sensitive to Ca2+. We further showed that Ca2+ entry through the L-type VSCC, but not Ca2+ release from intracellular stores, is important to this response. PGE2 release in response to shear seems to be mediated by secreted ATP, suggesting that purinergic signaling may be an important component of the mechanotransduction response of bone.

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Figure FIG. 6. Proposed model for data presented herein. Fluid shear stress activates the L-type VSCC calcium channel through some unknown mechanism. Calcium influx through the L-type VSCC mobilizes vesicular stores of ATP to the plasmalemma, where vesicular fusion releases ATP into the extracellular environment. ATP diffuses to P2 receptors to function in an autocrine or paracrine manner, activating either ionotropic P2X or metabotropic P2Y receptors to increase PGE2 release.20

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  1. Top of page
  2. Abstract
  7. Acknowledgements

This work was supported by National Institutes of Health NIAMS Grants P01 AR45218 and R01 AR43222 (RLD), NIA Grant 13087-09 (HJD), and the National Aeronautics and Space Administration Predoctoral Fellowship Grant NGT5-50366 (DCG). We thank Dr Suzanne M Norvell for assistance with prostaglandin assays and Dr Robert M Bigsby for assistance with the luciferin/luciferase assay.


  1. Top of page
  2. Abstract
  7. Acknowledgements
  • 1
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