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J. Biol. Chem., Vol. 280, Issue 52, 42952-42959, December 30, 2005

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The P2X₇ Nucleotide Receptor Mediates Skeletal Mechanotransduction^*

Jiliang Li1, Dawei Liu1, Hua Zhu Ke¶, Randall L. Duncan, and Charles H. Turner||2

From the Department of Anatomy and Cell Biology and Department of Orthopaedic Surgery, Indiana University School of Medicine, Indianapolis, Indiana 46202, the ^¶Pfizer Global Research and Development, Groton Laboratories, Groton, Connecticut 06340, and the ^||Biomechanics and Biomaterials Research Center, Department of Biomedical Engineering, Indiana University-Purdue University Indianapolis, Indianapolis, Indiana 46202

Received for publication, June 13, 2005 , and in revised form, September 29, 2005.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The P2X₇ nucleotide receptor (P2X₇R) is an ATP-gated ion channel expressed in many cell types including osteoblasts and osteocytes. Mice with a null mutation of P2X₇R have osteopenia in load bearing bones, suggesting that the P2X₇R may be involved in the skeletal response to mechanical loading. We found the skeletal sensitivity to mechanical loading was reduced by up to 73% in P2X₇R null (knock-out (KO)) mice. Release of ATP in the primary calvarial osteoblasts occurred within 1 min of onset of fluid shear stress (FSS). After 30 min of FSS, P2X₇R-mediated pore formation was observed in wild type (WT) cells but not in KO cells. FSS increased prostaglandin (PG) E₂ release in WT cells but did not alter PGE₂ release in KO cells. Studies using MC3T3-E1 osteoblasts and MLO-Y4 osteocytes confirmed that PGE₂ release was suppressed by P2X₇R blockade, whereas the P2X₇R agonist BzATP enhanced PGE₂ release. We conclude that ATP signaling through P2X₇R is necessary for mechanically induced release of prostaglandins by bone cells and subsequent osteogenesis.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mechanical loads applied to bone tissue increase bone formation and improves bone strength (1). Previous studies have suggested that several osteogenic factors, including insulin-like growth factors, transforming growth factor-, nitric oxide (NO), and prostaglandins (PGs),³ mediate mechanically induced bone formation (2–5). PGs in particular have been intensively investigated. Numerous cell culture studies have demonstrated an increased production of PGs in osteoblasts and osteocytes subjected to fluid shear stresses (for review, see Ref. 6). Several PGs have independent anabolic effects on bone tissue. In particular PGE₂ greatly enhances the synthetic activities of osteoblasts (7).

In addition to PGs, nucleotides, such as ATP and UTP, are released from cells in response to mechanical stimulation (8–11), including osteoblasts (12, 13). These nucleotides can act as autocrine and paracrine factors through activation of purinergic (P2) receptors. Both osteoblasts and osteoclasts express several types of P2 receptors (for review, see Ref. 14). Activation of P2 receptors in bone cells by ATP has been shown to increase [Ca²⁺]_i (15, 16). P2 signaling also modulates cyclo-oxygenase-2 expression (17), PG release (18, 19), and c-fos expression (20) in osteoblasts and other cell types.

Based on their molecular structure and activated signal pathways, P2 purinergic receptors are divided into two classes: P2X and P2Y (21). The P2X receptors are ligand-gated ion channels that, in general, are nonselective for monovalent cations. However, some of these receptors are also permeable to Ca²⁺ or even anions. P2Y receptors are G protein-linked receptors that in many cases are coupled through phospholipase C to the release of Ca²⁺ from intracellular stores. Seven P2X subtypes and six P2Y subtypes have been identified in mammalian cells (22). The P2X₇ receptor (P2X₇R) appears to be the most divergent member among P2X family. P2X₇Rs have the unique ability to form large aqueous pores in the membrane, permeable to hydrophilic molecules as large as 900 Da with prolonged exposure to agonists (21). Activation of P2X₇Rs stimulates the release of the inflammatory cytokines such as interleukin-1 in immune cells (23). P2X₇R activation also results in cell membrane blebbing (24) and changes in cellular morphology, which eventually leads to cell death by both necrotic and apoptotic mechanisms (25).

The P2X₇R forms a complex with several proteins including 2 integrin, receptor-like tyrosine phosphatase (RPTP), -actinin, phosphatidylinositol 4-kinase, membrane-associated guanylate kinase, and several heat shock proteins (26). Several of these proteins are associated with mechanotransduction. For instance, RPTP is involved in force-dependent formation of focal adhesion complexes (27), and -actinin links integrins with the actin cytoskeleton promoting transduction of mechanical forces (28). Recently, mice with a null mutation of the P2X₇R were shown to have reduced total bone mineral content, smaller periosteal circumference in the femur, increased trabecular bone resorption, and reduced periosteal bone formation (29). This phenotype of suppressed periosteal bone formation coupled with increased trabecular bone resorption resembles the effects of disuse on the skeleton. When a weight-bearing limb is placed in disuse, e.g. with a plaster cast, periosteal bone formation is suppressed and trabecular bone resorption increases (30). Considering the nature of the protein complex interacting with P2X₇R and the observation that P2X₇R null mice resemble the skeletal disuse phenotype, we hypothesized that the P2X₇R may be necessary for the proper skeletal response to mechanical loading.

To answer the question whether mice lacking the P2X₇R have decreased skeletal sensitivity to mechanical loading, we measured the skeletal response to loading in P2X₇R knock-out (KO) mice. To further study the role of P2X₇R in the osteoblastic response to mechanical loading, calvarial cells isolated from newborn KO and wild type (WT) mice were subjected to a steady laminar fluid flow (FSS). We found that 30 min FSS increased the cellular uptake of 4-[(3-methyl-2(3H)-benzoxazolylidene)methyl]-1[3-(trimethylammonio)propyl]iodide (YO-PRO-1) (molecular mass = 630 daltons) mediated by P2X₇R mediated pore formation in WT cells but not in KO cells. Application of FSS for 15–60 min significantly increased PGE₂ release in WT cells but not in KO cells. In addition, we demonstrated that mechanical signals induced ATP signaling in MC3T3-E1 osteoblasts and MLO-Y4 osteocytes and that ATP activation of P2X₇ is necessary for the subsequent release of PGE₂.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Animals—The P2X₇R KO mouse was generated as reported by Solle et al. (31). The P2X R KO (P2X₇R^–/–) mice as well as the WT (P2X₇R+^/+) mice (C57BL/6) were purchased from Taconic (Germantown, NY) at 4–5 weeks of age. The animals were housed at Indiana University School of Medicine Animal Care Facility for 13–14 weeks (acclimation period) before the experiment began. All animals were allowed free access to standard mouse chow and water during the whole experimental period. All procedures performed were in accordance with the Indiana University Animal Care and Use Committee Guidelines.

In Situ Mechanical Properties of the Ulna and Radius—When the animals reached 18 weeks of age, five mice from male and female WT and KO mice were chosen at random, anesthetized and killed by cervical dislocation. Immediately after killing, each animal was weighed, and the forearm was tested using a miniature materials testing machine (Vitrodyne V1000; Liveco, Inc., Burlington, VT), which has a force resolution of 0.05 newton. The right arm was minimally dissected to expose the medial surface of the midshaft ulna. A single element strain gauge (model EA-06–015DJ-120; Measurements Group, Inc., Raleigh, NC) was fixed to the exposed medial ulnar surface with cyanoacrylete (M-Bond 200; Measurements Group, Inc.) at a point 3.35 mm distal to the brachialis insertion based on the previous report in our laboratory (32). Once fitted with a strain gauge, the voltage was zeroed, and the forearm was loaded in cyclic axial compression using an electromagnetic actuator with force feedback control. Using a 2-Hz haversine waveform, the forearms were laded at 0.95, 1.40, 1.85, and 2.3 newtons, during which peak-to-peak voltage was measured on a digital oscilloscope. Voltage measurements were converted to strain as reported previously (32).

In Vivo Ulnar Loading—Animals in each gender WT or KO mice were subdivided randomly into 3 groups for in vivo loading: low magnitude, medium magnitude, or high magnitude loading groups. Under anesthesia with inhalation of isoflurane (2%), the right forearm of each mouse was loaded at 120 cycles per day for 3 consecutive days with a 2-Hz haversine waveform using the electromagnetic actuator. The left forearms were not loaded and served as an internal control for loading effects. All mice were allowed normal cage activity between loading sessions and afterward. Intraperitoneal injections of calcein (30 mg/kg body weight; Sigma) and alizarin (50 mg/kg body weight; Sigma) were administered 5 and 11 days after the first loading day. All animals were killed 18 days after the first loading day.

Tissue Processing, Histomorphometry, and Ulnar Strain Calculations—Right and left ulnae were processed as reported previously (32). Transverse thick sections (70 µm) were cut at the ulnar midshaft and further ground to a final thickness of 20 µm and then mounted unstained on microscope slides. One section per limb was read on a Nikon Optiphot flurescence microscope (Nikon, Inc., Garden City, NJ) using the Bioquant digitizing system (R&M Biometrics, Nashville, TN). The following primary data were collected from the periosteal surface at x250 magnification: total perimeter (B.Pm); single label perimeter (sL.Pm); double label perimeter (dL.Pm), and double label area (dL.Ar). From these primary data, the following derived quantities were calculated: mineralizing surface (MS/BS = [1/2sL.Pm + dL.Pm]/B.Pm x100; %); mineral apposition rate (MAR = dL.Ar/dL.Pm/6 days; µm/day) and bone formation rate (BFR/BS = MAR x MS/BS x 3.65; µm³/µm² per year). A new set of relative (r) values: rMS/BS, rMAR, and rBFR/BS using the left ulna (nonloaded) values subtracted from the right ulna (loaded) values showed purely mechanically induced bone formation.

Each ulnar cross-section used for histomorphometry was digitally captured through the microscope and imported into SCIONIMAGE, in which total area (Tt.Ar; mm²), cortical area (Ct.Ar; mm²), maximum second moment of inertia (I_MAX; mm⁴); I_MIN and Se.Dm were calculated. Mechanical strain was calculated for the medial periosteal surface of each histological section using methods described in Robling and Turner (32).

Cell Culture—Calvarial osteoblasts were obtained from 3–5-day-old neonatal calvariae from WT and KO mice. Calvariae halves of the same genotype were grouped and subjected to four sequential 20-min digestions with enzyme mixture of 1.5 units/ml collagenase (Sigma) in PBS and 0.05% trypsin, 1 mM EDTA (Invitrogen) at room temperature on a rocking platform. The first digest was discarded. The second to the fourth digest were pooled, and cell pellet was collected after centrifugation at 2,000 rpm for 10 min. Cells were resuspended in -minimal essential medium (-MEM) and passed through a 40-µm cell strainer (Falcon, BD Biosciences). Cell numbers were counted, and cells were plated at an initial density of 2,000 cells/cm² in a T25 culture flask (Costar, Corning, NY) in -MEM containing 10% fetal bovine serum (FBS) (Atlanta Biologicals, Norcross, GA), 100 units/ml penicillin G (Sigma), 100 µg/ml streptomycin (Sigma), and maintained in a 95% air/5% CO₂-humidified incubator at 37 °C. The cells were subcultured every 72 h. MC3T3-E1 cells, a preosteoblastic cell line (passage 10–20), were also cultured in -MEM containing 10% FBS under the same conditions as the primary cells. MLO-Y4 osteocytes were cultured in -MEM supplemented with 5% FBS and 5% calf serum.

Fluid Shear—Primary cells, MC3T3-E1 osteoblasts, and MLO-Y4 osteocytes were seeded at a density of 2,000/cm² and grown on 75 x 38-mm² glass slides (Fisher Scientific) coated with 10 µg/cm² collagen (BD Biosciences). Upon reaching 90% confluence (2–3 days), the cells were serum-starved for 24 h in 0.2% FBS-supplemented culture medium (the same medium used for flow). During the experiment, fluid flow was applied to the cell monolayer in a parallel plate flow chamber using a closed flow loop. This system subjected the cells to a steady laminar flow thus producing a 12 dynes/cm² FSS. The apparatus was maintained at 37 °C, and the medium was aerated with 95% air/5% CO₂ during experiment. In functional studies, the agonists were added immediately prior to flow, while the antagonists were applied to the cells for 30 min prior to flow. The cells were fluid-sheared for either 30 (for pore formation) or 15–60 min (for PGE₂ assay). Control groups were set under the same conditions without flow or drug treatments.

Agonists and Antagonists—To determine the functions of P2X₇ ion channel, several agonists and antagonists were used. BzATP (300 µM, which has been shown to have a high affinity for the P2X₇R (33), was used to activate P2X₇ ion channel and pore formation. The P2X₇R-selective inhibitor, Brilliant Blue G (BBG) (1 µM), was used to block the P2X₇ ion channel and pore formation. To determine the roles of FSS-induced ATP release in activation of P2X₇R, apyrase (5 units/ml) was used. All the above reagents were purchased from Sigma. The P2X₇ antibody that binds to the extracellular portion of the receptor to inhibit activation was obtained from Alomone Laboratories.

Pore Formation Assay—Fluid shear stress-induced pore formation was determined by measuring cellular uptake of YO-PRO-1, molecular mass = 629 daltons; Molecular Probes, Eugene, OR) in primary cells as well as MC3T3-E1 cells. As described above, the cells were fluid sheared for 30 min. After shear, the cells were rinsed once in PBS without Mg²⁺ and Ca²⁺ ions, which have been shown to inhibit pore formation (34). The YO-PRO-1 iodide dye (1 mM in Me₂SO) was diluted to a final concentration of 2 µM in PBS (without Mg²⁺ and Ca²⁺) and then placed on the cells immediately and incubated for 10 min. Afterward, the cells were washed again with PBS without Mg²⁺ and Ca²⁺. The stain-positive cells were observed under an UV-lighted microscope. YO-PRO-1-positive cells were normalized to total cells in the same vision field. The ratios of YO-PRO-1-positive cells over total cells were compared between different treatment groups, and a statistical difference was considered significant when p value was less than 0.05 (5% level). The pore formation assays were repeated three times for each of the experimental conditions.

Measurement of ATP—An ATP bioluminescence assay containing luciferin/luciferase reagent is used to detect ATP (ATP bioluminescence assay kit HS II, Roche Applied Science). This assay utilizes 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 (Pharmingen), reflects ATP concentration. Media samples were taken 1 or 5 min after beginning fluid shear stress and immediately frozen at –80 °C for further analysis.

Prostaglandin E₂ Assay—Cells were subjected to fluid shear stress for either 15 or 60 min. After 60 min of fluid shear flow, 2 ml of 0.2% FBS -MEM media was added onto the monolayer of cells and incubated for 30 min at 37 °C in the incubator. After 30 min, the conditioned media were collected for PGE₂ assay using a commercially available, competitive binding enzyme immunoassay kit (BioTrak, Amersham Biosciences). PGE₂ release to the conditioned media was normalized to total cell protein. The protein assay was accomplished by Amino Black method as used in Western blot.

Apoptosis Assay—To obtain a positive control for apoptosis, MC3T3-E1 cells were treated with TNF- and cyclohexamide (CHX) for 6 h, then lysed for Western blot analysis (35). Together with this positive control, cell lysates from different treatment groups of FSS, BzATP, as well as static control were resolved through 10% SDS-PAGE and blotted with caspase-3 antibody (1:1,000, Cell Signaling, Beverly, MA) as described below.

Western Blot—Following treatment, cells were washed with cold PBS (1x), lysed with 2x sample buffer on ice, and immediately boiled for 5 min. The lysis buffer contained 150 mM NaCl, 26% glycerol (v/v), 1.5 mM MgCl₂, 0.2 mM EDTA, 0.5 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride and buffered with 5 mM Na⁺-HEPES (pH 7.9). The protein samples were centrifuged at 14,000 x g for 10 min at room temperature to remove cellular debris and then quantified using the Amido Black method. Whole cell lysate (20 µg) and a prestained molecular weight marker (Bio-Rad) were boiled for 5 min and separated by 10% SDS-polyacrylamide gel electrophoresis and electrotransferred to a nitrocellulose membrane. The membrane was blocked in Tris-buffered saline containing 5% nonfat dry milk and 0.1% Tween 20 (TBST) and incubated with 1 µg/ml (1:250 in blocking buffer) rabbit anti-P2X₇R antibody (Calbiochem) overnight at 4 °C. The membranes were then washed three times with TBST and incubated with goat anti-rabbit IgG hydroperoxidase-conjugated secondary antibodies (1:5,000 in blocking buffer) for 1 h at room temperature. After washing with TBST, immunodetection was accomplished using the ECL method (Fuji machine).

Statistical Analysis—All the data were expressed as mean ± S.E. Differences between parameters in the loaded (right) and nonloaded (left) ulnae were tested using a paired t test. Dose response to different load magnitudes and mechanical strain within each gender WT or KO mice were tested for significance with least square regression. Differences in slope and x intercept (bone formation versus mechanical strain) between same gender WT and KO mice were tested for significance by analysis of covariance. For all tests, = 0.05. In cellular studies, Western blots were made from either three separate isolations for primary cells or three different passages of cell lines. Significance was established from densitometry measurements using Dunnett's comparison to the static controls or Bonferroni post hoc test for multiple comparisons.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
P2X₇R Null Mice Were Grossly Normal—In general, P2X₇R KO mice appeared normal. At 18 weeks of age, body weights for male KO mice were higher (+5%; p = 0.02) than male WT controls. There were no differences in body weights between KO and WT female mice (p = 0.62). No differences in femoral length were found between KO and WT in female (p = 0.63) or male (p = 0.62) mice.

The Osteogenic Response to Mechanical Loading Was Suppressed in P2X₇R Null Mice—We hypothesized that the long bones of P2X₇R null mice would be less responsive to mechanical loading. A verified experimental approach involving axial loading of the mouse ulna induced new bone formation, mainly at the medial and lateral quadrants of the periosteal surface (Fig. 1). The load-induced bone was lamellar based on the observation under polarized light microscope. Bone formation parameters rMS/BS, rMAR, and rBFR/BS were increased in a magnitude-dependent manner with mechanical loading. All bone formation parameters were significantly greater in WT mice than KO mice (p < 0.01). Of the bone formation parameters rBFR/BS best represents the new bone formation rate, which was closely correlated with peak mechanical strain calculated at the midshaft ulna (Fig. 2). Analysis of the correlations between mechanical strain and rBFR/BS revealed significant differences between WT and KO female mice (p < 0.01; Fig. 2A). Sensitivity to loading (the slope of rBFR/BS versus strain) in female KO mice was reduced by 61% compared with female WT mice, demonstrating that female WT mice exhibited significantly higher rBFR/BS per unit mechanical strain than female KO mice. Sensitivity to mechanical loading in male KO mice was reduced by 73% compared with male WT mice (p < 0.0001; Fig. 2B). In addition, the x intercept of the dose-response curve for the male KO mice was about 31% greater (p = 0.02) in comparison with male WT mice, indicating that more mechanical strain was required to initiate a response in KO mice.

ATP Release Is Not Suppressed in Osteoblasts from P2X₇R Null Mice—To study the role of P2X₇R in osteoblastic responsiveness to mechanical loading, we completed a series of experiments using cultured bone cells from newborn mouse calvaria osteoblasts. Western blot analysis revealed that the P2X₇R receptor was present in WT but not KO cells, whereas MC3T3-E1 osteoblasts and MLO-Y4 osteocytes expressed high levels of P2X₇R (Fig. 3). We have shown that FSS produces a 10-fold increase in ATP release in MC3T3-E1 osteoblasts within 1 min of the onset of FSS (36). When primary osteoblasts from WT mice were subjected to steady laminar fluid flow at 12 dynes/cm² FSS, release of ATP was also observed within 1 min. Primary bone cells from KO mice released similar amounts of ATP compared with WT cells (Fig. 4).

P2X₇R Null Mutation Eliminates Membrane Pore Formation and Prostaglandin Release after a Mechanical Stimulus—Activation of the P2X₇R has been linked with formation of pores in the cell membrane, capable of conducting large molecules (37). To examine whether activation of P2X₇R induced pore formation, we measured uptake of YO-PRO-1, a 629-dalton protein that becomes fluorescent when it binds to nucleic acids, in WT and KO primary bone cells and MC3T3-E1 osteoblasts. FSS induced P2X₇R-mediated pore formation in WT and MC3T3-E1 osteoblasts and MLO-Y4 osteocytes but not in KO osteoblasts (Fig. 5A). Activation of P2X₇R with BzATP also produced pore formation in MC3T3-E1 and WT osteoblasts and MLO-Y4 osteocytes, but not in KO osteoblasts (Fig. 5B). When apyrase was added to the flow media to degrade ATP, pore formation was almost completely inhibited in MC3T3-E1 and MLO-Y4 cells (Fig. 6).

View larger version (36K): [in this window] [in a new window]   FIGURE 1. Periosteal bone formation at the ulnar midshaft was increased with mechanical loading that induces 2,500-µ strain. Mechanical loading of the right ulna activated the formerly quiescent medial and lateral periosteal surfaces, as illustrated by the double fluorochrome labeling (bright bands near the bone surface). Bone formation induced by loading exhibited lamellar organization. The responses in WT mice were more robust (illustrated by the clear double labels) compared with KO mice in which double labels are hard to discern. Note the lack of response in the caudal and cranial periosteal surfaces, which straddle the neutral bending axis, and are consequently subjected to very low strains.

View larger version (11K): [in this window] [in a new window]   FIGURE 2. Relative (right minus left) bone formation rate was normalized to peak mechanical strain. A, female P2X7R knock-out mice exhibited lower sensitivity than female wild type mice; B, male P2X7R knock-out mice exhibited a higher osteogenic threshold and a lower rBFR/BA versus strain slope (lower sensitivity).

View larger version (10K): [in this window] [in a new window]   FIGURE 3. Western blot of P2X7R for calvarial osteoblasts from WT and KO mice, MC3T3-E1 osteoblasts, and MLO-Y4 osteocytes.

View larger version (14K): [in this window] [in a new window]   FIGURE 4. Release of ATP was observed within 1 min following steady laminar fluid flow at 12 dynes/cm2 FSS. Bone cells from KO mice released similar amounts of ATP compared with WT cells.

View larger version (66K): [in this window] [in a new window]   FIGURE 5. A, 30 min of FSS induced membrane pore formation, as determined by measuring cellular uptake of YO-PRO-1 (molecular mass = 630 daltons) in WT and MC3T3-E1 osteoblasts and MLO-Y4 osteocytes but not in KO osteoblasts. B, the P2X7R agonist BzATP induced pore formation in WT and MC3T3-E1 osteoblasts and MLO-Y4 osteocytes but not in KO osteoblasts. Scale bars represent 20 µm.

View larger version (10K): [in this window] [in a new window]   FIGURE 6. MC3T3-E1 osteoblasts and MLO-Y4 osteocytes were subjected to FSS for 30 min and incubated with YO-PRO-1 (1 µM) for 10 min. FSS induced pore formation and adding apyrase, which rapidly hydrolyzes ATP, to the media blocked pore formation.

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
ATP is an early mediator of mechanotransduction in osteoblasts that is released from cells within 1 min after applying mechanical loading (36). ATP initiates intracellular calcium release in osteoblasts through P2Y receptor pathways, which has been the main focus of several previous studies (15–16). However, our results suggest that a novel pathway involving P2X₇R activation is a major contributor to osteoblast mechanotransduction. This pathway culminates in PG release that promotes osteogenesis. Indeed, as much as 73% of the osteogenic response to loading in vivo appears to be linked to the P2X₇R. Previous studies have shown that 50–90% of mechanically induced osteogenesis can be suppressed using blockers of prostaglandin synthesis (39–41), demonstrating the importance of this pathway for bone formation.

View larger version (21K): [in this window] [in a new window]   FIGURE 7. A, the P2X7R antagonist BBG reduced PGE2 release from MC3T3-E1 osteoblasts in a dose-dependent manner. B, the P2X7R antibody (extracellular) significantly reduced flow-enhanced PGE2 release from MC3T3-E1 osteoblasts or MLO-Y4 osteocytes. C, the P2X7R agonist BzATP increased PGE2 release from MC3T3-E1 osteoblasts and MLO-Y4 osteocytes. *, significantly greater than negative control; **, significantly less than positive control.

View larger version (10K): [in this window] [in a new window]   FIGURE 8. PGE2 release was significantly increased after 15 and 60 min of FSS (p = 0.0001) in WT cells, but PGE2 release in KO cells was not significantly increased by FSS at either time point (p = 0.9). *, significantly greater than negative control.

View larger version (7K): [in this window] [in a new window]   FIGURE 9. As a positive control, MC3T3-E1 osteoblasts treated with TNF- and cyclohexamide (Apop) possessed not only an inactive for of caspase-3 (34 kD), but also an active form of caspase-3 (17 kD), demonstrating that apoptosis was induced. However, compared with the static control group, neither FSS nor P2X7R agonist BzATP induced the active form of caspase-3 (17kD).

Fluid shear induced pore formation in cell membranes of osteoblasts and MLO-Y4 osteocytes. This observation is consistent with another study in which pore formation in MLO-Y4 cells was seen after fluid shear (44). It has been proposed that membrane pores form by opening connexin 43 (Cx43) hemichannels. While this mechanism has some experimental support (44), our data suggest that the membrane pore formation is mediated by ATP and the major mechanism is the P2X₇R. We observed pore formation after treatment with the P2X₇R agonist BzATP and pore formation in osteoblasts, and MLO-Y4 osteocytes was blocked with apyrase, which hydrolyzes extracellular ATP thus interrupting its signaling. Others have suggested that Cx43 and P2X₇R might have interacting functions (45). In peritoneal macrophages, Cx43 and P2X₇R are co-localized at the cell membrane. However, we were not able to demonstrate membrane co-localization of Cx43 and P2X₇R in MLO-Y4 osteocytes.⁴

When subjected to mechanical loading, osteoblasts from P2X₇RKO mice did not release prostaglandin E₂.PGE₂ is an anabolic factor in bone (7), and the lack of PGE₂ response in KO cells is probably the cause of the suppressed bone formation response to loading observed in P2X₇RKO mice. Using a cell culture assay of bone formation, others have shown that primary osteoblasts from P2X₇R KO mice do not form mineralized nodules as well as WT osteoblasts (46). Moreover, nodule formation has been shown to be enhanced by the P2X₇R agonist BzATP and suppressed by blockade of prostaglandin synthesis with ibuprofen (46). These findings support our supposition that P2X₇R mediates new bone formation through release of prostaglandins.

It has been shown previously that P2X₇R forms a complex with several putative mechanotransduction proteins such as RPTP or -actinin, so the genetic mutation of P2X₇R could have disrupted the mechanotranduction complex and interfered with other related mechanotransduction pathways. However, since we were able to suppress PGE₂ release in osteoblasts and osteocytes using a neutralizing antibody for P2X₇R, our data suggest that blockade of P2X₇R is effective for suppressing mechanotransduction when the P2X₇R and associated proteins are intact.

Although the absence of a functional P2X₇R resulted in significantly lower sensitivity to loading, the response was not completely abolished. Thus there is the possibility that other P2 receptors are also involved in osteoblastic response to loading. For instance, one previous study demonstrated that P2Y₂R antisense oligodexoxynucleotides decreased the percentage of MC3T3-E1 osteoblastic cells responding to fluid flow with an increase intracellular calcium [Ca²⁺]_i (16). Others (15, 16) have shown that ATP can induce [Ca²⁺]_i mobilization in osteoblasts. In endothelial cells, the addition of apyrase, which rapidly hydrolyzes 5'-nucleotide triphosphates to monophosphates, prevents shear stress induced Ca²⁺ influx (11). This ATP-mediated [Ca²⁺]_i pathway appears to be independent of P2X signaling (16) suggesting that ATP works through two distinct signaling pathways during osteoblast mechanotransduction: 1) P2X activation leading to PG release and 2) P2Y activation and [Ca²⁺]_i mobilization (Fig. 10). Previous studies suggest that these two pathways are largely independent (47). The present study demonstrates that the majority of the osteogenic response is linked to nucleotide signaling through P2X₇R and subsequent PG release from bone cells.

View larger version (9K): [in this window] [in a new window]   FIGURE 10. Release of ATP after FSS in osteoblasts causes signaling through two pathways: P2YR, resulting in intracellular calcium mobilization, and P2X7R, resulting in prostaglandin release.

The linkage between the P2X₇R and osteogenesis provides possible therapeutic options for improving fracture healing and for skeletal diseases. Since the P2X₇R plays a significant role in mechanotransduction, agonists for this receptor may induce the more bone formation where mechanical stresses are highest. This provides a means for optimizing skeletal structure by placing bone only where it is most needed. Our previous study showed that mechanical loading of rat forelimbs can improve bone strength by almost 2-fold, even with only modest (5%) increases in bone mineral density (1). Consequently, selective targeting the P2X₇R may provide an efficient means for strengthening bone to treat diseases of bone fragility like osteoporosis or osteogenesis imperfecta. Also several PGs, particularly PGE₂, substantially increase bone formation and improve bone strength when administered to rats (7, 53). Since the P2X₇R is linked to load-induced release of PGE₂, manipulation of this receptor may provide a means to augment PGE₂ release locally. This strategy might prove useful for accelerating fracture healing or for improving bone regeneration around implants.

	FOOTNOTES

 
^* This work was supported by National Institutes of Health Grants R01AR046530 (to C. H. T.), RO1AR43222 (to R. L. D.), and P01AR045218 (to R. L. D.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ Both of these authors contributed equally.

² To whom correspondence should be addressed. Tel.: 317-274-3226; Fax: 317-274-3702; E-mail: turnerch{at}iupui.edu.

³ The abbreviations used are: PG, prostaglandin; P2X₇R, P2X₇ nucleotide receptor; RPTP, receptor-like tyrosine phosphatase; KO, knock-out; FSS, fluid shear stress; WT, wild type; PBS, phosphate-buffered saline; MEM, -minimal essential medium; FBS, fetal bovine serum; BzATP, 2',3'-O-(4-benzoyl)benzoyl-ATP; BBG, Brilliant Blue G; YO-PRO1, 4-[(3-methyl-2(3H)-benzoxazolylidene)methyl]-1[3-(trimethylammonio-)propyl]iodide; TNF, tumor necrosis factor; CHX, cyclohexamide; Cx43, connexin 43.

⁴ A. B. Castillo and C. H. Turner, unpublished data.

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