1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References

ATP release and subsequent activation of purinergic receptors has been suggested to be one of the key transduction pathways activated by mechanical stimulation of bone. The P2Y13 receptor, recently found to be expressed by osteoblasts, has been suggested to provide a negative feedback pathway for ATP release in different cell types. Therefore, we hypothesized that the P2Y13 receptor may contribute to the mediation of osteogenic responses to mechanical stimulation by regulating ATP metabolism by osteoblasts. To test this hypothesis, wild-type (WT) and P2Y13 receptor knockout (P2Y13R−/−) mice were subject to non-invasive axial mechanical loading of the left tibiae to induce an osteogenic response. Micro-computed tomography analysis showed mechanical loading induced an osteogenic response in both strains of mice in terms of increased total bone volume and cortical bone volume, with the P2Y13R−/− mice having a significantly greater response. The extent of the increased osteogenic response was defined by dynamic histomorphometry data showing dramatically increased bone formation and mineral apposition rates in P2Y13R−/− mice compared with controls. In vitro, primary P2Y13R−/− osteoblasts had an accumulation of mechanically induced extracellular ATP and reduced levels of hydrolysis. In addition, P2Y13R−/− osteoblasts also had a reduction in their maximal alkaline phosphatase (ALP) activity, one of the main ecto-enzymes expressed by osteoblasts, which hydrolyzes extracellular ATP. In conclusion, deletion of the P2Y13 receptor leads to an enhanced osteogenic response to mechanical loading in vivo, possibly because of the reduced extracellular ATP degradation by ALP. The augmented osteogenic response to mechanical stimulation, combined with suppressed bone remodeling activities and protection from OVX-induced bone loss after P2Y13 receptor depletion as previously described, suggests a potential role for P2Y13 receptor antagonist-based therapy, possibly in combination with mechanical loading, for the treatment of osteoporosis.


  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References

Bone integrity is maintained throughout life via bone remodeling, where the balance between bone resorption and formation is critical. Altered coupling of resorption and formation leads to bone disorders such as osteoporosis, which is characterized by higher resorption and lower formation.[1] Most current treatment strategies for osteoporosis have focused on antiresorptive therapies such as bisphosphonates and, more recently, antibodies to receptor activator of NF-κB ligand (RANKL; Denosumab), which can successfully reduce the risk of osteoporotic vertebral fractures.[2, 3] However, the only current anabolic agent for osteoporosis treatment available at the moment is parathyroid hormone (PTH) (either as PTH1-34/teriparatide or full-length PTH1-84). Because of the relatively poor antifracture efficacy at some skeletal sites with these current agents, the need for new anabolic targets is paramount. Mechanical loading of bone is widely accepted as a potent anabolic stimulus for bone formation,[4] and its use as a preventative measure or treatment for osteoporosis is becoming increasingly attractive,[5, 6] especially in combination with drugs that target the osteogenic response pathway.[7, 8] Bone osteogenic adaptation to mechanical loading is performed by regulating the activities of both osteoblasts and osteoclasts,[9] mediated by the osteocytes and bone lining cells that are thought to act as the principal mechanosensors.[10] At the cellular level, mechanical loading-induced osteogenic response is initiated via the release of intracellular molecules such as nitric oxide and prostaglandins (PG), which are anabolic to osteoblasts.[11, 12] Mechanical stimuli can also induce extracellular ATP release from a variety of cells, including osteoblasts.[13-15] This mechanism is now widely believed to be one of the transduction pathways by which mechanical stimulation initiates a cellular response. Upon stimulation, ATP not only mediates the secretion of other intracellular molecules such as PGs[16] but also activates the purinergic receptors such as the P2X7 receptor, which acts as fluid flow sensor for ATP-dependent phosphorylation of ERK in osteoblasts in vitro,[11, 17] stimulating proliferation.[18] In vivo, P2X7 receptor knockout mice have been shown to have ∼70% reduction in the skeletal sensitivity to mechanical loading.[19] Other purinergic receptors are activated by extracellular ATP and have been demonstrated to play a role in integrating local and systemic responses in the activation of bone remodeling.[20] More recently the P2Y13 receptor has been shown to be involved in the regulation of bone remodeling and protection of mice from estrogen deficiency–induced bone loss.[21] In addition, the P2Y13 receptor was also found to provide a negative feedback pathway to inhibit ATP release from human red blood cells in response to low oxygen level.[22] These findings suggest a role for P2Y13 receptors in ATP metabolism and potentially in the response to mechanical loading via other purinergic receptors such as the P2X7 receptor. Indeed, there is evidence showing P2Y13 and P2X7 receptors co-mediate intracellular calcium responses to BzATP in rat cerebellar astrocytes.[23] In addition, it was recently shown that blocking the P2Y13 receptor can mediate ERK1/2 involvement in β-cell apoptosis.[24] Interestingly, ERK1/2 signaling was demonstrated to be involved in osteoblastic response upon mechanical strain and fluid flow.[17, 25]

Given the expression of P2Y13 receptor by osteoblasts and the observed negative feedback pathway for ATP release in red blood cells, we hypothesized that the P2Y13 receptor would play a role in the osteogenic response to mechanical stimulation via regulating ATP metabolism in osteoblasts. To test this hypothesis, we examined the osteogenic response of P2Y13 receptor knockout (P2Y13R−/−) mice to mechanical stimuli in vivo. Non-invasive controlled axial mechanical loading was performed on left tibiae of 4-month-old P2Y13R−/− and wild-type (WT) mice in vivo.[26, 27] Micro-computed tomography (µCT) analysis and dynamic histomorphometry were used to determine the osteogenic response. ATP release and hydrolysis by primary osteoblasts was determined.

Materials and Methods

  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References


P2Y13R−/− mice[28] were backcrossed onto the C57BL/6J background as previously described. Sixteen-week-old P2Y13R−/− and WT mice were housed in the same environmentally controlled conditions with a 12-hour light/dark cycle at 22°C and free to access 2018 Teklad Global 18% Protein Rodent Diet containing 1.01% calcium (Harlan Laboratories, Bicester, UK) and water ad libitum in RB-3 cages. All procedures complied with the UK Animals (Scientific Procedures) Act 1986 and were reviewed and approved by the local Research Ethics Committee of the University of Sheffield (Sheffield, UK).

Mechanical loading in vivo

In this study, the non-invasive axial loading tibial model[26] was used to examine responses to mechanical loading in 16-week-old WT and P2Y13R−/− mice. The peak load (15N) was selected to induce bone formation in the loaded tibiae because evidence showed that similar peak load can induce osteogenic response in female C57BL/6 mice.[26, 29, 30] Briefly, a 14.5-N dynamic load was superimposed onto a 0.5-N preload at rate of 160,000 N/second. Forty trapezoidal-waveform load cycles (0.2-second hold at 15 N) with a 10-second interval between each cycle were applied to mice tibiae, three times a week for 2 weeks. Mice were injected intraperitoneally with calcein (30 mg/kg) on the first (day 1) and last day (day 12) of loading. Mice were then euthanized on day 14.[27] Both tibiae were dissected and fixed in 70% ethanol for µCT and dynamic histomorphometry analysis. The contralateral non-loaded limb (right tibia) was treated as internal control for loading (the functional adaptation in both cortical and trabecular bone being controlled locally and confined to the loaded bones),[27, 31] and the osteogenic responses were expressed as percentage change based on the non-loaded limb data ([parameters of loaded tibia (left)/parameters of own nonloading tibia (right)] × 100%).[32]


Fixed tibiae were scanned using a SkyScan 1172 desktop µCT machine at a resolution of 4.3 µm for the tibia proximal end and 17.3 µm for the whole tibia, with the X-ray source operating at 50 kV, 200 µA, and using a 0.5-mm aluminium filter. Two-dimensional µCT images were captured and reconstructed by Skyscan NRecon (Bruker microCT, Kontich, Belgium) software at a threshold of 0.0 to 0.16 and 0.0 to 0.14 for tibia proximal end and whole tibia scan, respectively. For the tibia proximal end scan, trabecular morphometry was characterized by measuring structural parameters in a 1.0-mm-thick trabecular region, which is 0.2 mm below the growth plate. Cortical morphometry was quantified from the cortical regions locating in the proximal 20% (1.0 mm thick, 1.0 mm below the growth plate) and the midshaft of tibiae (1.0 mm thick, 7.0 mm below the growth plate). Bone tissue mineral densities (TMD) equal to grams of hydroxylapatite per cubic centimeter were calculated based on image greyscale with the following equation: TMD = (0.012 × greyscale value) – 0.296.[21] Nomenclature and symbols were used to describe the µCT-derived bone morphometries according to Bouxsein and colleagues.[33]

Linear-elastic finite element analysis (FEA)

Linear-elastic finite element models of the tibiae were generated to simulate compression of the tibia and to verify strains induced by the 15-N loading force in representative bones from WT and P2Y13R−/− mice scanned post-mortem. Briefly, cement blocks were added to the ends of the tibia to facilitate even application of compressive force at the bone ends. Models were generated directly from voxels of the whole contralateral non-loaded tibial µCT scans using a cube-shaped, 8-node brick element with a side length of 0.0349 mm. Isotropic material properties were assigned to the bone elements using the following empirical equations of Somerville and colleagues:[34]

  • display math

where ρash and ρCT are ash density and bone density from µCT, respectively, and E is modulus of elasticity of bone. The modulus of elasticity for cement was assigned to 2 GPa. The Poisson's ratio was set to 0.35 for bone and cement. The models were solved by a commercial FE package ANSYS (ANSYS Inc., Canonsburg, PA, USA) for stress and strain at each element. The loading-induced average strain in the cortical and trabecular compartment were calculated on a 1.0 mm in length region, 0.2 mm below the growth plate in tibia. An overall strain through the whole length of the tibia was defined as the compressive displacement derived from the FEA (L1 – L'1) divided by the original tibial length (L1) in the non-loading state (Fig. 1A).


Figure 1. Whole bone response to mechanical loading. (A) Finite element models of the mice tibia showing the loading and constraint conditions and length changes before (L1) and under compressive load (L'1). (B) Percentage change in whole tibial bone volume of the loaded compared with unloaded internal control. All values are mean ± SEM, P2Y13R−/− n = 12; WT n = 9. ap < 0.05 (unpaired t test). (C) The 3D models of whole tibia from P2Y13R−/− and WT loaded and non-loaded animals were constructed from µCT images; scale bar = 2.0 mm. (D) The overall strain based on compressive displacement of the whole tibia was analyzed by FEA and compared between WT and P2Y13R−/−. The average strain in the (E) trabecular and (F) cortical compartment were also calculated from a 1.0 mm in length region, 0.2 mm below the growth plate in tibia. n = 5 (unpaired t test).

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Bone dynamic histomorphometry

After µCT analysis, tibiae were embedded into LR White resin (Taab Laboratory Equipment Ltd., Aldermaston, UK). Sections were cut (at 10 μm) longitudinally using a Leica Microsystems Microtome and were examined under UV illumination using a DMRB microscope (Leica Microsystems, Milton Keynes, UK). The bone histomorphometry software Osteomeasure (Osteometrics, Decatur, GA, USA) was used to measure the double-labeled surface (dLS), single-labeled surface (sLS), the separation width between the two fluorescent labels (Ir.L.Th), and total bone surface (BS) on a 3-mm length of both endocortical and periosteal surface, 0.25 mm from the growth plate.[35] The time separating the two labels (Ir.L.t) was the interval between the two IP injects of calcein and was 12 days in all animals. Based on these measurements, mineralizing surface (MS), mineral apposition rate (MAR), and bone formation rate (BFR/BS) were calculated and reported in the results using nomenclature based on the report of the ASBMR Histomorphometry Nomenclature Committee.[36]

Primary osteoblast isolation

Primary osteoblasts were isolated from neonatal mouse calvariae (less than 72 hours old, 5 to 7 pups per culture) as described before.[21] Calvariae were dissected and the attached soft tissue were digested in 1 mg/mL Collagenase 1A (Sigma, St. Louis, MO, USA) for 15 minutes. Calvariae were then subjected to serial digestions in 1 mg/mL Collagenase 1A for 30 minutes; 0.25% Trypsin/EDTA (Gibco, Life Technologies, Paisley, UK) for 15 minutes; and 1 mg/mL Collagenase 1A for 30 minutes at 37°C. All cells were harvested from the digestion suspensions and seeded into a T75 flask and cultured until confluent in DMEM + GLUTAMAX medium with sodium pyruvate (Gibco), 100 units/mL penicillin, 100 µg/mL streptomycin (Gibco), and 10% fetal bovine serum (FBS) (Gibco).

Endogenous ATP release

Fluid flow-induced shear stress is a known stimulator for endogenous ATP release from cells including osteoblasts.[11, 16] The mechanical disturbances caused by simple medium displacement or replacement in vitro are widely accepted methods to induce fluid flow-induced shear stress and stimulate ATP release[37, 38] from cells including osteoblasts.[39] Therefore, medium replacement on primary osteoblast was used to mimic mechanical loading in vitro. First passage primary osteoblasts were seeded into 24-well plates at the density of 5 × 103 cell/well and cultured until 70% confluence in growth medium: DMEM + GLUTAMAX medium with sodium pyruvate (Gibco), 100 units/mL penicillin, 100 µg/mL streptomycin (Gibco), and 10% FBS (Gibco). The cells were washed three times with serum-free medium: DMEM + GLUTAMAX medium with sodium pyruvate, 100 units/mL penicillin, 100 µg/mL streptomycin, and 25 mM HEPES buffer and replenished with 500 μL serum-free medium. Samples were collected from four replicate wells at time points 0, 5, 10, 20, 30, 40, 50, and 60 minutes. ATP concentration was then determined using the HS ViaLight Kit (Lonza, Slough, UK) as previously described.[15] To confirm that ATP release was not caused by cell death, the cell lysis marker lactate dehydrogenase was measured from non-heat-inactivated medium samples using the CytoTox 96-well Non-Radioactive Cytotoxicity Assay (Promega, Southampton, UK) on a SpectraMax M5e microplate reader (Molecular Devices, Sunnyvale, CA, USA) at 492 nm. Samples showing increased LDH release were removed from analysis. Samples for luciferase assay were heated at 98°C for 2 minutes to inactivate soluble ATPases and frozen down immediately in liquid nitrogen and stored at −80°C. Samples for LDH assay were directly frozen down in liquid nitrogen and stored at −80°C.

Exogenous ATP hydrolysis

After sample collection for endogenous ATP release measurement, the media were removed completely from the wells. Fresh serum-free medium (500 µL) was carefully added into each well and the plate incubated for 60 minutes at 37°C to return the medium pH and extracellular ATP concentration to basal levels. Medium samples were collected from four replicate wells per time point for both luciferase and LDH assay before (t = −1 minute) the addition of 300 nM ATP (Sigma; 99.9% pure by HPLC, reconstituted in 25 mM HEPES buffer) and at time points t = 0 (immediately after addition), 5, 10, 20, and 30 minutes.

Alkaline phosphatase (ALP) assay

First-passage primary osteoblast cells isolated from P2Y13R−/− and WT neonatal calvariae were seeded at 1.5 × 104 cells per well in 12-well cell culture plates and cultured for 6 days. At the end of this time period, cells were washed with PBS and harvested by addition of nuclease-free water into each well and samples were snap-frozen at −80°C. Cell lysates were obtained after three freeze-thaw cycles. ALP activity was measured using p-nitrophenyl phosphate (pNPP) (Sigma) as the chromogenic ALP substrate in the presence of Mg2+ ions in a buffered solution. The absorbance was read at 405 nm using the SpectraMax M5e microplate reader. The ALP activity was then normalized to DNA content quantified using Quant-iT PicoGreen dsDNA Assay Kit (Invitrogen, Life Technologies, Paisley, UK) according to the manufacturer's instructions.

Statistical analysis

All data are expressed as mean ± SEM. Statistical significance was tested for using either univariate analysis of variance (PASW Statistics, Armonk, NY, USA) or a t-test (Prism 5, GraphPad, La Jolla, CA, USA).


  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References

Osteogenic response of whole tibia

After 2 weeks of axial loading of the left tibiae of 16-week-old mice, µCT analysis at the level of the whole bone demonstrated that the loaded tibia of the P2Y13R−/− mice had a significantly greater increase in total bone volume (BV) than WT in response to mechanical loading, when compared with the BV of the non-loaded control (126.7% ± 1.2 versus 121.6% ± 1.4, p = 0.0140) (Fig. 1B). The morphological changes were compared on the loaded and non-loaded tibia of WT and P2Y13R−/− mice using µCT 3D models of the whole bone (Fig. 1C). The FEA showed that there was no significant difference in the simulated loading-induced strain through the full length of the tibia between WT and P2Y13R−/− mice (5081 ± 254.4 versus 5048 ± 258.8 microstrain, p = 0.9306) (Fig. 1D). The FEA-based average strain across the trabecular (696.0 ± 60.0 versus 693.4 ± 94.5 microstrain, p = 0.9820) and cortical compartments (757.8 ± 20.3 versus 758.2 ± 20.2 microstrain, p = 0.9894) were also not significantly different between WT and P2Y13R−/− mice (Fig. 1E, F).

Osteogenic response of trabecular bone

Analysis of the trabecular bone structure of the tibial region by µCT demonstrated that both P2Y13R−/− and WT mice had significantly increased trabecular bone volume (BV/TV), trabecular thickness (Tb.Th), trabecular number (Tb.N), and trabecular pattern factor (Tb.Pf) in loaded tibia compared to internal non-loaded controls. The quantitative data are summarized in Table 1, and thicker trabeculae were clearly visible in images of 3D models of the loaded tibia trabecular bone from both P2Y13R−/− and WT mice (Fig. 2A).

Table 1. Quantitative Results of Tibia Trabecular and Cortical Bone After Mechanical Loading Using µCT Analysis
 WT n = 9P2Y13R−/− n = 12
LoadedNon-loadedp valueLoadedNon-loadedp value
  1. TMD = tissue mineral density; BV/TV = trabecular bone volume; BS/BV = bone surface; Tb.Th = trabecular thickness; Tb.N = trabecular number; Tb.Pf = trabecular pattern factor; Tb.Sp = trabecular separation; SMI = structure model index ; DA = degree of anisotropy; Ct.V = cortical bone volume.

  2. Values are mean ± SEM,

  3. a

    p < 0.05,

  4. b

    p < 0.01,

  5. c

    p < 0.001 (paired t test).

TMD (g/cm3)1.12 ± 0.001.13 ± 0.01b1.12 ± 0.001.13 ± 0.00a
BV/TV12.30 ± 0.398.42 ± 0.19c8.60 ± 0.375.80 ± 0.24c
BS/BV (1/mm)65.00 ± 1.0585.41 ± 1.33c67.80 ± 0.6091.50 ± 1.24c
Tb.Th (mm)0.064 ± 0.0010.051 ± 0.001c0.065 ± 0.0010.048 ± 0.001c
Tb.N (1/mm)1.93 ± 0.071.66 ± 0.04a1.33 ± 0.061.20 ± 0.05a
Tb.Pf (1/mm)16.93 ± 0.9525.62 ± 0.57c24.38 ± 0.8930.72 ± 0.94c
Tb.Sp (mm)0.25 ± 0.010.26 ± 0.01 0.29 ± 0.010.31 ± 0.01 
SMI2.06 ± 0.062.16 ± 0.03 2.53 ± 0.052.37 ± 0.05a
DA2.04 ± 0.082.28 ± 0.10 1.66 ± 0.051.90 ± 0.06a
Proximal 20% Ct.V (mm3)1.16 ± 0.020.91 ± 0.01c1.17 ± 0.020.86 ± 0.01c
Midshaft Ct.V (mm3)0.98 ± 0.020.71 ± 0.01c1.00 ± 0.020.67 ± 0.01c

Figure 2. Trabecular bone response to mechanical loading. (A) Three-dimensional images of a region of 1.0-mm-thick trabecular bone 0.2 mm below the growth plate of mechanical loaded and non-loaded tibiae; scale bar = 0.5 mm. The contralateral non-loaded right tibiae were used as internal controls. The percentage change of (B) trabecular thickness (Tb.Th), (C) trabecular bone volume (BV/TV), (D) trabecular pattern factor (Tb.Pf), and (E) structure model index (SMI) for loaded tibia compared with non-loaded controls. All values are mean ± SEM, P2Y13R−/− n = 12; WT n = 9. ap < 0.05 (unpaired t test).

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When compared with the parameters from the contralateral non-loaded tibia, P2Y13R−/− mice showed a significantly higher Tb.Th increase compared with the increase in WT mice (134.1 ± 1.9% versus 126.3 ± 3.0%, p = 0.0316) (Fig. 2B), whereas the increase of BV/TV of P2Y13R−/− was not significantly higher than WT (149.1 ± 5.1% versus 146.4 ± 4.1%, p = 0.6982) (Fig. 2C). P2Y13R−/− mice had almost 21% lower Tb.Pf decreases in the loaded tibia (80.1 ± 3.7% versus 66.2 ± 3.8%, p = 0.0185) (Fig. 2D). More interestingly, the P2Y13R−/− trabecular bone had positive changes to the structure model index (SMI) compared with negative changes in the WT (107.0 ± 2.8% versus 95.4 ± 3.7%, p = 0.0189) (Fig. 2E).

Osteogenic response of cortical bone

Cortical bone volume of the tibia at 20% proximal and at the mid-shaft (Fig. 3A) was measured by µCT and demonstrated that both P2Y13R−/− and WT had significantly increased cortical bone volume (Ct.V) in the loaded tibia (Table 1). Compared with the osteogenic response of WT, P2Y13R−/− mice showed significantly greater responses in both regions (Fig. 3B, C), including significantly increased Ct.V response in both the proximal 20% region (136.4 ± 2.3% versus 128.2 ± 1.5%, p = 0.0130) (Fig. 3D) and the midshaft region (148.3 ± 4.1% versus 136.6 ± 2.8%, p = 0.0362) (Fig. 3E).


Figure 3. Cortical bone response to mechanical loading. (A) Mouse tibial 3D models indicating the two regions analyzed for determining cortical bone parameters, including proximal 20% and the midshaft of tibiae (1.0 mm in thickness, 1.0 mm., and 7.0 mm below the growth plate, respectively). The cross section µCT images of loaded and non-loaded tibiae were compared between WT and P2Y13R−/− at (B) 2.0 mm and (C) 8.0 mm below the growth plate. The Ct.V in loaded tibiae normalized to contralateral non-loaded right tibiae at (D) the proximal 20% region and (E) the midshaft region. All values are mean ± SEM, P2Y13R−/− n = 12; WT n = 9. ap < 0.05 (unpaired t test).

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Rate and extent of mineralization induced by mechanical loading of the tibia

Two distinctive calcein labels (14 and 2 days before euthanization, respectively) on both 20% proximal and midshaft endocortical surfaces of tibiae can be visualized using a fluorescent microscope and confirmed the endocortical lamellar bone formation (Fig. 4A). Calcein labels on both endocortical and periosteal surfaces were measured to calculate the parameters including MAR, BFR/BS, and MS. P2Y13R−/− mice showed a significant increase in all three parameters in both endocortical and periosteal surfaces of loaded tibiae compared with non-loaded control tibiae. However, WT mice only showed significant changes in periosteal BFR/BS and MAR on both periosteal and endocortical surfaces. All of the quantitative data are summarized in Table 2.


Figure 4. Rate and extent of mineralization induced by mechanical loading of the tibia. Double calcein labeling was used to determine the bone formation activities on both endocortical and periosteal surface. (A) Clear double labeling of calcein on endocortical surfaces confirmed lamellar bone formation at this site. The percentage change of loaded tibia compared with contralateral non-loaded right tibiae. (B) Mineral apposition rate (MAR), (C) bone formation rate (BFR/BS), and (D) mineralizing surface (MS%) on the endocortical surface. (E) MAR, (F) BFR/BS, and (G) MS on the periosteal surface. All values are mean ± SEM, P2Y13R−/− n = 5, WT n = 6, ap < 0.05 (unpaired t test).

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Table 2. Quantitative Results of Endocortical and Periosteal Tibia Dynamic Histomorphometry
 WT n = 6P2Y13R−/− n = 5
LoadedNon-loadedp valueLoadedNon-loadedp value
  1. MS = mineralizing surface; MAR = mineral apposition rate; BFR/BS = bone formation rate.

  2. Values are mean ± SEM,

  3. a

    p < 0.05,

  4. b

    p < 0.01,

  5. cp < 0.001 (paired t test).

Endocortical MS (%)85.68 ± 6.2678.20 ± 6.85 88.83 ± 0.8751.97 ± 6.77b
Endocortical MAR (µm/day)1.87 ± 0.161.38 ± 0.11a2.60 ± 0.590.80 ± 0.15a
Endocortical BFR/BS (µm3/µm2/day)1.58 ± 0.141.11 ± 0.16 2.32 ± 0.550.44 ± 0.13a
Periosteal MS (%)94.91 ± 3.4778.83 ± 11.01 92.09 ± 3.7152.53 ± 9.12b
Periosteal MAR (µm/day)3.62 ± 0.560.65 ± 0.06b3.28 ± 0.390.34 ± 0.03b
Periosteal BFR/BS (µm3/µm2/day)3.45 ± 0.570.54 ± 0.01b3.05 ± 0.420.19 ± 0.05b

To determine if the response of the P2Y13R−/− mice was different to WT, the loaded tibia data were compared to contralateral non-loaded tibia. In the endocortical surfaces, loaded tibiae of P2Y13R−/− showed more than a two-fold increased response in MAR (355.4 ± 88.4% versus 140.5 ± 16.4%, p = 0.0276) (Fig. 4B), a five-fold increased response in BFR/BS (714.7 ± 235.4% versus 171.1 ± 41.1%, p = 0.0338) (Fig. 4C), and almost a two-fold higher response in MS (186.6 ± 30.8% versus 115.6 ± 16.1%, p = 0.0599) (Fig. 4D). The same trend was found on the periosteal surface, but only the increased response in MAR by P2Y13R−/− mice reached statistical significance (973.7 ± 108.2% versus 586.6 ± 116.4%, p = 0.0402) (Fig. 4E).

Endogenous ATP release from primary osteoblasts in vitro

Endogenous ATP release after medium change from primary osteoblasts was examined using the luciferase assay. LDH assay was used to exclude ATP release owing to cell lysis. After medium change (t0), the initial extracellular ATP released from P2Y13R−/− osteoblasts into the medium showed no significant difference compared with WT cells (18.6 ± 3.6 nM versus 20.5 ± 3.4 nM, p = 0.7063). The extracellular ATP concentration in the medium of WT osteoblast cultures gradually returned to basal level 60 minutes (t60) after medium change (t0 = 20.5 ± 3.4 nM versus t60 = 9.6 ± 1.6 nM, p = 0.0227). However, the extracellular ATP concentration in the medium of P2Y13R−/− cells did not return to baseline and demonstrated a trend toward accumulation instead of degradation, with the ATP concentration being significantly higher than the initial concentration from 50 minutes onward (t0 = 18.6 ± 3.6 nM versus t50 = 32.7 ± 4.2 nM, p = 0.0182). The extracellular ATP concentration in the medium of P2Y13R−/− osteoblast cultures was also significantly higher than that of WT cultures from 50 minutess after medium change (32.7 ± 4.2 nM versus 15.6 ± 2.6 nM, p = 0.0023) (Fig. 5A).


Figure 5. Regulation of extracellular ATP levels in osteoblast cultures. (A) A time course of ATP release and degradation in osteoblast cultures after medium change. P2Y13R−/− osteoblasts showed a trend of extracellular ATP accumulation compared with the gradual degradation seen in WT cultures. All values are mean ± SEM, n = 4 per experiment, with three independent experiments, ap < 0.05, bp < 0.01 (unpaired t test). (B) Exogenous ATP (300 nM) was hydrolyzed to half the amount within 5 minutes in WT osteoblast cultures. However, the degradation of exogenous ATP in P2Y13R−/− osteoblasts was slower than WT, with extracellular ATP concentration in the P2Y13R−/− cultures being significantly higher than WT from 5 minutes onward. All values are mean ± SEM, n = 4 per experiment, with three independent experiments, bp < 0.01, cp < 0.001 (unpaired t test). (C) ALP activity of WT and P2Y13R−/− osteoblast cultures was measured using pNPP assay and normalized to dsDNA content. All values are mean ± SEM, n = 3 repeat experiments with 12 replicates per experiment, cp < 0.001 (univariate analysis of variance).

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Exogenous ATP hydrolysis by primary osteoblasts

After measuring endogenous ATP release, primary osteoblast cells were incubated in serum-free medium to let ATP concentration and pH settle back to basal levels. Exogenous ATP (300 nM) was added into each well and the concentration of ATP in the medium determined over a time course. The hydrolysis of exogenous ATP in P2Y13R−/− osteoblast cultures was slower than that in WT cultures. The ATP concentration in WT osteoblast cultures reduced by 50% within 5 minutes, whereas the ATP concentration of P2Y13R−/− cultures was significantly higher than WT from 5 minutes and remained at the 200 nM level even after 30 minutes (Fig. 5B).

ALP activity of primary osteoblasts

ALP is a nucleotidase highly expressed by osteoblasts that is capable of hydrolyzing extracellular ATP. The basal level of ALP activity was measured in primary osteoblast cultures using the pNPP assay. P2Y13R−/− mice showed a 15% reduction in ALP activity compared with osteoblasts from WT mice when normalized to DNA content (0.72 ± 0.02 U/μg versus 0.85 ± 0.03 U/μg, p = 0.0002, Fig. 5C).


  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References

The P2Y13 receptor has been suggested to be involved in ATP metabolism in different cell types, and ATP release and purinergic signaling is one of the main transduction pathways of mechanical stimulation. Therefore, we hypothesized that the P2Y13 receptor would play a role in regulating ATP metabolism by osteoblasts and in mediating the osteogenic response upon mechanical stimulation. To test this hypothesis, we examined the osteogenic response of P2Y13R−/− mice subject to mechanical stimuli both in vivo and in vitro. The results provide compelling evidence for a role for the P2Y13R in bone homeostasis. Although the effect of the deletion of the P2Y13R on the normal bone phenotype is modest, the response to loading in vivo is dramatically enhanced in the knockout mice, possibly because of the lack of a P2Y13R regulated negative feedback pathway for ATP release, as demonstrated in vitro.

Non-invasive axial mechanical loading at peak loading force of 15 N was performed on left tibiae of both P2Y13R−/− and WT mice in vivo using a method as described before.[26, 27] Compared with the contralateral non-loaded right tibia, the total bone volume of loaded tibia demonstrated significant increases in both WT and P2Y13R−/− mice, although bone length did not change. This indicated that mechanical loading successfully induced osteogenic response mainly in the tibia cross-sectional dimensions.[40] High-resolution µCT analysis showed that trabecular bone in both WT and P2Y13R−/− loaded tibia had significantly increased BV/TV, Tb.Th, and Tb.N. Similar increases in Ct.V were also found in cortical bone. Therefore, the total BV increase was a combined result of new bone formation activities from both trabecular and cortical bone. This was confirmed with the increased BFR and MAR in both WT and P2Y13R−/− loaded tibiae using dynamic histomorphometry analysis, especially the lamellar bone formation on the endocortical bone surfaces. In addition, increased bone remodeling activities led to a coarse surface, which was observed specifically in the periosteal surface of tibial proximal end 3D µCT image. This result was consistent with previous findings that there was a greater osteogenic response in the corticocancellous proximal metaphysis[41] and periosteal formation surface was predominantly woven bone.[42, 43]

To compare the extent of the osteogenic response between P2Y13R−/− and WT mice, the parameters from loaded tibiae were compared with those from the corresponding contralateral non-loaded tibia controls. The P2Y13R−/− mice had a further 20% response in total BV increase in the loaded tibiae compared with WT. This was mainly the result of the increased osteogenic response of cortical bone because P2Y13R−/− had a significantly greater response in the increases in Ct.V but not in trabecular BV/TV over that of WT. The higher osteogenic response in P2Y13R−/− mice under mechanical stimulation mainly involved osteoblastic bone-forming activities. This was confirmed by the results of fluorochrome double labeling in the cortical compartment, which showed dramatically higher MAR and BFR increases in P2Y13R−/− bones compared with WT, indicating enhanced activities of osteoblasts.[36]

The trabecular structure of P2Y13R−/− mice after loading did not alter toward the ideal load-bearing architecture as the WT mice did; the P2Y13R−/− mice showed less of a decrease in Tb.Pf and significantly increased SMI, indicating that the trabecular did not improve connectivity in any great extent and remained a rodlike structure.[44, 45] However, WT mice showed better structure alteration with significantly decreased Tb.Pf and slightly reduced SMI. The reduced change in Tb.Pf could be the result of a weaker primary trabecular structure in P2Y13R−/− bones, whereas the possible reason for an increased SMI could be because of a failure in osteoclast resorption of the P2Y13R−/− mice as demonstrated previously.[21] This would lead to an abnormal capacity to remodel the trabecular structure because osteoclasts are suggested to control the conversion of trabeculae from plate elements to rod elements.[45]

One possible explanation for the different osteogenic response could have been that the lower bone volume in the P2Y13R−/− mice led to an increase in the strains engendered by the 15 N loading. However, our FEA studies, a widely recognized method to predict loading-induced strain,[46] demonstrated that this is not the case because the bones of the WT and P2Y13R−/− mice experienced the same overall strains and average strain across trabecular and cortical compartments under modeled loading. The overall strains calculated were in the region of 5000 microstrain and are relatively higher than previous studies using strain gauge to measure strain[26, 27, 30] but are consistent with other new findings using FEA.[43] This is because applying the 15 N loads to the tibia in silico is not the same as loading tibia in vivo, where several layers of other tissues including skin, subcutaneous tissues, and at least two thicknesses of cartilage are compressed as well. The important issue is therefore not the absolute values derived from the FEA measurement but the lack of strain difference between WT and P2Y13R−/− bones and, hence, the observed enhanced osteogenic response to mechanical loading in P2Y13R−/− mice is real.

Another possible cause of the different osteogenic response could have been the result of enhanced woven bone formation owing to an increased inflammatory response.[47] However, our dynamic histomorphometry results clearly show lamellar bone formation on the endocortical bone surface, where the increases in both MAR and BFR/BS in loaded tibiae were significantly higher in P2Y13R−/− than those in WT mice. On the periosteal surface, where woven bone formation was predominant, there is a similar trend of enhanced bone formation in P2Y13R−/− mice, but it is not as dramatic as on the endocortical bone surfaces and only the MAR reached statistical significance at this site. Therefore, there may be an element of an inflammatory response, but we believe it is not the main cause of the different adaptation to mechanical loading between WT and P2Y13R−/− mice.

Many mechanisms have been suggested to be involved in the alteration of osteogenic response to mechanical loading in mice, including aging and changes in other signaling pathways such as Wnt, ER, and BMP/TGFβ pathways.[48, 49] The in vitro findings in this study may provide a possible explanation for the reason why P2Y13R−/− mice had enhanced osteogenic response to mechanical loading. The constitutive endogenous ATP release was investigated in the primary osteoblasts isolated from neonatal mice calvariae using luciferase assay. After medium change, the extracellular ATP concentration in the medium of P2Y13R−/− osteoblast cultures showed a trend toward accumulation of ATP instead of gradually degrading ATP as in WT osteoblast cultures. As a result, P2Y13R−/− osteoblasts showed three-fold higher extracellular ATP concentration than WT cells 1 hour after medium change. This confirms that the deletion of P2Y13R results in a lack of the negative feedback pathway for ATP release in P2Y13R−/− osteoblasts. Interestingly, when a higher concentration of exogenous ATP was added to the primary osteoblasts, P2Y13R−/− cells had a decreased capacity to hydrolyze ATP, whereas WT osteoblasts degraded the exogenous ATP back to basal levels within 5 minutes. Thirty minutes after exogenous ATP treatment, extracellular ATP concentration of P2Y13R−/− osteoblasts was double that of WT cells. Osteoblasts are known to have numerous membrane-bound nucleotidases, which are responsible for breaking down ATP to adenosine and are critical in the ATP turnover process.[50] One particular nucleotidase, ALP, is highly expressed by osteoblasts and, interestingly, the ALP activity in vitro was found to be 15% lower in P2Y13R−/− osteoblasts than WT under basal conditions, possibly because of the downregulation of RhoA/ROCK I signaling pathway as a consequence of P2Y13R deletion.[21, 51] Therefore, one possible mechanism leading to the observed higher osteogenic response to mechanical loading in P2Y13R−/− mice may be a result of a reduction in nucleotidase activity. Under basal conditions, it appears that the reduced level of ATP hydrolysis to ADP is still sufficient to provide a negative feedback pathway to regulate ATP release. However, under mechanical stimulation, increased and sustained ATP release may not be matched by hydrolysis to ADP because of basal-reduced ALP levels, and therefore, a lack of the negative feedback loop leads to extracellular ATP accumulation. This extracellular ATP accumulation may in turn trigger other P2 receptor signaling pathways and cause an increased osteogenic response possibly via ATP-dependent phosphorylation of ERK,[11, 17] which then stimulates osteoblastic proliferation and drives the osteogenic response.[18]

In conclusion, this study examined the role of P2Y13 receptor in bone osteogenic response to mechanical loading in vivo and in vitro. Deletion of the P2Y13R leads to higher bone formation, mainly in cortical compartment, than WT upon mechanical loading in vivo, possibly because of the lack of P2Y13R regulated negative feedback pathway for ATP release. This was further supported by our in vitro findings of abnormal extracellular ATP accumulation from primary osteoblast after mechanical stimulation. Reduced ALP activity caused by P2Y13R gene deletion and the following reduction in extracellular ATP degradation might be one reason for this phenomenon. This augmented osteogenic response to mechanical stimulation, combined with suppressed bone remodeling activities and protection from OVX-induced bone loss after P2Y13R depletion as recently described,[21] suggests a potential role for P2Y13R antagonist-based therapy, possibly in combination with mechanical loading, for the treatment of osteoporosis in the future.


  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References

The authors thank the staff of the Bone Analysis Laboratory, The University of Sheffield, for tissue processing. This study was supported by the European Commission under the 7th Framework Programme (proposal #202231) performed as a collaborative project among the members of the ATPBone Consortium (Copenhagen University, University College London, University of Maastricht, University of Ferrara, University of Liverpool, University of Sheffield, and Université Libre de Bruxelles), and is a substudy under the main study “Fighting osteoporosis by blocking nucleotides: purinergic signaling in bone formation and homeostasis.”

Authors' roles: AG, TS, and JMB conceived the project. NW and RR performed the experiments. NW and LY performed the FEA. BR generated the P2Y13R−/− mice. Data analysis and interpretation: NW, RR, LY, TS, JMB, and AG. NW and AG wrote the draft manuscript, with input from all authors.


  1. Top of page
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
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