Strontium-substituted hydroxyapatite nanoparticles as dual-function regulators of osteoblast mineralization and RANKL-induced osteoclastogenesis

Abstract

This study investigated the regulatory effects of strontium-substituted hydroxyapatite (SrHAp) on bone cell regulation, with a specific focus on osteoblast and osteoclast activities. X-ray diffraction verified the successful incorporation of strontium into the hydroxyapatite lattice, confirming the high phase stability despite the presence of a minor amount of CaO. SrHAp nanoparticles with a Sr/Ca molar ratio of 0.417 and a (Ca+Sr)/P ratio of 1.73 were synthesized. The resulting material exhibited a zeta potential of -14.9 mV and an average particle size of 712.6 nm. Cell viability assays revealed that SrHAp concentrations of 100 μg/mL for HOS cells and up to 1000 μg/mL for RAW 264.7 cells were not cytotoxic. Furthermore, SrHAp treatment significantly reduced basal reactive oxygen species levels in HOS cells, suggesting its antioxidant capacity. Our results demonstrated that SrHAp significantly promoted osteoblast differentiation and mineralization, as evidenced by increased calcium deposition detected using Alizarin Red S staining and the upregulation of osteogenic markers, including RUNX2 and osteocalcin. Moreover, SrHAp effectively inhibited RANKL-induced osteoclastogenesis. The morphological analysis of RANKL-treated RAW 264.7 cells revealed a reduction in TRAP-positive multinucleated cells, and this result was supported by decreased TRAP activity. Mechanistic investigations revealed that SrHAp interfered with the RANKL/TRAF6/NF-κB signaling pathway, leading to the downregulation of the master transcription factor NFATc1. In conclusion, SrHAp nanoparticles exhibit dual functions, as they promote osteoblastic mineralization while concurrently arresting osteoclast differentiation. This balanced regulation highlights the potential of SrHAp as a bioactive ceramic for the treatment of osteoporosis and for advanced bone regeneration.

Introduction

Bone tissue, a metabolically active and highly dynamic organ, is a complex composite material comprising primarily an extracellular matrix, an inorganic mineral phase, and diverse cell populations [1]. This remarkable tissue plays critical physiological roles, providing essential skeletal support, safeguarding the internal viscera, facilitating movement, and acting as a vital reservoir for essential minerals such as calcium and phosphate, which are crucial for systemic homeostasis [2]. The bone extracellular matrix itself is composed primarily of type I collagen (approximately 90% of the organic matrix), alongside various noncollagenous proteins, proteoglycans, and growth factors such as osteonectin, osteopontin, bone sialoprotein, and bone morphogenetic proteins that regulate cellular behavior and mineralization [3]. The principal cell types within bone are osteoblasts, which mediate bone matrix synthesis; osteoclasts, which are responsible for bone resorption; osteocytes, which are involved in mechanosensing and matrix maintenance; and bone lining cells, which regulate mineral flux and osteoblast activity. Dynamic equilibrium between osteoblast-mediated bone formation and osteoclast-mediated bone resorption is essential for maintaining skeletal homeostasis; dysregulation of this balance can result in pathological conditions such as osteoporosis [4].

Osteoporosis is a systemic skeletal disease characterized by a reduced bone mineral density and the deterioration of the bone microarchitecture, consequently increasing fracture risk [5]. This condition tends to occur in postmenopausal women, affecting approximately one-third of women over 50 years of age, compared with approximately one-fifth of age-matched men [6]. This increasing prevalence imposes substantial healthcare burdens, increasing medical expenditures related to treatment and long-term management. The underlying pathophysiology of osteoporosis is largely attributed to an imbalance in the functional activities of osteoblasts and osteoclasts [7]. Osteoblasts are the primary effectors of bone matrix deposition, whereas osteoclasts are the principal cells that mediate bone matrix degradation. A sustained imbalance in which osteoclast-mediated bone resorption surpasses osteoblast-mediated bone formation leads to progressive decrease in bone density and mass, ultimately compromising skeletal integrity and increasing the likelihood of fracture.

As a treatment for osteoporosis, strontium ranelate directly promotes the proliferation and differentiation of osteoblasts while concurrently suppressing osteoclast activity. This action effectively tips the balance in favor of bone formation [8]. The mechanism by which strontium exerts its dual effects is multifaceted and involves the activation of calcium-sensing receptors on mesenchymal stem cells, modulation of the Wnt/β-catenin signaling pathway [9], and induction of osteoprotegerin expression, which in turn inhibits osteoclastogenesis [10]. Strontium ions are essential for bone health, as they promote osteoblast differentiation by activating Cbfa1 signaling, a crucial regulator of bone formation [11]. Lin et al. demonstrated that ionic extracts of Sr-substituted calcium silicate induced bone marrow mesenchymal stem cell differentiation through the ERK and p38 signaling pathways [12]. Bonnelye et al. showed that Sr promotes osteoclast apoptosis, inhibits osteoclast proliferation and differentiation, and reduces bone resorption [13]. Due to its similarity to Ca, Sr acts as a CaSR agonist, likely influencing osteoclast apoptosis [14]. Sr also regulates inflammation, enhances BMSC osteogenesis, and suppresses the differentiation of RAW264.7 macrophages [15]. Zeng et al. found that an extract of strontium-substituted calcium phosphate silicate bioactive ceramic promotes osteogenesis by upregulating Wnt/β-catenin signaling and inhibiting osteoclastogenesis through the downregulation of the NF-κB signaling pathway [16].

The inorganic component of bone tissue, bioapatite, is similar to stoichiometric hydroxyapatite (HAp) and has the chemical formula Ca₁₀(PO₄)₆(OH)₂; it imparts rigidity and compressive strength to bone [17]. Synthetic HAp is extensively employed in bone grafting applications due to its favorable biocompatibility and osteoconductive properties [18]. Nevertheless, pure HAp has low osteoinductive properties and slow degradation rates, which can hinder osseointegration. Various strategic modifications involving the substitution of specific ions into its crystal lattice have been explored to address these shortcomings and improve the biological performance of HAp. Ions, such as strontium, magnesium, and zinc, have been incorporated to partially replace calcium within the HAp structure. Each of these substitutions aims to confer specific biological advantages, such as enhanced degradation kinetics, improved antibacterial properties, or direct cellular stimulation [19, 20]. Strontium ions are incorporated into the HAp lattice to increase its bioactivity and therapeutic potential, resulting in the formation of strontium-substituted hydroxyapatite (SrHAp). The substitution of strontium ions within the HAp crystal lattice alters its physicochemical characteristics, which may contribute to improved biological performance [21, 22]. The increased solubility of SrHAp generates a more bioactive material, supporting faster integration and resorption within the body [23]. Ge et al. reported increased alkaline phosphatase (ALP) activity in MC3T3-E1 cells grown on porous poly(L-lactic acid) scaffolds with SrHAp compared with those cultured on poly(L-lactic acid) alone [24]. Ren et al. demonstrated that SrHAp facilitates the osteogenic differentiation of bone marrow mesenchymal stem cells via the CaSR–JAK2/STAT3 signaling pathway [25].

In our previous study, we successfully fabricated strontium-substituted hydroxyapatite (SrHAp) nanofibers and identified that a 30 mol% substitution ratio is critical for maintaining high phase purity and minimizing secondary impurities, such as CaO or CaCO₃, in the resulting material [26]. While we subsequently showed that these 30 mol% SrHAp nanofibers significantly increase the expression of key osteogenic markers like RUNX2 and OCN in MG-63 cells compared with pure hydroxyapatite [27], this study was primarily focused on the combined effects of the chemical composition and fibrous surface topography on osteoblasts alone. In the present study, we utilized the same optimized 30 mol% formulation but omitted the electrospinning process to synthesize SrHAp in a nanoparticle form. This strategic shift allows us to isolate the effects of strontium substitution on biochemical signaling mediated by the physical topographic cues of a fibrous matrix, ensuring that the biological responses are driven specifically by the high-substitution bioceramic itself.

Panzavolta et al. reported that compared with HAp-containing scaffolds, the viability and activity of osteoclasts cocultured with osteoblasts are negatively affected in SrHAp-containing scaffolds [28]. Capuccini et al. observed that osteoclasts cultured on HAp display a characteristic ruffled border and polymorphic morphology, indicating active motility and bone resorption. In contrast, osteoclasts cultured on SrHAp exhibit fewer filopodia and ruffled borders, suggesting that SrHAp inhibits their bone-resorbing activity [29]. However, the molecular mechanisms underlying the effects of SrHAp on osteoclastogenesis remain unexplored. By transitioning from material fabrication to an in-depth exploration of the bone remodeling cycle using a dual-cell model, this study represents the first systematic evaluation of how 30 mol% SrHAp nanoparticles concurrently promote osteoblast mineralization and arrest RANKL-induced osteoclast activity. This study aimed to investigate the effects of SrHAp on osteoblast and osteoclast activities, focusing on cell viability, differentiation, and the molecular mechanisms involved, specifically the RANKL/TRAF6/NF-κB/NFATc1 signaling axis, to provide insights for osteoporosis treatment and the development of effective bone substitutes.

Materials and methods

Reagents

Calcium nitrate tetrahydrate, poly(ethylene glycol)-block-poly(propylene glycol)-block-poly(ethylene glycol) (Pluronic P123; MW = 5800) and polyvinylpyrrolidone (PVP; MW = 40,000) were purchased from Sigma-Aldrich Chemical Company (St. Louis, MO, USA). Triethyl phosphite (TEP) was purchased from Merck (Darmstadt, Germany). Unless specified otherwise, all other chemicals used were of reagent grade.

Synthesis of strontium-substituted hydroxyapatite (SrHAp)

SrHAp was synthesized as described in our previous study [30]. Briefly, 0.5 g of cetyltrimethylammonium bromide (CTAB) and 3.086 mL of TEP were mixed in 5 mL of a 50% v/v ethanol–water mixture and stirred continuously until a clear solution was obtained. Additionally, calcium nitrate was dissolved in 95% ethanol, and strontium nitrate was dissolved in deionized water at room temperature. The strontium nitrate and calcium nitrate solutions were subsequently added dropwise to the TEP/CTAB solution to prepare the precursor solution. The Sr/(Ca+Sr) molar ratio was 30 mol%. The precursor solution was directly calcined at 800 °C under a nitrogen atmosphere to form the strontium-substituted hydroxyapatite powder.

Characterization of SrHAp

The crystalline structure of SrHAp was analyzed using X-ray diffraction (XRD) with a Bruker D2 Phaser diffractometer with Cu-Kα1 radiation (λ = 1.5406 Å). The XRD patterns were recorded over the 2θ range of 20° to 60° with a step size of 0.04°. The hydrodynamic diameter and zeta potential of the SrHAp powders were measured using the dynamic light scattering (DLS) technique with a Zetasizer Nano-ZS90 (Malvern Instruments).

Cell culture

Osteoblast-like HOS cells (BCRC 60308) and the murine macrophage RAW 264.7 cell line (BCRC 60001) were purchased from the Bioresource Collection and Research Center (BCRC, Hsinchu, Taiwan). Both cell lines were maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% (v/v) penicillin/streptomycin in a humidified incubator at 37 °C with 5% CO₂. Upon reaching approximately 90% confluence, the HOS cells were passaged by trypsinization, whereas the RAW 264.7 cells were detached by gentle scraping.

Cytotoxicity assessment

HOS cells and RAW264.7 cells were seeded in 96-well plates at a density of 5000 cells per well with 100 μL of medium and cultured at 37 °C with 5% CO₂. After the cells were cultured overnight, the medium was discarded, and SrHAp was added at various concentrations (0, 100, 250, 500, and 1000 μg/mL). Following a 24-h incubation period, cell viability was evaluated using the MTT assay. The optical density (OD) was measured at 570 nm using a microplate reader (Take3, BioTek). The viability and cytotoxicity of the nonchallenged cells were considered 100%.

Measurement of intracellular ROS Levels

HOS cells were seeded at a density of 2.5 × 10⁴ cells/well in 96-well plates with 100 μL of culture medium and incubated overnight at 37 °C in a humidified atmosphere containing 5% CO₂. Following the incubation, the culture medium was replaced with fresh medium supplemented with SrHAp (100 μg/mL). After a 24-h incubation period, intracellular reactive oxygen species (ROS) levels were quantified using a DCFDA/H2DCFDA Cellular ROS Assay Kit (Ab113851; Abcam, UA) according to the manufacturer’s protocol. According to the manufacturer’s instructions, the cells were washed with buffer solution, after which the DCFDA solution was added and incubated with the cells for 45 min at 37 °C in the dark. The fluorescence intensity was measured using a plate reader at excitation and emission wavelengths of 485 nm and 535 nm, respectively.

Reverse transcription polymerase chain reaction (RT-PCR) to analyze gene expression

RT‒PCR was performed to determine the expression of the RUNX2, Osteocalcin (OCN), and GAPDH mRNAs. HOS cells were seeded at a density of 1 × 10⁶ cells per well in 6-well plates. After culture for 14 and 21 days, total RNA was extracted from the samples using Tri-isolation reagent according to the manufacturer’s instructions. Briefly, 1 mL of Tri-isolation reagent was added and incubated for 5 min at room temperature. The cells were scraped, and the solution was transferred to an Eppendorf tube. After 200 μL of chloroform was added and incubated for 5 min, the mixture was centrifuged at 12,000 rpm for 15 min at 4 °C, and the upper aqueous phase that contained the RNA was collected. Two hundred microliters of the aqueous phase was transferred to a new tube, followed by the addition of 200 μL of isopropanol, an incubation for 15 min, and centrifugation at 12,000 × g for 10 min at 4 °C to precipitate the RNA. One milliliter of 75% ethanol was added, and the centrifugation process was repeated twice to remove the residual salts. The RNA pellet was air-dried for 10 min, resuspended in 20 μL of DEPC-treated H₂O, and stored at −80 °C. The RNA concentration and purity were determined using a Nano 300 instrument, and the O.D. 260/280 ratio was measured before the use of the RNA in subsequent experiments. The RNA solution was adjusted to an appropriate concentration for reverse transcription using the HiScript-I First Strand cDNA Synthesis Kit to convert RNA to cDNA. PCR amplification was performed by mixing the Master Mix, cDNA, sense and antisense primers, and ddH₂O and placing the mixture in a PCR machine. Table 1 shows the sequences of the oligonucleotides that were used as PCR primers. Each PCR product was analyzed by electrophoretic separation on a 1.5% agarose gel, and the bands were stained with ethidium bromide for visualization.

Table 1.

Oligonucleotide primers for RT-PCR amplification

Gene		Primer sequence (5’ -3’)	Annealing Temp (°C)	Cycles
GAPDH	Sense	GCTCTCCAGAACATCATCCCTGCC	59	30
	Antisense	CGTTGTCATACCAGGAAATGAGCTT
RUNX2	Sense	ATGCTTCATTCGCCTCACAAAC	58	35
	Antisense	AGTCCCTCCTTTTTTTTTCAG
OCN	Sense	CAGTCTAACCACCTTGTTGCAG	59	35
	Antisense	CATCAGGTCCATCCTCATACCT

Quantification of mineralization using Alizarin Red S staining

HOS cells were seeded at a density of 1 × 10⁴ cells/well in 48-well plates and incubated overnight at 37 °C with 5% CO₂. The culture medium was replaced with fresh medium supplemented with SrHAp (100 μg/mL) and this process was repeated every three days. On days 14 and 21, the medium was removed, and the cells were washed with PBS. They were fixed with 3.7% formaldehyde (diluted in PBS) for 15 min, washed with distilled water, and stained with 2% Alizarin Red S (pH 4.1–4.3) for 30 min. After the samples were washed and air dried, ARS staining was quantified using the method described by Gregory et al. [31] by adding 800 μL of 10% acetic acid, shaking the plate for 30 min, scraping the monolayer into an Eppendorf tube, heating the sample at 85 °C for 10 min, centrifuging the sample at 12,000 rpm for 15 min, and measuring the absorbance of the supernatant mixed with 200 μL of 10% ammonia at 405 nm.

RANKL-mediated differentiation of RAW 264.7 cells into osteoclasts

RAW 264.7 cells were seeded at a density of 1 × 10⁴ cells/cm² in cell culture dishes and cultured with DMEM for 24 h. Following the initial incubation, the culture medium was replaced with α-Minimum Essential Medium (α-MEM) supplemented with 10% FBS, 1% penicillin/streptomycin, 1.5 g/L sodium bicarbonate, and 30 ng/mL recombinant murine RANKL. The cells were then cultured for three days, after which the formation of multinucleated osteoclasts was assessed.

TRAP staining to identify osteoclasts

Tartrate-resistant acid phosphatase (TRAP) staining was performed to identify differentiated osteoclasts. RAW 264.7 cells were treated with α-MEM containing RANKL in the presence or absence of SrHAp (500 μg/mL) for three days and then subjected to TRAP staining using an acid phosphatase, leukocyte (TRAP) kit (Sigma-Aldrich, 387A). Briefly, the cells were washed with 200 μL of phosphate-buffered saline (PBS) and fixed with 50 μL of fixative for 60 s. Following three washes with water, 50 μL of staining solution was added, and the cells were incubated at 37 °C in the dark for 60 min. After three washes, the cells were observed under a light microscope. Osteoclasts were identified as multinucleated (≥3 nuclei) cells that exhibited positive TRAP staining.

Determination of osteoclast differentiation using a TRAP activity assay

TRAP activity was quantified using a TRACP & ALP assay kit (TAKARA Biomedical, MK301) on day three of culture. Cells cultured in 6-well plates were washed with 1 mL of PBS and lysed with 200 μL of radioimmunoprecipitation (RIPA) buffer (Sigma, R0278) on ice for 20 min, followed by sonication. Cellular debris was removed by centrifugation at 13,000 × g for 15 min at 4 °C. The protein concentration in the supernatant was determined using a BCA assay, with the absorbance measured at 592 nm. For the enzymatic assay, 10 μg of total protein was mixed with 50 μL of reaction buffer and incubated at 37 °C in the dark for 15 min. The reaction was terminated by the addition of 50 μL of 0.5 N NaOH, and the absorbance was measured at 405 nm using a microplate reader.

Western blotting to analyze protein expression

Cells were lysed with 200 μL of RIPA buffer on ice for 20 min, followed by sonication in cycles of 10 s on and 10 s off on ice for a total of 5 min. Cellular debris was removed by centrifugation at 13,000 × g at 4 °C for 15 min. The protein concentration in the supernatant was determined using a BCA assay. Prior to gel electrophoresis, the proteins were denatured by mixing the samples with 5× loading buffer at a 4:1 ratio and heating at 95 °C for 10 min. Electrophoresis was performed at 120 V for 55 min. The separated proteins were then transferred onto polyvinylidene difluoride (PVDF) membranes at 145 mA for 40 min, with the transfer conditions optimized based on the molecular weight of the target protein. The expression levels of the TRAF6, NF-κB, TRAP and NFATc1 were detected. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a loading control for normalization. PVDF membranes were blocked with blocking buffer (F1 western, F1BLOK500P) to block nonspecific binding. The membranes were then washed with TBS-T for 1 min and incubated with rabbit anti-mouse primary antibodies overnight at 4 °C. After three washes with TBS-T, the membranes were incubated with goat anti-rabbit secondary antibodies at room temperature for 1 h, followed by three washes with TBS-T. The protein bands were visualized using a SuperLight Chemiluminescent HRP Kit and detected with a UPV ChemStudio Plus imaging system. The band intensity was quantified using ImageJ software.

Statistical analysis

Quantitative data are presented as the means±standard deviations. Statistical analyses were performed using SPSS v.10. Comparisons involving multiple groups were performed using the Kruskal‒Wallis test with Dunn’s post hoc test. p values < 0.05 were considered to indicate statistically significant differences.

Results

Characterization of SrHAp

The synthesized SrHAp was characterized using X-ray diffraction (XRD) to confirm its crystal structure. The phase composition of the material was analyzed by interpreting the diffraction patterns using DIFFRAC.SUITE EVA software (Bruker v.4.2) and comparing the results with reference PDF card #34-0480 (strontium-substituted hydroxyapatite). The XRD patterns showed distinct peaks corresponding to strontium-substituted hydroxyapatite. While minor peaks corresponding to calcium oxide (CaO) were observed (Fig. 1), the predominant phase remained SrHAp, indicating the successful incorporation of strontium ions into the lattice with high phase stability. The Sr/Ca and (Ca+Sr)/P ratios of the SrHAp from the ICP‒OES analysis were 0.417 and 1.73, respectively. SrHAp was characterized using a nanoparticle size and zeta potential analyzer. Zeta potential measurements revealed a surface charge of −14.9 mV, and the dynamic light scattering (DLS) analysis indicated an average particle size of 712.6 nm for the SrHAp powder.

Fig. 1.

X-ray diffraction (XRD) patterns of SrHAp. The blue circles correspond to the strontium-substituted hydroxyapatite phase (Ca_10-xSr_x(PO₄)₆(OH)₂), while the red triangles represent calcium oxide (CaO)

Effects of SrHAp on cell viability

Cell viability was assessed to determine the non-cytotoxic concentration range that was not cytotoxic to both cell lines. Based on the ISO 10993-5 standard, a viability >70% is considered noncytotoxic. HOS cell viability decreased significantly below 70% in a dose-dependent manner when the SrHAp concentration exceeded 100 μg/mL (Fig. 2A). In contrast, RAW 264.7 cells maintained their viability well above 100% across all the tested concentrations up to 1000 μg/mL, indicating not only a markedly higher tolerance but also potentially a stimulatory effect on macrophage-like cell proliferation (Fig. 2B). Consequently, SrHAp concentrations of 100 μg/mL for HOS cells and 500 μg/mL for RAW 264.7 cells were selected for subsequent experiments designed to assess the regulatory effects of SrHAp, as these dosages represented the maximum noncytotoxic levels.

Fig. 2.

Viability of A HOS cells and B RAW264.7 cells after treatment with various concentrations of SrHAp. The dashed red line represents the 70% cell viability threshold for noncytotoxicity as defined by ISO 10993-5

Regulation of reactive oxygen species (ROS) by SrHAp

To evaluate the impact of SrHAp on the redox state of the osteoblastic environment, intracellular ROS levels were measured in HOS cells using H₂O₂ as a positive control. As shown in Fig. 3, HOS cells treated with 100 μg/mL SrHAp alone exhibited a significant reduction in ROS levels compared with the untreated control, indicating that SrHAp effectively suppresses basal ROS production. Furthermore, the induction of oxidative stress by H₂O₂ resulted in a substantial increase in ROS levels (approximately 300% of the control). However, cotreatment with SrHAp and H₂O₂ markedly attenuated this effect, yielding significantly lower ROS concentrations compared to the H₂O₂ alone group. These results demonstrate that the strontium-substituted lattice possesses ROS-scavenging activity, allowing it to mitigate both endogenous and exogenous induced oxidative stresses within the osteoblastic microenvironment.

Fig. 3.

Intracellular reactive oxygen species (ROS) levels in HOS cells following 1 h of treatment with SrHAp, H₂O₂, or a combination of H₂O₂ and SrHAp are reported as a percentage relative to those in the control group. The data are presented as the means ± SDs; n = 3. Different letters represent significant differences at p < 0.05

Qualitative and quantitative evaluation of HOS cell mineralization

Mineralization, a critical aspect of osteoblast function, was evaluated by performing Alizarin Red S (ARS) staining, a histochemical method that selectively stains calcium deposits within the extracellular matrix. The qualitative assessment of ARS-stained HOS cell cultures revealed a marked increase in mineralization in the SrHAp-treated groups at both 14 and 21 days (Fig. 4A, B). This increased mineralization was evidenced by the presence of extensive, intensely red-stained areas, indicating significant calcium nodule formation. Furthermore, a temporal increase in both the intensity and spatial distribution of ARS staining was observed in the SrHAp-treated cultures between days 14 and 21, signifying progressive mineral accrual. Conversely, control cultures exhibited only minimal ARS staining and were characterized by sparse and faintly stained punctate deposits. The quantitative analysis of the extracted ARS dye corroborated these qualitative observations, indicating a statistically significant increase (p < 0.05) in mineral deposition in the SrHAp-treated groups compared with the untreated controls at both time points (Fig. 4C). Collectively, these findings demonstrate that SrHAp treatment effectively promotes the mineralization of HOS cells in vitro.

Fig. 4.

Alizarin Red S (ARS) staining of HOS cells after SrHAp treatment, showing calcium deposition at (A) 14 days and (B) 21 days. C Quantitative analysis of ARS staining by measuring the absorbance at 405 nm, confirming increased mineralization with SrHAp treatment for 14 and 21 days. The data are presented as the means ± SDs; n = 3. The asterisk (*) denotes a significant difference (p < 0.05). The scale bars in (A) and (B) represent 2 mm

Expression of the RUNX2 and OCN osteogenic differentiation markers

We examined the expression of key osteogenic markers to better understand the effect of SrHAp on gene expression during the differentiation of HOS cells. After 14 days of SrHAp treatment, the expression level of Runt-related transcription factor 2 (RUNX2) in HOS cells was significantly increased compared with that in the untreated control group (p < 0.05) (Fig. 5A, B). This increase suggests that SrHAp promotes RUNX2 expression, thereby supporting the initiation of osteogenic differentiation. Compared with that in the control group, the expression of osteocalcin (OCN) was markedly increased after 21 days of SrHAp treatment (p < 0.05) (Fig. 5A, C). Overall, these findings indicate that SrHAp regulates the expression of osteogenic genes, supporting the progression of HOS cell differentiation from early commitment to late-stage maturation.

Fig. 5.

RT-PCR analyses of the expression of bone-associated genes in HOS cells after SrHAp treatment for 21 days. Images of the gels of osteogenic differentiation markers, RUNX2 and osteocalcin (OCN), are shown in (A), and the expression of B RUNX2 and C OCN was normalized to that of the corresponding control GAPDH. The data are presented as the means ± SDs; n = 3. The asterisk (*) denotes a significant difference (p < 0.05)

Morphological assessment of osteoclast differentiation via TRAP staining

The inhibitory effect of SrHAp on RANKL-induced osteoclastogenesis was evaluated using tartrate-resistant acid phosphatase (TRAP) staining. Microscopic evaluation revealed that the negative control group (Control) consisted primarily of mononuclear macrophages lacking TRAP expression, appearing yellowish-brown (Fig. 6A). In the positive control group (RANKL), treatment with 30 ng/mL RANKL induced the formation of numerous large, multinucleated cells, with characteristic purplish-red TRAP staining, confirming successful osteoclast differentiation (Fig. 6A). Compared with the RANKL-only group, the experimental group treated with RANKL + SrHAp exhibited a marked reduction in the number and size of these TRAP-positive multinucleated cells (Fig. 6A). Quantitative analysis of relative TRAP activity corroborated these morphological findings, showing that SrHAp significantly attenuated the RANKL-induced increase in enzyme activity (p < 0.05) (Fig. 6B). These results indicate that SrHAp effectively suppresses osteoclast differentiation and maturation.

Fig. 6.

A Optical micrographs illustrating the effect of SrHAp on the differentiation of RAW 264.7 cells into osteoclasts. Cells were treated with RANKL or RANKL+SrHAp and stained for TRAP on day 3. Top row: Low-magnification views (scale bar: 400 μm). Bottom row: High-magnification views (scale bar: 200 μm). TRAP-negative, undifferentiated RAW 264.7 cells appear yellowish-brown and are mononuclear. TRAP-positive cells, which appear as purplish red, indicate preosteoclasts (mononuclear) and mature osteoclasts (multinucleated). B Quantification of relative TRAP activity. The data are presented as the means ± SDs; n = 3. Different letters represent significant differences at p < 0.05

Mechanism of SrHAp-mediated inhibition of osteoclast differentiation

The mechanism of SrHAp inhibition of osteoclast differentiation was assessed through the protein expression time course of TRAF6 and NF-κB. RANKL treatment significantly upregulated TRAF6 and NF-κB expression in RAW 264.7 cells (Fig. 7). While early expression levels at 10 and 20 min were comparable between groups, significant reductions in TRAF6 and NF-κB expression levels were observed at 30 and 60 min in the RANKL+SrHAp group (Fig. 7).

Fig. 7.

Effect of SrHAp on RANKL-induced TRAF6 and NF-κB expression in RAW264.7 cells. RAW264.7 cells were treated with RANKL alone (A, D) or RANKL combined with SrHAp (B, E) for the indicated durations (10, 20, 30, and 60 min). Western blot analysis was conducted to evaluate the expression levels of TRAF6 (A, B) and NF-κB (D, E), with GAPDH serving as the loading control. The results of the quantitative analysis of the protein expression levels normalized to those of GAPDH and relative to those of the RANKL group at 10 min are shown for TRAF6 (C) and NF-κB (F). The data are presented as the means ± standard deviations. The asterisk (*) denotes a significant difference (p < 0.05)

Downstream protein expression analysis revealed that NFATc1 levels significantly increased after two days of RANKL treatment but were decreased in the RANKL+SrHAp group, approaching control levels (Fig. 8A). Similarly, TRAP protein expression was significantly upregulated after three days of RANKL treatment (Fig. 8B). However, cotreatment with RANKL+SrHAp significantly decreased TRAP expression levels, which approached control levels (Fig. 8B). These findings demonstrate that SrHAp inhibits the RANKL-induced signaling pathway at multiple stages.

Fig. 8.

Western blot analysis of NFATc1 and TRAP protein expression in RAW264.7 cells. A NFATc1 protein expression after 2 days of treatment. C TRAP protein expression after 3 days of treatment. B, D Densitometric quantification of NFATc1 and TRAP protein expression, respectively, normalized to that of GAPDH and presented as the fold change relative to the control group. The data are shown as the means ± standard deviations. Different letters represent significant differences at p < 0.05. In Western blot panels (A) and (C), Lane (1) represents the control group. Lane (2) shows RAW264.7 cells treated with RANKL. Lane (3) indicates RAW264.7 cells treated with RANKL+SrHAp. GAPDH served as an internal loading control

Discussion

The primary objective of this study was to evaluate the biological effects of strontium-substituted hydroxyapatite particles with a high substitution level of 30%. While previous research has established the influence of strontium-containing compounds on bone metabolism, most studies have focused on lower substitution levels (typically <10%) and utilized terminal enzymatic markers, such as TRAP, activity for evaluation [32–34]. This study expanded upon these observations by characterizing 30% SrHAp particles and elucidating their role in modulating the RANKL-induced signaling cascade and the local redox environment.

Reactive oxygen species (ROS) are byproducts of cellular metabolism and play complex roles in biological systems [35]. While physiological concentrations of ROS are essential for cellular signaling and cellular homeostasis, their excessive production leads to oxidative stress, damaging vital biomolecules, such as DNA, proteins, and lipids [36, 37]. Increased levels of ROS in MSCs have been reported to promote lipogenic differentiation while inhibiting osteogenic differentiation by disrupting osteogenic signaling pathways [38]. Additionally, ROS act as second messengers in the RANKL/RANK signaling pathway, thereby facilitating the differentiation and maturation of osteoclasts [39, 40]. Therefore, modulating ROS levels is critical in the design of biomaterials for bone regeneration. The present results demonstrated that SrHAp plays a protective role by significantly attenuating both basal and induced ROS (Fig. 3). Liu et al. reported that strontium-containing materials, such as bioactive glasses (BGs), function as biomaterials with modulated natural enzymes that can increase the activities of SOD, CAT, and GPx. Such strontium-containing biomaterials exert a protective effect on ROS in the body, decreasing the inflammatory response induced by ROS and playing an important role in the repair and regeneration of bone [41]. Qi et al. reported that strontium-containing compounds, such as strontium fructose 1,6-diphosphate (FDP-Sr), protect osteoblastic cells from oxidative damage through direct ROS scavenging and the activation of Wnt/β-catenin signaling. Under oxidative stress, the Wnt/β-catenin pathway is typically repressed, as β-catenin is diverted to the FoxO pathway, inhibiting the transcription of bone differentiation genes, such as Runx2 and ALP [42].

SrHAp treatment enhances mineralized nodule formation by modulating key osteogenic genes. As a pivotal transcription factor, RUNX2 initiates differentiation and regulates downstream matrix proteins, such as OCN. Inducing RUNX2 expression is a recognized mechanism for promoting matrix mineralization and bone homeostasis [43]. The subsequent upregulation of OCN, which occurs before the onset of mineralization, confirms the progression into late-stage maturation [44]. OCN facilitates hydroxyapatite crystal nucleation and growth, indicating that SrHAp creates a microenvironment conducive to phenotypic maturation and controlled extracellular matrix mineralization.

Tartrate-resistant acid phosphatase (TRAP) is a protein that is abundantly expressed during osteoclast differentiation. TRAP-positive staining was observed in a subset of mononuclear cells within the RANKL-treated group (Fig. 6A), reflecting the stages of differentiation involving cytoskeletal remodeling and the upregulation of fusion-related protein expression prior to complete cell fusion. The significant reduction in TRAP-positive multinucleated cells indicates that SrHAp interferes with critical stages of osteoclastogenesis. Osteoclast differentiation requires the fusion of mononuclear precursors into mature cells capable of bone resorption, a process regulated by complex cytokine networks [45]. The observed decrease in both the number and size of TRAP-positive cells suggests that SrHAp may specifically impede these late-stage fusion processes or the signaling pathways required for precursor maturation [46]. By effectively attenuating TRAP activity, SrHAp modulates bone remodeling, potentially balancing osteoblastic formation and osteoclastic resorption.

A critical signaling cascade involving TRAF6, NF-κB, NFATc1, and TRAP orchestrates the RANKL-induced differentiation of RAW264.7 cells into osteoclasts [47, 48]. RANKL binds to its receptor RANK, recruiting TRAF6 and activating the IKK complex, which subsequently phosphorylates IκB for proteasomal degradation, leading to the release of NF-κB. This liberated NF-κB translocates to the nucleus to induce the expression of key transcription factors, including NFATc1 [49–51]. Concurrently, RANKL signaling increases intracellular calcium levels, activating calcineurin activity and further promoting NFATc1 activation [52–54]. This study indicates that SrHAp interferes with the sustained activation of these early signaling molecules rather than their initial recruitment, a critical finding that explains the downstream suppression of osteoclast maturation.

The biological performance of SrHAp is linked to its crystallographic characteristics; incorporating Sr ions (1.12 Å) into the HA lattice increases lattice parameters and chemical solubility compared to Ca ions (0.99 Å). This creates a strontium-rich microenvironment that facilitates molecular intervention in the RANKL/TRAF6/NF-κB pathway. By simultaneously increasing osteoblast activity and suppressing RANKL-induced osteoclastogenesis, SrHAp maintains bone homeostasis, supporting its potential for bone regeneration and osteoporosis treatment.

Conclusions

This study shows that strontium-substituted hydroxyapatite (SrHAp) exerts a dual regulatory effect on bone remodeling by promoting osteoblast differentiation and mineralization while inhibiting osteoclast activity. Our findings indicate that SrHAp increases the deposition of mineralized matrix by osteoblasts, as evidenced by increased Alizarin Red S staining and the upregulation of key osteogenic markers such as RUNX2 and osteocalcin. Concurrently, SrHAp suppresses osteoclastogenesis in RAW 264.7 cells by interfering with the RANKL signaling pathway, specifically through the downregulation of TRAF6, NF-κB, and NFATc1. The ability of SrHAp to coordinate bone formation and resorption highlights its potential for treating osteoporosis and other bone loss conditions. By shifting the cellular balance toward bone anabolism and modulating key signaling pathways, SrHAp represents a promising strategy to enhance bone regeneration. Future in vivo studies will be essential to validate these findings and explore the clinical translation of SrHAp in regenerative therapies.

Abbreviations

BCA

bicinchoninic acid

CTRL

negative control

DMEM

Dulbecco’s modified Eagle’s medium

FBS

fetal bovine serum

GAPDH

glyceraldehyde-3-phosphate dehydrogenase

HRP

horseradish peroxidase

MTT

3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide

NFATc1

nuclear factor of activated T-cells, cytoplasmic 1

NF-κB

nuclear factor kappa-B

RANKL

receptor activator of nuclear factor kappa-B ligand

RIPA

radioimmunoprecipitation

ROS

reactive oxygen species

SrHAp

strontium-substituted hydroxyapatite

TRAF6

TNF receptor-associated factor 6

TRAP

tartrate-resistant acid phosphatase

Author contributions

Shiao-Wen Tsai: Conceptualization, Data curation, and Writing-original draft. Cheng Li: Data curation, Formal analysis, and Methodology. Shu-ting Lee: Data curation, Formal analysis, and Methodology. Pai-An Hwang: Conceptualization, Methodology, and Writing-review & editing. Fu-Yin Hsu: Conceptualization, Funding acquisition, Project administration and Writing-review & editing. I certify that the above information is true and correct. All the authors contributed to the study and the manuscript. If the manuscript is accepted for publication, I agree to transfer all copyright ownership of the manuscript to the Journal of Materials Science: Materials in Medicine, which covers the rights to use, reproduce, or distribute the article.

Funding

This research was funded by the Ministry of Science and Technology of Taiwan (Grant Nos. NSTC 111-2221-E-019-012 and NSTC 112-2221-E-019-011-MY2).

Data availability

All the data generated or analyzed during this study are included in the published article. The data that support the findings of this study are available upon reasonable request from Fu-Yin Hsu, the corresponding author.

Compliance with ethical standards

Conflict of interest

The authors declare no competing interests.

Ethics approval and consent to participate

Not applicable.

Consent for publication

All the authors approve of the content of this manuscript.