The influence of soluble epoxide hydrolase inhibition and their PUFA-derived epoxides in osteoblast bone metabolism: an in vitro study

Abstract

EpFAs are crucial mediators in resolving inflammation and regulating various biological processes. However, their activity is constrained by the rapid metabolism mediated by the soluble epoxide hydrolase (sEH), which converts EpFAs into inactive or even pro-inflammatory diols. Nevertheless, the specific effects of soluble epoxide hydrolase inhibition (sEHI) and EpFAs on osteogenic cell metabolism remain unclear. Cultures of the human immortalized osteoblast-like cell line (SAOS-2) were treated with varying concentrations (0.1 −10 μM) of the potent sEHI TPPU or EpFAs (epoxyeicosatrienoic acids [EETs], epoxydocosapentaenoic acids [EDPs], and epoxyeicosatetraenoic acids [EEQs], derived from arachidonic acid [ARA], eicosapentaenoic acid [EPA], and docosahexaenoic acid [DHA], respectively). Cellular metabolic activity and proliferation were evaluated. Osteogenic potential was assessed through alkaline phosphatase activity, mineral nodule formation, and the expression of osteogenic markers, including Runx-2, Osx, Col1, Bsp, Opg, Ocn, Opn, and sEH. Treatment with TPPU and EpFAs enhanced cellular metabolic activity during the first 48 hours without affecting proliferation. Alkaline phosphatase activity and mineral nodule formation assays revealed that TPPU significantly stimulated osteogenic differentiation, while EpFAs, particularly EETs, EEQs, and EDPs, promoted osteogenesis predominantly at later stages. Furthermore, TPPU modulated the expression of key osteogenic markers, enhancing differentiation. Notably, EDPs were found to disrupt the synergistic effects between sEHI and EpFAs during the mineralization process. These findings suggest that sEHI enhances mineralization and may facilitate tissue regeneration in vitro. The differential effects of EpFAs and their interplay with sEHI provide insights into potential therapeutic strategies for bone tissue engineering and regeneration.

Graphical Abstract

INTRODUCTION

Globally, there is a significant demand for tissue regeneration, particularly in addressing hard tissue deficiencies in the oral and maxillofacial regions. These deficiencies often arise from pathological conditions such as periodontal disease and oral cancers. Periodontitis, the sixth most prevalent disease worldwide, induces a hyperinflammatory state that results in the destruction of both soft and hard tissues, ultimately leading to tooth loss (Peres et al., 2019; Van Dyke & Sima, 2020). Similarly, malignant tumors in the oral region often necessitate surgical resection, which invariably causes substantial tissue loss and bone defects in the affected area (Muzaffar et al., 2021). Additionally, the rapidly growing aging population contributes significantly to the need for regenerative therapies. Currently, 8.5% of the global population is aged 65 or older, and by 2050, this figure is projected to reach 83.7 million in the United States alone (He et al., 2015; Ashammakhi et al., 2019). Furthermore, traumatic injuries, such as those resulting from motor vehicle accidents, falls, and other incidents, represent another critical driver of the demand for effective tissue regeneration strategies (Manodh et al., 2016).

Tissue engineering has emerged as a promising approach to meet this demand, utilizing a triad of cells, scaffolds, and signaling molecules to mimic the biological cascades of wound healing and the resolution phase of inflammation (Tavelli et al., 2023). Among these components, stem cells are extensively studied due to their remarkable ability to self-renew and differentiate into multiple cell types, offering advantages over autogenous grafts or somatic cells like fibroblasts and keratinocytes (Pagni et al., 2012; Shanbhag et al., 2019). Scaffolds play a pivotal role in guiding new bone formation by maintaining the biological space required for regeneration. Recent advances in scaffold technology have focused on personalized, three-dimensional multilayer scaffolds designed to enhance cell migration and differentiation (Yu et al., 2019; Latimer et al., 2021). Signaling molecules, including growth factors and pharmacological agents, further optimize osteogenesis by promoting cell adhesion, migration, differentiation, and mineralization.

Polyunsaturated fatty acids (PUFAs) are essential nutrients obtained through dietary intake. Key PUFAs, such as arachidonic acid (ARA), docosahexaenoic acid (DHA), and eicosapentaenoic acid (EPA), are metabolized by cytochrome P450 (CYP) enzymes into bioactive lipids known as epoxy fatty acids (EpFAs), which are critical regulators of inflammation (Frömel et al., 2023). EpFAs derived from ARA (epoxyeicosatrienoic acids, EETs), EPA (epoxyeicosatetraenoic acids, EEQs), and DHA (epoxydocosapentaenoic acids, EDPs) exhibit potent biological effects, particularly in anti-inflammatory and pro-resolving processes (Newman et al., 2005; Panezai & Van Dyke, 2022). However, these bioactive epoxides are rapidly degraded into inactive diols by the soluble epoxide hydrolase (sEH), often resulting in diminished biological activity or even promoting inflammatory responses (Wagner et al., 2017; Wagner et al., 2020; Hammock et al., 2020). Given the critical role of EpFAs in resolving inflammation, the pharmacological inhibition of sEH offers a promising avenue to enhance bone metabolism and optimize the healing process.

This study aims to evaluate the effects of pharmacologically inhibiting sEH in a human osteoblast cell line, focusing on its potential to promote tissue mineralization. Additionally, the study investigates the individual contributions of epoxy fatty acids—EETs, EEQs, and EDPs—to elucidate their specific roles in osteogenic processes. To the best of our knowledge, this research represents the first comprehensive investigation of the impact of EpFAs and sEH inhibition (sEHI) on osteoblast function in vitro.

2. METHODS

2.1. Cell Line and Cell Culture

SAOS-2 cells, a human osteoblastic lineage (isolated from the bone of an 11-year-old, White, female) acquired from the American Type Culture Collection (ATCC, Manassas, VA, USA), were thawed and concentrated in centrifuge tubes containing 5 mL of McCoy’s 5A (Sigma, St. Louis, MO, USA), and then centrifuged at 336 g for 3 min. The supernatant was discarded, and the cells were cultured in 25 cm² culture flasks (Sarstedt, Hildesheim, Germany) containing McCoy’s 5A (Sigma) supplemented with 15% fetal bovine serum (Nutricell Nutrientes Celulares, Campinas, SP, Brazil), 100 IU/mL of penicillin (Sigma) and 50 μg/mL of streptomycin (Sigma). The culture medium was changed every two days. The cells were kept at 37ºC during all the experimentation in a humidified atmosphere containing 5% CO₂ and 95% atmospheric air. Once cell confluence was reached, a solution with 1mM of ethylenediaminetetraacetic acid (EDTA) (Gibco/Invitrogen Life Technologies, Grand Island, NY) and 2.5 mg/ml of trypsin (Gibco) was used to detach the adherent cells. Subsequently, cells were seeded in 96-well polystyrene plates or 21.5 cm² petri dishes at 110 cell/mm² cell density.

To induce osteogenic differentiation, the cells were cultured in an osteogenic medium containing McCoy’s 5A (Sigma) supplemented with 10 % (v/v) fetal bovine serum (LGC Biotechnology), 100 U/mL penicillin and 100 μg/mL streptomycin (Sigma), 10 mM of β-glycerophosphate (Sigma) and 50 μg/ml of ascorbic acid (Gibco). During the culture period, cells were incubated at 37 °C in a humidified atmosphere of 5% CO₂ and 95% air, and the medium was changed every 3 days.

2.2. Drug Supplementation

All the drugs, including the sEH inhibitor 1-(1-propanoylpiperidin-4-yl)-3-[4-(trifluoromethoxy)phenyl]urea (TPPU), the mixture of EET regioisomers (5,6-EET, 8,9-EET, 11,12-EET, and 14,15-EET), the EDPs, and EEQs were synthesized at the Department of Entomology and Nematology, University of California-Davis (USA), as previously published (Rose et al., 2010; Morisseau et al., 2010). TPPU was weighed and dissolved in a glass container in polyethylene glycol 400 (PEG400; Sigma; St. Louis, MO, USA). The glass container was sealed with Parafilm M (Sigma) tape and submerged in a sonicator chamber until complete TPPU dissolution. The EET, EDP, and EEQ stock solution (10mM diluted in DMSO) was kept at −80^◦C. On the day of the experiment, the compounds were thawed and dissolved in cell culture media at the final concentration of 0, 1, or 10nM.

2.3. Cell Viability (MTT assay)

SAOS-2 was seeded in 96-well plates at 2×10⁵ cells/well density and incubated for 24 hours. Subsequently, cells were primed with different treatments (TPPU, EETs, EDPs, EDQs, or different combinations) and concentrations (ranging from 1 to 10 nM). Cell viability was assessed by 3-[4,5-dimethylthiazol-2-yl]− 2,5-diphenyl tetrazolium bromide (MTT; Sigma) assay after 24, 48, and 72 h. Then cells were incubated with 10% MTT (5 mg/ml) in a culture medium at 37^◦C for 4 h. The MTT solution was then aspirated from the well, and 100 μL of Dimethyl Sulfoxide (DMSO, Sigma) was added to solubilize the blue-purple crystals formed. The plates were placed on a plate shaker for 5 min, and 150 μL of this solution was transferred to a new 96-well plate. The optical density was read at 570–650 nm on the plate reader (Epoch; Bio-Tek, Winooski, VT). Cell viability was calculated as: (absorbance of wells with TPPU treatment/absorbance of control well) × 100%. All experiments were performed blindly in triplicate for each time point.

2.4. Cell Proliferation

The culture medium was removed at 24, 48, and 72 hours, and the wells were rinsed with PBS. Subsequently, a solution with 1 mM of EDTA (Gibco) and mg/mL of trypsin (Nutricell Nutrientes Celulares) was added by pipette. Cultures were incubated for 5 min at 37°C. After that, the cells were homogenized by gently pipetting up and down. Finally, an aliquot was utilized to count the number of cells on a hemocytometer (Hausser Scientific, Horsham, PA, USA). Cell proliferation data were expressed as cell number × 10⁴.

2.5. Alkaline Phosphatase Activity

Alkaline phosphatase (ALP) activity was estimated by the release of thymolphthalein from thymolphthalein monophosphate using a commercial kit (Labtest Diagnostica SA, Lagoa Santa, MG, Brazil). Briefly, the culture medium was removed, and the wells were washed three times with PBS (Gibco) at 37 ºC. Subsequently, they were filled with 2 ml of 0.1% sodium lauryl sulfate (Sigma) and left for 30 min at room temperature. Then, 50 μL of thymolphthalein monophosphate was blended with 0.5 mL of diethanolamine buffer, 0.3 mmol/mL, pH 10.1, and left for 2 min at 37ºC. After this period, 50 μL of the sample was taken from each well. This stood for 10 min at 37ºC, and then 2 mL of a solution of Na₂CO₃ 0.09 mmol/mL and NaOH 0.25 mmol/mL was added to allow color development. After 30 min, absorbance was measured at 590 nm, and ALP activity was calculated from a standard curve. Results were calculated, and data were expressed as ALP activity normalized by the total protein content (Pierce BCA Protein Assay Kit).

2.6. Alizarin Red Stain

The Alizarin Red stain (Sigma) assessed the mineralized matrix formation. The mineralization process was quantified on days 10, 14, and 21. For that, the cells were initially fixed in 1.33 M formalin at room temperature for two hours. Then, the cells were dehydrated through a graded series of alcohol and stained with 0.02 g/ml of Alizarin Red S (pH 4.2) for 10 min. Subsequently, the cells were thoroughly washed with deionized water to remove excess dye, and the microplates were left to dry. Once parched, 280 μL of 10% acetic acid was added to each well and placed in the microplate shaker for 30 minutes at room temperature. Thereupon, the contents of each microplate were transferred to polypropylene tubes and heated at 85ºC for 10 min, followed by cooling on ice for 5 min. After that, the tubes were centrifuged at 20,000 g for 15 min. Supernatants were collected, and 100 μL was transferred to 96-well plates, along with 40 μL of 10% ammonium hydroxide to neutralize the acid. The absorbance was measured using a spectrophotometer (Epoch; Bio-Tek, Winooski, VT) at a wavelength of 405 nm. The formation of the mineralized matrix was expressed as absorbance and was calibrated by those obtained from non-exposed cultures. The experiments were conducted independently two times with triplicate samples each time.

2.7. Reverse Transcriptase-Polymerase Chain reaction (RT-PCR)

In brief, real-time polymerase chain reactions were carried out using the SYBR Green System (Thermo Fisher Scientific, Waltham, MA, USA), on the 7500 Fast Real Time PCR System (Applied Biosystems, Foster City, CA). At 1, 3, and 5 days, the culture medium was discarded from the wells, and the cells were collected for RNA removal using the Illustra RNAspin Mini Kit (GE Healthcare, Milwaukee, WI, USA), according to the manufacturer’s specifications. Then, total RNA was quantified at different wavelengths (260, 280, 230, and 320 nm) in a spectrophotometer (GE Healthcare). Subsequently, a cDNA strand was made from 1 μg of total RNA. This procedure was carried out in the Veriti^® Thermal Cycler (Applied Biosystems) using the reverse transcriptase enzyme for a final 40 ng of cDNA, using the RevertAid First Strand cDNA Synthesis kit (Thermo Fisher Scientific). The reaction product was quantified with the Relative Quantification tool, using GAPDH as the reference gene. The data are presented as a fold-change of relative quantity using the 2^-ΔΔCt method. All the genes sequencing are summarized in Table 1.

Table 1.

PCR primers for genes encoding Runx2, Col1, Bsp, Opg, Opn, sEH, and GAPDH.

Gene Target	Forward Sequence	Reverse Sequence
Runx2	GAGGCGGTCAGAGAACAAAC	GGAAGTGGACGAGGCAAGAG
Col1	CCAGAAGAACTGGTACATCAGCAA	GGACATCAGGCGCAGGAA
Bsp	ACGATTTCCAGTTCAGGGCAG	GCCCAGTGTTGTAGCAGAAAG
Opg	CCCTGTGTGAGGAGGCATTCTTCA	CACTCTCTGCGTTTACTTTGGTGCCA
Opn	GCCGAGGTGATAGTGTGGTT	AACGGGGATGGCCTTHTATG
sEH	CTTCATACCAGCAAATCTC	CCTCCCGCTTCACAGAC
Gapdh	ACCCACTCCTCCACCTTTGA	TGTTGCTGTAGCCAAATTCGTT

2.8. Statistical Analysis

The data were initially tested for normality. Thereafter, data were analyzed by two-way ANOVA, followed by Tukey’s post hoc test for multiple comparisons, using GraphPad 10.2. All data are presented as mean ± SD. A P-value less than 0.05 was considered significant.

3. RESULTS

3.1. Metabolic activity and proliferation of SAOS-2 cultured with sEHI and or EpFAs.

We initially examined whether different concentrations of soluble epoxide hydrolase inhibitor, TPPU, and its derivative epoxides could affect the mitochondrial activity of human osteoblast cells. For that, MTT assay was performed at three periods (24, 48, and 72 hours) (Figure 1). Our results demonstrate that all treatments, in all dosages, significantly increased the cell viability at 24 hours (Figures 1A – D). These findings persist at 48 hours, where the dosages of 1 and 10nM of sEHI, EETs, EDPs, or EEQs sustain these higher values (Figures 1A – D). However, at 72 hours post-treatment, there is no change in any of the treatments compared to the vehicle.

Figure 1. Metabolic activity with MTT assay.

The effect of different concentrations of (A) sEHI (TPPU), (B) EETs, (C) EDPs, and (D) EEQs on SAOS-2 cell viability. Cell viability was calculated by MTT assay. Data are expressed as mean ± SD; Experiments were performed in triplicate and 2 independent experiments. At 24 hours, * means significantly differences between 0.1, 1, and 10nM vs vehicle. At 48 hours, * means significantly differences between 1, and 10nM vs vehicle.

Subsequently, the proliferation was assessed by counting the number of cells on a hemocytometer (Figure 2). Herein, no significant difference in the proliferation rate of osteoblast treated with sEHI, EETs, EDPs, or EEQs was found compared to the vehicle after 24, 48, and 72 hours (p < 0.05; Figures 2A – D).

Figure 2. Proliferation assay.

Human osteoblastic lineage cells (SAOS-2) were treated with different concentrations of (A) sEHI (TPPU), (B) EETs, (C) EDPs, and (D) EEQs for 24, 48, and 72 hours. The cell proliferation assay was performed by counting with a hemocytometer and expressed as the number of cells ×10^⁴. Data are presented as mean ± SD. The experiment was conducted in technical duplicate and with two biological replicates. n.s. means non-significant.

3.2. Functional test of alkaline phosphatase activity and mineralization.

Because differentiated SAOS-2 cells are capable of forming mineral nodules in vitro, herein, SAOS-2 were cultured with sEHI, and EpFAs-homologous and regioisomers (EETs, EDP, and EEQ), and alkaline phosphatase activity was estimated by the release of thymolphthalein from thymolphthalein monophosphate on days 5 and 10 (Figure 3A and B). Moreover, calcium deposition was assessed by the Alizarin red assay on days 10, 14, and 21 in order to access the potential of sEHI and EpFAs-derivates to promote SAOS-2 differentiation. Our results demonstrated that sEHI enhances the ALP activity on days 5 and 10 (P<0.05; Fig.3A and B). Similarly, EEQ improves ALP activity in the early stages, while EETs augment ALP function later (P<0.05; Fig.3A and B). On the other hand, the SAOS-2 cells cultured with EDPs did not change the ALP activity over time (P>0.05; Fig.3A and B). These data indicate that the homologs of the EpFA are not biologically equivalent.

Figure 3. The impact of sEHI and EpFAs in the mineralization process.

(A – B) Alkaline phosphatase activity assay. SAOS-2 cells were cultured with or without osteogenic induction (Differentiation (+) and Differentiation (−), respectively) and treated with sEHI (10 μM), EETs (1 μM), EDPs (1 μM), or EEQs (1 μM) for 5 and 10 days. (C) Representative image of the mineralization assay using Alizarin Red staining. (D – F) Quantification of Alizarin Red staining absorbance at 10, 14, and 21 days. Data are presented as mean ± SD. The experiment was conducted in technical duplicates with two biological replicates. Different letters indicate statistically significant differences between groups.

Regarding the mineral nodule formation, the sEHI (TPPU) clearly increased the calcium deposition from day 10 toward day 21 (P<0.05; Fig.3C - F). The EpFAs-derivates EETs showed an increased calcium deposition at the late stages, reaching the same parameters as sEHI (P<0.05; Fig.3F). Likewise, the EpFA EEQs raised the calcium deposition since day 14, although in lower scales comparing to sHEi and EETs (P<0.05; Fig.3G and F). Corroborating the ALP activity findings, the SAOS-2 cells cultured with EDPs negatively impact the calcium deposition and, consequently, the mineral nodule formation (P>0.05; Fig.3C - F). These in vitro results demonstrate the potential of sHEi to promote SAOS-2 cell differentiation, leading to increased calcium deposition, as evidenced by the alizarin red assay.

3.3. Gene expression modulation through sEHI and their EpFA substrates.

The impact of sEHI and EpFAs on osteoblast cell markers was quantitatively assessed by measuring the gene expression of Runx-2, Osx, Col1, Bsp, Opg, Ocn, Opn, and sEH. The results showed that sEHI significantly increased the gene expression of Runx-2, Osx, and Bsp on day 1 (P<0.05; Fig.4A, B, and D). Similarly, EEQ significantly enhances the expression of Opg (P<0,05; Fig.4E). On day 3, sHEi significantly increased the gene expression of Col1, Ocn, and Opn (P<0,05; Fig.4C, F, and G). The levels of Ocn were also upregulated in SAOS-2 cells cultured with EEQs (P<0,05; Fig.4F). Finally, on day 5, the sSEHI maintained the augmented gene expression of Opn, with EETs and EEQ (P<0,05; Fig.4G). The levels of Runx2 at day 5 were found enhanced in SAOS-2 cells cultured with EETs and EEQs (P<0,05; Fig.4A), as well as Bsp expression (P<0,05; Fig.4D). Lastly, the treatment with EDPs significantly increased the mRNA expression of sEH for up to three days, followed by a reduction observed on the fifth day (P<0,05; Fig.4H). These results provide a possible explanation for the inhibitory effect of EDPs on the mineralization process.

Figure 4. Changes in gene expression induced by sEHI and EpFAs.

Runx2 (A), Osx (B), Col1 (C), Bsp (D), Opg (E), Oc (F), Opn (G), and sEH (H) gene expression in SAOS-2 cell cultures after treatment with sEHI (10 μM), EETs (1 μM), EDPs (1 μM), or EEQs (1 μM) for 1, 3 and 5 days by Real-Time PCR. The results are presented as the expression of the target mRNA normalized by GAPDH. Data were reported as mean ± SD of two independent experiments in triplicate. p-values were determined by 2-way analysis of variance (ANOVA) with post hoc contrasts using Tukey’s test. Different letters indicate statistically significant differences between groups.

3.4. Cumulative interaction between sEHI and EpFA in the mineralization process.

Subsequently, we aimed to investigate a potential cumulative interaction between sEH inhibition and the effects induced by EpFAs during the mineralization process and ALP activity. Cell viability and proliferation from all combinantions are presented in Supplementary Figure 1. Herein, we demonstrate that sEH inhibition by TPPU enhanced the effects of EETs and EEQs but not those of EDPs (Fig. 5A – D). Specifically, mineralization levels were higher and achieved in a shorter time (exhibiting high mineralization within 10 days) when EETs and EEQs were combined with TPPU (P<0,05; Fig. 5B, C and D). Conversely, the presence of sEHI did not enhance the effects of EDPs in any period (P>0,05Fig. 5A – D). Furthermore, it inhibited the natural mineralization process induced by osteogenic media (OM+), suggesting that either EDPs or their corresponding diols exert a negative influence on the mineralization process observed in this study.

Figure 5. Cumulative interaction between sEHI and EpFA in the mineralization process.

SAOS-2 cells were culture with or without osteogenic induction (Differentiation (+) and Differentiation (−), respectively) and treated with different combinations of sHEI (10 μM), EETs (1 μM), EDPs (1 μM), and EEQs (1 μM). (A) Representative image of the mineralization assay using Alizarin Red staining. (B – D) Quantification of Alizarin Red staining absorbance at 10, 14, and 21 days. (E – F) Alkaline phosphatase assay using Fast Red staining. Absorbances were measured at 5 and 10 days. Data are presented as mean ± SD. The experiment was conducted in technical duplicates with two and three biological replicates. Different letters indicate statistically significant differences between groups.

Likewise, when distinct combinations of sEHI and epoxide metabolites were evaluated, the pairing of sEHI with EETs exhibited the highest alkaline phosphatase activity during the initial 5-day period, followed closely by the combination of sEHI with EEQs (P<0.05; Fig. 5E and F). Conversely, consistent with its previously observed effects in isolation, EDP significantly inhibited the mineralization process (P < 0.05; Fig. 5E and F).

4. DISCUSSION

Osteoblasts play a crucial role in maintaining bone physiology, being responsible for bone formation and mineralization. These specialized cells are essential for regulating the balance between bone formation and resorption, a dynamic process necessary for bone growth, development, and repair throughout life. Osteoblasts synthesize and secrete components of the bone matrix, such as type I collagen, non-collagenous proteins, and proteoglycans, which form the bone structure and provide its mechanical strength. Additionally, osteoblasts produce growth factors and cytokines that regulate the activity of other bone cells, such as osteoclasts, which are responsible for bone resorption (Stegen & Carmeliet, 2024). Therefore, the function of osteoblasts is crucial for maintaining bone homeostasis and preserving the structural and functional integrity of the skeleton.

In the present study, we demonstrated that both the sEHI (TPPU) and its EpFA substrates increased cellular metabolic activity within the initial 48 hours but did not affect cell proliferation. Functional assays revealed that sEHI enhanced osteogenic events, while EETs and EEQs showed improvement mainly in later stages. When combined, the sEHI accelerated the effects of EETs and EEQs in the mineralization process. We hypothesize that the sEHI are acting to preserve the EpFAs storred or being produced in the cells, although this hypothesis has not yet been tested. Furthermore, SAOS-2 cultured with sEHI demonstrated modulation in the expression of key markers associated with osteoblast differentiation and activity.

To better understand how sEH inhibition and its epoxy fatty acid substrates influence osteoblast biology, we selected a panel of well-established genes that represent different stages of osteogenic differentiation. Runx2 and Osx are key transcription factors that drive early osteoblast commitment and regulate the genetic programs required for matrix production and mineralization (Stegen & Carmeliet, 2024). Col1, which encodes type I collagen, is one of the first markers to be expressed during osteoblast maturation and reflects the onset of extracellular matrix synthesis (Wang et al., 2006).

As osteoblasts progress toward mineralizing the matrix, they increase expression of non-collagenous proteins such as Bsp and Opn, which are crucial for matrix organization and mineral deposition (Wang et al., 2006; Trindade-da-Silva et al., 2017). Later in this process, Ocn (osteocalcin) becomes prominent, signaling terminal differentiation and active mineralization (Gao et al., 2023). Lastly, Opg plays an essential role in bone homeostasis by preventing excessive bone resorption, acting as a natural inhibitor of osteoclastogenesis (Trindade-da-Silva et al., 2020; Abdalla et al., 2024).

In line with their known functions, we observed that TPPU upregulated Runx2, Osx, and Bsp as early as day 1, suggesting a rapid activation of the osteogenic transcriptional program. The early increase in Col1 and later elevation of Ocn and Opn expression further indicate that sEH inhibition promotes not only commitment but also maturation of the osteoblast phenotype. Interestingly, EEQs and EETs also increased Runx2, Bsp, and Ocn, although to a lesser extent and predominantly at later time points. These findings are consistent with the idea that epoxy fatty acids act synergistically with sEHI to support osteoblast differentiation, possibly by preserving endogenous pro-resolving lipid mediators that enhance bone formation (Gao et al., 2023; Trindade-da-Silva et al., 2020). Conversely, EDPs showed limited ability to stimulate osteogenic gene expression and even increased sEH mRNA levels in the early phase, potentially reflecting a compensatory upregulation due to weak substrate affinity or distinct signaling dynamics. This may partially explain the lack of mineralization enhancement observed with EDPs, as compared to EETs and EEQs. Altogether, our gene expression data support the hypothesis that sEH inhibition, particularly when combined with select EpFAs, can prime osteoblasts for effective differentiation and matrix mineralization, revealing a promising axis for bone regenerative approaches.

The sEH inhibitor TPPU enhances cellular osteogenic differentiation in a co-culture of human umbilical vein endothelial cells (HUVECs) and human dental pulp stem cells (hDPSCs) in vitro, and also facilitates long bone growth (femur), contributing to bone repair (Gao et al., 2023). Research indicates that sEH plays a protective role against vascular calcification by controlling the availability of epoxyeicosatrienoic acids and other long chain PUFAs (not all have same chain length or olefin number or position, especially EPA and DHA), which are essential in the calcification process. Thus, inhibition of sEH increases vascular calcification by presumably by protecting EpFA from hydrolysis and thus enhancing calcium deposition and osteochondrogenic marker expression, indicating a protective role in bone mineralization (Varennes et al., 2020). Additionally, TPPU-mediated sEH inhibition diminishes bone resorption and inflammatory responses, showcasing immunomodulatory effects in models of bone resorption (Abdalla et al., 2023; Abdalla et al., 2024). Overall, the inhibition of soluble epoxide hydrolase (sEH), as evidenced in the current study, affects bone mineralization by fostering osteogenesis and augmenting cellular metabolic activity.

In osteoblast cells, soluble epoxide hydrolase (sEH) inhibition offers several potential benefits. Firstly, sEH inhibitors have been shown to enhance osteogenic differentiation and bone repair by promoting angiogenesis and cell proliferation (Gao et al., 2023). Additionally, inhibiting sEH prevents the metabolism of anti-inflammatory epoxyeicosatrienoic acids, which can reduce pain associated with conditions like arthritis (Trindade-da-Silva et al., 2020; Phillips et al., 2022). Moreover, targeting the EET/sEH axis may help in resolving chronic inflammatory disorders by activating endogenous regulatory pathways, including the production of specialized pro-resolving mediators (Abdalla et al., 2024).

Epoxyeicosatrienoic acids (EETs) play a crucial role in promoting bone formation through various molecular mechanisms. EETs have been shown to inhibit osteoclastogenesis by modulating multiple pathways involved in osteoclast formation and activity, including the NF-κB, AP-1, and MAPK signaling pathways, as well as preventing the production of reactive oxygen species (Guan et al., 2015). Additionally, EETs have been found to attenuate bone loss by decreasing the RANKL:osteoprotegerin ratio and serum levels of inflammatory cytokines like TNF-α and IL-1β (Trindade-da-Silva et al., 2017). Furthermore, EETs have been demonstrated to enhance chondrogenic responses during bone repair by promoting chondrocyte proliferation and differentiation through the EPO/EPOR signaling pathway (Wan et al., 2014). Overall, these findings suggest that EETs exert their beneficial effects on bone formation by regulating key molecular pathways involved in osteoclastogenesis and chondrogenesis.

As demonstrated in the present study, only the sEH inhibitor TPPU was initially capable of increasing collagen type I gene expression. A previous study demonstrated that combined inhibition of cyclooxygenase (COX) and soluble epoxide hydrolase (sEH) demonstrated superior efficacy in controlling joint pain, regulating prostanoid responses, and maintaining the balance of collagen synthesis and degradation compared to individual compound inhibition (Tucker et al., 2021). Furthermore, Bsp is a key non-collagenous protein found in bone extracellular matrix, playing a critical role in regulating bone mineralization and remodeling (Wang et al., 2006). Herein, sEHI augmented the expression of Runx2 and Osx in the first 24 hours, whereas EETs and EEQ enhanced Runx2 levels at five days post-treatment. These findings on osteoblast differentiation markers are associated with the enhancement of Col1, Bsp, Oc, and Opn by sEHI, which are all enrolled in osteogenic activity.

Clinical studies have investigated the potential benefits of using ω−3 as a supplementary therapy in periodontal treatment. For instance, adjunctive ω−3 (3 g of fish oil/daily) associated with low doses of aspirin (100mg/daily) after periodontal debridement provides clinical improvements, like attachment gain in moderate and deep pockets (Castro-dos-Santos et al., 2020). It was observed that patients receiving this treatment regimen showed a decrease in the release of inflammatory cytokines in their crevicular fluid. Morever, a mixture of PUFAs with EPA and DHA were provided as a dietary supplementation, combined with scaling and root planing (SRP). The researchers found that ω −3 treatment significantly reduced bleeding during probing, showed a higher gain of clinical attachment loss, and a higher number of closed pockets within three months. Additionally, pathogenic bacteria were found to have decreased compared to the control group (Stańdo-Retecka et al., 2023).

Unlike ω−6 fatty acids, such as arachidonic acid (ARA), ω−3 fatty acids exhibit low affinity as substrates for inflammatory pathways. The epoxides located at the ω−3 position of EPA and DHA are less efficient substrates for soluble epoxide hydrolase (sEH). In contrast, ARA-derived epoxide metabolites can serve as substrates for cyclooxygenase (COX), potentially leading to the formation of pro-inflammatory products. (Rand et al., 2017). Notably, ω−3 epoxides, such as those derived from EEQs and EDPs, are poor substrates for sEH due to their unique structural characteristics, including a C2H5 group that slows their turnover compared to other isomers (Morisseau et al., 2010; Morisseau et al., 2021). Consequently, ω−3 epoxides circulate in the blood as markers of ω−3-rich diets, potentially elucidating their role in promoting inflammation-resolving processes (Castro-dos-Santos et al., 2020; Stańdo-Retecka et al., 2023). This metabolic distinction may contribute to the anti-inflammatory benefits associated with ω−3 fatty acid consumption.

These findings indicate that sEHI and their EpFA substrates significantly influence the gene expression of osteoblast cell markers, highlighting their potential for the development of novel therapies targeting bone-related disorders. The data further indicate that the effects of EETs and EEQs, but not EDPs, have similar but not identical effects qualitatively and quantitatively. Additionally, the sEHI, TPPU, may stabilize pre-existing epoxy fatty acids within cells, potentially as phospholipid-bound forms, thereby prolonging and enhancing their biological activity. This interaction appears to be synergistic, further amplifying their therapeutic potential.

5. CONCLUSION

Altogether, our results demonstrated that sEHI is a highly promising approach for improving osteoblast functions in vitro. Not only does it enhance mineralization, but it also has the potential to promote tissue regeneration in vitro. Therefore, it can be speculated that sEHI can significantly contribute to the improvement of bone tissue engineering and regenerative medicine.

HIGHLIGHTS.

• sEHI and EpFAs boost osteogenesis, enhancing activity and mineral formation.

• sEHI regulates osteogenic markers, promoting mineralization and differentiation.

• EDPs disrupt sEHI-EpFAs synergy, breaking the guidance to bone regeneration.

Data Availability

The data that support the findings of this study are available from the corresponding author upon reasonable request.