Synthesis and characterization of 3D gelatin/calcium hydroxide/dentin matrix proteins/propolis scaffolds for hard tissue engineering

Abstract

This paper explores the development of three-dimensional porous scaffolds composed of gelatin, calcium hydroxide, dentin matrix proteins, and propolis (signified as G, Ca, DP, and P, respectively). First, the scaffolds were synthesized using freeze-drying technique, then their physicochemical properties, mechanical characteristics, cytocompatibility with dental pulp stem cells, and cell adhesion, differentiation and mineralization were evaluated. The results showed that the scaffolds were porous with asperity. The surface roughness analysis identified higher roughness value for the G-Ca-P and G-Ca-DP-P scaffolds (mean Rq of 38.16 and 21.69 nm, respectively). All the scaffolds exhibited hydrophilic behavior (contact angles < 90°). However, those having propolis, calcium hydroxide and protein showed higher wettability. Furthermore, all scaffolds degraded over time with a synergistic effect between calcium hydroxide and dentin matrix proteins in making a more robust scaffold and enhancing the alkalinizing effect. The G and G-P scaffolds had elastic-plastic behavior, while G-Ca-P and G-Ca-DP-P scaffolds showed brittleness. Moreover, all scaffolds revealed no cytotoxicity (cell viability >80%). The alkaline phosphatase (ALP) concentration and matrix deposition, as measures of osteogenic differentiation, were highest for G-Ca-DP-P scaffolds, followed by G-Ca-P. Besides, the cells attached to the scaffold surfaces with a flattened morphology, and the highest cell number adhered on G-Ca-DP-P and G-Ca-P scaffolds, respectively.

Altogether, the G-Ca-DP-P scaffolds acted as hydrophilic, rough, and biodegradable frameworks supporting cell viability, adhesion, differentiation and mineralization.

Introduction

Hard tissue engineering (HTE) focuses on the restoration of mineralized tissues such as bone and dentin, which exhibit limited natural healing potential, especially in cases of critical-sized defects. This field plays a vital role in addressing challenges in dentistry, orthopedics, and craniofacial reconstruction through the application of biomaterials, cells, and bioactive signals. A wide range of regenerative approaches have been developed to overcome the limitations and risks associated with autografts, allografts, and xenografts, and to enhance tissue regeneration effectively [1]. The three-dimensional (3D) porous structure scaffolds can provide a suitable matrix needed in tissue engineering (TE) for efficient mass transport, that is the exchange of nutrients and wastes [2, 3]. Furthermore, the pores and their interconnectivity enable cell migration, tissue ingrowth, and vascularization [4]. These scaffolds in TE are supportive structures that facilitate post-transplant cell proliferation in the patient’s body at the defect site [5].

Despite the advances in the development of scaffolds, HTE still faces significant challenges and there is need for structures with improved function. The challenges include inadequate mechanical adaptation of scaffolds to the host tissue, early or late degradation of materials, and unwanted inflammatory responses. An ideal scaffold, therefore, should possess several characteristics including adequate mechanical strength, high biocompatibility, appropriate biodegradability rate, and proper interactions with cells and tissues. Indeed, the design and fabrication of a scaffold is an application-specific process, which should be done judiciously based on the composition and structure of the repairing tissue to achieve optimal functions [6]. Therefore, the scaffolds used in dental tissue engineering (DTE) may require different properties rather than the materials used in other tissues. DTE seeks to regenerate damaged or lost tooth components such as enamel, dentin, and pulp [7]. However, in DTE, limitations in providing a suitable environment for the growth and differentiation of dental stem cells and inefficiency in regenerating dentin or real pulp are considered major obstacles [8].

DTE also deals with bone tissue engineering (BTE), when developing 3D scaffolds to regenerate lost bone tissues for craniofacial defects or in oral cavity. In BTE, similarly, the 3D scaffolds are engineered with appropriate composition, architecture and additional biological agents to boost bone regeneration in osteonecrosis and trauma-induced defects [9]. Dental and bone tissue engineering (DTE/BTE) face unique mechanical and biochemical challenges. DTE scaffolds must withstand occlusal forces while promoting oral tissue (like pulp-dentin) regeneration, whereas BTE materials require load-bearing capacity and osteoconductivity. The scaffolds in BTE and DTE can further be enriched with specific biomolecules such as polypeptides or proteins that attach to the certain receptors on the surface of target cells. For example, Bone Morphogenetic Protein (BMP) receptors affect a wide range of cellular activities such as proliferation, differentiation, and apoptosis of dental pulp cells [10].

Various tissue scaffolds with a variety of compounds and structures have been developed for tissue engineering in medicine and dentistry, with their benefits and drawbacks [11]. The scaffolds can be made up of different materials (natural or synthetic polymers, metals, ceramics, or their composites), in various forms (hydrogels, particles, nanofibers, and porous solids), and by different manufacturing methods (bioprinting, 3D printing, freeze-drying, casting, etc.) [12–17]. Hydrogels and 3D solid scaffolds have been widely developed to promote effective tissue regeneration. Injectable hydrogels, for instance, enable minimally invasive administration and in situ gelation, making them particularly suitable for treating irregularly shaped defects. Meanwhile, 3D porous solid scaffolds with interconnected porosity facilitate cell migration, nutrient and oxygen diffusion, and waste elimination, critical factors for successful tissue integration. Despite the significant progress in this field, considerable challenges remain. These challenges drive ongoing research into novel formulations and biomimetic materials that better replicate the native tissue’s structure and function, ultimately aiming to enhance hard tissue repair. A combination of polymeric materials and minerals might be an effective strategy as the natural extracellular matrix (ECM) of the hard tissues is comprised of organic-inorganic (mainly collagen and hydroxyapatite) components. In this regard, we proposed a composite scaffold comprising of gelatin and calcium hydroxide (Ca (OH)₂) in equivalence to collagen and hydroxyapatite in native ECM of hard tissues. This composite may be reinforced with biological factors like propolis and dentin matrix proteins (DMPs).

Gelatin is a molecular collagen derivative, that is obtained by the irreversible denaturation of collagen proteins. Gelatin has a very close molecular structure and function to collagen, and it is often used in cellular and tissue culture to replace collagen for biological and medical purposes [18]. Furthermore, gelatin-based materials have been widely employed to support bone tissue regeneration, in various clinical applications [19]. Despite its desired characteristics including favorable biocompatibility, low toxicity, biodegradability, increased cell adhesion, differentiation and proliferation, no immunogenic response, and being cost-effective [20], it lacks mechanical strength and thermostability. Therefore, cross-linking is done to overcome the latter, and combining with stiffer materials like ceramics and minerals is accomplished to provide improved mechanical properties, which is fruitful for hard tissue repair [21].

Calcium hydroxide (Ca (OH)₂) has been widely used to direct pulp cap in modern clinical dentistry. It has the potential to induce hard tissue repair, provide disinfection (initially bactericidal, then bacteriostatic), cause intra canal medicament, and stimulate fibroblasts [22]. Therefore, the combination of this material with gelatin as a composite scaffold may provide advantages for hard tissue engineering.

Propolis, a resinous substance gained from the bees salivary secretions, recently has been proposed as a bioactive material that contains mostly resin, wax, natural aromatic oils, pollen, flavonoids, amino acids, phenolic components and other inorganic ingredients [23, 24]. Flavonoids are the main bio-components of propolis extract, which indicate the biological capability of the propolis products [25]. Besides, propolis has strong antimicrobial and anti-inflammatory properties [26–28]. In experimental studies, propolis has shown the ability to reduce inflammation, accelerate the tissue repair process, and inhibit the growth of bacteria and fungi, making it a suitable option for application in DTE and BTE. Therefore, this natural biomaterial, has recently attracted the attention of researchers in the field of tissue engineering. Specifically, this material has been used in combination with other compounds for BTE, and the results were promising [29–32].

Furthermore, the use of bio-factors including growth factors, or extracellular vesicles along with scaffold can aid in better regeneration process [33]. The DMPs have the ability to inherently bind to calcium in the ECM, which leads to tissue calcification, act as intracellular signaling proteins inducing differentiation of stem cells, and also serve as ECM nucleating proteins [34]. These make the DMPs an attractive choice for BTE and DTE.

The purpose of this study is to develop a 3D porous scaffold using biocompatible and bioactive materials including gelatin, calcium hydroxide, and propolis. These materials were selected due to their known biological properties, such as the ability to stimulate differentiation of dental pulp stem cells (DPSCs) and exerting anti-inflammatory effects, which make them suitable candidates for the hard tissue regeneration process [35]. Additionally, DMPs were incorporated as a bio-factor to further enhance the biological performance of the synthesized scaffolds. Given the existing limitations in HTE, there is a critical need for novel composite scaffolds that can concurrently offer bioactivity, and mechanical, and anti-inflammatory properties. This study not only explores the feasibility of fabricating such a scaffold but also assesses its physicochemical properties and the influence in the proliferation, differentiation and mineralization of DPSCs. By combining the main regenerative requirements in a single composite formulation, the findings of this study may contribute meaningfully to the development of advanced biomaterials for practice in clinical procedures. Finally, this research seeks to advance the translation of experimental scaffold design to real and practical therapeutic applications in regenerative medicine.

Materials and methods

Preparation of gelatin/calcium hydroxide/dentin matrix proteins/propolis scaffolds

First, a 10 w/v% gelatin solution in distilled water was prepared at 40 °C. Once the solution became uniform, the heating was stopped to reduce the temperature to 25℃, and calcium hydroxide (5 w/v%) was gradually added. Finally, the DMPs was added to the solution (20 µg for each scaffold with the dimensions of 2.5 × 2.5 × 10 mm³) before pouring the solution into the plastic molds, which were then frozen at −20 °C for 24 h and freeze-dried. Then, the freeze-dried samples were immersed in glutaraldehyde 1% for 6 h to be crosslinked. After several washing and drying, the crosslinked solid scaffolds were coated (immersion for 15 min) with 2.5 w/v% propolis in distilled water. To prepare 2.5 w/v% propolis solution, 250 mg of propolis was first dissolved in 10 mL of 96% ethanol and stirred with a magnetic stirrer for two days. The ethanol was allowed to evaporate. Next, 10 mL of distilled water was added to the residue and the resulting milk-like solution was filtered by a filter paper to obtain a 2.5 w/v% propolis solution in distilled water. For comparison, different scaffolds with and without Ca (OH)₂, DMPs and propolis were also synthesized (Table 1). Figure 1 shows the fabrication process and the scaffolds made.

Table 1.

Studied groups and their corresponding compositions

No.	Scaffold codes	Scaffold compositions
1	G	Gelatin
2	G-P	Gelatin/Propolis
3	G-Ca-P	Gelatin/Calcium Hydroxide/Propolis
4	G-Ca-DP-P	Gelatin/Calcium Hydroxide/Dentin Matrix Protein/Propolis

Fig. 1.

Schematic of the fabrication process and the resulting scaffolds

Evaluation of surface morphology and porosity of the scaffolds

Surface morphology and porosity of the scaffolds were examined using scanning electron microscopy (SEM, ZEISS). First, the scaffolds were sputter-coated with a conductive material (gold), and SEM images were obtained in various magnifications.

Surface topography and measurement of surface roughness

For surface topography and roughness evaluation, an atomic force microscope (AFM, Ara-Research Company, Iran) was utilized. The surface of the samples was imaged in three regions with the scanning area of 10 × 10 µm², at the ambient temperature, employing non-contact mode. The surface roughness (Root Mean Square (RMS) roughness (Rq) and average roughness (Ra)) was measured via the Imager software version 1.0 supplied by Ara-Research Company.

Wettability/contact angle test

To obtain the wettability of the scaffolds, which indicates the degree of hydrophilicity, static contact angle (CA) test (sessile drop method) was performed at room temperature. In this experiment, a drop of distilled water was automatically dropped on the scaffold surface using a syringe pointed perpendicular down onto the scaffold surface. The water contact angle with the surface of the samples was measured using a special microscope equipped with a video camera to record the water drop images instantly once it was deposited on the surface. Three regions were imaged for each scaffold, and the contact angle was measured from right and left sides.

Biodegradability evaluation

The biodegradability assessment provided important information on the degradation behavior of the scaffolds, which is a critical factor in determining their suitability for tissue engineering applications [36]. The evaluation of biodegradability of the scaffolds was studied in phosphate-buffered saline (PBS) for 42 days. The scaffolds were immersed in PBS at 37 °C, and the solution was replaced twice a week. At the predetermined time intervals (1 h, 3 h, 6 h, 24 h, 72 h, and up to 42 days), the scaffolds were removed from the PBS solution, washed with distilled water and dried at room temperature. The weight loss of the scaffolds was measured with a digital balance and the degradation rate was calculated using Eq. 1.

1

Where W_i and W_f are the initial and final weights of the scaffolds, respectively.

Measurement of pH changes

The scaffolds were individually immersed in containers having PBS at 37℃ in a water bath for 21 days and during the soaking period, the pH changes of the PBS solutions were recorded (every day) using a pH meter (Shimifan. CO, Iran).

Evaluation of mechanical properties

The mechanical properties of the scaffolds were evaluated using compression testing. The ultimate compressive strength (UCS) and Young’s modulus (E) were measured using a universal testing machine (UTM, Santam Co., Iran) at a crosshead speed of 1 mm/min. The compressive strength was calculated as the maximum stress before failure, and the Young’s modulus was calculated as the slope of the linear region of the stress-strain curve.

Investigation of biological properties

Cell culture and scaffold sterilization

In this study, dental pulp stem cells (DPSCs, purchased from the Royan Institute for Biotechnology, Isfahan, Iran) were cultured in a high-glucose Dulbecco’s Modified Eagle Medium (DMEM) medium enriched with 10% fetal bovine serum (FBS), 1% L-glutamine, and 1% penicillin/streptomycin antibiotics, and incubated at 37 °C with 5% CO₂ and humidity. After synthesizing the scaffolds, to do the cellular tests, they were sterilized using UV radiation (20 min on each side).

Cell adhesion assay

The cell adhesion was analyzed using field emission SEM (FE-SEM, Sigma 300-HV, Ziess, Germany). DPSCs (1 × 10³ cells/scaffold) were seeded on the surface of each scaffold in a 24-well plate and incubated for 72 h. Next, the adherent cells were fixed with ethanol at concentrations of 30%, 50%, 70%, and 96% (each for 15 min). After drying, the samples were coated with gold and the SEM images were obtained at various magnifications.

Cell viability assay

First, the cells (5 × 10³) were seeded onto the scaffolds in a 24-well plate and incubated for 3 and 7 days. After the incubation time, the culture medium was removed and replaced with 900 µL/well of incomplete culture medium and 100 µL/well of Alamar Blue (resazurin) in the dark. The cells were then incubated for 3 hours at 37 °C in the incubator. Finally, the culture medium (supernatant) was transferred into a 96-well plate and the absorbance was measured at 570 and 600 nm wavelengths using a microplate reader equipped with a fluorescence reader (Synergy H1 Hybrid MultiMode Microplate Reader, BioTek, USA). The same scaffolds and cells were analyzed over time. Specifically, after the measurement at day 3, the scaffolds and cells were washed with PBS to remove the resazurin dye, complete growth medium was added, and the plate was incubated until day 7. In this assay, a control group including DPSCs in complete culture medium with no scaffold was considered, where the same cells were studied at both time points, similar to the scaffold groups. The cell viability was calculated based on Eq. 2.

2

Cell differentiation assay

Measurement of alkaline phosphatase concentration

To assess the differentiation of dental pulp stem cells by measuring the alkaline phosphatase (ALP) concentration, the scaffold medium extracts were used. To prepare the extracts, 3 samples from each group were placed in a 24-well cell culture plate for 72 h in complete cell culture medium. After 72 h, the scaffolds were discarded, and the extracts were used for the test. To do the ALP assay, 5 × 10³ DPSCs were seeded in a 96-well plate for 24 h to form a complete monolayer on the bottom of the plate. Then, the medium was replaced by the scaffold medium extracts along with differentiating factors (ascorbic acid at 50 µg/ml and β-glycerophosphate at 10mM). ALP concentration was evaluated at days 5 and 10 of incubation using an alkaline phosphatase kit (DELTA DARMAN PART Co.). The cells were lysed with 100µL of diluted Triton X-100, and the supernatant was transferred into another 96-well plate and incubated with pNPP substrate for 1 min at 37℃. The ALP concentration was analyzed by measuring the absorbance at 405 nm in 1, 2 and 3 minutes, and using Eq. 3.

The ∆A is the difference between the measured absorbances at different time points.

3

Alizarin red staining

Alizarin red staining (ARS), which identifies the calcium deposition by cells, was performed after DPSCs incubation for 7 and 14 days. First, 5 × 10³ cells/well were cultured in 24-well plates, which were kept in an incubator at 37 °C with 5% CO₂ and 95% humidity to form a monolayer of the cells. Next, the scaffold media along with osteogenic agents (50 µg/mL ascorbic acid, and 10mM β-glycerophosphate, Sigma-Aldrich, USA) replaced the complete growth medium (except for control groups). The osteogenic factors were added twice a week. In this test, two control groups were also considered including positive control (C⁺, DPSCs with no scaffold medium and with osteogenic agents) and negative control (C⁻, with no scaffold medium and no osteogenic agents). After 7 and 14 days of incubation, the cells were fixed using 4% paraformaldehyde for 15 min in a refrigerator, then washed with PBS, and stained with 1% (w/v) alizarin red dye for around 40 min at ambient temperature in dark conditions. The stained cells were rinsed with PBS several times to remove unreacted alizarin red. Lastly, they were observed under loop and optical microscopes and photographed.

The mineralized/calcific red area (A _Ca) in each well and the total area of the well (A _total) were measured using ImageJ Software. Then, the mineralization% was calculated based on Eq. 4.

4

Statistical analysis

The normality and homoscedasticity were checked. Analysis of variance (ANOVA) and non-parametric tests were carried out based on the normality. The significance level of α = 0.05 was applied to do the statistical analyses. Post hoc pairwise comparisons were done using Tukey’s or Dunn’s or Sidak’s multiple comparisons tests. All data were reported as mean ± standard error, and all analyses and graph generation were conducted using GraphPad Prism software (Version 8).

Results

Surface morphology

The surface morphology of the scaffolds was examined using scanning electron microscopy. The scaffolds, as shown in Fig. 2, exhibited a porous structure, which allow for the penetration of nutrients and the escape of waste products. Furthermore, they had rough surfaces with asperity, which may enable the cells to anchor to their surfaces, flattening, spreading and structural organization. The pores were more visible in G and G-P scaffolds, and they are partly filled by addition of calcium and proteins.

Fig. 2.

SEM images of the as-fabricated scaffolds

Surface topography and roughness

The surface topography and roughness results for the scaffolds are presented in Figs. 3 and 4, respectively. Figure 3 displays the AFM images of the scaffolds, while Fig. 4 provides the corresponding roughness values. It was found that the surface roughness of the scaffolds varied depending on the composition. The statistical analysis also indicated that the scaffold composition is an influential factor in the surface roughness values (both Ra and Rq with P < 0.0001). The G-Ca-P and G-Ca-DP-P scaffolds exhibited higher roughness value (mean Rq of 38.16 and 21.69 nm, respectively), indicating a rougher surface compared to the other scaffolds, as shown in Fig. 3c and d. The G-P scaffolds had a lower roughness value (mean Rq of 7.18 nm), suggesting a smoother surface than the G-Ca-P and G-Ca-DP-P scaffolds but rougher than the G scaffolds (mean Rq of 2.47 nm).

Fig. 3.

AFM images of the scaffolds; (a) G, (b) G-P, (c) G-Ca-P, and (d) G-Ca-DP-P scaffolds

Fig. 4.

Surface roughness of the studied scaffolds; (a) Ra, and (b) Rq. Kruskal-Wallis test was used with Dunn's multiple comparisons test. N=24-49, P<0.05: *, P<0.01: **, P<0.001: ***, and P<0.0001: ****

Wettability

The results of the wettability test are presented in Fig. 5. The contact angles (CA) for the G, G-P, G-Ca-P, and G-Ca-DP-P scaffolds were found to be 82.55°, 63.06°, 59.21°, 77.73°, respectively. The lower contact angle indicates higher wettability and hydrophilicity, which is desirable for cell attachment and tissue engineering applications. The results obtained here demonstrated higher wettability for the scaffolds with propolis, calcium hydroxide and protein content, compared to the G scaffolds. The wettability results indicated that all the scaffolds exhibited hydrophilic behavior, as their CAs were < 90°. The statistical analysis also revealed that the scaffold composition is an influential factor in the surface wettability (P = 0.0422). However, pairwise comparisons showed that there was no significant difference between groups.

Fig. 5.

The results of wettability test. One-way ANOVA was used with Tukey pairwise comparison. There was no significant difference between the groups. N=5-11

Biodegradability and pH changes

The biodegradability of the scaffolds was assessed over a period of 42 days in PBS solution (Fig. 6a). The degradability percentage of each scaffold was recorded at different time intervals including short-term (from 1 to 72 h) and long-term (up to 42 days). The in vitro degradation behavior of the fabricated scaffolds showed that adding the bioactive components including calcium hydroxide and DMPs influences their stability in both short- and long-term. In the short-term (up to 72 h), all samples exhibited minimal degradation (< 2%), with G and G-P showing the lowest rates (0.67% and 0.77% after 72 h, respectively). This indicates limited early water uptake and structural breakdown. G-Ca-P exhibited the highest short-term degradation, possibly as a result of ionic dissociation of calcium hydroxide. In contrast, G-Ca-DP-P maintained a moderate in vitro degradation rate, suggesting that DMPs may contribute to a more stabilized matrix, reducing early structural destruction. Over a longer period (up to 42 days), degradation increased substantially in all tested groups, particularly in G and G-P (> 70% by day 42). The incorporation of calcium hydroxide in G-Ca-P slowed the degradation process, but it was the presence of DMPs in G-Ca-DP-P that most efficiently reduced the degradation (limiting to < 15% by day 42). This designates a synergistic effect between calcium and DMPs in making a more robust and bioactive scaffold.

Fig. 6.

(a) In vitro degradability of the scaffolds, and (b) pH changes of PBS solution caused by scaffolds. N=3

Figure 6b displays the pH changes occurred in PBS medium over 21 days for four different scaffold formulations including G, G-P, G-Ca-P, and G-Ca-DP-P. The G and G-P scaffolds (pink and light purple lines with circles and squares, respectively, in Fig. 6b) showed rather stable pH values throughout the 21-day period, keeping values about 7.45–7.55. This indicates minimal interaction between these scaffolds and the surrounding medium, and limited bioactivity in terms of pH modulation. In contrast, G-Ca-P (purple triangles) and G-Ca-DP-P (dark purple diamonds) exhibited an obvious progressive rise in pH, reaching values around 7.8 and 7.9, respectively, by day 21. This gradual alkalinization is likely due to the dissolution of Ca (OH)₂, which releases hydroxide ions (OH⁻) from the scaffold, leading to a more basic environment. The G-Ca-DP-P group revealed the most noticeable increase in pH, suggesting enhanced ion release, which may be beneficial for osteogenic activity or biomineralization, as slightly alkaline conditions are often favorable for bone tissue engineering. The presence of Ca and dentin matrix proteins in these scaffolds appeared to enhance this alkalinizing effect possibly by influencing ion release dynamics by dentin matrix proteins, indicating potential synergistic effects in the composite.

Mechanical properties

The mechanical properties of the scaffolds were evaluated using a universal testing machine. The scaffolds were subjected to compressive loading, and the stress-strain curves were analyzed to determine the Young’s modulus (E), which is a measure of the scaffold’s rigidity, and the ultimate compressive strength (UCS), which is the stress required to fail a specimen. As shown in Fig. 7a, G and G-P had elastic-plastic behavior (linear and plateau regions in stress-strain curves), known as ductile behavior. However, G-Ca-P and G-Ca-DP-P showed brittle characteristics as they only had linear elastic deformation and were broken without any plastic strain. This result was expected, as the Ca (OH)₂ is a mineral and its addition to the scaffold composition causes brittleness. The G-Ca-P and G-Ca-DP-P scaffolds had comparable UCS (31.07 vs. 44.83 KPa) and E (485.8 vs. 401.6 KPa), as shown in Figs. 7b an 7c. The G-P scaffolds had the highest compressive strength (88.81 KPa), and the G scaffolds showed the lowest Young’s modulus (152.4 KPa). The statistical analysis also indicated that the scaffold composition is an influential factor in both scaffold UCS (P = 0.0085), and E (P = 0.0004). The UCS of G-P scaffolds was significantly higher than that of G-Ca-P and G-Ca-DP-P scaffolds, and the E of G scaffolds was significantly lower than that of the other groups.

Fig. 7.

Mechanical properties of the scaffolds; (a) stress-strain curves, (b) ultimate compressive strength, and (c) Young's modulus. One-way ANOVA was used with Tukey pairwise comparison. N=3, P<0.05: *, P<0.01: **, P<0.001: ***, and P<0.0001: ****

Cell adhesion

The FE-SEM images, in Fig. 8, revealed that the cells attached to the surfaces of the scaffolds with a relatively flattened morphology. As it can be seen, the number of cells attached to the surfaces of the scaffolds was the highest for G-Ca-DP-P scaffolds, followed by G-Ca-P scaffolds.

Fig. 8.

The SEM images of the scaffolds

Cell viability

Cell viability was assessed using the Alamar Blue staining method. The results showed that the cell viability was > 80% in all scaffold groups, meaning that there was no cytotoxicity associated with the scaffolds (Fig. 9). After 3 days, the mean cell viability percentages for G-Ca-DP-P, G-Ca-P, G-P, and G scaffolds were 115.18%, 113.85%, 139.88%, and 136.27%, respectively. However, after 7 days, the cell viability was decreased (but still higher than 80%); it was 80.32%, 81.37%, 89.63%, and 94.09% for G-Ca-DP-P, G-Ca-P, G-P, and G scaffolds, in respect. The statistical analysis indicated that the scaffold composition (P < 0.0001), time (P < 0.0001), and their interaction (P = 0.0004) are influential factors in cell viability.

Fig. 9.

Alamar Blue test results for DPSCs after 3 and 7 days. Repeated measure ANOVA was used with Sidak's multiple comparisons test. The dash line identifies the 80% viability, which is usually considered as a limit for non-cytotoxicity of biomaterials. N=6, P<0.05: *, P<0.01: **, P<0.001: ***, and P<0.0001: ****

Cell differentiation

ALP concentration

Cell differentiation was assessed through measuring the alkaline phosphatase concentration. This method allows monitoring the differentiation of precursor cells into mature cells (DPSCs toward osteoblasts), which is a key aspect of tissue regeneration. Figure 10 shows the ALP concentration at two time points. The results showed that the ALP concentration increased significantly over time for all scaffolds, particularly G-Ca-DP-P scaffolds. At day 5, the ALP concentration was only detected in G-Ca-P (1.65 U/mL) and G-Ca-DP-P (0.73 U/mL) groups. There was no detection of ALP for G, G-P and control groups. The mean ALP concentration for G-Ca-P scaffolds was slightly higher than that of the G-Ca-DP-P scaffolds at day 5. However, after 10 days, the ALP concentration for G-Ca-DP-P scaffolds increased to 5.87 U/mL, while for G-Ca-P scaffolds, it increased only to 3.85 U/mL. For G-P and G scaffolds, the ALP activity was observed at day 10, however, the mean ALP concentration was low (2.2 U/ml and 0.73 U/ml, respectively). The statistical analysis indicated that the scaffold composition (P = 0.0051) and time (P < 0.0001) are influential factors in ALP concentration, but their interaction is not (P = 0.1568).

Fig. 10.

ALP concentration as a measure of DPSCs differentiation after 5 and 10 days (C is control group (cells in complete culture medium with no scaffold extract)). Two-way ANOVA was used with Tukey's multiple comparisons test. N=3, P<0.05: *, P<0.01:**, P<0.001: ***, and P<0.0001: ****

Matrix mineralization

Figures 11 and 12 show the Alizarin red staining of DPSCs exposed to the scaffold media after osteogenic induction at days 7 and 14 (both the photographs from wells and microscopic images a long with quantitative data). As it is observed in these figures, the red nodules were highly formed by DPSCs exposed to G-Ca-P and G-Ca-DP-P scaffold media at both time points, indicating the positive effect of Ca and dentin matrix proteins in mineralization. Furthermore, the G-P group induced more mineralized matrix compared to G group, as the slight red stain was observed in G-P scaffold group. This means that propolis could help in the cell mineralization, but in a lower extent than that caused by incorporation of Ca (OH)₂ and DMPs. Similar observation was evident at days 7 and 14. Nevertheless, on the 14th day of differentiation, the red nodules had more robust intense red color compared to day 7, particularly in G-Ca-P and G-Ca-DP-P groups. This means that the calcium deposition was highest in these groups compared to other groups. Furthermore, as shown in these figures, no red stains were found on C⁻, as it was expected. The quantitative results in Fig. 12b confirms these findings, identifying the superiority of G-Ca-P and G-Ca-DP-P scaffolds in inducing matrix mineralization (> 70%) over the other groups. The statistical analysis indicated that the scaffold composition (P < 0.0001) is an influential factor in matrix mineralization, but time (P = 0.2498) and their interaction (P = 0.5991) are not.

Fig. 11.

ARS as a measure of DPSCs mineralization after 7 days. Scale bars in microscopic images are 250 µm

Fig. 12.

(a) ARS images as a measure of DPSCs mineralization after 14 days, and (b) quantitative mineralization (%) after 7 and 14 days. Two-way ANOVA was used with Tukey's multiple comparisons test. N=2, P<0.05: *, P<0.01:**, P<0.001: ***, and P<0.0001: ****. Scale bars in microscopic images are 250 µm

Discussion

Hard tissue engineering deals with restoring the structure and function of mineralized tissues like bone and dentin, which have restricted reparative capacity after injury or disease, particularly when the tissue defects have critical sizes. This field addresses critical needs in dentistry, orthopedics, and craniofacial reconstruction by utilizing biomaterials, cells, and signaling molecules. Numerus substitutes (hydrogels and 3D scaffolds) have been developed to induce efficient regeneration. For example, the injectable hydrogels offer minimally invasive delivery and in situ forming, which are beneficial for irregularly shaped defects, and the 3D porous scaffolds with interconnected porosity allows for the cell migration, diffusion of nutrients, oxygen, and waste removal [37, 38]. Despite recent advances in this field, there is still important challenges, which impose researchers to develop different formulations and substitutes, trying to mimic the native tissue structure and function and to enhance hard tissue repair.

The aim of the current study was to examine the effect of incorporation of different materials including calcium hydroxide, propolis and dentin matrix proteins on physicochemical, mechanical and cellular behavior of gelatin scaffolds. First, the gelatin, gelatin-calcium hydroxide, and gelatin-calcium hydroxide-dentin matrix proteins scaffolds were fabricated by freeze-drying technique. Subsequently, they were immersed in propolis to provide G-P, G-Ca-P, and G-Ca-DP-P, in addition to G scaffolds. The freeze-drying (lyophilization) technique is a well-established approach for making highly porous 3D constructs with interconnected pore networks, particularly appropriate for TE applications. Freeze-drying makes porosity by freezing a polymer solution and the subsequent sublimating the solvent (usually water) under vacuum condition, creating a solid porous structure. Indeed, ice crystals form throughout the scaffold structure during the freezing step. These ice crystals act as space-holders or porogens. During the sublimation step, the space previously occupied by the ice becomes interconnected pores. As it was expected, the fabricated scaffolds had porosity with surface roughness, which are necessary for cell migration, vascularization and transport of nutrients and waste products. The rough surfaces of scaffolds also play a role on cell anchorage, flattening, spreading and structural organization [39]. Here, it was found that the surface roughness of the scaffolds varied depending on the composition. The G-Ca-P and G-Ca-DP-P scaffolds exhibited rougher surfaces compared to the other scaffolds. The G-P scaffolds had a lower roughness value; a smoother surface than the G-Ca-P and G-Ca-DP-P scaffolds, but rougher than the G scaffolds. Furthermore, the wettability results indicated that all the scaffolds exhibited hydrophilic behavior, (CA < 90°). Hydrophilicity is desirable for cell attachment, and when the CA is in the range of 60°−80°, the adhesion of fibroblasts is the highest [39, 40]. The surface wettability is greatly influenced by factors including the surface functional groups, and the material surface roughness. Hydrophilic surfaces (θ < 90°) enhance early protein adsorption, which in turn promotes cell migration, integrin-mediated adhesion, and proliferation [37, 41]. In tissue regeneration contexts, the hydrophilic surfaces can provide a balance between favorable surface energy for protein binding and structural retention of the scaffold. Notably, the G-Ca-P (θ = 59.21°) and G-Ca-DP-P (θ = 77.73°) scaffolds fall within the ideal wettability range, suggesting they offer a favorable biointerface for DPSCs attachment, migration, and eventual matrix mineralization.

Biodegradability of the scaffold is an important aspect when developing tissue engineered scaffolds. Following the scaffold implantation, it must degrade in a timely manner to coincide the remodeling of repairing tissue [42]. The in vitro biodegradation test, here, showed that all scaffolds degraded over time. Overall, G-Ca-DP-P offered the lowest long-term structural degradation. The incorporation of dentin matrix proteins into the scaffold (G-Ca-DP-P) effectively delayed degradation, suggesting enhanced structural integrity and potential for prolonged in vivo function. This makes it highly suitable for applications like bone tissue engineering, where sustained scaffold presence is critical. In hard tissue engineering, the biodegradation rate of scaffolds plays a significant role in balancing structural support with tissue regeneration. Too fast biodegradation rate may collapse the porous structure of the scaffold, hindering mass transfer and resulting in tissue necrosis, while too slow biodegradation, may lead to the fibrous capsule formation and absence of scaffold integration with the host tissue [43]. Faster-degrading scaffolds, including those incorporating natural polymers like gelatin and bioactive agents like calcium hydroxide, can provide early-stage space maintenance and release of bioactive cues. These are particularly advantageous where rapid cellular infiltration and remodeling are desirable such as pulp-dentin complex regeneration or non-load-bearing bone defects. Conversely, slower-degrading scaffolds are often more appropriate for load-bearing applications, where prolonged mechanical stability is essential to support the developing tissue until it fully matures.

Gelatin, a denatured form of collagen protein, degrades either enzymatically for example by collagenases, or matrix metalloproteinases, or hydrolytically [44, 45]. The degradation products of gelatin include amino acids, and peptides, which are biocompatible, and can even promote angiogenesis and bone regeneration [46]. Furthermore, DMPs include non-collagenous proteins such as dentin matrix protein 1 (DMP1), dentin sialophosphoprotein (DSPP), and Osteopontin (OPN), and it can degrade enzymatically, producing peptides and other bioactive fragments, some of which may support odontogenic or osteogenic differentiation, making them beneficial degradation products [47]. Propolis is a complex resin-like material produced by honeybees comprising of flavonoids, phenolic acids, and essential oils [8]. It releases these components, which can have antioxidant, anti-inflammatory, and antibacterial effects [48]. However, the non-toxicity and bioactivity may depend on release concentration. Calcium hydroxide also dissociates in aqueous solutions, particularly under physiological conditions, and produces calcium (Ca²⁺) and hydroxide (OH⁻) ions, causing pH increase (as observed here), supporting mineralization and enhance antibacterial effects [49]. However, excessive alkalinity may cause cytotoxicity.

When the scaffolds immersed in PBS, an alkaline shift was observed in the G-Ca-P and G-Ca-DP-P groups, which can be indicative of bioactive behavior, contributing positively to osteogenic differentiation (as observed here in osteogenic differentiation assays), antibacterial activity, and remineralization processes [50, 51]. The increase in pH is attributed to the dissolution of calcium hydroxide, which releases hydroxide ions [52]. The more pronounced pH increase in the medium of G-Ca-DP-P scaffolds suggests that the presence of dentin matrix proteins may synergize with Ca (OH)₂, probably through the interactions influencing the release of calcium ions. In contrast, the G and G-P groups displayed negligible pH change, demonstrating no ion exchange and limited bioactivity.

Among the studied scaffolds, G and G-P showed an elastic-plastic behavior. While G-Ca-P, and G-Ca-DP-P scaffolds were brittle, because the Ca (OH)₂ is a mineral and its addition to the scaffold composition causes brittleness. The highest strength was related to G-P scaffolds. However, the G-Ca-P and G-Ca-DP-P scaffolds had comparable UCS and E. The G scaffolds showed the lowest Young’s modulus with highest flexibility. Other studies also measured the mechanical properties of gelatin and gelatin-based scaffolds. For example, Wu et al. [53] prepared aligned porous gelatin scaffolds using unidirectional freeze-drying method and measured their compressive strengths. Their results showed that the compressive strength increased from 27.5 to 227.4 KPa in the longitudinal direction and from 8.5 to 45.6 KPa in the transverse direction by increase in gelatin concentration from 1 to 5%. To make these scaffolds glutaraldehyde (5%) was added to the gelatin solution. In another study, gelatin foams and their composite with hydroxyapatite were fabricated by freeze-drying and crosslinking [54]. The authors found the compressive strength of around 200 KPa for 5% gelatin solution crosslinked by immersing into acetone-water-based solution (10%) containing 0.7% w/v water-soluble carbodiimide derivative. Khajavi et al. also fabricated nano-hydroxyapatite/gelatin scaffolds by freeze-drying technique and obtained UCS and E of 16.01 KPa and 303.98 KPa, respectively [17]. The differences in the strength of freeze-dried gelatin scaffolds can be related to the crosslinker material, concentration of crosslinker and gelatin solution, and the way the crosslinker is added to the gelatin solution. The mechanical properties of these scaffolds suggests that they may assist bone repair at low load-bearing locations.

The cell viability at early and late exposure to the scaffolds showed that all scaffolds were not cytotoxic with high viability% (> 80%) at all time points. Indeed, cell viability was notably elevated at day 3 (over 100%), which suggests a strong initial cell proliferation and metabolic activation in response to the scaffolds. By day 7, viability declined to > 80%, indicating a reduction in cell metabolic activity over time. This decrease can be attributed to cell adaptation to the culture environment, that is scaffold. It is believed that the bioactive substances in the scaffold compositions elevated the cell viability at day 3, which may be associated with the stimulatory effects of these bioactive agents on cell growth and metabolism during the early stages. These materials regularly promote accelerated cellular responses shortly after exposure, resulting in enhanced viability. Nevertheless, as the cells adapt to the 3D scaffold environment, the early stimulatory effect might subside. This could explain the observed reduction in cell viability by day 7, indicating a change from a proliferative state to a more stabilized or plateaued phase of cellular activity. Similar trend in the viability of different cells exposed to various biomaterials has been reported in the literature [55, 56]. Furthermore, several studies analyzed the cell viability for gelatin and gelatin-based scaffolds, and found no cytotoxicity associated with these scaffolds [54, 57–59], which confirms the results obtained here. Regarding the cell differentiation, only the G-Ca-P and G-Ca-DP-P scaffolds exhibited early (5 days) cell differentiation. However, in long-time culture (10 days), all scaffolds showed a degree of osteogenic differentiation. Nevertheless, it appeared that G-Ca-DP-P scaffolds could better maintain its support for differentiating of DPSCs. Furthermore, ARS and matrix mineralization results indicated that G-Ca-P and G-Ca-DP-P were highly effective in DPSCs mineralization process (observed intense red stain in large area) owing to their Ca and dentin matrix proteins content. The G-P group induced more mineralized matrix compared to G group, as the slight red stain was observed in G-P scaffold group, suggesting that propolis could help in cell mineralization, but in lower extent than Ca (OH)₂ and DMPs. The elevated ALP concentration observed in the G-Ca-DP-P scaffolds is likely due to the synergistic biological effects of calcium hydroxide, DMPs, and propolis, which together create a favorable microenvironment for osteo/odontogenic differentiation of DPSCs. Several factors may account for the enhanced ALP activity in this group. Ca(OH)₂ steadily releases Ca²⁺ ions, which are known to upregulate the expression of ALP and initiate mineral deposition through early osteogenic differentiation and various intracellular signaling pathways, such as MAPK [60]. DMPs are rich in DMP-1 and DSPP, which are potent osteoinductive factors. These have been reported to induce ALP expression and matrix mineralization through activating signaling cascades that regulate osteoblast- and odontoblast-specific gene expression, including ALP [61, 62]. Propolis contains polyphenolic compounds, which can upregulate early osteogenic markers (like Runt-related transcription factor 2 (RUNX2) and ALP) and enhance matrix mineralization [63, 64]. Askari et al. fabricated a combined matrix of poly(lactide-co-glycolide) (PLGA) and propolis extract, which showed higher calcium content, ALP activity, and expression of bone-related genes including RUNX-2, type I collagen, osteocalcin, and osteonectin in human adipose-derived mesenchymal stem cells (AD-MSCs) differentiated at different time points, in comparison with PLGA scaffold [65]. Therefore, the combination of these three components in the G-Ca-DP-P scaffold possibly provided a sustained release of biologically active cues and enhanced the interactions between DPSCs and scaffold material, causing an improved differentiation profile over time. This was also confirmed by the ARS staining results, showing more extensive mineral deposition in the G-Ca-DP-P group compared to others. In a study by Soares et al. [66], a calcium hydroxide-mediated porous chitosan-calcium scaffold was fabricated for dentin tissue engineering. In their study, the chitosan powder was dissolved in 2% acetic acid aqueous solution and calcium hydroxide (1% w/v) was prepared in deionized water. After dropwise mixing of the calcium hydroxide solution with the chitosan solution and pouring the final solution into Teflon molds, freeze-drying was done overnight. The results showed higher viability, and ALP activity of dental pulp cells exposed to chitosan-calcium scaffolds in comparison with chitosan scaffolds. Similar results obtained in our investigation. In another study, 3D-printed porous scaffolds were fabricated using propolis-modified wollastonite [28]. The scaffolds were assessed for cell proliferation and osteogenic differentiation using Human Bone Marrow Stem Cells (bmMSCs). Promising cell proliferation and metabolic activity along with osteogenic capacity after 21 days were observed. The G-Ca-P and G-Ca-DP-P scaffolds, in our study, also showed acceptable cell (DPSCs) viability, higher ALP activity and mineralization.

Cell adhesion is a crucial factor in success of the developed scaffolds in tissue engineering applications, as it determines the ability of cells to attach and proliferate on the scaffold surface. Our findings suggested that the studied scaffolds have good biocompatibility and support cell adhesion, particularly G-Ca-DP-P scaffolds with higher number of adhered cells. This can be attributed to its chemical composition, presence of dental matrix proteins, and higher surface roughness of G-Ca-DP-P scaffolds, as these factors affect the interaction of the cells with a biomaterial surface and cell adhesion [67]. The observation that DPSCs viability was higher in the G and G-P groups after three days, despite lower cell adhesion, can be ascribed to differences in metabolic activity versus physical attachment of these cells to the biomaterial surface. These are measured through different biological processes. Here, MTT viability assay was used to primarily evaluate the mitochondrial metabolic activity of DPSCs, which does not always directly correlate with the number of cells physically adhered to the surface of scaffold. The G and G-P groups (composed of gelatin and gelatin–propolis) promoted early metabolic activation because of their biocompatibility and the antioxidant properties of propolis, leading to raised viability signals. However, their surface chemistry, surface topography and roughness could not enhance cell binding in the early phase. This could describe the lower number of cells physically adhered in these scaffolds despite high cell viability. G and G-P scaffolds showed lower surface roughness compared to G-Ca-P and G-Ca-DP-P scaffolds. Therefore, these scaffolds possibly had less favorable microstructural features for stable initial adherence, which could delay the DPSCs adhesion, while still enabling metabolic activity of loosely adhered or non-adherent cells.

The present study was a preliminary investigation on the development of 3D porous scaffolds using biocompatible and bioactive materials including gelatin, calcium hydroxide, propolis, and DMPs. The feasibility of the scaffold fabrication, and assessment of their physicochemical and mechanical properties along with their effect on the viability and differentiation of dental pulp stem cells were done. The use of G-Ca-DP-P scaffolds can be promising in dental and bone tissue engineering. Collectively, the G-Ca-DP-P scaffolds showed hydrophilicity, which supported the cell adhesion, and biodegradability beyond 42 days. Furthermore, these scaffolds caused the growth, osteogenic differentiation, and matrix mineralization of dental pulp stem cells. The mechanical properties of the G-Ca-DP-P scaffolds could balance the degradation with structural support, to some extent. However, their mechanical property may limit their use in load-bearing applications, necessitating further optimization for clinical translation. Furthermore, long-term mechanical stability and integrity beyond 42 days would be beneficial to study in future research, as the tissue remodeling in dental and bone contexts generally requires several weeks to months. To overcome these, one way is the incorporation of biomaterials with higher mechanical strength and lower biodegradation rates. Examples are synthetic polymers such as poly lactic acid (PLA) and poly caprolactone (PCL) with pore design via advanced manufacturing techniques like three-dimensional printing [68–70]. Furthermore, it would be interesting to analytically study the degradation products of the developed scaffolds to ensure about the safe concentrations of calcium ions, pH-modulating species, and residual propolis polyphenols. Another point that should be considered, is the sterilization process, an important step in the biomaterial preparation for clinical use, which may influence the physicochemical and biological properties of scaffolds. Here, we used UV radiation. It would be beneficial to assess the impact of different sterilization techniques (such as gamma irradiation, ethylene oxide, or plasma) on the scaffold integrity and functionality, in future work. Furthermore, we believe that further studies are needed to investigate the differentiation ability of the studied scaffolds through osteogenic gene expression and by molecular analyses to identify the underlying mechanisms. In vivo studies using orthotopic dental or craniofacial models are also necessary to evaluate tissue regeneration and potential systemic immune and inflammatory responses. Regarding potential clinical translation, the scaffolds can be employed either without cells or by recruitment of cells, depending on the clinical scenario. The inclusion of bioactive agents such as calcium hydroxide, DMPs, and propolis supports cell growth, differentiation, and mineralization. However, for more controlled and enhanced outcomes in complex or compromised cases, cell-seeded G-Ca-DP-P scaffolds with autologous or allogenic stem cells may alternatively offer improved regenerative capacity. Preliminary cost considerations for the scaffolds fabricated here suggest that the fabrication process is affordable and easy, and the biomaterials used (gelatin, calcium hydroxide, and propolis) are relatively low-cost and widely accessible. Only DMPs preparation involves multi-step extraction processes from dental tissues, which involves complexity, cost, variability, and regulatory compliance. This may cause challenges in commercial or clinical scalability. Nevertheless, a thorough cost-benefit analysis and economic practicality assessment will be required to support clinical translation.

Conclusion

This study reports an approach to modifying the chemical composition of gelatin scaffolds by incorporating a calcium hydroxide, propolis and dental matrix proteins using freeze-drying technique. The fabricated G-Ca-DP-P scaffolds were porous, rough, hydrophilic and biodegradable. The mechanical properties of these scaffolds suggests that they may assist bone repair at low load-bearing locations. The G-Ca-DP-P scaffolds are non-toxic as they induced human dental pulp stem cell viability at early (> 100%) and late (> 80%) stages. Furthermore, these scaffolds supported cell adhesion and provided an environment, which was able to positively modulate osteogenic differentiation of human dental pulp stem cells evidenced by higher ALP concentration and matrix mineralization. Overall, the porous G-Ca-DP-P scaffolds developed here, seems to be a viable alternative for hard tissue (tooth and bone) repair.

Abbreviations

AD-MSCs

Adipose-derived mesenchymal stem cells

AFM

Atomic Force Microscope

ALP

Alkaline Phosphatase

ANOVA

Analysis of Variance

ARS

Alizarin Red Staining

bmMSCs

Bone Marrow Stem Cells

BMP

Bone Morphogenetic Protein

BTE

Bone Tissue Engineering

CA

Contact Angle

DMEM

Dulbecco’s Modified Eagle Medium

DMPs

Dentin Matrix Proteins

DMP1

Dentin Matrix Protein 1

DPSCs

Dental Pulp Stem Cells

DSPP

Dentin Sialophosphoprotein

DTE

Dental Tissue Engineering

ECM

Extracellular Matrix

FBS

Fetal Bovine Serum

FE-SEM

Field Emission Scanning Electron Microscopy

OPN

Osteopontin

HTE

Hard Tissue Engineering

PBS

Phosphate-buffered Saline

PCL

Poly Caprolactone

PLA

Poly Lactic Acid

PLGA

Poly(lactide-co-glycolide)

RMS

Root Mean Square

RUNX2

Runt-related transcription factor 2

SEM

Scanning Electron Microscopy

TE

Tissue Engineering

UCS

Ultimate Compressive Strength

UTM

Universal Testing Machine

3D

Three-dimensional

Author’s contributions

•M. J. S: conceptualization, design of experiments, provision of study materials, reviewing•M. T: data analysis, draft preparation, reviewing •E. S: provision of study materials•M. B: conceptualization, design of experiments, data analysis, experimentation, draft preparation, and writing and editing.

Funding

Not applicable.

Data availability

The datasets used and/or analyzed during the current study available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

Human dental pulp stem cells (DPSCs) used in the present study were purchased from the Royan Institute for Biotechnology (Isfahan, Iran), a leading institution in stem cell research in Iran. The Royan Institute complies with the ethical guidelines established by the Iran National Committee for Ethics in Biomedical Research, which align with the principles of the Declaration of Helsinki, ensuring that all human-derived materials, including DPSCs, are obtained with informed consent from donors. This study was conducted in accordance with the Declaration of Helsinki and all protocols was approved by the institutional ethics committee at Semnan University of Medical Sciences (IR.SEMUMS.REC.1402.019).

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.