Addition of Bone‐Marrow Mesenchymal Stem Cells to 3D‐Printed Alginate/Gelatin Hydrogel Containing Freeze‐Dried Bone Nanoparticles Accelerates Regeneration of Critical Size Bone Defects

Abstract

A 3D‐printed biodegradable hydrogel, consisting of alginate, gelatin, and freeze‐dried bone allograft nanoparticles (npFDBA), is developed as a scaffold for enhancing cell adhesion, proliferation, and osteogenic differentiation when combined with rat bone marrow mesenchymal stem cells (rBMSCs). This composite hydrogel is intended for the regeneration of critical‐sized bone defects using a rat calvaria defect model. The behavior of rBMSCs seeded onto the scaffold is evaluated through scanning electron microscope, MTT assays, and quantitative real‐time PCR. In a randomized study, thirty rats are assigned to five treatment groups: 1) rBMSCs‐loaded hydrogel, 2) rBMSCs‐loaded FDBA microparticles, 3) hydrogel alone, 4) FDBA alone, and 5) an empty defect serving as a negative control. After 8 weeks, bone regeneration is assessed using H&E, Masson's trichrome staining, and immunohistochemistry. The 3D‐printed hydrogel displays excellent adhesion, proliferation, and differentiation of rBMSCs. The rBMSCs‐loaded hydrogel exhibits comparable new bone regeneration to the rBMSCs‐loaded FDBA group, outperforming other groups with statistical significance (P‐value < 0.05). These findings are corroborated by Masson's trichrome staining and osteocalcin expression. The rBMSCs‐loaded 3D‐printed hydrogel demonstrates promising potential for significantly enhancing bone regeneration, surpassing the conventional clinical approach (FDBA).

The substantial upregulation of the osteocalcin marker indicates a notable enhancement in new bone regeneration facilitated by the presence of rat bone marrow mesenchymal stem cells within the 3D printed hydrogel, particularly in critical‐sized bone defects. These results underscore the promising potential of this construct as a reliable tissue‐engineered scaffold with promising clinical applicability.

1. Introduction

3D printing has emerged as a prominent technology, facilitating the advanced development of bio‐scaffolds made from hydrogels for applications such as drug screening,^{[ 1 ]} tissue engineering,^{[ 2 ]} and regenerative medicine.^{[ 3 ]} This process involves using a computer‐aided design model to guide the layer‐by‐layer deposition of bio‐ink onto a platform, resulting in the creation of intricate 3D tissue structures characterized by a porous architecture.^{[ 4 ]} Tissue‐engineered scaffolds produced through 3D printing offer numerous advantages, including the ability to achieve intricate and customizable designs, regulate pore sizes and degradation rates, and facilitate personalized scaffold construction.^{[ 5 ]} As a result, there has been a notable surge of interest in employing this technique for bone tissue engineering applications.^{[ 6 , 7 ]}

Hydrogels offer numerous advantages for tissue engineering due to their favorable physicochemical properties. These include the potential for drug loading, suitable biodegradability, high water content (typically exceeding 90%), and excellent biocompatibility.^{[ 8} , ⁹ , ^{10 ]} The microstructure of the native extracellular matrix (ECM), which provides a suitable pattern for cellular ingrowth, can be effectively mimicked by the 3D hydrophilic cross‐linked structure of hydrogels. Additionally, the hydrophilic properties of hydrogels can influence cell adhesion, proliferation, and differentiation processes.^{[ 11} , ^{12 ]}

Alginate, a natural polysaccharide, has garnered significant attention in biomedical engineering and 3D printing due to its unique properties, such as biocompatibility, biodegradability, ease of printing, and cross‐linking feasibility.^{[ 13 ]} However, one limitation of alginate is its inability to facilitate cell attachment, primarily attributed to the absence of mammalian cell‐adhesive ligands. To address this, the incorporation of gelatin, another biocompatible and biodegradable polymer, has proven beneficial. This addition not only enhances printability but also augments cell‐ink interactions through the introduction of RGD sequences.^{[ 14 ]} The combination of gelatin and alginate has been widely explored in prior research. Distler et al. utilized a gelatin/alginate composition laden with mouse myoblast cells (C2C12) for tissue engineering purposes.^{[ 15 ]} They posited that this composition possesses favorable attributes for tissue regeneration; however, they emphasized that the correct selection of nozzle size and extrusion pressure during 3D printing significantly influences cell attachment and viability. Similarly, Pan et al. employed a gelatin/alginate mixture and addressed hydrogel‐related challenges by implementing cross‐linking techniques to enhance mechanical properties.^{[ 16 ]} Furthermore, Gharacheh et al. demonstrated that augmenting a methacrylate alginate bio‐ink with human bone allograft particles leads to improved stem cell osteogenesis. This enhancement was evidenced by elevated alkaline phosphatase activity, calcium deposition, and osteocalcin expression levels.^{[ 17 ]}

In this present study, we embarked on 3D printing two distinct hydrogel bio‐inks containing freeze‐dried bone allograft nanoparticles (npFDBA), each exhibiting varying proportions of gelatin and alginate. These compositions were subjected to comprehensive physicochemical characterization. Subsequently, rat bone marrow‐derived mesenchymal stem cells (rBMSCs) were isolated and cultivated on nanocomposite hydrogel scaffolds. Rigorous in vitro analyses were performed to assess attachment, proliferation, and differentiation. Furthermore, to evaluate osteogenesis capacity, a rat calvaria defect model was employed. This entailed investigating the performance of rBMSC‐loaded 3D printed scaffolds in comparison with standard treatment approaches utilizing human bone allograft, both with and without rBMSCs. The outcome of this study introduces a novel 3D printed hydrogel that exhibits the potential for crafting personalized 3D constructs. This innovation, coupled with the utilization of autogenous mesenchymal stem cells, presents a promising avenue to supplant autogenous bone grafting—the current gold standard treatment for craniomaxillofacial region critical‐sized bone defects such as alveolar clefts, atrophic ridges, and traumatic bone defects.

2. Experimental Section

2.1. Materials

Gelatin (CAS 9000‐70‐8) and calcium chloride dihydrate (CaCl₂, CAS 10043‐52‐4) were procured from Merck Co. (USA), while sodium alginate (A2033, medium viscosity) was sourced from Sigma Co. Ltd. (USA). Human freeze‐dried bone allograft (FDBA) was obtained from Regen Co. (Iran). For all aqueous solutions, deionized (DI) water was employed in the preparation process.

2.2. 3D Printed Hydrogel Scaffold Fabrication

2.2.1. Inks Preparation

The desired bio‐inks, namely Alg10Gel3 and Alg6Gel6, were formulated within two distinct groups as outlined in Table  1 . Despite the differing weight ratios, the preparation procedures remained consistent across both groups.

Table 1.

The bio‐inks compositions.

Inks	Alginate [w/v%]	Gelatin [w/v%]	npFDBA [w%]	DI water [mL]
Alg10Gel3	10	3	25	10
Alg6Gel6	6	6	25	10

In the initial step, the alginate solution was created by dissolving the alginate powder in deionized (DI) water at a temperature of 40 °C, allowing it to stand for 1 h. Subsequently, the freeze‐dried bone allograft (FDBA) micro‐particles were subjected to milling using a planetary ball mill device (Amin Asia Fanavar Pars Co, Iran) to obtain nanoparticles (npFDBA). The nanoparticle size was determined utilizing dynamic light scattering technique employing a Zetasizer Nano (Malvern Instruments, Westborough, MA). The npFDBA was then individually dispersed by incorporating the powder into DI water under the same conditions. Following a 1‐h duration, gelatin powder was introduced into the npFDBA solution and stirred for an additional hour. The prepared solutions from both components were amalgamated and thoroughly mixed for a period of 2 h. Subsequently, the final prepared ink was transferred into a sterile syringe and centrifuged for 1 min at 1000 revolutions per minute (RPM). The resulting solution was then placed in a refrigerator set at 4 °C for storage.

2.2.2. Attenuated Total Reflection–Fourier Transform Infrared

The prepared ink samples underwent an initial drying process, followed by their introduction into the attenuated total reflectance Fourier‐transform infrared (ATR‐FTIR) device (Nicolet 6700, Thermo Fisher Scientific, Waltham, MA, USA). This analytical step enabled the assessment of the functional groups present within the hydrogel.^{[ 18 ]}

2.2.3. Rheological Analysis

Rheological analysis was conducted using an MCR Rheometer (Anton Parr, Germany) operating in a rotational shear mode. The samples were introduced between two parallel plates (PP‐50) set at a gap size of 0.1 mm. Subsequently, measurements were taken at shear rates spanning from 0.1 to 1000 S−1, all maintained at a consistent temperature of 37 °C.

2.2.4. 3D‐Printing

The 3D printing process was executed using a 3DPL designed 3D printer (N2+, Tehran, Iran).^{[ 19 ]} The scaffold designs were tailored to incorporate macropores measuring 1000 × 1000 µm. Printing operations were conducted at a temperature of 25 °C and a pressure of 2.45 bar, employing a 21‐G nozzle. In order to cross‐link the fabricated scaffolds, the printed structures were immersed in a solution of CaCl₂ (100 mm) for a duration of 3 h.

2.3. Physicochemical Characterization of the 3D Printed Scaffolds

2.3.1. Morphology of the 3D Printed Scaffold by FE‐SEM Analysis

The morphology and microstructure of the 3D printed scaffolds were analyzed using a field emission scanning electron microscope (FE‐SEM, Vega, Tescan, Philadelphia, PA). To facilitate this analysis, a thin layer of gold was sputtered onto the surface of the samples using an EMITECH K450X sputtering device (UK).

2.3.2. Swelling Test

The swelling capacity of hydrogels in contact with biological fluids is a crucial characteristic for their application in tissue engineering. To assess this property, the 3D printed scaffolds produced from Alg10Gel3 and Alg6Gel6 were initially dried in a 60 °C incubator. Subsequently, they were immersed in phosphate‐buffered saline (PBS) with a pH of 7.4 and maintained at 37 °C in the incubator. After specific time intervals of 20, 40, 60, 80, 100, and 120 min, the hydrogels were taken out from the solution and their weights were measured after gently removing surface water with a filter paper.^{[ 20 ]} The swelling ratio was then calculated using the following equation, where W0 and W1 represent the initial and swollen weights of the hydrogels at different time points.^{[ 21 ]} This test was performed three times for each scaffold.

$$\mathrm{Swelling}\mathrm{ratio}\left(\%\right)=\left(\mathrm{W}1-\mathrm{W}0\right)/\mathrm{W}0\times 100$$ (1)

2.3.3. Degradation and pH Changes

To assess the in vitro degradation rate of the 3D printed scaffolds, a PBS medium maintained at 37 °C was employed. For each group, three samples of equal volume were submerged in 10 mL of PBS medium with a pH of 7.4. At various time intervals, the samples were taken out of the medium, filtered, rinsed with distilled water, and weighed after carefully eliminating surface moisture using a filter paper. The weight loss percentage was determined using the subsequent formula, where W0 represents the initial weight of the samples and W1 signifies the dry weight after removal from the environment.^{[ 22 ]}

$$\mathrm{Weight}\mathrm{loss}\left(\%\right)=\left(\mathrm{W}1-\mathrm{W}0\right)/\mathrm{W}0\times 100$$ (2)

Concurrently with the sample weighing process, the pH alterations of the medium were gauged.

2.4. In Vitro Biocompatibility and Osteogenesis of the 3D Printed Scaffolds

2.4.1. Isolation, Culture, and Characterization of rBMSCs

All procedures involving the isolation and cultivation of rBMSCs followed established protocols from previous clinical studies.^{[ 23 ]} Initially, anesthetization of the donor rat (8 weeks old, weighing 200 g) was carried out using an injection of 10% ketamine (100 mg k⁻¹ g) and 2% xylazine (5 mg k⁻¹ g). The bone marrow tissue was extracted and washed with normal saline within 100 mL plates to eliminate blood. The tissue was then placed on a new plate and sectioned into small fragments using a sterile scalpel. Subsequently, the divided tissue pieces were transferred to a 50‐mL Falcon tube containing collagenase I. This Falcon tube was placed inside a CO₂ incubator and gently agitated. After 90 min, the tube was removed from the incubator, the tissue was neutralized using collagenase serum, and the digested tissue was subjected to centrifugation at 1500 rpm for 5 min at room temperature. The supernatant was discarded, and the pellet of adherent cells was rinsed thoroughly with PBS. Finally, the isolated cells were introduced into a T75 flask with alpha‐MEM as the culture medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (Gibco, USA). The flask was maintained at 37 °C with 5% carbon dioxide. Following a 2‐day incubation for mesenchymal cell adhesion, the medium on the cell surface was replaced, and the cells were incubated until they reached an appropriate passage number. Cells at Passage 3 were employed for subsequent analyses.

2.4.2. Cell Attachment Assay

The scaffolds were examined for morphology and microstructure with a scanning electron microscope (FE‐SEM, Vega, Tescan, Philadelphia, PA). The scaffolds were soaked into 5 mL of culture medium supplemented with 10% FBS. After 4 h of incubation in the incubator, 60 000 cells were cultured on each scaffold. After 48 h of incubation at 37 °C, 5% CO₂, and 95% humidity, the medium was slowly removed and the following procedure was carried out. In brief, the samples were immersed respectively in a solution of 2.5% glutaraldehyde dissolved in 0.1 m PBS for 4 h, and 0.1% solution of osmium tetroxide (OsO₄) was dissolved in 0.1 m PBS for 30 min. Between each step, the scaffolds were washed three times with PBS solution. In the last step, dehydration was done with the ethanol percentages of 50%, 60%, 70%, 80%, 90%, 100% (one time each for 20 min). Then, the samples were put in a freezer at −80 °C for 12 h, and then, they were immediately placed in a freeze dryer with a pressure of 0.06 mbar and a temperature of 57 °C overnight. The samples were placed in the vicinity of silica gel to avoid moisture absorption. Before SEM analysis, first, the surface of the samples was covered by a thin layer of gold with a sputtering device (EMITECH K450X, UK).

2.4.3. Cell Viability and Proliferation Assay

The impact of the nanocomposite hydrogel on rBMSCs proliferation was assessed utilizing the standard colorimetric MTT assay (3‐(4,5‐dimethylthiazol‐2‐yl)−2,5‐diphenyltetrazolium bromide).^{[ 24 ]} To this end, cells at Passage 3 were cultured at a density of 100 000 cells on each scaffold. Viability and proliferation of rBMSCs were gauged after 1, 3, and 7 days of incubation. MTT solution (Sigma Aldrich) was mixed with fresh culture medium at a ratio of 1:10, and the resulting solution was added to the well containing the samples. The culture was maintained in the incubator for a 2‐h duration. Subsequently, the solution was replaced with dimethyl sulfoxide (DMSO) solution (Sigma Aldrich), which was agitated for 10 min. The optical density (OD) of the samples was measured at a wavelength of 570 nm using a microplate reader (Synergy HT, BioTek, USA). Additionally, a control analysis was conducted on cell cultures without the presence of scaffolds for comparative purposes.

2.4.4. Osteogenic Differentiation Study

The expression levels of Runx2, ALP, and OCN genes were assessed through quantitative real‐time PCR. Scaffolds loaded with rBMSCs were cultured in an osteogenic medium for a span of 14 days. To extract RNA from the cells cultivated on the 3D printed scaffolds, a total RNA isolation kit (Takara, Xi'an, China) was utilized. Reverse transcription (RT)‐PCR primers for osteogenic genes are outlined in Table  2 . The preparation of cDNA was carried out using a reverse transcription kit (Takara, Xi'an, China). Subsequently, a quantitative real‐time PCR detection system (Takara, Xi'an, China) was employed for q‐rtPCR with Fast Start Universal SYBR Green Master Mix (Takara, Xi'an, China), following the temperature profile of 95 °C for 10 min, 95 °C for 15 s, and 60 °C for 1 min. The specificity of PCR products was confirmed via melting curve analysis. For gene expression quantification, the 2^−ΔΔCt method was applied, using glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) as the internal control. Each experiment was performed in triplicate with three samples.

Table 2.

Primer sequences for target genes.

Gene	Forward primer	Reverse primer
OCN	GAGGGCAGTAAGGTGGTGAATAG	AGAGAGAGGGAACAGGGAGGGT
ALP	CTTTTGGACAGCAGGGTGGG	AAGGAGGGTTGGGTTGAGGGA
Runx‐2	GCCGTAGAGAGTAGGGAAGAC	CCACAAGGTGCCAGGAATG
GAPDH	AAGTTCAACGGCACAGTCAAGG	CATACTCAGCACCAGCATCACC

2.5. In Vivo Implantation

2.5.1. Ethics Considerations

The present study was duly registered with the Ethics Committee of the Faculty of Dentistry, Shahid Beheshti University of Medical Sciences, under the registration code IR.SBMU.DRC.REC.1398.123. Additionally, the study was conducted in strict accordance with the “Guideline for the Care and Use of Laboratory Animals in Iran.”^{[ 25 ]} The animals were maintained in an environment adhering to all principles governing the housing of laboratory animals.^{[ 26 ]} In essence, the animal housing facility was outfitted with automated air conditioning, heating, cooling systems, adjustable lighting, and sound insulation. Notably, the section dedicated to rat housing was segregated from the surgical area, and regular monitoring of humidity and temperature was upheld. The animals enjoyed unrestricted access to water and food. Surgical procedures were executed under rigorous aseptic conditions.^{[ 27 ]}

2.5.2. Surgical Procedure

A total of thirty male albino rats weighing between 200 and 250 grams were subjected to general anesthesia through the administration of Ketamine hydrochloride (100 mg k⁻¹ g, Alfasan, The Netherlands) and Xylazine hydrochloride (10 mg k⁻¹ g, Alfasan, The Netherlands).^{[ 28 ]} Following a surgical incision on the skin and periosteum, a critical‐sized bone defect measuring 8 mm in diameter was meticulously created on the calvaria bone. The rats were then randomly divided into five distinct groups as follows: 1) Hydrogel + Cell, involving rBMSCs seeded Alginate (6 w/v%)/gelatin (6 w/v%)/npFDBA (25 w%) hydrogel; 2) FDBA + Cell, involving rBMSCs seeded with FDBA microparticles (250–500 µm, Regen co, Tehran, Iran); 3) Hydrogel, consisting of the hydrogel scaffold; 4) FDBA microparticles, comprising only FDBA particles; and 5) negative control, represented by an empty defect as a model. Both scaffolds and FDBA microparticles were soaked in DMEM 48 h prior to surgery, and cell seeding was performed in accordance with the protocol. The surgical site was meticulously sutured in a bilayer manner with 3‐0 absorbable Vicryl suture (Vicryl, ETHICON) for the periosteum and 3‐0 non‐absorbable nylon suture (SUPA, Iran) for the skin. To prevent postoperative hypothermia, the rats were placed on a heating mat during the recovery phase. Following the surgery, all animals received subcutaneous doses of antibiotics (Enrofloxacin (Baytril), 10 mg k⁻¹ g per day for 3 days) and analgesics (Ketoprofen (Ketofen), 5 mg k⁻¹ g, administered once every 24 h for 5 days).

2.5.3. Histology and Immunohistochemistry

After a duration of 8 weeks, the animals were humanely euthanized by intracardial injection of gallamine triethiodide (1 mg k⁻¹ g, Specia, Paris, France) subsequent to the induction of general anesthesia.^{[ 7 ]} The calvaria bones were then harvested and promptly immersed in a 10% formalin solution for a period of 24 to 72 h. Following this, the samples underwent decalcification in a 10% EDTA solution over a span of 4 weeks. Subsequently, the samples, embedded in paraffin, were meticulously sectioned. Serial cross‐sections of the decalcified samples were subjected to both H&E staining and Masson's trichrome staining in accordance with the instructions of the manufacturer. The resulting images were captured utilizing a light microscope (Olympus BX51; Olympus, Tokyo, Japan). The extent of new bone formation was quantified through the utilization of ImageJ software. The number of osteoblasts, osteocytes, and osteoclasts, as well as the proportion of newly formed bone tissue, were meticulously calculated and subsequently expressed as a percentage of the tissue area.

Furthermore, immunohistochemical analysis (IHC) was undertaken to evaluate the expression of the mineralization marker, osteocalcin (OCN). Briefly, antigen retrieval was achieved by subjecting deparaffinized sections to citrate buffer in an oven for a duration of 10 min. The slides were subsequently placed within a TBS 1X solution (T5912‐Sigma) and subjected to heating until the boiling point was reached. Following this, the oven was deactivated, and the samples were left in the solution for a duration of 20 min. The slides were then exposed to mouse monoclonal anti‐OCN antibody (OCN; sc‐365797) at a temperature of 4 °C overnight. Subsequent to this incubation, the samples underwent washing with PBS for a total of three times, each lasting for 5 min. The tissue slides were then incubated with Goat Anti‐Mouse (orb688924) for a duration of 1 h at 37 °C. After another round of washing with PBS, a solution of Linker (Diagnostic BioSystems‐PVP1000D) was applied to the samples for a duration of 15 min. Following this, the samples underwent washing with PBS, and a polymer solution (Diagnostic BioSystems‐PVP1000D) was added to the samples for a duration of 30 min. Subsequent to yet another round of washing with PBS, the samples were subjected to a DAB solution (ScyTek‐ACV999). Following a period of 5 min, the samples were washed with deionized water, and subsequently immersed in hematoxylin dye for a duration of 10 s. Post washing with deionized water, and following the steps of dehydration and clarification, the slides were evaluated utilizing a light microscope (BX53; Olympus, Japan).

2.6. Statistical Analysis

The quantitative data were subjected to statistical analysis using SPSS software (version 18; SPSS Inc., Chicago, IL). The data were expressed as mean ± standard deviation. To assess statistical differences among the groups, a one‐way ANOVA was employed, followed by post hoc Tukey's Multiple Comparison Test. A p‐value of less than 0.05 was deemed to be statistically significant.

3. Results

3.1. Physicochemical Characteristics of the Prepared Inks

The npFDBA exhibited an average particle size of 371.3 ± 53.7 nm. The 3D printed scaffolds are depicted in Figure  1A, displaying appropriate height and flexibility to replicate the contours of the alveolar bone defect. The microstructure of the 3D printed composite hydrogel scaffold surface is illustrated in Figure 1B, revealing a consistent pattern with micron‐sized pores. The ATR‐FTIR spectrum of the nanocomposite hydrogel is presented in Figure 1C. Notably, the bands at 3332 cm⁻¹ correspond to free − OH groups. Alginate displays a characteristic carboxyl peak at 1418 cm⁻¹, while gelatin exhibits a distinct amine peak at 1628 cm⁻¹. Significantly, the gelatin hydrogel lacks the characteristic peak at 1543 cm⁻¹ in the spectrum, indicating its involvement in the crosslinking reaction.

Figure 1.

A) The 3D printed Alg6Gel6 scaffold according to a form of bone defect of an alveolar cleft, B) morphology analysis of the Alg6Gel6 scaffold by FE‐SEM, C) ATR‐FTIR of the Alg6Gel6 hydrogel, D) rheological analysis, E) swelling test, F) in vitro degradation, and G) pH changes.

The hydrogel demonstrates thermos‐sensitive properties, and its rheological characteristics were analyzed at 37 °C, the physiological temperature of the human body. The outcomes indicate that the hydrogel inks possess a lower viscous component (loss modulus, G′′) compared to their elastic component (storage modulus, G′) at 37 °C (Figure 1D).

3.2. Physicochemical Characteristics of the 3D Printed Scaffolds

The swelling behavior of the 3D printed scaffolds was explored in PBS medium at 37 °C, and the results are displayed in Figure 1E. After 120 min, the swelling ratios of Alg10Gel3 and Alg6Gel6 scaffolds were recorded as 305.80 ± 4.63% and 349.40 ± 3.19%, respectively. This increase in swelling for Alg6Gel6 compared to Alg10Gel3 can be attributed to the greater extent of alginate cross‐linking in the former.

The in vitro degradation of the 3D printed scaffolds in PBS at 37 °C over a span of 60 days is depicted in Figure 1F. Up to the fourth week, both groups exhibited similar degradation profiles, but after 60 days, the degradation rates for Alg10Gel3 and Alg6Gel6 scaffolds were determined as 74.20% ± 1.84 and 50.08% ± 1.27, respectively. Notably, this difference in degradation rates was statistically significant (P‐value < 0.001).

Concurrently with the assessment of degradation, changes in the pH of the PBS medium were evaluated (Figure 1G). The pH changes observed for both scaffolds fell within the normal physiological range for bodily tissues. The pH value for the Alg10Gel3 scaffold ranged from 7.20 to 7.40, while that for the Alg6Gel6 scaffold ranged from 7.40 to 7.52.

3.3. In Vitro Biocompatibility and Osteogenesis of the 3D Printed Scaffolds

Figure  2A presents rBMSCs observed under a light microscope. The flow cytometry analysis highlighted that the isolated cells expressed the CD105 (endoglin) and CD90 (Thy‐1) markers, indicative of mesenchymal stem cells, while they did not exhibit expression of the CD45 (hematopoietic surface antigens) and CD34 markers (Figure 2B–E).

Figure 2.

A) rBMSCs under light microscopy. Flowcytometric analysis showing rBMSCs were positive for surface markers of B) CD105 and C) CD90, and negative for surface markers of D) CD45 and E) CD34. The red line is the negative control.

The adhesion of rBMSCs to the hydrogel scaffold after 48 h is visualized through FE‐SEM in Figure  3A. Notably, rBMSCs demonstrated favorable attachment on the scaffolds and adopted a well‐oriented configuration. Furthermore, Figure 3B illustrates the viability and proliferation of rBMSCs on the scaffolds on days 1, 3, and 7. Alg6Gel6 displayed a significant increase in attached cell populations in comparison to Alg10Gel3 on the third day (P‐value = 0.015) and the seventh day (P‐value = 0.011). However, there was no significant difference observed on the first day, even though the results for Alg6Gel6 were numerically higher (P‐value = 0.077).

Figure 3.

A) The adhesion of rBMSCs on the scaffold using FE‐SEM analysis, B) the viability and proliferation of rBMSCs seeded on the scaffolds by MTT analysis; * shows significant difference from the Alg10Gel3 scaffold, ** shows significant difference from both Alg10Gel3 and Alg6Gel6 scaffolds. C) The quantitative real‐time PCR analysis for the expression of OCN, ALP, and RUNX2; * shows P‐value was in the range of 0.01–0.05, ** shows P‐value was in the range of 0.001–0.01.

The outcomes of the quantitative real‐time PCR analysis are portrayed in Figure 3C. The rBMSCs seeded on the scaffolds created from Alg6Gel6 exhibited significantly higher messenger RNA expression levels of OCN, Runx2, and ALP compared to ink 1 (P‐value < 0.05). Notably, the OCN, Runx2, and ALP markers were all significantly upregulated for cells cultured on both inks on day 14.

3.4. In Vivo Results

In the evaluation of in vivo bone regeneration efficiency of the 3D printed hydrogel scaffold created from Alg6Gel6, a rat calvaria critical‐sized defect model was employed over a period of 8 weeks. The study proceeded without any observed complications among the rats. Figure  4 showcases H&E staining of the study samples at two magnifications, with arrows indicating the presence of osteoblasts, osteoclasts, and osteocyte cells. This figure also presents areas of healthy and new bone formation.

Figure 4.

Histologic evaluation of the in vivo groups using H&E staining. A) Hydrogel + Cell, B) Hydrogel, C) FDBA + Cell, D) FDBA, E) empty defect (the magnification increases from left to right).

Figure  5A–C provides diagrams indicating the number of osteoblasts, osteocytes, and osteoclasts in the study groups. Notably, the Hydrogel + Cell group demonstrated significantly greater quantities of osteoblasts (P‐value < 0.001), osteocytes (P‐value < 0.001), and osteoclasts (P‐value < 0.01) compared to the Hydrogel and Model groups. In comparison to the FDBA group, Hydrogel + Cell group exhibited significantly higher amounts of osteoblasts (P‐value < 0.05), while no significant differences were noted between these two groups regarding osteocytes and osteoclasts. Additionally, there were no significant differences in the number of cells between the Hydrogel + Cell and FDBA + Cell groups.

Figure 5.

The results of histological evaluation with H&E staining. A) Number of osteoblasts, B) number of osteocytes, C) number of osteoclasts, D) percentage of newly formed bone.

• The model is the same as the empty control group.

• The columns with the same symbols (a, b, c, d) have no statistically significant difference (P‐value > 0.05), but different symbols indicate a significant difference between the columns (P‐value < 0.05).

The percentage of newly formed bone was as follows: Hydrogel + Cell group 32.44 ± 2.67%, FDBA + Cell group 28.23 ± 1.88%, FDBA group 24.18 ± 3.96%, Hydrogel group 15.82 ± 1.16%, and the control group 12.29 ± 1.00%. A diagram illustrating the amount of newly formed bone is presented in Figure 5D. Significantly higher new bone formation was observed in the Hydrogel + Cell group compared to the FDBA (P‐value < 0.05), Hydrogel (P‐value < 0.001), and control groups (P‐value < 0.001). There were no significant differences between the Hydrogel + Cell and FDBA + Cell groups in terms of the percentage of new bone formation, while the FDBA + Cell group showed a near‐significant increase in bone formation compared to the FDBA group (P‐value = 0.059).

Figure  6 displays the Masson's trichrome staining of the study groups, while the osteofibrosis percentages are shown in Figure  7 . The extent of osteofibrosis tissue in the Hydrogel + Cell group was 73.07 ± 2.96%, FDBA + Cell group 74.64 ± 2.49%, FDBA group 62.72 ± 4.73%, Hydrogel group 95.9 ± 9.24 47%, and the control group 38.69% ± 4.90%. Significantly higher osteofibrosis tissue amounts were evident in the Hydrogel + Cell group compared to the Hydrogel (P‐value < 0.01) and control (P‐value < 0.001) groups. Notably, no significant difference was noted between the Hydrogel + Cell and FDBA + Cell groups, which demonstrated similar levels. Additionally, the new bone formation in the FDBA group was slightly lower than the FDBA + Cell group, reaching near statistical significance (P‐value = 0.052).

Figure 6.

Masson's trichrome staining. A) Hydrogel + Cell, B) Hydrogel, C) FDBA + Cell, D) FDBA, E) empty defect (the magnification increases from left to right).

Figure 7.

The osteofibrosis area calculated by Masson's trichrome staining for the study groups.

• The model is the same as the empty control group.

• The columns with the same symbols (a, b, c, d) have no statistically significant difference (P‐value > 0.05), but different symbols indicate a significant difference between the columns (P‐value < 0.05).

Figure  8 illustrates IHC staining for the OCN protein in the study groups, while the expression levels of OCN are depicted in Figure  9 . OCN expression in the Hydrogel + Cell group reached 42.15 ± 2.50%, FDBA + Cell group 46.86% ± 4.25, FDBA group 32.51% ± 2.61%, Hydrogel group 22.97% ± 2.69, and the control group 7.98% ± 3.27%. OCN expression levels in the Hydrogel + Cell and Allograft + Cell groups were nearly equal with no significant difference, and both were notably higher than the FDBA (P‐value < 0.05), hydrogel (P‐value < 0.001), and control (P‐value < 0.001) groups. Furthermore, OCN expression in the Hydrogel group was significantly greater than in the control group (P‐value < 0.01), yet it was significantly lower compared to the FDBA group (P‐value < 0.05).

Figure 8.

IHC staining for OCN protein. A) Hydrogel + Cell, B) Hydrogel, C) FDBA + Cell, D) FDBA, E) empty defect (images from left to right: staining with OCN antibody, DAPI staining and superimposition of the previous images).

Figure 9.

The results of IHC staining for OCN protein.

• The model is the same as the empty control group.

• The columns with the same symbols (a, b, c, d) have no statistically significant difference (P‐value > 0.05), but different symbols indicate a significant difference between the columns (P‐value < 0.05).

4. Discussion

In the context of this study, a 3D printed hydrogel scaffold was successfully engineered and demonstrated its potential for effectively regenerating critical‐sized bone defects in conjunction with the presence of mesenchymal stem cells. Initial efforts involved the fabrication of two types of alginate/gelatin hydrogel compositions containing nanoparticles from human bone allografts, which were subsequently compared based on their in vitro characteristics. Notably, the 3D printed scaffold comprising alginate (6 w/v%)/gelatin (6 w/v%)/npFDBA (25 w%) exhibited more favorable outcomes, particularly in terms of promoting the viability and differentiation of rat bone marrow‐derived mesenchymal stem cells (rBMSCs). Furthermore, this particular scaffold showed remarkable bone regeneration effects when evaluated in vivo.

The conventional approach of utilizing autogenous bone grafting has been associated with various limitations, such as significant bone resorption rates (43.1% within one year),^{[ 29 ]} limited availability of donor bone, and notable complications.^{[ 30 ]} In light of these challenges, there has been a surge of interest in devising strategies to mitigate the need for substantial harvested bone or to minimize secondary resorption, and to explore innovative methods to obviate the necessity for autogenous bone grafts altogether. In this context, the strategic application of regenerative techniques involving cells or growth factors has emerged as a promising avenue to enhance the body's natural healing capacity and reduce donor site complications. As a result, these endeavors have paved the way for the translation of tissue engineering principles from laboratory research to practical clinical applications.^{[ 31 ]}

In the realm of tissue engineering, the selection of appropriate biomaterials for crafting scaffolds holds paramount importance. These biomaterials should possess key attributes, including biocompatibility, biodegradability, and bioactivity. The scaffold assumes a pivotal role in dictating the final architecture and performance of the engineered tissue. Numerous investigations have underscored the favorable qualities of alginate/gelatin hydrogel, particularly in the context of bone tissue engineering. This scaffold combination has exhibited noteworthy potential owing to its capacity to emulate the natural environment required for successful bone regeneration.^{[ 32} , ^{33 ]} To further enhance the mechanical characteristics of the alginate/gelatin hydrogel, the incorporation of nanoparticles has emerged as a promising strategy. Nanoparticles like silicate nanoparticles,^{[ 34 ]} cellulose nanocrystals,^{[ 35 ]} and attapulgite nanoparticles^{[ 36 ]} have been explored for this purpose. In the present study, the addition of npFDBA to the gelatin/alginate composite was aimed at not only bolstering mechanical properties but also fostering an environment conducive to heightened interactions with cells, thereby mimicking a bone tissue microenvironment. Earlier investigations by Khojasteh et al. highlighted the positive responses of mesenchymal stem cells when cultivated on FDBA blocks, emphasizing their potential for bone regeneration.^{[ 37 ]} Furthermore, the inclusion of gelatin, a protein derivative of collagen, which forms a vital structural component of skin, tendons, bones, and vertebrates, can significantly contribute to improved biocompatibility and bioactivity within the construct. Gelatin's presence can particularly enhance the initial adhesion of cells onto the scaffold, promoting a favorable milieu for cellular interactions.^{[ 38 ]}

The ATR‐FTIR results are consistent with the expected absorption peaks of alginate and gelatin. In the case of pure alginate, the absorption peaks at 3203 and 1636 cm⁻¹ signify the presence of hydroxyl (─OH) and carbonyl (─C═O) groups, respectively, within its molecular structure.^{[ 39 ]} Gelatin's absorption peaks at 1633, 1534, and 1236 cm⁻¹ correspond to carbonyl (─C═O), amide (─N─H), and amide III (─C─N) groups, respectively, within the gelatin molecule.^{[ 39} , ^{40 ]} Additionally, the broad absorption peak at 3280 cm⁻¹ in gelatin is indicative of hydroxyl (─OH) and amine (─NH₂) groups within its structure.

The porosity of hydrogel inks plays a pivotal role in influencing cellular behaviors such as attachment, proliferation, and migration.^{[ 41 ]} Pore size and distribution have a significant impact on the ability of cells to penetrate and distribute themselves throughout the hydrogel network.^{[ 42 ]} In our hydrogel scaffold structure, micro‐pores were observed. It is worth noting that non‐cross‐linked alginate/gelatin hydrogels tend to have more surface pores compared to their cross‐linked counterparts.^{[ 36 ]} The introduction of cross‐linking, whether ionic or chemical, tends to reduce the porosity of the composite hydrogel, leading to a significant increase in its mechanical strength. Moreover, electrostatic interactions between nanoparticles and gelatin or alginate may contribute to variations in pore size.^{[ 43 ]}

The rheological properties of the ink state were successfully controlled, allowing for printing at a temperature of 37 °C. Similar findings were observed by Liu et al., who demonstrated that the addition of nanoparticles led to a situation where the loss modulus (G′′′′) of inks was lower than the storage modulus (G′′), regardless of temperature.^{[ 36 ]} This can be attributed to the fact that nanoparticles tend to hinder the movement of polymer chains, resulting in higher G′′ values compared to pure polymer hydrogels.^{[ 35 ]} Furthermore, Gao et al. also reported results in line with our observations. They found that the addition of alginate to the gelatin hydrogel composite led to a decrease in G', and they were able to correlate the values of G' and G′′′′ of alginate/gelatin inks to the required extrusion pressure during the printing process.^{[ 32 ]}

The swelling of hydrogel scaffolds is a critical factor in the context of their biomedical applications. Swelling efficiency directly pertains to the capability of absorbing moisture and maintaining stability within biological systems.^{[ 2} , ^{44 ]} It is firmly established that factors such as the extent of crosslinking, the presence of hydroxyl groups, amorphous regions, and the level of crystallinity profoundly influence the swelling potential of scaffolds. Moreover, the presence of an additional cross‐linking agent enhances the efficacy of cross‐linking in scaffolds, subsequently leading to a reduction in swelling efficiency.^{[ 39 ]} Additionally, in line with the anticipated outcome, the crosslinking effect of calcium chloride on alginate contributes to a higher swelling potential in Alg6Gel6 due to its lower alginate concentration.

The degradability of engineered bone scaffolds stands as a pivotal factor in bone tissue engineering. In this regard, Alg6Gel6 exhibited significantly prolonged degradation, suggesting its potential for maintaining robust mechanical integrity throughout the course of bone regeneration. Giuseppe et al. have previously demonstrated that elevating the alginate concentration up to 5% leads to a substantial enhancement in the mechanical properties of alginate/gelatin hydrogel.^{[ 33 ]} Concurrently, higher alginate concentrations enhance print accuracy while necessitating increased printing pressure. Similarly, a higher concentration of gelatin contributes to improved mechanical attributes of the hydrogel. Moreover, viscosity assumes a critical role in preserving hydrogel stability during the extrusion process, as an increased layering of hydrogel induces augmented pressure.^{[ 45 ]} Giuseppe et al. have indicated that extending the crosslinking duration also constitutes a vital aspect in augmenting the mechanical properties of hydrogel scaffolds.^{[ 33 ]} In our study, the Alg6Gel6 concentration closely approximated the 5% alginate/6% gelatin hydrogel concentration in Giuseppe et al.’s investigation. The optimization index in our case was significantly higher for this concentration, aligning with their study, where it was identified as the optimal hydrogel formulation.^{[ 33 ]}

The cells extracted from rat bone marrow exhibited positive expression of specific markers indicative of mesenchymal stem cells, namely CD90 and CD105. These findings align with the outcomes of a study by Yusop et al., where regardless of cell origin and isolation methodology, rat bone marrow‐derived cells demonstrated the expression of mesenchymal stem cell markers.^{[ 23 ]} Additionally, these isolated cells were confirmed to originate from the mesenchymal lineage and fulfilled the stipulated criteria put forth by the International Society for Cell Therapy (ISCT).^{[ 46 ]} Notably, these cells tested negative for CD45, signifying the successful isolation of rat bone marrow mesenchymal stem cells from the pool of hematopoietic stem cells that are abundantly present within the bone marrow stromal environment. Such a distinction is crucial as it underscores their association with the intraosseous milieu and underscores their differentiation from hematopoietic stem cells, a critical characteristic in defining MSCs.^{[ 47 ]}

Cell adhesion to the ECM is a pivotal determinant influencing the proliferation and differentiation of mesenchymal stem cells. As such, the modification of ECM and the utilization of biomimetic scaffolds can play a pivotal role in enhancing cell adhesion properties, ultimately promoting the process of bone regeneration.^{[ 48 ]} Our SEM analysis has indicated the compatibility of the formulated inks in our study with rBMSCs. This observation is in concordance with the findings of Dutta et al., who illustrated the adhesion of MSCs sourced from human bone marrow to alginate/gelatin hydrogel scaffolds incorporating cellulose nanocrystals.^{[ 35 ]} Furthermore, Dutta et al. demonstrated the proliferation of BMSCs within the scaffold from day 3 to day 7. Similarly, in our study, we also discerned the proliferation of rBMSCs from day 1 to day 3 and subsequently from day 3 to day 7 on both bio‐inks. It is worth highlighting that the proliferation of rBMSCs adhered to Alg6Gel6 exhibited a noteworthy increase compared to Alg10Gel3 at both the 3‐day and 7‐day time points.

During the development of osteoblasts, a diverse array of genes undergoes dynamic activation and deactivation, contributing to the complex process of osteoblastogenesis. These genes encompass a range of factors such as the pivotal transcription factor Runx2, bone morphogenic protein‐2 (BMP‐2), ALP, OCN, osteopontin (OPN), bone sialoprotein (BSP), osteoblast‐specific transcription factor (OSX), and various types of collagen (COL).^{[ 35 ]} In our study, we have effectively demonstrated that the gelatin/alginate/npFDBA hydrogel has the capacity to induce the osteogenic differentiation of rBMSCs in an in vitro environment. This observation is in alignment with earlier investigations^{[ 34} , ³⁵ , ^{36 ]} that have highlighted the potential of similar composite hydrogels in facilitating osteogenic differentiation. Moreover, we observed a significant upregulation in the relative expression of OCN, RUNX2, and ALP genes in rBMSCs seeded onto the Alg6Gel6 scaffold when compared to the Alg10Gel3 scaffold. This enhanced expression of osteoblastic markers provides compelling evidence for the enhanced differentiation of rBMSCs on the Alg6Gel6 scaffold, which contains double the concentration of gelatin as compared to Alg10Gel3. This finding resonates with the outcomes of the study conducted by Lee et al.,^{[ 49 ]} where they illustrated that the concentration of gelatin plays a pivotal role in directing the differentiation of mesenchymal stem cells.^{[ 49 ]} Specifically, they demonstrated that higher gelatin concentrations are conducive to osteogenic differentiation, whereas lower concentrations are favorable for adipogenic differentiation.

The selection of the rat calvaria model for our in vivo study was deliberate, as it offers several advantages that align well with the goals of our research. This model was chosen due to its relevance to craniofacial regeneration, which involves neural crest‐derived cells. By inducing a critical‐sized defect within the skull and facial region, particularly within the calvaria bone, we establish an optimal context for studying in vivo bone regeneration in a rat model.^{[ 50 ]} In this context, a critical‐sized defect refers to a dimensionally significant injury that cannot naturally heal and regenerate with the original tissue over the animal's lifespan.^{[ 51 ]} This type of defect serves as a stringent testbed for assessing the regenerative potential of our engineered hydrogel scaffold. The use of such a model provides insights into the ability of our scaffold to stimulate and facilitate bone regeneration under challenging circumstances, ultimately contributing to the evaluation of its clinical viability for addressing craniofacial bone defects.

Our research highlights the successful in vivo bone regeneration achieved through the strategic combination of rBMSCs with a 3D printed hydrogel scaffold designed for critical‐sized calvaria bone defects. The incorporation of rBMSCs into the 3D printed scaffold yielded significant improvements in key regenerative parameters. Notably, the scaffold seeded with rBMSCs exhibited a substantial increase in the number of osteoblasts, osteocytes, and osteoclasts. Moreover, the percentage of newly formed bone was markedly higher in this group compared to the scaffold alone. The comparable results between the Hydrogel + Cell and FDBA + Cell groups validate the benefits of utilizing hydrogel as a bone regenerative strategy. FDBA microparticles have been established as a standard treatment for bone defect regeneration,^{[ 52} , ⁵³ , ^{54 ]} further emphasizing the promising potential of our hydrogel scaffold in clinical applications. Our findings are in line with existing literature that underscores the enhanced bone formation observed when mesenchymal stem cells are integrated into polymeric scaffolds enriched with hydroxyapatite nanoparticles and gelatin, or gelatin/bioactive glass.^{[ 55} , ^{56 ]} Notably, recent studies have also elucidated the paracrine effects of mesenchymal stem cells, which contribute significantly to tissue healing through intricate interactions with the local microenvironment.^{[ 57 ]} Among these effects, the modulation of macrophage polarization toward the M2 phenotype and favorable reprogramming events has emerged as pivotal factors in tissue engineering.^{[ 58 ]} Thus, in addition to the direct osteogenic capacity of rBMSCs, their paracrine effects could potentially contribute to the observed osteogenesis in our study.

Masson's trichrome staining further corroborated the substantial extent of new bone formation facilitated by rBMSC‐loaded scaffolds. This trend in bone regeneration was consistently observed across all experimental groups. Remarkably, the presence of rBMSCs significantly elevated the amount of osteofibrosis in comparison to the other groups. Furthermore, every treated group displayed noteworthy bone formation when compared to the control group. Our results align with prior investigations that have explored similar themes.^{[ 56} , ⁵⁹ , ^{60 ]} For instance, the study by Liu et al. unveiled the presence of osteoblasts and fully developed bone tissues in defects treated with alginate/gelatin/nanoparticle composite hydrogel 8 weeks post‐surgery.^{[ 36 ]} This congruence in outcomes underscores the reproducibility of our findings within the existing body of related research.

In order to gain a comprehensive insight into the intricacies of the bone healing process, we conducted an in‐depth evaluation of the bone marker Osteocalcin (OCN) through Immunohistochemical (IHC) staining. OCN, a non‐collagenous protein found within bone tissue, serves as a late‐stage marker of osteogenic differentiation.^{[ 36 ]} Thus, the assessment of OCN expression provided us with a deeper understanding of the progress of bone healing across the study groups. In concurrence with our histological observations, the rBMSCs‐loaded hydrogel prominently exhibited significant OCN marker expression in comparison to the Hydrogel, FDBA, and control groups. This concordance between the histological evidence and the OCN expression further underscores the robustness of our findings and the noteworthy potential of the rBMSCs‐loaded hydrogel for enhancing the bone healing process.

In the context of bone formation, it is noteworthy to mention that the FDBA + Cell group exhibited a trend toward a significant increase in bone formation compared to the FDBA group, as indicated by the results of H&E and Masson's trichrome staining (P‐value = 0.059 and 0.052, respectively). This observation gains further significance considering the OCN analysis, which revealed a significant reduction in OCN expression when FDBA was used without cells. This consistent pattern underscores the beneficial impact of incorporating cells, particularly bone marrow mesenchymal stem cells, onto the FDBA scaffold. Our findings align with previous researches wherein the favorable attachment of bone marrow mesenchymal stem cells to FDBA has been demonstrated.^{[ 31} , ^{61 ]} This consistency underscores the potential of FDBA as a substrate for cell attachment and its synergistic effects in promoting bone regeneration.

5. Conclusion

We have successfully developed a biocompatible, biodegradable, and bioactive hydrogel by incorporating npFDBA into an alginate/gelatin ink for 3D printing, with the aim of its application in bone tissue engineering. The 3D printed scaffolds exhibited promising physicochemical properties suitable for tissue engineering applications. Importantly, the presence of the 3D‐printed hydrogel did not negatively impact the viability of rBMSCs, demonstrating its biocompatibility. The hydrogel's ability to promote osteogenesis was evident through a significant upregulation of key osteogenic gene markers, including ALP, OCN, and Runx2. Histological and IHC analyses further confirmed the enhanced regeneration of new bone within critical‐sized bone defects when the 3D printed scaffold was loaded with rBMSCs. Consequently, this novel nanocomposite hydrogel holds great potential as a biomaterial for bone tissue engineering applications. Furthermore, it has the potential to replace the current standard treatment involving autogenous grafting for non‐load‐bearing critical‐sized craniofacial bone defects, such as alveolar clefts. This promising approach should be validated in future human clinical trials.

Conflict of Interest

The authors declare no conflict of interest.

Data Availability Statement

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