Decellularized Bone Matrix-Enriched 3D-Printed GelMA Scaffold as a Cell-Homing Platform: Analysis Using an Artificial Pulp Chamber Model
Short title: 3D-Printed GelMA/Decellularized Bone Matrix
IsabelaSanchesPompeo
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Silva1Vitor
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Toledo Stuani1EsterAlvesFerreiraBordini2
FernandaBalestreroCassiano1
ErikaSoaresBronze-Uhle1
PriscilaToninattoAlves
de
Toledo1CarlosAlberto
de
Souza Costa3DianaGabrielaSoares1,4✉Emaildianasoares@fob.usp.br
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Department of Operative Dentistry, Endodontics, and Dental MaterialsUniversity of São Paulo – USP, Bauru School of Dentistry2Department of Prosthodontics and Dental MaterialsUniversity of São Paulo – USP, Ribeirão School of Dentistry
3Department of Physiology and PathologyAraraquara School of Dentistry, São Paulo State University – UNESPAraraquaraSPBrazil
4Department of Operative Dentistry, Endodontics and Dental Materials, Bauru School of DentistryUniversity of São Paulo - USPAlameda Doutor Octávio Pinheiro Brisolla, 9-75BauruSPBrazil
Isabela Sanches Pompeo da Silva1, Vitor de Toledo Stuani1, Ester Alves Ferreira Bordini2, Fernanda Balestrero Cassiano1, Erika Soares Bronze-Uhle1, Priscila Toninatto Alves de Toledo1, Carlos Alberto de Souza Costa3, Diana Gabriela Soares1
1 Department of Operative Dentistry, Endodontics, and Dental Materials, University of São Paulo – USP, Bauru School of Dentistry
2 Department of Prosthodontics and Dental Materials, University of São Paulo – USP, Ribeirão School of Dentistry;
3 Department of Physiology and Pathology, Araraquara School of Dentistry, São Paulo State University – UNESP, Araraquara, SP, Brazil;
Corresponding author:
Diana Gabriela Soares
Department of Operative Dentistry, Endodontics and Dental Materials
Bauru School of Dentistry, University of São Paulo - USP
Alameda Doutor Octávio Pinheiro Brisolla, 9–75, Bauru - SP, Brazil
Email: dianasoares@fob.usp.br
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ABSTRACT
Objective
The aim of the study was to develop and evaluate bio-printed hydrogels based on gelatin methacrylate (GelMA) combined with different proportions of decellularized bovine bone matrix microparticles (BMdc) used as a bioactive factor.
Methods
GelMA hydrogels were synthesized and incorporated with decellularized bovine bone matrix (BMdc) at 1% by weight. 3D scaffolds were fabricated through extrusion, with varying infill densities (40%, 50%, and 60%), followed by photoactivation. Biological analyses included cell viability (Live/Dead assay), and proliferation (Alamar Blue assay), as well as osteogenic differentiation (ALP activity and Alizarin Red staining) over a 21-day period in HDPCs. Porosity and pore size were assessed with Rhodamine B staining, and cell migration to scaffolds was evaluated in a biomimetic artificial pulp chamber model. Data were analyzed with one-way ANOVA and Tukey's test (p < 0.05).
Results
Scaffolds with the highest porosity and the largest pore size in comparison with other groups was detected in the 40% infill group (p < 0.05). Cells in the 50% and 60% infill groups exhibited higher viability, proliferation, and osteogenic differentiation, especially when BMdc particles were incorporated (p < 0.05). The greatest cell migration at the artificial pulp chamber model was observed in the 60% infill group in association with BMdc particles (p < 0.05).
Conclusions
In summary, 3D-printed GelMA-BMdc hydrogel with 60% infill is a cytocompatible biomaterial capable of inducing cell adhesion, odontogenic differentiation, and mineralization. This innovative biomaterial shows potential for future direct pulp capping applications.
GRAPHICAL ABSTRACT
Keywords:
Tissue Engineering
Hydrogels
Decellularized Extracellular Matrix
Dentin
Gelatin
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1. INTRODUCTION
The cell-homing strategy represents a promising approach for pulp-dentin regeneration, particularly in direct capping therapies. Biomaterials containing chemotactic signaling molecules can facilitate the migration and differentiation of pulp cells into odontoblast-like cells, thereby promoting dentin matrix synthesis and deposition.1–4 Gelatin Methacryloyl (GelMA), a natural polymer derived from collagen, offers numerous advantages for tissue engineering applications due to its biocompatibility, low immunogenicity, and excellent biodegradability.2,5–9 Post-implantation, GelMA degrades enzymatically, enabling the controlled release of bioactive molecules and gradual replacement with newly formed tissue.10,11
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A key feature of GelMA is its ability to form a stable hydrogel network through photoinitiator-induced crosslinking and light exposure, allowing precise control over the gelation process. This property of GelMA supports 3D printing techniques and enables the fabrication of complex structures.6,12,13 Furthermore, GelMA incorporates cell-responsive gelatin-derived molecules, such as matrix metalloproteinases (MMPs) and RGD (Arg-Gly-Asp) sequences, which promote cell attachment, spreading, and proliferation within the hydrogel matrix.12,14 Due to its close similarity to the natural extracellular matrix (ECM), GelMA can be modified or functionalized with bioactive molecules, such as growth factors, peptides, and drugs, to enhance specific cellular responses.15–19Decellularized extracellular matrices (dECMs) have emerged as potent signaling factors in tissue regeneration. These matrices preserve essential proteins, such as collagen, fibronectin, and laminin, as well as growth factors that provide crucial signals for cell adhesion, migration, proliferation, and differentiation.20–22 A study by Paduano et al. (2016)23 demonstrated that a hydrogel scaffold derived from bone dECM significantly increased the expression of dentin sialophosphoprotein (DSPP), dentin matrix protein 1 (DMP-1), and extracellular matrix phosphoglycoprotein (MEPE), as well as mineral deposition in dental pulp stem cells (DPSCs) after 21 days of culture. In a current study from our group, Da Silva et al. (2024)24 assessed the bioactivity of GelMA combined with decellularized extracellular bovine bone matrix (BMdc). The authors showed that such combination produces a porous and stable hydrogel that enhances odontoblast differentiation and mineral deposition in human dental pulp cells (HDPCs), demonstrating its potential for pulp-dentin regeneration
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The optimal pore size and porosity of scaffolds for mineralized tissue regeneration can be achieved through 3D printing, an innovative technique that allows for the modulation of hydrogel microstructures to create porous scaffolds. This method uses a computer-aided design (CAD) model to guide the 3D printer in depositing bioink layer by layer, enabling precise scaffold construction.25–27 Recently, Yang et al. (2023)28 combined porcine dental follicle-derived dECM with methacrylate gelatin (GelMA) to form a GelMA/dECM cell-laden 3D bioprinted scaffold, which exhibited excellent mechanical properties, printability, biocompatibility, and capacity for inducing fibrogenesis and osteogenic differentiation in vivo. The potential for pulp-dentin regeneration was further explored by Cunha et al. (2023),29 using a 3D-printed GelMA microgel supplemented with dentin matrix molecules, which successfully induced odontoblastic differentiation and mineral deposition.In this study, a bioactive hydrogel was developed based on GelMA, combined with BMdc, as a bioink for the fabrication of 3D-printed scaffolds, with the aim of creating a biomaterial suitable for direct pulp capping. To assess the potential of the 3D-printed hydrogel in a cell-homing strategy, an artificial pulp chamber (APC) biomimetic model was used.
2. MATERIALS AND METHODS
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2.1. Gelatin Methacryloyl (GelMA) Hydrogel Synthesis: Gelatin methacryloyl (GelMA) was prepared by dissolving type A porcine gelatin (Sigma-Aldrich, St. Louis, MO, USA) in phosphate-buffered saline (PBS; pH 7.4; Gibco, Invitrogen, Carlsbad, CA, USA) at a concentration of 10% (w/v) at 50°C. Methacrylic anhydride was then added to the solution, and the mixture was stirred for 2 hours. Following this, 100 mL of PBS (Gibco, Invitrogen) was added, and the resulting solution was dialyzed in deionized water for 5 days with two daily water changes. After dialysis, the final solution was filtered (0.22 µm), frozen at -80°C for 2 days, and lyophilized (150 × 10⁻³ Mbar; FreeZone, Labconco Corporation, Kansas City, MO, USA) at -80°C for 5 days. The lyophilized GelMA was then dissolved in PBS (Gibco, Invitrogen) at a concentration of 15% (w/v), and 0.075% (w/v) lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP; Sigma-Aldrich) photoinitiator was added to formulate the hydrogel.
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2.2. Decellularized Bovine Bone Matrix (BMdc) Obtention: Samples of the trabecular area of clavicular bovine bone (Protocol no. 002/2023, Ethics Committee on the Use of Animals, Bauru School of Dentistry, Bauru, SP, Brazil) were subjected to a decellularization protocol described by Da Silva et al. (2024).24 The samples were incubated in the following sequence of reagents: 1) PBS (Gibco, Invitrogen) + 0.1% EDTA (Sigma-Aldrich) at 25°C for 1 hour; 2) PBS (Gibco, Invitrogen) + 10 mM Tris (Invitrogen, Thermo-Fisher Scientific, Eugene, OR, USA) + 0.1% EDTA (Sigma-Aldrich) at 4°C overnight; 3) PBS (Gibco, Invitrogen) + 10 mM Tris (Invitrogen) + 0.5% sodium dodecyl sulfate (SDS; Sigma-Aldrich) for 12 hours; 4) Seven successive washes in PBS (Gibco, Invitrogen); 5) Enzymatic solution containing 50 U/mL of DNase (Thermo-Fisher Scientific) + 1 U/mL of RNase (Thermo-Fisher Scientific) + 10 mM Tris (Invitrogen) for 3 hours at 37°C. Afterward, the samples were frozen at -20°C and subjected to overnight lyophilization at -80°C (150 × 10⁻³ Mbar; FreeZone, Labconco Corporation). To achieve a particle size of 75 µm, the samples were macerated using a crucible and pestle, then sieved. To prepare the GelMA-BMdc hydrogel, the bone particles were added and mixed into the GelMA hydrogel at 1% by weight. This concentration was selected based on Silva et al. (2024), who demonstrated that at this concentration, GelMA-BMdc supported dental pulp cell viability and proliferation, exhibiting bioactive potential.
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2.3. Three-Dimensional Printed Scaffolds: The 3D-printed hydrogel scaffold was obtained using the extrusion method with the TissueStart™ 3D Bioprinter (TissueLabs Ltda, Manno, Switzerland) (Fig. 1a and b). The printing temperature of the hydrogel was 30°C, and the printing speed was set at 6 mm/s with a flow rate of 200%, using a 410 µm inner diameter stainless steel blunt needle. Each printed layer was photoactivated with LED light from the bioprinter at 405 nm. The pore diameter of the hydrogel scaffold were modulated by adjusting the internal filling of the 3D structure (infill), as determined using the Repetier-Host software (Hot-World GmbH & Co. KG, Willich, Germany). The evaluated parameters were 40%, 50%, and 60% infill. A rectangular shape of 30 mm × 15 mm × 1 mm was printed (Figs. 1c and d). Additional post-printing photoactivation was performed for 30 seconds using a light-emitting diode (LED) curing unit at a wavelength of 385–515 nm (1,200 mW/cm²; Bluephase N - Ivoclar Vivadent), covering 10 regions and distributing the light across the rectangular mesh with a distance of 10 mm. Finally, 10 cylindrical samples were obtained from each mesh using a 6 mm-diameter dermatological punch (Fig. 1e-f).
Fig. 1
Representative images of the process of extrusion printing. (A) Extrusion printer used to obtain the samples; (B) Printing process onto a glass slide; (C) Rectangular shape with 50% infill of 30mm X 15 mm X 1mm printed on the glass slide; (D) Printed rectangular mesh suspended by tweezers; (E) Cylindrical samples cut from the rectangular mesh; (F) 10 cylindrical samples obtained from 1 rectangular impression; (G) Cylindrical sample with 6 mm in diameter.
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2.4. Porosity of Printed Hydrogels: The hydrogels were immersed in a 0.5% (w/v) Rhodamine B solution (Sigma-Aldrich) in deionized water for 5 minutes. Afterward, the hydrogels were analyzed using an inverted microscope (Leica DM IRBE - INV-100). Ten images of each sample (n = 5) were captured at 16× magnification and analyzed with ImageJ software (National Institutes of Health, Bethesda, MD, USA) to determine the pore diameter.
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2.5. Biological Analysis: For the biological analysis, the hydrogels, printed under aseptic conditions, were placed in wells of 96-well plates. One drop of α-MEM culture medium (Minimum Essential Medium Eagle Alpha; Gibco, Invitrogen) containing 1 × 10⁴ human dental pulp cells (HDPCs; CAEE: 53489421.9.0000.5417. The participant donating the tooth for cell culture gave Informed Consent) was added onto the surface of the scaffolds, and the HDPC/scaffold constructs were cultivated in α-MEM culture medium (Gibco, Invitrogen), supplemented with fetal bovine serum (FBS; Gibco, Invitrogen), 1% L-glutamine, and 1% penicillin-streptomycin (Gibco, Invitrogen). The cells were then incubated at 37°C in a 5% CO₂ environment. As a negative control (representing 100% of cell parameters), plain GelMA hydrogel (100 µL) was injected into 96-well plates and photoactivated for 30 seconds using LED light (385–515 nm, 1,200 mW/cm²; Bluephase N - Ivoclar Vivadent), as previously described by Silva et al. (2024).24
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2.5.1. Cell Viability and Proliferation Over Time: After 1, 3, 7, 14, and 21 days (n = 2), the samples were washed in PBS (Gibco, Invitrogen) and incubated with α-MEM culture medium (Gibco, Invitrogen) supplemented with the Live/Dead cell viability/cytotoxicity kit (Invitrogen, San Francisco, CA, USA). Live (Calcein AM-positive) and dead (Ethidium homodimer-1-positive) cells on the hydrogels were observed under a fluorescence microscope (FLoid®, Life Technologies, Carlsbad, CA, USA). To quantify metabolically viable cells, the Alamar Blue assay was performed after 1, 3, 7, 14, and 21 days of cultivation (n = 6). In each analysis period, the hydrogels were incubated for 3 hours at 37°C and 5% CO₂ in an α-MEM (without FBS) + 10% Alamar Blue® reagent (Invitrogen) solution. Afterward, the supernatant was transferred to 96-well plates, and fluorescence was measured at 540 nm excitation and 590 nm emission (Synergy H1, Biotek, Winooski, USA). After this assay, GelMA and GelMA-BMdc groups with 50% and 60% infill were further evaluated, with injected GelMA hydrogel (non-printed) serving as control.
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2.5.2. Cell Adhesion and Spreading (F-actin): After 1, 3, 7 and 14 and 21 days of cultivation (n = 2), the HDPCs/scaffold constructs were rinsed with PBS (Gibco, Invitrogen), fixed in 4% paraformaldehyde (PFA; Sigma-Aldrich), and then exposed to the fluorescent probe Alexa Fluor Phalloidin 555 (1:50; Life Technologies) for 20 minutes. Subsequently, nuclear counterstaining was performed using DAPI (ProLong, Thermo Fisher Scientific, Waltham, MA, USA), and images of the samples were obtained using confocal microscopy at 20x magnification (Leica TCS SPE, Confocal Microscope).
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2.5.3. Alkaline Phosphatase (ALP) Activity: The samples seeded with HDPCs (n = 6) in osteogenic medium (α-MEM supplemented with FBS, 50 µg/mL ascorbic acid, and 10 mmol/L β-glycerophosphate) were incubated and analyzed using the Alkaline Phosphatase (ALP) Activity Final Point Assay kit (Labtest Diagnostic S.A.) after 14 days of cultivation. First, the samples were lysed in a 0.1% sodium dodecyl sulfate (SDS) solution (Sigma-Aldrich) under piston maceration, followed by centrifugation at 4,000 rpm for 10 minutes. Next, the supernatant was collected and transferred to tubes containing the substrate thymolphthalein monophosphate (22 mmol/L, pH 10.1; Labtest Diagnostica S.A.; Lagoa Santa, MG, Brazil). The color reagent (94 mmol/L sodium carbonate and 250 mmol/L sodium hydroxide; Labtest Diagnostica S.A.; Lagoa Santa, MG, Brazil) was added, and absorbance was measured at 590 nm (Synergy MX, Biotek). The ALP activity value was obtained by dividing the ALP dosage value by the total protein value, which was quantified by adding Folin's Solution and Ciocalteau's Phenol Reagent (Sigma-Aldrich) for 30 minutes. Absorbance was read at a wavelength of 655 nm.30
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2.5.4. Evaluation of Mineral Deposition (Alizarin Red): The Alizarin Red test was performed after 21 days of cultivation of the HDPCs-hydrogel constructs in osteogenic medium. To evaluate mineral deposition by the HDPCs, the samples (n = 6) were fixed in 70% ethanol at 4°C for one hour, washed with deionized water, and incubated with an Alizarin Red solution (40 mM, pH 4.2; Sigma-Aldrich) under agitation for 15 minutes. Following this, the hydrogels were washed five times with deionized water, and digital photographs were captured. Next, a solution of cetylpyridinium chloride (10 mM, pH 7.0; Sigma-Aldrich) was applied for 15 minutes to dissolve the mineral nodules, and the absorbance of the resulting solution was measured at 560 nm (Synergy MX, Biotek). Hydrogel samples without cells served as the background control (n = 6).
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2.6. Artificial Pulp Chamber Assay: To evaluate the cell-hydrogel interaction in a cell-homing strategy within a biomimetic model, the artificial pulp chamber (APC) developed by our group and previously described by Soares et al. (2021)31 was used. Its mechanism simulates the normal fluid pressure of human pulp tissue. The APC consists of a lid, a central chamber (CC), and a base (Fig. 2). A human dentin disc containing a 1 mm diameter central perforation is placed between upper and lower CC compartments (Fig. 3A and B). A HDPC 3D cell culture31 is placed inside the lower compartment of the CC in direct contact with pulpal side of dentin disc (Fig. 3C and D). The 3D culture was established by dissolving a 3.7 mg/mL type 1 collagen solution (Corning Inc., Somerville, MA, USA) in a 4:1 ratio with a 10× concentrated α-MEM culture medium (Gibco, Invitrogen). To achieve a pH of 7.2, 5 M sodium hydroxide (Sigma-Aldrich) was used for neutralization in a 4°C environment.31 Next, 1 × 10⁶ HDPCs in 5 µL of medium were added to 200 µL of the collagen solution. The resulting mixture was applied to the central chamber (CC) of the APC (Fig. 3C) and incubated at 37°C and 5% CO₂ for 30 minutes to allow the collagen matrix to solidify into a gel (Fig. 3D). The 3D culture was incubated without pressure for 48 hours to ensure stable culture conditions. After this period, the selected hydrogels were placed into the dentin perforation (Fig. 3E and F), stablishing direct contact with the 3D culture, and then covered with polystirene. A silicon ring is placed onto the polystyrene (Fig. 3G) and all parts of APC are threaded (Fig. 3H). To stablish simulated pressure the lateral connections are coupled to the pressure system (Fig. 3I), that includes a syringe with culture medium to create the liquid column (15 cm of liquid), coupled to the APC inlet connection, providing hydrostatic pressure of 20 cm/H2O (i.e., 14.7 mm Hg), which represents the natural pulp (Fig. 3J).32,33 The pressurized system was incubated at 37°C and 5% CO₂, with daily perfusion of 5 mL culture medium to renew the medium in the system for up to 14 days. For this assay, the following groups were stablished: GelMA – injected GelMA; GelMA-BMdc – injected GelMA-BMdc; GelMA 60% − 3D printed GelMA with 60% infill; GelMA-BMdc 60% − 3D printed GelMA-BMdc with 60% infill.
Fig. 2
Representative images of Artificial Pulp Chamber (APC) and its parts.
Fig. 3
Representative images of APC experimental design. (A) Upper view of lower CC with dentin disc (*) containing a central perforation; (B) Dentin disc pulpal surface (*) view from inside lower CC chamber; (C) Collagen solution containing HDPCs application inside lower CC chamber; (D) 3D culture view after gelification; (E) GelMA (*) being placed onto dentin disc; (F) Upper view of GelMA (*) adapted onto dentin; (G) Silicon ring placed to promote adequate sealing; (H) APC view after thearing all parts and with CC filled with culture medium; (I) APC with lateral connections to pressure system; (J) Pressurized system.
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2.6.1. Cell Viability and proliferation: After 1, 3, 7, and 14 days (n = 4), the Live/Dead assay was performed as described previously, and images of the scaffold and 3D culture were captured using a fluorescence microscope (FLoid®, Life Technologies). Cell metabolism of 3D culture was monitored at 3, 7, and 14 days (n = 4) using Alamar Blue Assay.
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2.6.2. Cell Migration to Scaffold: The migrating cells on the surface of the hydrogels were counted after 14 days of cultivation. Ten images per sample (n = 4) were obtained using a fluorescence microscope (FLoid®, Life Technologies) after incubation with the Hoechst fluorescent probe (1:10,000; Life Technologies). The number of migrating cells was quantified using ImageJ software (National Institutes of Health).
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2.7. Statistical Analysis: After assessing normality and homoscedasticity using the Shapiro–Wilk test, data were analyzed using Prism 8 software (GraphPad; San Diego, CA, USA). One-way ANOVA, followed by the Tukey post hoc test, was used to identify significant differences between groups. A p-value of < 0.05 was considered statistically significant.
3. RESULTS
3.1. Porous Architecture: All the 3D-printed hydrogels exhibited a macroporous structure, as observed in the Rhodamine B-stained samples (Fig. 4A). The pore size was proportional to the infill percentage, as anticipated. However, at 40% infill, the samples were more fragile due to the larger printed pores, resulting in a more disorganized printed pattern (Fig. 4B).
Fig. 4
Scaffold porosity and biological characterization. (A) Rodamin B-stained 3D printed scaffolds according to hydrogel composition and infill %, at 2.6x and 5x magnification; (B) Bar graph of mean values of pore size from 3D printed scaffolds. Numbers are mean values. Different letters allow for statistic comparisons (one-way ANOVA; Tukey’s test. P < 0.05). (C) Alamar Blue assay. Representative graph of the evolution of cell proliferation for 21 days. (D) Live/Dead assay. Representative images of the surface of hydrogels (20x). Green: Viable cells; red: dead cells.
3.2. Biological Characterization: As shown in Figs. 5C and 5D, all groups supported cell viability and proliferation. Notably, among the printed groups, the 40% infill demonstrated a lower number of viable cells compared to the 50% and 60% infill groups, particularly at the 14- and 21-day time points, as observed in the Live/Dead images (Fig. 4D). Consequently, the 40% infill group was excluded from subsequent experiments.
The F-actin assay allowed for the observation of HDPCs actin filaments (Fig. 5A), demonstrating cell adhesion and spreading on all hydrogels throughout the 21-day analysis period. However, a greater quantity of filaments was observed in the GelMA-BMdc 60% infill group at the 3- and 7-day time points. The differentiation rate was significantly influenced by the modulation of porosity through 3D printing and the addition of the bone matrix, in comparison to plain GelMA (Figs. 5B-E). The 3D-printed samples exhibited higher alkaline phosphatase (ALP) activity and greater mineralized matrix deposition compared to the injected GelMA, with the 60% infill group showing statistical difference (p < 0.05). The incorporation of BMdc also increased the expression of odontogenic markers compared to the injected BMdc samples, with significant difference only detected in the 60% infill group (p < 0.05). Digital images of Alizarin Red-stained samples at 21 days (Figs. 5D-E) indicate that the 50% infill samples appeared more degraded, while the 60% infill remained stable over time.
Fig. 5
Biological characterization - Cell spread and differentiation. (A) Representative images of the cell adhesion and spreading assay (F-Actin; 20x). Green: actin filaments; Blue: cell nuclei; (B) Representative bar graph of percentage of ALP activity at 14 days. Bars represent mean and standard deviation (n = 6); (C) Bar graph representing mineral deposition marked by the Alizarin Red assay at 21 days. Bars represent mean and standard deviation (n = 6). Values represent average and different letters indicate a statistical difference between groups (One-way ANOVA; Tukey´s test. p < 0.05); (D) Images of samples of plain GelMA groups stained by Alizarin Red and (E) Images of GelMA-BMdc groups stained by Alizarin Red.
3.3. APC assay: For this assay, the 60% infill pattern was selected based on previous analysis. Living cells were observed in the 3D culture of all groups throughout the 14-day period (Fig. 6A). Regarding the hydrogels, viable cells were detectable in the non-printed control groups only after 7 days of analysis, whereas in the printed groups, live cells were already observed 3 days after being seeded. At 14 days, the GelMA-BMdc 60% group exhibited a notably higher number of live cells on the hydrogel surface. The presence of BMdc and the application of 3D printing increased the number of metabolically active cells and promoted greater cell migration compared to the control group (plain GelMA), as observed in the Fig. 6B. Finally, Fig. 7 shows the quantification of migrating cells from the 3D culture to the hydrogel surface. Cell migration was significantly influenced by both 3D printing and the incorporation of BMdc. The GelMA-BMdc 60% infill group demonstrated a significantly greater ability to stimulate cell migration compared to the other groups (p < 0.05).
Fig. 6
Cell viability and proliferation assay. (A) images of Live/Dead assay performed on hydrogels and 3D cultures (20x) grown in pAPC for up to 14 days. Green: living cells; Red: dead cells; (B) Representative graph of the percentage of cellular metabolism (Alamar Blue).
Fig. 7
Cell migration assay. (A) Images obtained by fluorescence microscopy representing cells migrating to the surface of the hydrogels (20x). (B) Representative graph of the number of cells migrating to the surface of the hydrogels (One-way ANOVA; Tukey´s test. p < 0.05).
4. DISCUSSION
GelMA hydrogel has demonstrated suitability as a bioink for 3D printing applications.34,35 In this study, a decellularized bone matrix (BMdc) was incorporated as a signaling factor within the GelMA hydrogel to develop a bioactive bioink for 3D printing, specifically aimed at dentin tissue engineering. The composition of the bioink and the BMdc particles used in this study were previously characterized by Silva et al. (2024).24 BMdc particles were obtained using a protocol adapted from Graysson et al. (2008),36 containing both organic and inorganic components, with a 99.6% efficiency in DNA removal. Silva et al. (2024)24 also demonstrated the absence of cells in the BMdc particles following decellularization, as confirmed by fluorescence microscopy. The incorporation of BMdc particles enhanced the bioactive potential of GelMA, surpassing that of bovine bone ceramic, which is derived through a conventional physical-chemical process that ensures the complete removal of organic matter. At a 1% (w/v) concentration, the addition of BMdc to 15% GelMA allowed for the viability and proliferation of HDPCs within the hydrogel structure, with these cells showing increased expression of odontoblastic markers. The injectability, porosity and pore size, as well as the degradability and density of the hydrogel were unaffected by the incorporation of BMdc. However, the increased compressive modulus of the biomaterial indicated that its resistance was enhanced.
Firstly, GelMA and GelMA-BMdc hydrogels were 3D printed with three different internal filling densities—40%, 50%, and 60%—to modulate pore size and porosity. The viability and proliferation of cells in contact with the scaffolds were assessed by Live/Dead and Alamar Blue assays. Cytocompatibility was observed in all experimental groups, in which adhesion, spreading, and proliferation of HDPCs occurred over a 21-day period. Notably, the 60% infill groups exhibited a higher cells proliferation rate at day 21. On the other hand, decreased cell viability was noted in the 40% infill groups at days 14 and 21, that was attributed to the faster degradation of the biomaterials due to its lower infill density. Oversized pores and thinner walls observed in the 40% infill scaffolds, as determined by the Rhodamine B assay, may compromise the mechanical integrity of the hydrogel and lead to structural collapse,37,38 thereby interfering with the support necessary for continued cell growth. This hypothesis warrants further investigations through degradation analysis. Consequently, the 40% infill group was excluded from subsequent experiments.
The incorporation of BMdc into the GelMA hydrogel effectively stimulated odontogenic differentiation and mineral deposition by HDPCs, particularly at the 60% infill concentration, as evidenced their increased alkaline phosphatase (ALP) activity. These effects may be attributed to the synergistic interaction of BMdc incorporated to the biomaterial and its optimized porosity. Additionally, the improved mechanical resistance demonstrated by Silva et al. (2024)24 may contribute to an increase in scaffold stiffness, thereby making it a suitable platform for mineralized tissue regeneration, as previously suggested by Bottino et al. (2017).39 Similar results are consistent with the findings of Yang et al. (2023),28 who developed bioprinted scaffolds combining GelMA with a porcine dental follicle-derived decellularized extracellular matrix (dECM), loaded with dental follicle cells (GelMA/dECM). The authors demonstrated that such scaffolds exhibited excellent mechanical properties, printability, biocompatibility, and potential to induce fibrogenesis and osteogenic differentiation in vivo. Similarly, Cunha et al. (2023)29 reported that a 3D-printed GelMA microgel, supplemented with dentin matrix molecules, successfully induced odontoblastic differentiation and mineral deposition. This can be explained by the fact that ECM mediates signaling to resident cells, regulating their proliferation, migration, and differentiation. Decellularized ECM (dECM) biomaterials have the potential to stimulate tissue regeneration by providing a native-like environment. During decellularization, immunogenic cells and molecules are largely removed, while functional components such as glycosaminoglycans, glycoproteins, and cytokines are preserved, promoting cellular differentiation and, consequently, tissue regeneration.40
In the present study, the pore size and porosity of the 3D-printed hydrogels were evaluated after incubation the biomaterials with a Rhodamine B fluorescent marker. It was observed that the 3D-printed hydrogels containing GelMA with 50% infill and GelMA-BMdc with 50% infill exhibited mean pore diameters of 592 µm and 698 µm, respectively, while the GelMA 60% infill and GelMA-BMdc 60% infill groups had average pore diameters of 361 µm and 355 µm, respectively. These pore sizes seem to be aligned with the ideal parameters for mineralized tissue regeneration, which typically range from 100 to 400 µm.3,41–43 Due to their higher water content, hydrogels possess a microporous cross-linking structure that not only provides structural support but also facilitates the retention of water and nutrients. In this context, the use of GelMA-BMdc 3D-printed hydrogels has proven effective in optimizing the porous structure, creating mesoscale pores larger than 100 µm, which promotes nutrient circulation, stimulates cell migration, and provides adequate space for new tissue formation—key processes for cell proliferation.30,44–47 In support of these findings, Soares et al. (2020)30 demonstrated that increasing the pore diameter of chitosan scaffolds from 86.9 µm to 202.1 µm using Ca(OH)2 stimulated odontogenic differentiation and calcium-rich matrix deposition by dental pulp cells.
Based on the scientific data initially obtained in this laboratorial study, the 60% infill parameter was selected for further analysis employing a biomimetic strategy. The artificial pulp chamber (APC) model used in this study effectively simulated human intrapulpal pressure, and the inclusion of type I collagen in the 3D culture seemed to turn this methodology closer to the physiologic environment of the vital pulp tissue. In a previous investigation, Soares et al. (2021)31 demonstrated that chitosan-calcium scaffolds combined with simvastatin was capable of inducing chemotaxis to pulp cells, which showed enhanced expression of odontoblastic markers when in contact with a 3D culture adapted to the APC. These interesting results that were validated in vivo, highlight the potential of the biomimetic strategy used by the authors to replicate a natural pulp tissue environment effectively. In applying this contemporary methodology to the present study, HDPCs from the 3D culture in contact with the 3D-printed hydrogels remained viable and proliferated throughout the 14-day analysis period. All groups stimulated cells migration into the scaffolds, with the GelMA-BMdc 60% infill group showing the most noticeable results. Viable cells were observed on the surface of the printed hydrogels as early as 3 days, whereas in the non-printed hydrogels used as controls, viable cells were only detected after 7 days. This specific result suggests that modulating the porosity of 3D-printed hydrogels through 3D printing enhances early cell migration, with the GelMA-BMdc 60% infill group exhibiting superior outcomes under simulated pulp pressure. However, despite the artificial pulp chamber used to perform biomimetic laboratory strategies closely mimics the in vivo conditions, results from in vitro studies cannot be directly extrapolated to clinical situations. Even the innovative biomimetic protocol employed in this study presents critical limitations, such as the absence of immune cells associated with blood vessels and nerves network, which play a fundamental role in the mechanism of pulp-dentin tissue regeneration. Therefore, further research is needed to assess the responses of connective tissues exposed to the 3D-printed GelMA-BMdc hydrogel as well as to determine the biologic potential of this biomaterial to improve the pulp-dentin complex regeneration.
5. CONCLUSION
The 3D-printed GelMA-BMdc hydrogel with 60% infill resulted in a material with a pore size favorable for dentin regeneration. The hydrogel exhibited biocompatibility, facilitated cell adhesion and spreading, and promoted odontogenic differentiation along with mineralized matrix deposition. These findings suggest that this biomaterial holds significant potential for future applications in direct pulp capping.
ACKNOWLEDGMENTS
This study was funded by Fundação de Amparo a Pesquisa do Estado de São Paulo - FAPESP (grants 2021/09498-8; 2016/15674-5; 2022/05888-9), and Coordenação de Aperfeiçoamento de Pessoal de Nível Superior - CAPES (grant 001).
CONFLICT OF INTEREST STATEMENT
The authors declare no conflicts of interest.
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Data Availability
The data that support the findings of this study are available from the corresponding author upon reasonable request.
ORCID
Isabela Sanches Pompeo da Silva: 0000-0003-2000-2671
Vitor de Toledo Stuani: 0000-0001-5290-7614
Ester Alves Ferreira: 0000-0002-4178-5794
Fernanda Balestrero Cassiano: 0000-0002-2336-876X
Erika Soares Bronze-Uhle: 0000-0002-9273-9421
Priscila Toninatto Alves de Toledo: 0000-0003-0677-8895
Carlos Alberto de Souza Costa: 0000-0002-7455-6867
Diana Gabriela Soares: 0000-0002-1485-6104
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Author Contribution
I.S.P.S.: Conceptualization, Methodology, Investigation, Data curation, Writing – original draft.V.T.S.: Methodology, Formal analysis, VisualizationE.A.F.B.: Methodology, Validation, Data curation.F.B.C.: Investigation, Resources, Data analysis.E.S.B.: Conceptualization, Supervision, Data curation, Data analysis.P.T.A.T.: Data analysis, Writing – review & editing.C.A.S.C.: Conceptualization, Supervision, Writing – review & editing.D.G.S.: Conceptualization, Supervision, Project administration, Funding acquisition, Writing – review & editing.
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