Bioactive photo-crosslinkable hydrogel based on decellularized dental pulp matrix for functional pulp regeneration

Abstract

Dental pulp extracellular matrix (DPEM), as a naturally derived scaffold material for dental pulp regeneration, exhibits excellent biocompatibility. This study aims to develop a novel photo-cross-linked hydrogel composed of methacry- lated gelatin (GelMA) loaded with DPEM for application in pulp regeneration. In vitro experiments demonstrated that the GelMA–DPEM hydrogel, fabricated by mixing a 10 mg mL − 1 DPEM solution with a 10% (w/v) GelMA solution at a ratio of 1:1, formed a stable cross-linked network with a swelling ratio of approx- imately 343%. Frequency-sweep rheology indicated that incorporation of DPEM increased the storage modulus of the hydrogel compared with GelMA alone, and the composite hydrogel significantly promoted cell migration and angiogenic ca- pacity. RT–qPCR analysis revealed that the GelMA–DPEM hydrogel markedly upregulated the expression levels of neuroepithelial stem cell protein (Nestin), dentin sialophosphoprotein (DSPP), platelet endothelial cell adhesion molecule-1 (CD31), and runt-related transcription factor 2 (Runx2). Moreover, ectopic trans- plantation in nude mice indicated that, when combined with human dental pulp stem cells (hDPSCs), the GelMA–DPEM hydrogel preserved the DPEM’s capa- bility to support the formation of a cell-rich, vascularized pulp-like tissue con- sisting of collagen-rich connective tissue and CD31-positive blood vessels. In conclusion, the GelMA–DPEM hydrogel showed favorable cytocompatibility and preliminary pro-angiogenic effects in vitro and supported pulp-like tissue forma- tion in a small-animal ectopic treated dentin matrix (TDM) model. These findings suggest that GelMA–DPEM is a promising candidate scaffold for dental pulp re- generation.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12903-026-08001-6.

Introduction

Dental pulp regeneration represents a pivotal component of the broader field of tooth regeneration, aiming to restore the biological function of dental pulp through tissue engineering strategies [1]. Currently, the primary clinical approach for treating necrotic pulp remains root canal therapy, which involves the mechanical removal of necrotic tissue followed by obturation of the canal with synthetic materials to prevent reinfection [2, 3]. However, this approach fails to restore the physiological functions of the native pulp tissue. In contrast, dental pulp regeneration is increasingly recognized as a promising strategy for re-establishing the biological vitality of the pulp [4]. This regenerative approach typically involves the synergistic application of stem cells, biomaterials, and growth factors to re- construct a microenvironment that closely mimics native pulp tissue. Such an en- vironment facilitates stem cell proliferation, lineage-specific differentiation, and the formation of functional vasculature within the regenerated tissue [5].

In the context of dental pulp regeneration, DPEM serves as a biologically relevant scaffold with strong tissue specificity, providing both a physical substrate for cell attachment and a reservoir of bioactive cues [6]. The preserved extracellu- lar matrix microenvironment of DPEM comprises a complex mixture of proteins, glycosaminoglycans, growth factors, and other signaling molecules. In addition to offering a three-dimensional structural framework, it plays a critical role in mod- ulating cellular behaviors, including proliferation and differentiation [7–9]. Our previous studies have demonstrated that DPEM effectively supports the formation of pulp-like tissue [6]. However, the native macrostructure retained in DPEM lim- its its adaptability to the diverse morphologies of dental pulp chambers, thereby constraining its clinical applicability. To address this limitation, we aim to engineer DPEM into a syringe-deliverable form that can conform to the complex and irregular geometry of the pulp cavity.

In recent years, hydrogels have emerged as highly promising biomaterials and have been increasingly utilized in various biomedical applications, includ- ing tissue engineering [10], drug delivery [11], and wound healing [12]. Serving as carriers for cells or therapeutic agents, hydrogels provide a microenvironment that closely mimics native tissue architecture, thereby supporting cell adhesion, proliferation, and differentiation. Moreover, by incorporating growth factors, stem cells, and other bioactive molecules, hydrogels can significantly enhance localized tissue repair and regeneration [13, 14]. Among naturally derived hydro- gels, gelatin [15], hyaluronic acid [16], sodium alginate [17], and silk fibroin [18] are commonly employed. GelMA, a methacrylate-modified derivative of gelatin containing vinyl groups, can undergo rapid photocrosslinking under blue light or visible light in the presence of photoinitiators. This material forms a three-dimensional (3D) structure conducive to cellular growth and differentiation, exhibiting a unique combination of both natural and synthetic biomaterial characteristics [19].

Guo et al. demonstrated that GelMA can be injected into dental pulp defects in vivo and subsequently crosslinked via external light exposure [20]. Its injectability enables application within irregular or curved pulp cavities, while its photo-curing capability allows for rapid stabilization into a robust 3D construct upon illumination—offering structural integrity and a favorable microenvironment for pulp tissue regeneration [21–23]. Ju et al. proposed that extracellular vesicles can be encapsulated within hydrogels to achieve controlled release and targeted delivery, thereby enhancing their therapeutic efficacy in local tissue regeneration [24].

Inspired by the biomineralization process, a mineral-based hydrogel composed of polyacrylic acid, carboxymethyl chitosan, and dentin matrix has been developed, exhibiting self-healing properties, injectability, and mineralization-regulating capabilities, with promising outcomes in dentin and bone regeneration [25]. Yang et al. demonstrated that GelMA microspheres encapsulating hDPSCs effectively support cell adhesion, proliferation, and extracellular matrix secretion, and pro- mote the formation of highly vascularized pulp-like tissue in vivo [26]. Elnawam et al. fabricated an injectable hydrogel derived from bovine dental pulp tissue, which preserved glycosaminoglycans and collagen structure, and enabled sustained re- lease of multiple key regenerative growth factors. This hydrogel significantly maintained stem cell viability and induced the formation of organized pulp-like tissue [27]. These findings provide important theoretical support for developing GelMA-DPEM composite hydrogels and exploring their regenerative potential.

Taken together, the development of scaffold materials that combine the bioactivity of natural dental pulp matrix with favorable injectability and photo-crosslinkability has become a key focus in dental pulp regeneration research. Therefore, this study aims to develop a bioactive, photo-crosslinkable GelMA–DPEM composite hydrogel with a syringe-deliverable precursor solution and to evaluate its effectiveness in promoting dental pulp tissue regeneration.

A schematic illustration of the experimental design is shown below (Fig. 1):

Fig. 1.

Experimental design flow chart. A Isolation and culture of hDPSCs, preparation of DPEM and TDM from porcine teeth, followed by enzymatic digestion of DPEM to obtain a soluble DPEM solution. B The DPEM solution was mixed with GelMA to form a GelMA-DPEM hydrogel, which was then loaded with hDPSCs and injected into the TDM scaffold. The construct was photo-crosslinked under 405 nm blue light for 30 s and implanted subcutaneously into immunodeficient (nude) mice for 12 weeks

Materials and methods

Preparation of GelMA-DPEM hydrogel materials

Dental pulp tissue was extracted from permanent tooth germs of three fresh porcine mandibles (6–12 months) obtained from a local market. The tissue was sectioned into small fragments and washed three times with sterile Phosphate.

Buffered Saline (PBS, Coolaibo, China) to remove surface impurities. It was then stirred in 1% SDS (Bio-Rad, USA) for 12 h, followed by immersion in dis- tilled water containing 1% Triton X-100 (Sigma, USA) for 30 min. After de- cellularization, the samples were rinsed three times with sterile PBS (10 min each) and subsequently washed in distilled water for an additional 30 min. The decellularization and pepsin-digestion procedures used to prepare DPEM in this study were adapted from previously reported protocols with minor modifications [6, 28]. The resulting DPEM was sterilized by soaking in a penicillin–streptomycin solution (Hyclone, USA) for 96 h and stored at −20 。C for future use.

The prepared DPEM was first stored at − 80 。C for 24 h and then lyophilized for 8 h to obtain freeze-dried DPEM. The resulting material was ground into a fine and homogeneous powder using a mechanical grinder (Jingxin, China).

DPEM particles were enzymatically digested in a solution containing 0.1 M hy- drochloric acid (Kangde Chemical, China) and 1.5 mg/mL pepsin (Coolaibo, China) for 48 h. The digest was centrifuged at 2000 rpm for 10 min at room temperature to remove undissolved debris, yielding a DPEM solution at a fi- nal concentration of 10 mg/mL. The pH of the solution was subsequently adjusted to neutral by the dropwise addition of 10 M sodium hydroxide (Kelon Chemical, China).

Separately, freeze-dried GelMA (Engineering for Life, China) was dissolved in sterile PBS containing 0.5% (w/v) lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) (Engineering for Life, China) to prepare a 5% (w/v) GelMA solution, which was then photo-crosslinked under 405 nm blue light for 30 s to form a stable hydrogel. A 10% (w/v) GelMA solution was prepared following the same protocol and mixed in a 1:1 volume ratio with the 10 mg/mL DPEM solution. The resulting mixture, containing 5% GelMA and 5 mg/mL DPEM, was irradiated under blue light for 30 s to form a stable GelMA–DPEM hydrogel.

Characterizations of GelMA-DPEM hydrogel

Characterization of the DPEM

Native dental pulp tissue and DPEM samples were fixed in 4% paraformalde- hyde (Bo Pei, China) for 12h. The specimens were then dehydrated through a graded ethanol series (Sinopharm Group Chemical Reagent Co., LTD., China) and further dried using a freeze-dryer. Subsequently, the dried samples were affixed to conductive carbon adhesive tape and sputter-coated with gold for approximately 30 s using an ion sputter coater. Surface morphology was examined using scanning electron microscopy (SEM, Hitachi, Japan).

Native dental pulp tissue and DPEM samples were dehydrated, embedded, and sectioned into 5 µ m-thick slices.The tissue sections were then subjected to hematoxylin and eosin (H&E) staining(Seville, China) and Masson’s trichrome staining(Solaibao, China) to evaluate cellular removal and collagen distribution.

General observation and SEM

The macroscopic extrusion behavior and filling morphology of the GelMA–DPEM hydrogel were first examined using a transparent pulp-cavity model. Hydrogel precursor solutions (5% (w/v) GelMA–DPEM containing 0.2 mg/mL dye) were loaded into a 2 mL syringe fitted with a 23G needle and slowly injected into the model. After curing with blue light for 30 s, morphological changes were recorded.

The microstructure of the GelMA–DPEM hydrogel was assessed using SEM. GelMA and GelMA–DPEM solutions were injected into cylindrical molds (6 mm in diameter and 8 mm in height) and photo-crosslinked under 405 nm blue light for 30 s. Subsequent procedures were performed as described in Sect. 2.2.1.

Rheological analysis of GelMA–DPEM hydrogels

The rheological properties of the DPEM hydrogel and GelMA-DPEM hy- drogel were analyzed using a HAAKE rheometer. Frequency sweep tests were performed with a parallel-plate geometry at 37 °C. A constant strain of 1% was applied over a frequency range of 0.1–10 Hz. Each sample was tested in tripli- cate.For each formulation, three independent hydrogel samples were tested (n = 3), and the frequency-sweep curves were plotted as mean values.

Swelling and degradation analysis in vitro

For the swelling assay, GelMA and GelMA–DPEM hydrogels were first lyophilized to obtain completely dried samples, which were then weighed on a precision electronic balance (Mettler Toledo, Switzerland) to determine the initial dry weight (W0). Each sample was placed in an individual well of a 6-well plate, and 3 mL of PBS was added to each well. The plates were incubated at 37.5 ◦ C in a 5% CO2 atmosphere. At predetermined time points (0.5, 1, 1.5, 2, 2.5, and 24 h; n = 3), the samples were carefully removed, excess surface water was gently blotted with sterile filter paper, and the swollen weight (W1) was measured on the same balance. The swelling ratio was calculated as:

Using lyophilized hydrogels provided a standardized initial state and allowed a direct comparison of the relative swelling behavior of GelMA and GelMA–DPEM under identical conditions.

For the enzymatic degradation assay, photo-crosslinked hydrogels were lyophilized and weighed to obtain the initial dry weight (W0), and then immersed in 3 mL of 2% (w/v) type I collagenase solution (Thermo Fisher, USA) at 37 ◦ C. The col- lagenase concentration was selected based on previous studies on the accelerated enzymatic degradation of collagen-based/GelMA hydrogels [29, 30]. At designated time points (1, 3, 6, 12, and 24 h; n = 3), samples were retrieved, rinsed with PBS, lyophilized again, and weighed to determine the remaining dry weight (W1). The degradation rate was calculated as:

Cell isolation and culture

Human dental pulp tissue was obtained from extracted orthodontic or third molar teeth collected for therapeutic purposes at the First Affiliated Hospital of Dalian Medical University. Informed consent was obtained from all donors prior to sample collection. Under aseptic conditions, each tooth was decoronated at the cementoenamel junction using a sterile diamond disk under copious sterile saline irrigation to expose the pulp chamber and root canals. The dental pulp tissue was then gently extirpated from the crown and root canal space using ster- ile endodontic instruments (barbed broaches and K-files) and transferred into a centrifuge tube. Under sterile conditions, the pulp tissue was gently retrieved using forceps and transferred into PBS. The tissue was cut into small fragments, washed three times with sterile PBS to remove surface debris, and then transferred to centrifuge tubes containing 0.25% Trypsin–EDTA (Invitrogen, USA). Samples were incubated at 37.5 ◦ C with 5% CO2 for 30 min. The enzymatic digestion was terminated by adding an equal volume of complete culture medium, followed by centrifugation at 1000 rpm for 5 min. The supernatant was discarded, and the tissue fragments were resuspended and cultured in α-MEM (HAKATA, USA) supplemented with 10% fetal bovine serum (FBS) (Omega, USA) and 1% penicillin–streptomycin. Cells were incubated at 37.5 ◦ C with 5% CO2, and the medium was refreshed every 3 days. When hDPSCs reached 80–90% confluence, they were passaged for further expansion. Cells at passages 3 to 6 were used for all subsequent experiments. HDPSCs were isolated from human dental pulp using a collagenase digestion protocol and expanded in standard culture medium. Previous studies have confirmed that such DPSCs display typical MSC properties, with an MSC marker profile and the ability to differentiate into multiple mesenchymal lineages [31].

Cell proliferation assay

A total of 3,000 hDPSCs were seeded into each well of a 96-well plate and incubated at 37.5 ◦ C with 5% CO2. After 24 h, once cell adherence was confirmed, the culture medium was carefully aspirated. For the control group, 100 µL of fresh complete medium was added to each well.

For the experimental groups, materials were added at a final volume fraction of 20%. Specifically, for the GelMA and GelMA–DPEM groups, 80 µL of com- plete medium was mixed with 20 µL of the corresponding uncrosslinked pre-gel solution (10% (w/v) GelMA with or without DPEM) to obtain a total volume of 100 µL per well (final pre-gel fraction 20% (v/v)). For the DPEM group, small hydrated DPEM blocks were gently cut, and an amount corresponding to approx- imately 20% of the well volume was placed in each well, followed by addition of 80 µL complete medium.

Cells were cultured under standard conditions, and at days 1, 3, 5, and 7, the culture medium was aspirated and replaced with 100 µL of freshly prepared CCK-8 working solution (Biyuntian, China). The plates were incubated in the dark for 2 h, and absorbance at 450 nm was measured using a microplate reader.

The culture medium was refreshed every 2 days.

Cell viability

A live/dead cytotoxicity assay kit was used to evaluate cell viability. HDP- SCs were harvested by trypsinization and centrifugation, and 3,000 cells were seeded into each well of a 96-well plate. After 24 h of attachment, the cul- ture medium was replaced with the same treatment conditions as described in Sect. 2.2.6, including the control group (complete medium only), the GelMA group, the GelMA–DPEM group, and the DPEM group (materials added at a final volume fraction of 20%). At days 1, 4, and 7, live/dead staining was performed using PI and Calcein-AM working solutions (Biyuntian, China) prepared accord- ing to the manufacturer’s instructions. Cells were incubated at room temperature in the dark for 15 min and then immediately observed and imaged under a fluo- rescence microscope. Cell viability was quantified using ImageJ software.

Cell migration

The migratory ability of hDPSCs was evaluated using Transwell inserts (Thermo Fisher Scientific, USA) in a 24-well plate. A total of 2 × 104 cells were seeded into the upper chambers in serum-free medium.

The lower chambers were filled with 200 µL of material formulations as follows: 5% (w/v) GelMA pre-gel solution, 5% (w/v) GelMA–DPEM pre-gel solution, or a suspension of small hydrated DPEM blocks. The pre-gel solutions in the GelMA and GelMA–DPEM groups were photo-crosslinked in situ using 405 nm blue light for 30 s to form thin hydrogel layers at the bottom of the wells, after which 300 µL of complete medium was added to each lower chamber. For the DPEM group, the hydrated DPEM blocks settled at the bottom of the wells before adding 300 µL of complete medium.

After 12 h of incubation, cells on the upper side of the Transwell membrane were gently removed with a cotton swab, and cells that had migrated to the lower surface were fixed with 4% paraformaldehyde for 30 min. The inserts were then stained with 0.1% crystal violet (Biyuntian, China) for 20 min. Four random fields per well were imaged using an inverted microscope, and the number of migrated cells was quantified using ImageJ software.

Tube formation

A Matrigel-based tube formation assay was conducted to assess the angio- genic behavior of endothelial cells. GelMA hydrogel, GelMA–DPEM hydrogel and endothelial cell culture medium were thoroughly mixed to prepare the re- spective conditioned media. Immortalized human umbilical vein endothelial cells (HUVECs; Yakoin, China) were seeded into 6-well plates at a density of 8 × 104 cells per well and cultured in the corresponding conditioned media for 24 h.

Separately, 50 µL of Matrigel (Corning, USA) was gently added to each well of a 24-well plate and allowed to solidify at 37.5 ◦ C. After 24 h of conditioning, endothelial cells were harvested, centrifuged, resuspended in endothelial growth medium and counted. A total of 8 × 104 cells were seeded into each Matrigel- coated well. The plates were incubated at 37.5 ◦ C with 5% CO2 for 4 h.

For quantitative analysis, tube formation was evaluated by counting the num- ber of tube-like structures in several random microscopic fields per well using ImageJ software, and the mean tube count per well was used for statistical com- parison among groups.

RT-qPCR analysis

The odontogenic, angiogenic, and neurogenic gene expression profiles of hDPSCs cultured with different materials for 7 days were evaluated by RT-qPCR analysis. A total of 1 × 104 hDPSCs were seeded with either GelMA or GelMA- DPEM hydrogel in culture plates. After 7 days of incubation, the culture medium was removed, and total RNA was extracted from the cells for reverse transcription and RT-qPCR analysis. The specific primer sequences for the target genes are listed in Table 1.

Table 1.

Primer sequences used for RT-qPCR analysis

Gene	Primer Sequences (5’–3’)
Runx2	F-CTTTACTTACACCCCGCCAGTC R-AGAGATATGGAGTGTGCTGCTGTC
CD31	F-GTGCTGCAATGTGCTGTGTGAA R-TGCTAGCCTTCTGCTGGTGTC
DSPP	F-CTGTTGGGAAGAGGCCAAGATAAG R-CCAAGATCATTCCATGTTGTCCT
Nestin	F-AGGAATGCCGCTGAGTCTCTGA R-GGACTCTCTATCTCCCTTCCCTTG
GAPDH	F-CTTTGGTATCGTGGAAGGACTC R-GTAGAGGCAGGGATGATGTTC

GelMA-DPEM hydrogel for in vivo pulp regeneration

Preparation of TDM

TDM was used as a hard tissue scaffold to simulate the dental pulp cavity. The crowns and apical portions of porcine incisors were removed using a high- speed dental handpiece, preserving approximately 5 mm of the middle root seg- ment with a diameter of 5 mm.The dentin cylinders were sequentially treated with 17% ethylenediaminetetraacetic acid (EDTA) for 10 min, followed by a 10 min rinse with deionized water; then with 10% EDTA for 10 min, followed by another 10 min rinse; and finally with 5% EDTA for 5 min, followed by a final 10 min rinse with deionized water. The resulting dentin scaffolds were immersed in sterile PBS containing penicillin and streptomycin for 72 h, ultrasonically cleaned in dis- tilled water for 10 min, and stored at 4 ◦ C until further use [6].

Animal experiment

A total of 12 male BALB/c-nu mice were purchased from Beijing Vital River Laboratory Animal Technology (Beijing, China). All animal procedures were ap- proved by the Animal Ethics Committee of Dalian Medical University (Approval No. AEE22081) and complied with institutional and national guidelines.

Mice were randomly assigned to three groups (n = 4 per group) according to the scaffold loaded into the TDM: (1) GelMA + hDPSC; (2) GelMA–DPEM + hDPSC; and (3) DPEM + hDPSC.

For theGelMA + hDPSCand GelMA–DPEM + hDPSC groups, third-passage hDPSCs were digested, centrifuged and resuspended as a single-cell suspension. A total of 2 × 104 hDPSCs were mixed with 2 mL of GelMA or GelMA–DPEM pre-gel solution, and 0.5 mL of the cell–hydrogel mixture was injected into the lumen of each TDM cylinder and photocrosslinked in situ using 405 nm blue light for 30 s.

For the DPEM + hDPSC group, hydrated DPEM blocks (approximately 0.5 × 0.5 cm) were co-cultured with hDPSCs in complete medium at 1 × 104 cells/mL for 24 h to promote cell adhesion, and the resulting DPEM–cell complexes were gently packed into TDM cylinders with an internal length of about 0.5 cm.

All TDM constructs were implanted subcutaneously into the dorsal region of nude mice. At 12 weeks post-implantation, mice were deeply anesthetized by intraperitoneal injection of sodium pentobarbital (50 mg/kg; Solarbio, China).

Once the absence of reflexes was confirmed, cervical dislocation was performed to ensure painless death. The subcutaneous implants were harvested and fixed in 4% paraformaldehyde for subsequent histological analysis.

Histological and immunostaining analysis

The explants retrieved from the subcutaneous tissue of nude mice were fixed in 4% paraformaldehyde and subsequently decalcified in 10% EDTA buffer for 3 months. After decalcification, the samples were processed through standard his- tological procedures, including dehydration, clearing, paraffin infiltration, embedding, and sectioning. Tissue Sect. (5 μm thick) were prepared for H&E staining to evaluate the formation of pulp-like tissue and neovascularization within the re- generated constructs.

Immunohistochemical (IHC) staining (Eliret, China) was performed to as- sess the formation of odontoblast-like cell layers and vascular structures. Briefly, paraffin sections were deparaffinized, rehydrated and subjected to heat-induced antigen retrieval in citrate buffer (pH 6.0) using microwave heating (95–100 ◦ C, 8 min), followed by natural cooling. After blocking with 5% bovine serum albumin (BSA) for 60 min at room temperature, sections were incubated overnight at 4 ◦ C with primary antibodies against dentin sialophosphoprotein (DSPP, 1:200; Thermo Fisher, USA), dentin matrix protein 1 (DMP1, 1:300; Thermo Fisher, USA) and platelet endothelial cell adhesion molecule-1 (CD31, 1:100; Thermo Fisher, USA). PBS was used instead of the primary antibody in negative control sections. After washing with tris buffered saline (TBS, Seville, China), sections were incubated with the appropriate secondary antibody for 60 min at room temperature, developed with DAB solution(Eliret, China), counterstained with hema- toxylin, dehydrated through graded ethanol, cleared in xylene and mounted with neutral resin for microscopic observation.

Statistical analysis

All statistical analyses were performed using GraphPad Prism 9.5 software. Data are presented as mean ± standard deviation (SD). Depending on the experi- mental design, one-way ANOVA, two-way ANOVA, or Student’s t-test was used to evaluate statistical significance. A p-value of less than 0.05 was considered statistically significant.

Results

Characterization of DPEM

The dental pulp tissue was decellularized, freeze-dried to thoroughly remove moisture, and then ground into DPEM powder to facilitate storage and application (Fig. 2A). H&E staining, along with Masson’s trichrome staining, showed that native dental pulp tissue had evenly distributed nuclei and cytoplasm, while the DPEM retained only the extracellular matrix with most cellular components effectively removed. SEM images revealed that native pulp tissue exhibited a dense, compact structure, whereas the DPEM displayed a three-dimensional fibrous network (Fig. 2B). These results confirm that the prepared DPEM effec- tively eliminated cellular components from the dental pulp tissue.

Fig. 2.

Preparation and characterization of DPEM. A DPEM powder preparation included decellularization, freeze drying, and grinding. B H&E staining, Masson’s trichrome staining, and SEM characterization of native pulp and DPEM

Characterization of GelMA-DPEM hydrogel and hDPSCs

The prepared GelMA–DPEM hydrogel initially exhibited a liquid state. Upon exposure to 405 nm blue light for 30 s, it rapidly formed a structurally stable gel with a smooth surface. When the precursor solution was loaded into a syringe and slowly extruded through a 23G needle into a transparent pulp cavity model, a continuous filament filled the simulated pulp chamber without clogging, qualitatively demonstrating its ability to be delivered through the needle and conform to the cavity shape. Furthermore, when injected into cylindrical molds (6 mm in diameter, 8 mm in height) and photo-crosslinked, the hydrogel formed well-defined cylindrical constructs with good shape retention, indicating its suit- ability for filling confined spaces analogous to the dental pulp cavity (Fig. 3A).

Fig. 3.

Preparation and characterization of GelMA–DPEM hydrogel. A Macroscopic appearance of the GelMA–DPEM precursor solution extruded through a 23G needle into a pulp-cavity model and cylindrical molds, showing round and columnar hydrogel constructs after photo-crosslinking. B Morphology of primary and third-passage hDPSCs under phase-contrast microscopy (scale bar = 500 µ m). C SEM images of the internal microstructure of GelMA and GelMA–DPEM hydrogels. D Frequency-sweep rheological analysis of GelMA and GelMA–DPEM hydrogels at 37.5 ◦ C, showing storage modulus (G′) and loss modulus (G′′); curves represent mean values from three independent samples per group (n = 3). E In vitro enzymatic degradation profiles of GelMA and GelMA–DPEM hydrogels in 2% (w/v) type I col- lagenase at 37.5 ◦ C (mean ± SD, n = 3). F Swelling behavior of GelMA and GelMA–DPEM hydrogels in PBS at 37.5 ◦ C (mean ± SD, n = 3). T: tissue blocks. Red arrows indicate represen- tative particulate/granular deposits observed on the pore surfaces of the GelMA–DPEM samples, whereas the GelMA pore walls appear comparatively smooth

Observations of hDPSCs showed that, with successive passages, the cells gradually adapted to the in vitro culture environment, exhibiting improved morphological stability and enhanced proliferative capacity. All subsequent experiments were conducted using third-passage (P3) cells (Fig. 3B).

SEM images of freeze-dried GelMA and GelMA-DPEM hydrogels revealed typical porous structures with relatively uniform pore sizes and smooth pore walls.

The GelMA surface showed no visible particles or deposits, while the GelMA-DPEM hydrogel exhibited a rougher surface texture, likely due to the incorporation of DPEM altering the originally homogeneous structural morphology (Fig. 3C). It should be noted that SEM mainly reveals the fibrillar collagen network and does not fully reflect amorphous ECM components, which may still be present but are less visible under these imaging conditions.

The rheological properties of GelMA and GelMA–DPEM hydrogels were analyzed using a HAAKE rheometer. Frequency-sweep tests showed that the storage modulus (G′) of the GelMA–DPEM hydrogel was higher than that of GelMA within the tested frequency range, indicating relatively increased viscoelastic stiff- ness after incorporation of DPEM (Fig. 3D).

In the in vitro collagenase-mediated degradation assay, both GelMA and GelMA–DPEM hydrogels exhibited gradual degradation over 24 h, with similar degradation profiles (Fig. 3E). A degradation rate of 100% indicates complete mass loss of the hydrogel, with no visible solid gel remaining under the test conditions. Swelling experiments revealed that the GelMA–DPEM hydrogel reached a swelling ratio of approximately 343%, while the GelMA hydrogel exhibited a swelling ratio of approximately 334%. These results indicate that the incorporation of DPEM did not significantly alter the swelling behavior of the GelMA hydrogel, and both hydrogels exhibited comparable swelling capacities (Fig. 3F).

Effects of GelMA–DPEM on cell viability, proliferation, and migration

Live/dead cell staining results showed that during the entire culture period (Day 1, 4, and 7), all groups (Control, GelMA, GelMA–DPEM, and DPEM) ex- hibited a marked increase in cell number, with consistently high cell viability maintained in each group (all above 90%) (Fig. 4A, B).

Fig. 4.

Live/dead cell staining, CCK-8 assay, and Transwell cell migration assay. A Live/dead staining images of hDPSCs co-cultured for 1, 4, and 7 days with complete medium, GelMA hydro- gel, GelMA–DPEM hydrogel, and medium containing bulk DPEM. Live cells are stained green, and dead cells are stained red (scale bar = 200 µ m). B Quantitative analysis of cell viability from the live/dead staining assay. C CCK-8 assay showing the proliferation of hDPSCs under different culture conditions. D Transwell assay evaluating the migratory capacity of hDPSCs under different culture conditions (scale bar = 200 µ m). E Quantitative analysis of the number of hDPSCs that migrated to the lower side of the membrane after 12 h. Data are presented as mean ± SD (n = 4). ns indicates no significant difference; *p < 0.05, ****p < 0.0001 versus the control group

CCK-8 assay results showed that there were no significant differences in hDPSC proliferation between the groups at Day 1, 3, 5, and 7. Cell viability increased normally over time in all experimental groups, indicating that the injectable GelMA–DPEM hydrogel exhibits good biocompatibility (Fig. 4C).

The Transwell assay evaluated cell migration ability by counting the number of cells that migrated to the opposite side of the membrane. The results showed that the GelMA–DPEM group significantly enhanced cell migration, exhibiting the strongest promotive effect. These findings indicate that the DPEM solution, even after pepsin digestion, retained its ability to promote the migration of hDP- SCs (Fig. 4D, E).

Pro-angiogenic effects of GelMA-DPEM hydrogel on hDPSCs

The tube formation assay showed that the GelMA–DPEM hydrogel group significantly enhanced angiogenic capacity compared to the other groups, forming the most dense and well-organized capillary-like networks (Fig. 5A). Quan- titative analysis (Fig. 5B) indicated no significant difference between the DPEM and GelMA–DPEM groups, yet both exhibited markedly higher angiogenic activity than the control and GelMA groups. These findings suggest that the GelMA– DPEM hydrogel retains the pro-angiogenic potential of DPEM.

Fig. 5.

GelMA–DPEM enhances multilineage differentiation of hDPSCs. A Tube formation of HUVECs under different conditions after 4 h of culture (scale bar = 500 μm). B Quan- titative analysis of the number of tubes formed. Relative mRNA expression levels of Nestin (C), DSPP (D), CD31 (E), and Runx2 (F) in hDPSCs cultured under control, GelMA, and GelMA– DPEM conditions. Data are presented as mean ± standard deviation (n = 3). ns indicates no significant difference; *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001 versus the GelMA– DPEM group

RT-qPCR analysis showed that the GelMA–DPEM hydrogel significantly upregulated the expression of Nestin (~ 20-fold vs. control) and DSPP (~ 12-fold vs. control) compared with the GelMA and control groups (mean ± SD, n = 3; p < 0.01), indicating enhanced odontoblast-like differentiation and dentin matrix formation (Fig. 5C, D). In addition, the expression of Runx2, a key transcription factor for early odontogenic/osteogenic commitment (~ 1.8-fold vs. control), and CD31, an endothelial marker associated with angiogenesis (~ 18-fold vs. control), was also markedly increased in the GelMA–DPEM group (mean ± SD, n = 3; p < 0.01; Fig. 5E, F), suggesting that the composite hydrogel simultaneously pro- motes odontogenic programming and pro-angiogenic responses.

In vivo regeneration of pulp-like tissue by GelMA-DPEM hydrogel

To evaluate the feasibility of using the GelMA–DPEM hydrogel as a potential dental pulp regeneration material, TDM segments approximately 5 mm in length were prepared (Fig. 6A). Various material formulations containing hDP- SCs were injected into the TDM and subcutaneously implanted into nude mice. After 12 weeks, the subcutaneous implantation sites were harvested. The over- lying skin appeared intact without ulceration, necrosis or obvious inflammatory complications, and the implant areas showed a comparable macroscopic appear- ance among groups (Fig. 6B).

Fig. 6.

Subcutaneous regeneration of pulp-like tissue in the dorsal region of nude mice. A Pre- pared TDM. B Materials implanted subcutaneously in the dorsal region of nude mice. C H&E staining images of pulp-like tissue regenerated using GelMA, bulk DPEM, and GelMA–DPEM hydrogels; red arrows indicate representative blood vessel–like structures within the regenerated tissue. The main panels have a scale bar of 200 µ m, and the yellow boxed regions show higher- magnification views with a scale bar of 100 µ m. n = 4

H&E staining revealed the formation of a cell-rich soft connective tissue within the treated dentin cylinders in all three groups, accompanied by newly formed blood vessel–like luminal structures (Fig. 6C). In the bulk DPEM and GelMA–DPEM groups, small-caliber vessels appeared more frequently within the regenerated tissue compared with the GelMA group, suggesting a trend to- ward enhanced neovascularization when DPEM was present.

Immunohistochemical staining revealed positive expression of DSPP and DMP1 – key markers associated with dentin formation – in the regenerated tissue of the GelMA, DPEM and GelMA–DPEM groups, particularly along the inner dentin wall and within the pulp-like tissue region. The endothelial marker CD31 was also positively expressed in microvessel-like structures in all three groups. In contrast, the PBS group used as a negative control exhibited no specific positive staining. Taken together, these findings indicate that all three types of scaffolds were able to support the formation of pulp-like tissue expressing odontogenic- and angiogenic-related markers in this ectopic model (Fig. 7).

Fig. 7.

Immunohistochemical staining results of subcutaneous implants in nude mice. DSPP and DMP1, key markers of dentin formation, showed positive expression in the GelMA, DPEM, and GelMA–DPEM groups. CD31, an endothelial marker, was also positively expressed in all three groups. The PBS group, used as a blank control, showed negative staining results. These findings indicate the odontogenic and angiogenic activity of the tested materials. Scale bar = 500 µ m. n = 4

Immunohistochemical staining revealed positive expression of DSPP and DMP1 – key markers for dentin formation – in the GelMA, DPEM, and GelMA– DPEM groups. The endothelial marker CD31 also showed positive expression in all three groups. In contrast, the PBS group used as a negative control exhibited no positive staining. These findings indicate the odontogenic and angiogenic potential of the tested materials (Fig. 7).

Discussion

Both this study and previous research have demonstrated that DPEM effec- tively removes cellular components from dental pulp tissue while preserving the functional extracellular matrix (ECM) scaffold, making it suitable for applica- tions in dental pulp regeneration [32]. However, due to the complex and curved morphology of the root canal system, solid DPEM cannot achieve complete filling, leading to voids that hinder regeneration [33]. Injectable DPEM was obtained through physical grinding, demonstrating promising potential for tissue regenera- tion. However, its granular form limited its injectability, making it unsuitable for clinical application. Therefore, this study aims to develop a photo-crosslinkable GelMA–DPEM composite hydrogel with an extrudable precursor solution, designed as an ideal scaffold for dental pulp tissue engineering.

First, decellularization of dental pulp tissue was performed. Our experimen- tal results, consistent with findings by Bakhtiar et al., showed that the process effectively removed cellular components while retaining key ECM proteins such as type I collagen and laminin [34]. Yuan et al. further confirmed that after 12hours of decellularization, the ECM structure remained intact with minimal DNA content (47.60 ± 2.53 ng/mg), validating the effectiveness of the decellularization process [35]. Pepsin digestion is a commonly used method to convert natural tis- sues into soluble forms by enzymatically degrading collagen and other macro- molecules, resulting in materials that are easy to inject [36, 37]. Multiple studies have confirmed the feasibility and biosafety of this technique [38]. In our study, we combined pepsin-digested DPEM with GelMA to form a homogeneous, photo- crosslinkable hydrogel. Li et al. and Bakhtiar et al. have previously used similar approaches to prepare dental pulp matrix solutions, further verifying the method’s feasibility and biosafety [34, 39]. Bioactive components, making it more effective in promoting dental pulp regeneration [40–42].

Subsequently, we evaluated the effects of the GelMADPEM hydrogel on hDPSC proliferation, migration, and angiogenesis. Yue et al. reported that GelMA, a hydrogel synthesized from gelatin and methacrylic acid, could be stably photo-crosslinked under blue light to form a three-dimensional network [43]. Yuan et al. highlighted that GelMA’s hydrophilicity and biodegradability provide an ideal microenvironment for cell growth, supporting stem cell proliferation and differentiation [35]. To enhance GelMA’s mechanical properties and biological functionality, researchers often combine it with other materials. For example, Wang et al. demonstrated that GelMA’s mechanical strength improves when combined with calcium phosphate ceramics or hydroxyapatite [44]; Barati et al. showed that incorporating growth factors like BMP-2 and VEGF enhances cell proliferation and differentiation [45]; and Dorterler et al. reported that combining GelMA with nano-hydroxyapatite boosts dentin regeneration and mineralization [46].

In this context, the GelMA–DPEM system used in the present study combines the extrudability and in situ photocrosslinking of GelMA with the tissue-specific bioactivity of dental pulp ECM, offering a balance between mechanical support and biological signaling. Injectable GelMA-based hydro- gels have been widely employed as cell carriers for minimally invasive tissue engineering, and previous work has shown that encapsulated cells can maintain high viability after extrusion or bioprinting of GelMA constructs under optimized conditions [47]. For application in the pulp chamber, blue light/visible-light activated GelMA hydrogels allow minimally invasive delivery, in situ shaping and rapid gelation; however, light penetration in long or curved root canals and the choice of photoinitiator concentration and exposure parameters must be care- fully controlled to avoid potential phototoxic or thermal effects. Our findings suggest that GelMA–DPEM retains these practical advantages of GelMA while introducing pulp-derived ECM components that may contribute to the observed pro-angiogenic and odontogenic responses.

In this study, pepsin-digested DPEM was combined with GelMA to create a novel photo-crosslinkable hydrogel with an extrudable precursor solution. One of the major challenges in pulp regenera- tion is re-establishing a functional vascular network. Due to the highly vascu- larized nature of dental pulp, angiogenesis is critical for tissue metabolism and survival, especially in damaged or necrotic pulp [48]. Liang et al. found that DPEM significantly promoted hDPSC differentiation toward odontogenic, angio-genic, and neurogenic lineages by providing a bioactive microenvironment with essential exogenous signals [49]. Our results indicated that the GelMA-DPEM hy- drogel formed dense, well-defined vascular-like structures, demonstrating strong angiogenic potential. The composite hydrogel provided a three-dimensional scaf- fold and a conduit for growth factor delivery, further enhancing endothelial cell growth and vascular network formation. In our RT-qPCR panel, Nestin, DSPP, Runx2, and CD31 were chosen as representative markers of dentin–pulp complex regeneration. Nestin is expressed in dental pulp progenitor cells and newly dif- ferentiated odontoblasts, and is commonly used as a marker of odontoblast-like and neurogenic differentiation. DSPP is a dentin-specific matrix protein that is essential for dentin mineralization and proper dentinogenesis. Runx2 is a mas- ter transcription factor that governs early odontogenic/osteogenic differentiation and the expression of dentin/bone matrix genes, whereas CD31 reflects endothe- lial cell presence and angiogenic activity, which are crucial for supporting dentin formation. In this study, the GelMA–DPEM hydrogel significantly increased the expression of Nestin, DSPP, Runx2, and CD31 compared with GelMA alone, indicating that incorporation of DPEM enhances odontoblast-like differentiation and dentin matrix production while simultaneously promoting a pro-angiogenic environment conducive to dentin–pulp complex regeneration.

Furthermore, we evaluated the in vivo regenerative effects using a subcu- taneous ectopic implantation model in nude mice. HE staining revealed a higher density of neovascularization inthe GelMA-DPEM group compared tothe GelMA group. Li et al. emphasized the critical role of vascularization in supplying nutri- ents, removing waste, and supporting tissue viability during pulp regeneration [48]. Studies have shown that standalone GelMA may lack the bioactivity of natural.

ECM components such as collagen or glycosaminoglycans, limiting its angio- genic potential [50]. Goldberg et al. attributed DPEM’s angiogenic ability to its unique composition of peptides and glycosaminoglycans, which interact with angiogenic biomarkers to promote vessel formation [8, 49]. Our results suggest that the addition ofDPEM enhanced the biocompatibility and bioactivity of the hydro- gel, leading to improved angiogenesis.

Immunohistochemistry further validated these findings. DSPP and DMP1, markers of dentinogenesis, were positively expressed in newly formed pulp-like tissue, consistent with results from Yue et al. [42, 51, 52]. CD31, an endothelial marker, was also strongly expressed, indicating robust angiogenesis [53]. Collectively, these data support the in vivo efficacy of the GelMA–DPEM hydrogel in promoting both vascular and dentin regeneration, suggesting that it is a promising candidate scaffold for dental pulp regeneration, although further studies in ortho- topic and long-term models are required before clinical translation.

In summary, this study suggests that a bioactive photo-crosslinkable GelMA–DPEM hydrogel can support pulp-like tissue regeneration in an ectopic model and provides preliminary evidence of its potential use in clinical endodontic applications. Nevertheless, several important limitations should be noted. First, we relied on 2D assays for in vitro screening and an ectopic subcutaneous TDM model to assess 3D pulp-like tissue formation, but did not perform dedicated 3D in vitro encapsulation studies, and the ectopic model does not fully reproduce the native dental pulp microenvironment. Second, mechanical evaluation was restricted to small-amplitude rheology and short- to mid-term degradation; the compressive modulus, photo-crosslinking depth under clinically relevant light-curing conditions, and the influence of GelMA–DPEM stiffness and mechanical cues on hDPSC fate were not assessed. Third, although porcine DPEM was decellularized, its immunogenicity was not systematically examined and no cell-tracking techniques were used, so the relative contributions of transplanted versus host-derived cells remain unclear. In addition, we did not perform new proteomic characterization of DPEM; based on previous work, major structural ECM proteins are likely preserved whereas some soluble non-collagenous components and growth factors may be partially lost, and donor- and processing-related batch variability in ECM composition may influence bioactivity and hinder standardization for large-scale clinical use [48]. Finally, angiogenesis was evaluated only by tube formation, CD31 expression and histological identification of microvessel-like structures, without quantification of key pro-angiogenic mediators such as VEGF, and more comprehensive angiogenic profiling will be required in future studies to better characterize the vascular responses induced by GelMA–DPEM.

Conclusion

In this study, a photo-crosslinkable GelMA–DPEM hydrogel with a syringe-deliverable precursor solution was successfully developed as a scaffold for pulp regeneration. Preliminary char- acterization showed that the hydrogel formed a stable crosslinked network with appropriate swelling and degradation behavior. In vitro experiments confirmed that the hydrogel provided a supportive microenvironment for hDPSC growth, proliferation, and migration, indicating good cytocompatibility and potential for tissue repair applications. In a small-animal ectopic TDM implantation model, the hydrogel combined with hDPSCs supported the formation of cell-rich, vascularized pulp-like tissue with CD31-positive neovessels, suggesting preliminary pro-angiogenic effects and potential utility as a candidate scaffold for dental pulp regeneration. Future investigations should include more comprehensive mechanical and injectability testing, long-term in vivo evaluation in orthotopic and large- animal models, and further optimization of the hydrogel’s composition and functional properties to enhance its regenerative efficacy and facilitate its eventual clinical translation for endodontic applications. In conclusion, the GelMA–DPEM hydrogel showed favorable cytocompatibility and preliminary pro-angiogenic effects in vitro and supported pulp-like tissue formation in a small-animal ectopic TDM model.

Authors’ contributions

Y.Y.: Conceptualization, Methodology, Investigation, Data Curation, Formal Analysis, Writing – Original Draft. J.Z.: Methodology, Validation, Cell Experiments, Visualization, Writing – Review & Editing. B.Q.: Animal Experiments, Histological Analysis, Data Interpretation, Writing – Review & Editing.Z.X.: Data Curation, Software, Statistical Analysis. L.W.: Material Preparation, Mechanical Testing, Formal Analysis. W.S.: Resources, Supervision, Project Administration. Z.S.: Visualization, Manuscript Revision. R.Z.: Investigation, Language Polishing, Technical Support. H.H.: Resources, Conceptual Support. G.C.: Conceptualization, Supervision, Funding Acquisition, Writing – Review & Editing, Project Administration, Corresponding Author.

Funding

This research was funded by the National Natural Science Foundation of China (China, 81600818).

Data availability

The data underlying this article will be shared upon reasonable request to the corresponding author.

Declarations

Ethics approval and consent to participate

The pulp tissue was derived from orthodontic teeth or third molars extracted for therapeutic purposes from the First Affiliated Hospital of Dalian Medical University, and the pulp tissue used met the relevant ethical requirements of the First Affiliated Hospital of Dalian Medical University (PJ-KS-KY-2019-174). A total of 12 SPF male BALB/c-nu mice purchased from Beijing Weitong Lihua Technology Co., Ltd., 8 weeks old, mass 18–22 g, license number: SYXK (Beijing) 2021-0006. The animal experiment plan was approved by the Animal Experiment Ethics Committee of Dalian Medical University, and the approval number is AEE22081. The experimental process followed the International Association of Veterinary Editors Consensus on Authors’ Guidelines on Animal Ethics and Welfare and local and national regulations.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.