An innate defense regulator peptide modulates the inflamed pulp microenvironment and promotes pulp repair

Abstract

Persistent infection and unresolved inflammation remain major obstacles to successful vital pulp therapy. Strategies capable of simultaneously controlling bacterial burden and regulating the inflamed pulp microenvironment remain limited. In this study, we investigated the effects of the innate defense regulator peptide IDR-1002 on pulp repair under inflammatory conditions. IDR-1002 showed antibacterial activity against multispecies biofilms, Streptococcus mutans, and Enterococcus faecalis. In human dental pulp stem cells, IDR-1002 reduced LPS-induced expression of IL-6, IL-1β, and TNF-α, attenuated intracellular reactive oxygen species accumulation, and improved wound closure. It also enhanced odontogenic differentiation, as evidenced by increased DSPP and DMP1 expression and enhanced mineral deposition. Exploratory transcriptomic analysis identified enrichment of biological processes potentially associated with IDR-1002 treatment, including extracellular matrix organization, receptor signaling, and tissue repair. To develop a local, microenvironment-responsive delivery system, IDR-1002 was incorporated into an MMP-responsive hydrogel. Biologically active IDR-1002 was released following MMP-9-triggered hydrogel degradation. In a rat pulpitis model, IDR-1002 reduced inflammatory cell infiltration, decreased IL-6 and IL-1β expression, and improved tissue organization at the exposure site. Collectively, these findings demonstrate that IDR-1002 possesses antibacterial, anti-inflammatory, antioxidative, and pro-regenerative properties in the experimental models used in this study and support further investigation of host-regulatory peptides as potential adjunctive agents for biologically based vital pulp therapy.

Graphical abstract

IDR-1002 attenuates bacterial challenge, inflammation, and oxidative stress while promoting mineralization-related responses. An MMP-responsive hydrogel enables inflammation-triggered release of biologically active IDR-1002, supporting pulp repair and mineralized tissue formation during vital pulp therapy.

Highlights

• •IDR-1002 attenuated inflammatory cytokine expression and oxidative stress in hDPSCs.
• •IDR-1002 promoted wound healing and odontogenic mineralization under inflammatory conditions.
• •IDR-1002 exhibited antibacterial activity against oral biofilms and dentinal tubule infection.
• •An MMP-responsive hydrogel enabled inflammation-associated release of biologically active IDR-1002.
• •IDR-1002 reduced pulp inflammation and enhanced reparative dentin-related tissue formation in vivo.

1. Introduction

Dental caries is the most prevalent human disease worldwide and remains a major cause of pulpal inflammation [1]. As lesions progress, bacteria and their virulence factors penetrate the dentin–pulp complex, triggering host immune responses within the pulp tissue [2,3]. In addition to microbial invasion, pulpitis is characterized by dysregulated inflammatory signaling, oxidative stress, and extracellular matrix degradation, all of which can compromise tissue homeostasis and repair [4].

Vital pulp therapy aims to preserve pulp vitality and function under these conditions [5,6]. Current approaches, including indirect and direct pulp capping and pulpotomy, focus on removal of infected tissue and sealing with bioactive materials [[7], [8], [9]]. Although these procedures are often successful, treatment outcomes remain variable, particularly in teeth with moderate to severe inflammation [[10], [11], [12]]. Existing strategies primarily address bacterial control and tissue sealing but provide limited regulation of the inflammatory microenvironment, which may contribute to persistent inflammation and impaired repair.

Several factors contribute to this challenge [[5], [6], [7], [8], [9]]. Reliable molecular tools for assessing pulpal inflammation are not currently available in clinical practice, and treatment decisions often rely on indirect indicators. Consequently, inflamed tissue may be incompletely removed. In addition, although calcium silicate–based materials provide favorable sealing ability and bioactivity, their capacity to directly regulate inflammatory processes is limited. These considerations highlight the need for therapeutic approaches that combine antibacterial activity with modulation of the local microenvironment [13,14].

Host defense peptides (HDPs) are important components of innate immunity and exhibit both antimicrobial and immunomodulatory activities [15,16]. In addition to direct antibacterial effects, HDPs influence inflammatory responses and tissue homeostasis. Among synthetic innate defense regulator peptides, IDR-1002 has demonstrated anti-inflammatory and tissue-reparative activities in several experimental models [[17], [18], [19], [20], [21]]. However, its effects on inflamed dental pulp and pulp repair remain largely unexplored.

Localized delivery of bioactive agents is an important consideration for regulating the pulp microenvironment. Hydrogels provide a hydrated matrix that supports cell viability while enabling local delivery of therapeutic molecules [22]. Stimuli-responsive hydrogel systems can further modulate drug release in response to pathological cues such as pH, oxidative stress, or enzymatic activity [[23], [24], [25], [26]]. Among these stimuli, matrix metalloproteinases (MMPs), particularly MMP-9, are upregulated during pulp inflammation and contribute to extracellular matrix degradation [[27], [28], [29], [30], [31]]. This characteristic provides an opportunity to engineer responsive delivery systems that release therapeutic agents in response to inflammatory conditions [32,33].

In this study, we investigated the antibacterial, anti-inflammatory, antioxidative, and pro-regenerative effects of the innate defense regulator peptide IDR-1002 under inflammatory conditions. Its effects on bacterial viability, inflammatory responses, oxidative stress, wound closure, and odontogenic differentiation were evaluated in human dental pulp stem cells (hDPSCs). An MMP-responsive hydrogel was further employed to examine whether microenvironment-triggered release could preserve the biological activity of IDR-1002. Finally, the therapeutic potential of IDR-1002 was evaluated in a rat pulpitis model.

2. Materials and methods

2.1. Synthesis of host defense peptide IDR-1002

The host defense peptide IDR-1002 (VQRWLIVWRIRK; molecular weight: 1652.05 Da) was synthesized by GenScript Biotech Corp. (Piscataway, NJ, USA) using standard solid-phase 9-fluorenylmethoxy carbonyl (Fmoc) chemistry and purified to ≥95% by reverse-phase high-performance liquid chromatography (RP-HPLC). Amino acid analysis was performed to confirm peptide composition, and molecular weight was verified by mass spectrometry.

2.2. Evaluation of the cytocompatibility of IDR-1002 in hDPSCs

hDPSCs (Lonza, Walkersville, MD, USA) were cultured in α-MEM (Gibco, Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10% fetal bovine serum (FBS; Gibco) at 37 °C in a humidified incubator containing 5% CO₂. Cells (passage 4; ∼80% confluence) were seeded into 96-well plates and cultured for 24 h prior to treatment.

Experimental groups consisted of control medium (α-MEM supplemented with 10% FBS) or IDR-1002 at 20, 50, 80, or 100 μg/mL. The selected concentrations were based on previous studies demonstrating the immunomodulatory activity and low cytotoxicity of IDR-1002 in mammalian cells and hDPSCs [[17], [18], [19], [20], [21]]. At 24, 48, and 72 h, 10 μL of Cell Counting Kit-8 (CCK-8; Abcam, Cambridge, UK) reagent was added to each well and incubated for 2 h. Absorbance was measured at 450 nm using a microplate reader. Cell viability was expressed as a percentage of the corresponding control group after subtraction of background absorbance. Relative optical density (OD, %) values were normalized to the control group, which was set as 100%.

2.3. Evaluation of LPS-induced inflammatory cytokine expression in hDPSCs following IDR-1002 treatment

hDPSCs (passage 3; ∼80% confluence) were seeded into 6-well plates and cultured for 24 h to allow attachment. Cells were then assigned to the following groups (n = 3 per group): (i) control (α-MEM supplemented with 10% FBS); (ii) LPS (1 μg/mL; Escherichia coli LPS, Sigma-Aldrich, St. Louis, MO, USA); (iii) LPS + IDR-1002 (20 μg/mL); (iv) LPS + IDR-1002 (50 μg/mL); and (v) LPS + IDR-1002 (80 μg/mL). For inflammatory stimulation experiments, LPS and IDR-1002 were added simultaneously and cells were incubated for 24 h before analysis.

2.3.1. Quantitative real-time PCR (RT–qPCR)

Total RNA was extracted using TRIzol reagent (Invitrogen, Carlsbad, CA, USA). One-step RT–qPCR was performed using the Luna Universal One-Step RT–qPCR Kit (New England Biolabs, Pickering, ON, Canada) on a StepOnePlus Real-Time PCR System (Applied Biosystems, Waltham, MA, USA). The mRNA expression levels of IL6, IL1B, and TNFA were quantified using primers listed in Table S1. Relative gene expression was calculated using the 2^−ΔΔCt method with ACTB as the reference gene. All experiments were performed using three independent biological replicates.

2.3.2. Western blot analysis

Total protein was extracted using radioimmunoprecipitation assay (RIPA) lysis buffer (AS1004, ASPEN, USA), and protein concentrations were determined using a bicinchoninic acid (BCA) protein assay (AS1086, ASPEN). Equal amounts of protein (40 μg) were separated by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE; AS1012, ASPEN, USA) and transferred onto 0.45 μm polyvinylidene fluoride (PVDF) membranes (IPVH00010, Millipore).

Membranes were blocked with 5% bovine serum albumin (BSA; Sigma-Aldrich) for 1 h at room temperature and incubated overnight at 4 °C with primary antibodies. After washing with Tris-buffered saline containing 0.1% Tween-20 (TBST), membranes were incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies for 30 min at room temperature. Antibody information is provided in Table S2. Protein bands were visualized using enhanced chemiluminescence (ECL) reagent (AS1059, ASPEN, USA) and quantified using the AlphaEaseFC analysis system (Alpha Innotech, USA). Experiments were performed in triplicate (n = 3 per group).

2.4. Evaluation of the antibacterial activity of IDR-1002 against multispecies and S. mutans biofilms

2.4.1. Establishment of multispecies and S. mutans biofilm models on HA discs

Sterile hydroxyapatite (HA) discs (11.8 mm in diameter and 1.7 mm in thickness; Baiamon Bioactive Materials Co., Ltd., Chengdu, China) were used as biofilm substrates as previously described [34,35]. Discs were coated with bovine dermal type I collagen (PureCol®, Advanced Biomatrix, San Diego, CA, USA) overnight at 4 °C in 24-well plates (Corning, Corning, NY, USA).

Streptococcus mutans (ATCC 25175) was cultured on BHI sheep-blood agar plates under anaerobic conditions at 37 °C for 48 h. Colonies were harvested and suspended in brain–heart infusion (BHI) broth (Becton, Dickinson and Company, Franklin Lakes, NJ, USA). Supragingival plaque samples were collected from a healthy adult volunteer using sterile wooden toothpicks and suspended in BHI broth to generate a multispecies bacterial inoculum. Both suspensions were adjusted to OD405 = 0.10. Collagen-coated HA discs were placed in 24-well plates containing 1.8 mL sterile BHI and 0.2 mL bacterial suspension per well and incubated anaerobically at 37 °C for 7 days to establish biofilms.

2.4.2. IDR-1002 treatment and CLSM analysis

IDR-1002 was dissolved in sterile deionized water at final concentrations of 10, 20, 50, and 100 μg/mL. Biofilm specimens were rinsed with phosphate-buffered saline (PBS) and randomly assigned to five groups: (i) control (sterile deionized water); (ii) IDR-1002 (10 μg/mL); (iii) IDR-1002 (20 μg/mL); (iv) IDR-1002 (50 μg/mL); and (v) IDR-1002 (100 μg/mL).

Biofilms were treated with the assigned solutions and analyzed by confocal laser scanning microscopy (CLSM) as previously described [34,35]. A 50 μL aliquot of the designated solution was applied to the biofilm surface for 3 min. Following treatment, biofilms were rinsed with PBS and stained using the LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes, Eugene, OR, USA).

Biofilms were imaged using CLSM (FV10i-LIV, Olympus, Richmond Hill, Canada). Fluorescence signals were reconstructed into three-dimensional volumes using Imaris software (v7.4.2; Bitplane AG, Zurich, Switzerland). Live and dead bacterial volumes were quantified, and bacterial killing was expressed as the percentage of dead bacteria calculated from the ratio of red fluorescence volume to total fluorescence volume (red + green). Ten independent specimens were analyzed per group.

2.5. Evaluation of the antibacterial activity of IDR-1002 against E. faecalis within dentinal tubules

2.5.1. Dentin block preparation

Dentin blocks were prepared as previously described [36,37]. Human single-rooted premolars extracted for reasons unrelated to this study were collected and sectioned perpendicular to the long axis to obtain 4 mm root segments. Root canals were enlarged using a #3 Gates–Glidden drill.

Each segment was split longitudinally into two semi-cylindrical halves. External surfaces were polished to standardize specimen thickness and remove residual cementum. Additional trimming was performed using a slow-speed handpiece (300 rpm) to ensure compatibility with Nanosep centrifugal filter units (Pall Corporation, Ann Arbor, MI, USA). To remove the smear layer, specimens were ultrasonically treated in 5% NaOCl and 6% citric acid for 4 min each, rinsed with sterile water, neutralized with 5% Na₂S₂O₃ for 30 s, and sterilized by autoclaving at 121 °C for 30 min.

2.5.2. Establishment of an Enterococcus faecalis-infected dentinal tubule model

A dentinal tubule infection model was established as previously described [36,37]. E. faecalis (ATCC 29212) was cultured on BHI agar plates under anaerobic conditions at 37 °C for 24 h. Colonies were harvested and suspended in BHI broth to obtain a bacterial suspension of approximately 3 × 10⁶ CFU/mL.

Each dentin specimen was mounted within a Nanosep centrifugal filter unit with the canal surface facing upward. The specimen–filter interface was sealed using flowable composite resin (Filtek A2; 3M ESPE, St. Paul, MN, USA).

For bacterial inoculation, 500 μL of suspension was applied to the canal surface. Specimens were subjected to sequential centrifugation at 5000, 6000, 8000, and 10,000 rpm for 5 min each, repeated twice, with fresh bacterial suspension added before each cycle. Specimens were subsequently incubated in BHI broth under anaerobic conditions at 37 °C for 7 days to establish dentinal tubule infection.

2.5.3. IDR-1002 treatment and CLSM analysis

Infected dentin specimens were removed from the filter units, rinsed with PBS, and randomly assigned to treatment groups as described in Section 2.4.2.

Following treatment with IDR-1002, specimens (n = 10 per group) were analyzed by CLSM as previously described [36,37]. Briefly, a 50-μL aliquot of the designated solution was applied to the canal surface for 3 min. Specimens were then vertically fractured to expose fresh dentinal tubule surfaces. The exposed surfaces were stained using the LIVE/DEAD BacLight Bacterial Viability Kit (Molecular Probes) and imaged using CLSM (FV10i-LIV, Olympus). Three-dimensional image reconstruction and quantitative analysis were performed using Imaris software (v7.4.2; Bitplane AG). Antibacterial activity was expressed as the percentage of dead bacteria calculated from the ratio of red fluorescence volume to total fluorescence volume.

2.5.4. Penetration of IDR-1002 into dentinal tubules

5-FAM-labeled IDR-1002 (VQRWLIVWRIRK; molecular weight: 2123.54 Da; C-terminal amidation; N-terminal 5-FAM–Ahx modification) was synthesized by GenScript Biotech Corp. and purified to ≥95% by RP-HPLC.

Dentin specimens were prepared as described in Section 2.5.1. A freshly prepared solution of 5-FAM-labeled IDR-1002 (50 μg/mL) was used for the penetration experiment. Specimens were randomly assigned to a control group (deionized water) or an IDR-1002 group (n = 3 per group). Following drying of the canal surface, 50 μL of the designated solution was applied to the canal dentin surface for 3 min and subsequently removed.

Specimens were then split longitudinally to expose dentinal tubules and imaged using confocal laser scanning microscopy (FV10i-LIV, Olympus). Images were acquired using identical magnification and acquisition settings across all groups. Fluorescence signals were reconstructed using Imaris software (v7.4.2; Bitplane AG), and the penetration depth of 5-FAM-labeled IDR-1002 from the canal surface toward the outer dentin was measured using ImageJ.

2.6. Evaluation of the inhibitory activity of IDR-1002 against S. mutans and multispecies bacteria

S. mutans (ATCC 25175) and multispecies bacterial suspensions were prepared as described in Section 2.4.1. The initial OD600 was adjusted to 0.01.

Bacterial suspensions were treated with deionized water (control) or IDR-1002 at 10, 20, 50, and 100 μg/mL and incubated anaerobically at 37 °C. Bacterial growth was monitored by measuring OD600 at 6, 12, 24, and 48 h. Relative OD values were normalized to the control group at the corresponding time point and expressed as percentages (n = 3).

To evaluate inhibition of multispecies biofilm formation, bacterial suspensions were seeded into 96-well plates and treated with IDR-1002 (0, 10, 20, 50, and 100 μg/mL; n = 3 per group) for 72 h under anaerobic conditions. Biofilm biomass was quantified by crystal violet staining. Briefly, biofilms were stained with 0.1% crystal violet (Millipore Sigma), and bound dye was solubilized in ethanol. Absorbance was measured at 570 nm and normalized to the untreated control group.

2.7. Preparation of matrix metalloproteinase-responsive hydrogels

The MMP-cleavable peptide linker (CVPLS↓LYSGC; molecular weight: 1041.25 Da) was synthesized by GenScript Biotech Corp. using standard solid-phase Fmoc chemistry. Blank and IDR-1002-loaded hydrogels were fabricated according to a previously reported protocol [25,33]. The peptide sequence was selected based on its reported susceptibility to MMP-9-mediated cleavage [38,39].

Four-arm poly(ethylene glycol)-norbornene (PEG-NB, 10 kDa; Creative PEGWorks, Durham, NC, USA) and the peptide linker were dissolved in PBS containing 0.05% (w/w) lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP; Millipore Sigma). The precursor solution was prepared at a final polymer concentration of 4% (w/w) with a thiol-to-ene ratio of 1:1.

The precursor solution was transferred into flat-bottom 96-well plates and photocrosslinked under 365 nm UV irradiation (10 W) for 30 s. For peptide-loaded hydrogels, IDR-1002 (50 μg/mL) was incorporated directly into the precursor solution before crosslinking. Cylindrical hydrogel specimens approximately 6.5 mm in diameter and 3 mm in height (∼100 μL) were prepared.

2.8. Evaluation of the cytocompatibility of IDR-1002–loaded hydrogels

In our previous study, hydrogel degradation and IDR-1002 release in response to pre-activated MMP-9 were characterized using HPLC analysis [25]. However, the cytocompatibility and biological effects of hydrogel-derived products on hDPSCs were not evaluated. Therefore, the present study investigated the cytocompatibility of hydrogel-derived products and their effects on hDPSCs in the context of vital pulp therapy.

2.8.1. Preparation of hydrogel leachates and degradation products

Blank hydrogel leachates (BHL) and IDR-1002-loaded hydrogel leachates (PHL) were prepared by incubating hydrogels in α-MEM at 37 °C for 24 h under gentle shaking. Each hydrogel specimen was immersed in 2 mL of α-MEM.

Blank hydrogel degradation products (BHD) and IDR-1002-loaded hydrogel degradation products (PHD) were prepared by incubating hydrogels in α-MEM containing 20 nM pre-activated recombinant human MMP-9 (molecular weight: 66 kDa; Millipore Sigma) under identical conditions. Following incubation, supernatants were sterilized through a 0.2 μm membrane filter and stored at 4 °C for up to 24 h prior to use [33].

2.8.2. CCK-8 assay

hDPSCs (passage 4; ∼80% confluence) were seeded into 96-well plates and cultured for 24 h. Cells were then exposed to undiluted BHL, PHL, BHD, or PHD supplemented with 10% FBS. After 24 h, cell viability was evaluated using the CCK-8 assay as described in Section 2.2 (n = 3 per group).

2.8.3. CLSM evaluation of hDPSC viability after exposure to hydrogel-derived products

hDPSCs (passage 4; ∼80% confluence) were seeded onto glass-bottom dishes (MatTek Corporation, Ashland, MA, USA) and cultured for 24 h. Cells were then assigned to four groups (n = 3 per group): (i) control; (ii) BHD; (iii) PHL; and (iv) PHD.

Cell viability was evaluated using the LIVE/DEAD™ Viability/Cytotoxicity Kit (Invitrogen) and imaged by CLSM (FV10i-LIV, Olympus). Images were acquired using identical acquisition settings across all groups. Dead cells were quantified using ImageJ and normalized to the analyzed area.

2.8.4. CLSM evaluation of LPS-stimulated hDPSC viability

hDPSCs (passage 4; ∼80% confluence) were seeded and cultured as described above. Cells were assigned to four groups (n = 3 per group): (i) control; (ii) LPS (1 μg/mL); (iii) LPS + PHL; and (iv) LPS + PHD. Live/dead staining, CLSM imaging, and quantitative analysis were performed as described in Section 2.8.3.

2.9. Cyclo(-RGDfC) peptide modification of hydrogels

IDR-1002–loaded hydrogel precursor was prepared as described in Section 2.7. Following sterile filtration (0.22 μm), the precursor solution was transferred into hydrogel curing rings (EFL Group, Suzhou, China) and photocrosslinked under 365 nm UV irradiation (10 W) for 5 s.

To enhance cell adhesion, cyclo(-RGDfC) peptide (APExBIO Technology LLC, Houston, TX, USA) was dissolved at 5 mg/mL in PBS containing 0.05% (w/w) LAP. Following sterile filtration, the peptide solution was applied to the hydrogel surface and immobilized via a thiol–ene photoclick reaction under 365 nm UV irradiation (10 W) for 30 s [[40], [41], [42], [43]].

Modified hydrogels were incubated in α-MEM supplemented with 10% FBS for 4 h to remove unreacted molecules. Unmodified and cyclo(-RGDfC)-modified hydrogels (n = 3 per group) were placed in 24-well plates and seeded with hDPSCs (passage 4; ∼80% confluence).

Cell viability was evaluated by live/dead staining and CLSM after 1 and 3 days of culture. After 1 month, F-actin staining was performed using Alexa Fluor™ 488 Phalloidin (Thermo Fisher Scientific), followed by CLSM imaging.

2.10. Evaluation of the antibacterial activity of hydrogel-derived products against S. mutans and multispecies oral bacteria

S. mutans (ATCC 25175) and multispecies bacterial suspensions were prepared as described in Section 2.6. The initial OD600 was adjusted to 0.01. Bacterial suspensions were assigned to three groups: (i) control (sterile deionized water); (ii) PHL; and (iii) PHD. Samples were incubated under anaerobic conditions at 37 °C, and bacterial growth was monitored by measuring OD600 at 6, 12, 24, and 48 h. Relative OD values were normalized to the control group at the corresponding time point and expressed as percentages (n = 3).

2.11. Expression of inflammatory cytokines in hDPSCs following exposure to hydrogel-derived products

hDPSCs (passage 3; ∼80% confluence) were seeded into 6-well plates and treated as described in Section 2.8.4 (n = 3 per group). Total RNA was extracted and RT–qPCR was performed as described in Section 2.3.1. Relative mRNA expression levels of IL6, IL1B, and TNFA were calculated using the 2^−ΔΔCt method with ACTB as the reference gene.

2.12. Scratch-wound assay

hDPSCs (passage 3; ∼80% confluence) were seeded in 6-well plates and serum-starved for 24 h. A linear scratch was generated using a sterile 200 μL pipette tip, and detached cells were removed with PBS.

Cells were assigned to five groups (n = 3 per group): (i) control (α-MEM supplemented with 10% FBS); (ii) LPS (1 μg/mL in α-MEM supplemented with 10% FBS); (iii) BHD + LPS in α-MEM supplemented with 10% FBS; (iv) PHL + LPS in α-MEM supplemented with 10% FBS; and (v) PHD + LPS in α-MEM supplemented with 10% FBS. Phase-contrast images were captured at 0, 12, and 24 h using identical acquisition settings. Wound closure (%) was calculated using ImageJ according to [(S₀−S_t)/S₀] × 100, where S₀ and S_t represent the wound area at baseline and at the indicated time point, respectively.

2.13. Measurement of intracellular ROS

Intracellular reactive oxygen species (ROS) levels were measured using 2′,7′-dichlorofluorescein diacetate (DCFH-DA; Sigma-Aldrich). hDPSCs (passage 3; ∼80% confluence) were seeded in 6-well plates and cultured for 24 h, followed by treatment as described in Section 2.12 (n = 3 per group).

Cells were incubated with DCFH-DA (10 μM) for 30 min in the dark, rinsed with PBS, and imaged by CLSM. Fluorescence intensity was quantified using ImageJ. Integrated density (IntDen), defined as the product of area and mean gray value, was calculated for each image. Values were normalized to the control group and expressed as fold change.

2.14. Alizarin Red S staining

hDPSCs (passage 3; ∼80% confluence) were seeded into 6-well plates and cultured for 24 h. Cells were then cultured in mineralization induction medium containing β-glycerophosphate (5 mM), ascorbic acid (0.28 μM), and dexamethasone (100 nM) for 21 days, with medium changes every 3 days.

Cells were assigned to five groups (n = 3 per group): (i) blank; (ii) control (mineralization induction medium); (iii) IDR-1002 (50 μg/mL); (iv) BHD; and (v) PHD. Following induction, cells were fixed with 4% paraformaldehyde and stained with 1% Alizarin Red S solution (pH 4.2; Sigma-Aldrich) for 30 min. Mineralized nodules were imaged under identical acquisition settings. For quantitative analysis, bound dye was dissolved in 10% cetylpyridinium chloride (Sigma-Aldrich) at 37 °C for 60 min, and absorbance was measured at 562 nm.

2.15. Evaluation of odontogenic marker expression in hDPSCs following treatment with hydrogel-derived products

hDPSCs (passage 3; ∼80% confluence) were seeded into 6-well plates and cultured for 24 h. Cells were then treated under the same conditions as described in Section 2.14 (n = 3 per group).

Total RNA was extracted and RT–qPCR was performed as described in Section 2.3.1. Relative mRNA expression levels of DSPP and DMP1 were calculated using the 2^−ΔΔCt method with ACTB as the reference gene.

2.16. Exploratory transcriptomic analysis of hDPSCs following IDR-1002 treatment

hDPSCs (passage 3; ∼80% confluence) were seeded in 6-well plates and cultured for 24 h to allow cell attachment. Cells were then treated for 24 h with either (i) control medium (α-MEM supplemented with 10% FBS) or (ii) IDR-1002 (50 μg/mL) prepared in the same medium (n = 2 per group).

Total RNA was extracted using TRIzol reagent (Invitrogen), and mRNA was enriched using oligo(dT) magnetic beads (Invitrogen). RNA-seq libraries were prepared using the TruSeq™ RNA Sample Preparation Kit (Illumina, San Diego, CA, USA) according to the manufacturer's instructions, including mRNA fragmentation, cDNA synthesis, adaptor ligation, and amplification. Libraries were quantified using the Quantus™ Fluorometer and QuantiFluor® dsDNA System (Promega, Madison, WI, USA), purified using the Agencourt AMPure XP system (Beckman Coulter, Brea, CA, USA), and sequenced on an Illumina platform to generate paired-end reads.

Raw sequencing data were processed in R (v4.3.1) using the edgeR package. Libraries were normalized using the trimmed mean of M-values (TMM) method, and gene expression levels were expressed as counts per million (CPM) and log₂(CPM). Genes with low expression (CPM <1 in fewer than two samples) were excluded. Z-score–scaled log₂(CPM) values were visualized using the ComplexHeatmap package (v2.18.0).

Given the exploratory nature of the dataset and the limited number of biological replicates, formal differential expression analysis was not used as the primary basis for biological interpretation. Instead, genes showing consistent directional changes between groups and an average fold change >1.5 were selected for exploratory enrichment analysis. Gene Ontology (GO) enrichment analysis was performed using clusterProfiler (v4.10.0) and org.Hs.eg.db. Biological Process (BP), Molecular Function (MF), and Cellular Component (CC) categories were analyzed using the Benjamini–Hochberg method with a false discovery rate (FDR) < 0.05. Enrichment results were used to provide preliminary insight into biological processes potentially associated with IDR-1002 treatment.

2.17. In vivo evaluation of IDR-1002 in a rat molar pulpitis model

2.17.1. Establishment of a rat pulpitis model and vital pulp therapy procedure

All animal procedures were approved by the Institutional Animal Care and Use Committee (SYSU-IACUC-2025-001326) and conducted in accordance with relevant guidelines for laboratory animal welfare.

The experimental design was adapted from previously reported protocols [26,44]. Sample size estimation was performed using G∗Power software (version 3.1; Heinrich Heine University, Düsseldorf, Germany) to ensure adequate statistical power while minimizing animal use. A total of 36 rats were included, with four animals allocated to each group at each time point. Bilateral maxillary first molars from the same animal were assigned to different treatment groups to reduce inter-animal variability and compensate for potential specimen loss. The selected sample size was based on previous studies employing similar experimental designs [26,44].

Male Sprague–Dawley rats (7–8 weeks old) were randomly assigned to three groups: negative control, positive control, and experimental. Experimental pulpitis was induced in bilateral maxillary first molars using LPS derived from Porphyromonas gingivalis (InvivoGen, Toulouse, France). General anesthesia was achieved by intraperitoneal injection of 1% pentobarbital sodium. The operative field was disinfected with 3% hydrogen peroxide. Standardized occlusal access cavities were prepared using a sterile round bur under aseptic conditions with sterile saline irrigation. Following pulp exposure, the cavities were sequentially irrigated with 3% sodium hypochlorite, 17% EDTA, and PBS.

In the PBS control group, a gelatin sponge soaked in PBS (10 μL) was placed over the exposed pulp. In the LPS control group, 2 μL of P. gingivalis LPS (10 mg/mL) was applied to the exposed pulp for 30 min, followed by placement of a PBS-soaked gelatin sponge (10 μL). In the LPS + IDR-1002 group, 2 μL of P. gingivalis LPS (10 mg/mL) was applied for 30 min, followed by placement of a gelatin sponge impregnated with 10 μL of IDR-1002 solution (50 μg/mL).

All cavities were capped with a bioceramic material (iRoot BP Plus; Innovative BioCeramix Inc., Vancouver, Canada) and restored using a flowable composite resin. Animals were euthanized at 3, 7, and 28 days postoperatively, and the maxillae were harvested for subsequent analyses.

2.17.2. Micro-CT analysis

At 28 days postoperatively, specimens were analyzed using a high-resolution micro-computed tomography system (μCT-50, SCANCO Medical AG, Brüttisellen, Switzerland; 70 kVp, 114 μA, 8 W; voxel size 10 μm).

Three-dimensional reconstructions were generated, and a standardized region of interest (ROI) was defined beneath the pulp exposure site. Mineralized volume (MV) within the ROI was quantified and normalized to total volume (TV), and results were expressed as the MV/TV ratio to evaluate mineralized tissue formation. To minimize interference from the radiopaque capping material, the ROI was confined to the pulpal tissue space beneath the exposure site, and the clearly radiopaque iRoot BP Plus layer was excluded from quantitative analysis.

2.17.3. Histological and immunofluorescence analyses

Samples were dehydrated, embedded in paraffin, and sectioned longitudinally through the center of the pulp exposure site.

For histological evaluation, sections obtained at days 3, 7, and 28 were stained with hematoxylin and eosin (H&E). Five tooth samples per group at each time point were analyzed. For each sample, one section containing the central pulp exposure area was evaluated. The inflammatory response in the coronal pulp was assessed according to the degree of inflammatory cell infiltration and graded as follows: 0 (none), 1 (mild), 2 (moderate), and 3 (severe or abscess formation). Pulp inflammation was evaluated according to previously reported criteria (Table S3) [44,45]. Histological assessment was performed by a blinded examiner.

For immunofluorescence staining, four tooth samples per group were analyzed. Sections were deparaffinized, rehydrated, and subjected to antigen retrieval using citrate buffer. Samples were then permeabilized and blocked with 3% BSA and 0.3% Triton X-100 for 1 h. Primary antibodies against DSPP (1:200; Santa Cruz Biotechnology), IL-1β (1:1000; Servicebio), and IL-6 (1:500; Servicebio) were applied overnight at 4 °C. After incubation with fluorescent secondary antibodies (1:500), nuclei were counterstained with DAPI. Images were acquired using a CLSM (Olympus FV3000, Tokyo, Japan), and fluorescence intensity was quantified using ImageJ.

For DSPP analysis, two randomly selected fields adjacent to the pulp exposure site were captured from each section at 20× magnification. For IL-1β and IL-6 analyses, two randomly selected fields adjacent to the pulp exposure site were captured at 40× magnification. Fluorescence intensity was quantified using identical image acquisition and analysis settings across all groups by a blinded examiner.

2.18. Statistical analysis

Statistical analyses were conducted using GraphPad Prism (version 10.0.2; GraphPad Software Inc., San Diego, CA, USA). Data distribution was assessed for normality and homogeneity of variance prior to statistical testing. For comparisons among multiple groups, one-way analysis of variance (ANOVA) followed by appropriate post hoc multiple-comparison tests was used for normally distributed data. When assumptions of normality or equal variance were not met, nonparametric analyses were performed using the Kruskal–Wallis test followed by Dunn's multiple-comparisons test. For experiments involving multiple time points, including bacterial growth and wound closure assays, comparisons were performed separately at each time point using the appropriate statistical test. A two-sided p value < 0.05 was considered statistically significant.

3. Results

3.1. Cytocompatibility of IDR-1002 in hDPSCs

The cytocompatibility of IDR-1002 toward hDPSCs was assessed using a CCK-8 assay (Fig. 1A–C). Relative OD (%) values were normalized to the control group at each time point. IDR-1002 showed no apparent cytotoxicity toward hDPSCs at concentrations up to 80 μg/mL. A significant reduction in cell viability was observed only at 100 μg/mL across all evaluated time points (Fig. 1A–C).

Fig. 1.

IDR-1002 attenuates LPS-induced inflammation in hDPSCs at the cellular and molecular levels. (A–C) Cell viability of hDPSCs treated with IDR-1002 (0, 20, 50, 80, and 100 μg/mL) for 24 h (A), 48 h (B), and 72 h (C), assessed by CCK-8 assay. Relative OD (%) values were normalized to the control group at the corresponding time point (n = 9). (D–F) Relative mRNA expression levels of IL6 (D), IL1B (E), and TNFA (F) in hDPSCs following 24 h of treatment with LPS and IDR-1002 (0, 20, 50, and 80 μg/mL) (n = 3). (G–I) Quantitative analysis of IL-6 (G), IL-1β (H), and TNF-α (I) protein expression normalized to β-actin following 24 h of treatment with LPS and IDR-1002 (0, 20, 50, and 80 μg/mL) (n = 3). (J) Representative Western blot images showing IL-6, IL-1β, TNF-α, and β-actin expression. ns, not significant (p > 0.05); **p < 0.01; ***p < 0.001; ****p < 0.0001.

3.2. Effect of IDR-1002 on LPS-induced cytokine expression in hDPSCs

LPS stimulation significantly increased IL6, IL1B, and TNFA expression in hDPSCs compared with untreated controls (Fig. 1D–F). IDR-1002 reduced the expression of all three cytokines in a concentration-dependent manner, with the strongest inhibitory effect observed at 80 μg/mL.

Consistent with the RT–qPCR results, Western blot analysis demonstrated elevated IL-6, IL-1β, and TNF-α protein levels following LPS stimulation, which were progressively reduced by IDR-1002 treatment (Fig. 1G–J).

3.3. Antibacterial effects of IDR-1002 on multispecies, S. mutans, and E. faecalis biofilms

3.3.1. Effects of IDR-1002 on multispecies and S. mutans biofilm viability

CLSM analysis demonstrated concentration-dependent antibacterial activity of IDR-1002 against both multispecies and S. mutans biofilms (Fig. 2C). Increasing peptide concentrations resulted in a progressive increase in dead bacteria, with the greatest antibacterial effect observed at 100 μg/mL. Quantitative analysis confirmed a corresponding increase in bacterial killing in both biofilm models.

Fig. 2.

Antibacterial activity of IDR-1002 against multispecies and S. mutans biofilms. (A) Schematic illustration of the antibacterial activity of IDR-1002. The amino acid sequence of IDR-1002 is shown. IDR-1002 interacts with the bacterial cell surfaces through electrostatic attraction, inserts into the membrane structures, and has been reported to induce structural alterations in bacterial cell envelopes [46]. Created in BioRender. (B) Schematic diagram of biofilm formation on hydroxyapatite (HA) discs and subsequent treatment. Multispecies bacteria or S. mutans were cultured on HA discs for 1 week to establish biofilms, followed by treatment with IDR-1002 and analysis by CLSM. Created in BioRender. (C) Representative CLSM images of live/dead-stained multispecies and S. mutans biofilms treated with IDR-1002 (10, 20, 50, and 100 μg/mL). Green fluorescence indicates live bacteria, and red fluorescence indicates dead bacteria. Quantitative analysis of the percentages of dead bacteria is shown on the right (n = 10). Different uppercase letters indicate statistically significant differences among groups (p < 0.05).

3.3.2. Antibacterial effects of IDR-1002 on E. faecalis biofilms in dentinal tubules

IDR-1002 exhibited concentration-dependent antibacterial activity against E. faecalis biofilms within dentinal tubules (Fig. 3B). Treatment with 50 and 100 μg/mL resulted in greater bacterial killing than the control group, which was confirmed by quantitative CLSM analysis.

Fig. 3.

Antibacterial activity and dentinal tubule penetration of IDR-1002. (A) Schematic illustration of the dentin block infection model and experimental workflow. Dentin blocks were prepared, infected with bacteria, and treated with IDR-1002. Following longitudinal splitting, bacterial viability was evaluated by fluorescent staining and CLSM. For penetration analysis, 5-FAM-labeled IDR-1002 (50 μg/mL) was applied, and its distribution within dentinal tubules was visualized by CLSM. Created in BioRender. (B) Representative CLSM images showing the antibacterial activity of IDR-1002 against E. faecalis within dentinal tubules at concentrations of 10, 20, 50, and 100 μg/mL. Green fluorescence indicates live bacteria, and red fluorescence indicates dead bacteria. Scale bar: 50 μm. Quantitative analysis of the percentages of dead bacteria is shown on the right (n = 10). (C) CLSM images and quantitative analysis of the penetration depth of 5-FAM-labeled IDR-1002 within dentinal tubules compared with the control group (n = 3). Scale bar: 100 μm. ****p < 0.0001. (D) Antibacterial activity of IDR-1002 (10, 20, 50, and 100 μg/mL) against S. mutans at 6, 12, 24, and 48 h, expressed as relative OD (%) (n = 3). Different uppercase letters indicate statistically significant differences among groups (p < 0.05), whereas the same letter indicates no significant difference.

3.3.3. Penetration of IDR-1002 into dentinal tubules

Fluorescence imaging using 5-FAM-labeled IDR-1002 showed extensive penetration of the peptide into dentinal tubules, whereas no fluorescence signal was detected in the control group (Fig. 3C). Quantitative analysis confirmed greater penetration depth in the IDR-1002 group than in the control group.

3.3.4. Inhibitory effects of IDR-1002 on S. mutans and multispecies oral bacteria

IDR-1002 inhibited planktonic growth of both S. mutans and multispecies bacterial suspensions in a concentration-dependent manner across all evaluated time points (Fig. 3D and Fig. S1). Greater inhibitory effects were observed at higher peptide concentrations, particularly at 50 and 100 μg/mL. Consistent with these findings, crystal violet staining demonstrated reduced multispecies biofilm formation following IDR-1002 treatment, with the greatest reduction in biofilm biomass observed at 100 μg/mL (Fig. S2).

3.4. Cytocompatibility of blank and IDR-1002–loaded hydrogels

CCK-8 analysis showed comparable hDPSC viability among groups following exposure to hydrogel-derived leachates (BHL, PHL) or degradation products (BHD, PHD) compared with the control group (Fig. S3). Consistent with these findings, live/dead staining revealed low levels of cell death across all groups, with no significant differences in dead cell counts (Fig. 4B and C). Under inflammatory conditions, LPS significantly increased cell death, whereas treatment with hydrogel-derived products reduced LPS-induced cytotoxicity, with a greater protective effect observed in the PHD group (Fig. 4D and E).

Fig. 4.

Preparation of hydrogel-derived products and evaluation of cytocompatibility in hDPSCs. (A) Schematic illustration of hydrogel fabrication and preparation of hydrogel-derived products for cell experiments. Hydrogels were formed by UV-initiated crosslinking of four-arm PEG-NB and MMP-9–cleavable peptide linkers in the presence of IDR-1002. Peptide hydrogel leachate (PHL) and hydrogel degradation products (PHD) were collected and applied to hDPSCs or LPS-stimulated hDPSCs. LAP: lithium phenyl-2,4,6-trimethylbenzoylphosphinate. Created in BioRender. (B) Representative live/dead staining images of hDPSCs cultured with control medium, blank hydrogel degradation products (BHD), PHL, and PHD. Scale bar: 200 μm. (C) Quantitative analysis of dead cells in hDPSCs under different treatments (n = 3). Same uppercase letter indicates no significant difference (p > 0.05). (D) Representative live/dead staining images of LPS-stimulated hDPSCs treated with control medium, LPS, PHL, and PHD. Scale bar: 200 μm. (E) Quantitative analysis of dead cells in LPS-stimulated hDPSCs under different treatments (n = 3). ns, not significant (p > 0.05); ***p < 0.001.

3.5. Cyclo(-RGDfC) functionalization enhances hDPSC adhesion and spreading

CLSM analysis demonstrated limited cell attachment and spreading on unmodified hydrogels throughout the observation period (Fig. 5B). In contrast, cyclo(-RGDfC)-modified hydrogels supported early cell attachment and spreading, resulting in greater cell coverage over time. After 1 month of culture, hDPSCs formed a dense and interconnected cellular network on the modified hydrogel surface.

Fig. 5.

Fabrication of cyclo(-RGDfC)-functionalized hydrogels and evaluation of hDPSC adhesion and spreading. (A) Schematic illustration of hydrogel fabrication and cyclo(-RGDfC) functionalization. The hydrogel precursor was cast into curing rings and photocrosslinked under UV irradiation to generate control hydrogels. Cyclo(-RGDfC) peptide was subsequently immobilized onto the hydrogel surface through a thiol–ene click reaction under UV irradiation, generating cyclo(-RGDfC)-modified hydrogels. (B) Schematic diagram of hDPSC culture on control and cyclo(-RGDfC)-modified hydrogels and subsequent CLSM analysis. Representative live/dead staining images of hDPSCs cultured on hydrogels for 1 and 3 days, and representative F-actin staining images after 1 month of culture. Scale bar: 100 μm. Created in BioRender.

3.6. Inhibitory effects of IDR-1002–loaded hydrogels on S. mutans and multispecies oral bacteria

Hydrogel-derived products inhibited planktonic growth of both S. mutans and multispecies bacteria compared with the control group (Fig. S4 and Fig. S5). In general, PHD exhibited stronger antibacterial activity than PHL throughout the observation period, particularly against S. mutans.

3.7. IDR-1002–loaded hydrogels attenuate proinflammatory cytokine expression in hDPSCs

LPS stimulation significantly increased IL6, IL1B, and TNFA expression in hDPSCs compared with the control group (Fig. 6A–C). Treatment with hydrogel-derived products attenuated this response, with PHD generally exhibiting a greater inhibitory effect than PHL. While both PHL and PHD reduced IL1B and TNFA expression, a significant reduction in IL6 expression was observed in the PHD group but not in the PHL group.

Fig. 6.

IDR-1002-loaded hydrogel-derived products attenuate inflammatory responses and oxidative stress while enhancing hDPSC wound closure. (A–C) Relative mRNA expression levels of IL6 (A), IL1B (B), and TNFA (C) in hDPSCs treated with LPS, IDR-1002-loaded hydrogel leachate (PHL), and hydrogel degradation products (PHD) (n = 3). (D) Representative CLSM images showing intracellular reactive oxygen species (ROS) levels in hDPSCs treated with LPS, blank hydrogel degradation products (BHD), PHL, and PHD. Scale bar: 100 μm. (E) Quantitative analysis of intracellular ROS levels expressed as integrated density (IntDen) (n = 3). (F) Representative images from the scratch-wound assay showing wound closure in hDPSCs treated with LPS, BHD, PHL, and PHD at 0, 12, and 24 h. Scale bar: 500 μm. (G, H) Quantitative analysis of wound closure, expressed as relative wound area reduction at 12 h (G) and 24 h (H) (n = 3). ns, not significant (p > 0.05); *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

3.8. IDR-1002–loaded hydrogels attenuate intracellular ROS generation in hDPSCs

LPS stimulation significantly increased intracellular ROS levels in hDPSCs compared with the control group (Fig. 6D and E). Both PHL and PHD reduced ROS accumulation, with PHD exhibiting a greater inhibitory effect than PHL.

3.9. IDR-1002–loaded hydrogels improve hDPSCs wound closure under inflammatory conditions

LPS stimulation significantly reduced wound closure in hDPSCs at both 12 and 24 h compared with the control group (Fig. 6F–H). PHD enhanced wound closure at both time points, whereas the effect of PHL was less pronounced.

3.10. IDR-1002–loaded hydrogels promote odontogenic differentiation and mineralization in hDPSCs

Odontogenic differentiation was evaluated by Alizarin Red S staining and expression of the odontogenic markers DSPP and DMP1 (Fig. 7A–D). After 21 days of induction, mineral deposition was markedly increased in the control group compared with the blank group and was further enhanced in the PHD group, as demonstrated by Alizarin Red S staining and quantitative analysis (Fig. 7A and D). Consistent with these findings, DSPP and DMP1 expression was higher in the PHD group than in the blank and control groups, whereas BHD induced a more moderate increase (Fig. 7B and C).

Fig. 7.

IDR-1002 enhances odontogenic differentiation and mineralization of hDPSCs, while exploratory transcriptomic analysis identified enrichment of ECM- and signaling-related pathways. (A) Alizarin Red S staining of hDPSCs after 21 days of treatment with blank medium, mineralization induction medium (control), IDR-1002 peptide (50 μg/mL), blank hydrogel degradation products (BHD), and IDR-1002-loaded hydrogel degradation products (PHD). Scale bar: 200 μm. (B, C) Relative mRNA expression levels of odontogenic markers DSPP (B) and DMP1 (C) in hDPSCs after 21 days of treatment with blank, control, BHD, and PHD groups (n = 3). (D) Quantitative analysis of Alizarin Red S staining in hDPSCs after 21 days of treatment with blank, control, IDR-1002 peptide (50 μg/mL), BHD, and PHD groups (n = 3). Absorbance was measured at 562 nm. (E) Exploratory transcriptomic analysis of hDPSCs treated with IDR-1002 (n = 2). The heatmap shows distinct gene expression profiles between control and IDR-1002-treated groups, with red and green indicating relatively higher and lower gene expression levels, respectively. (F) GO enrichment analysis of genes selected for exploratory enrichment analysis, showing biological processes potentially associated with IDR-1002 treatment, including GAG binding, receptor regulation, receptor ligand activity, and ECM organization. (G) Bubble plot of selected enriched GO terms identified in the exploratory transcriptomic dataset. Bubble size represents gene count, and color indicates statistical significance (–log₁₀ FDR). ns, not significant (p > 0.05); *p < 0.05; ***p < 0.001; ****p < 0.0001.

3.11. Exploratory transcriptomic analysis of IDR-1002–treated hDPSCs

To obtain preliminary insight into biological processes associated with IDR-1002 treatment, exploratory RNA sequencing was performed in hDPSCs (Fig. 7E–G). A total of 12,703 genes were detected, of which 254 met the predefined criteria for upregulation in the IDR-1002 group. Hierarchical clustering revealed distinct transcriptional profiles between IDR-1002–treated and control cells (Fig. 7E).

GO enrichment analysis of upregulated genes identified enrichment of terms related to glycosaminoglycan (GAG) binding, receptor regulation, receptor ligand activity, heparin binding, and extracellular matrix organization (Fig. 7F). To facilitate data visualization, selected enriched GO terms were grouped into broader biological categories (Fig. 7G).

Given the limited number of biological replicates, these transcriptomic findings should be considered exploratory and hypothesis-generating. Collectively, the analysis identified enrichment of biological processes related to extracellular matrix organization and cellular signaling following IDR-1002 treatment.

3.12. IDR-1002 attenuates pulp inflammation and promotes reparative dentin formation in vivo

3.12.1. IDR-1002 reduces LPS-induced pulp inflammation and improves tissue organization

H&E staining revealed more extensive inflammatory cell infiltration and tissue disorganization in the LPS group than in the IDR-1002 group throughout the observation period (Fig. 8B and Fig. S6). In contrast, IDR-1002 treatment was associated with reduced inflammatory infiltration and improved preservation of pulp tissue architecture. Semi-quantitative analysis confirmed a lower degree of inflammation in the IDR-1002 group at days 3 and 7 (Fig. 8D and E).

Fig. 8.

IDR-1002 attenuates LPS-induced pulp inflammation and promotes reparative mineralized tissue formation in vivo. (A) Schematic illustration of the experimental design for evaluating IDR-1002 in an LPS-induced pulpitis model in SD rats undergoing vital pulp therapy. Created in BioRender. (B) Representative hematoxylin and eosin (H&E) staining images of dental pulp tissues from control, LPS, and LPS + IDR-1002 groups at Day 3 and Day 7. Arrows indicate inflammatory cell infiltration, and asterisks denote necrotic areas. (C) Representative immunofluorescence staining images of IL-6, IL-1β, and DSPP at the pulp exposure sites in control, LPS, and LPS + IDR-1002 groups at Day 7. The dashed lines indicate the pulp–dentin interface, and the solid triangles indicate the pulp exposure site. (D, E) Quantitative analysis of pulp inflammation scores showing the percentage distribution of different grades at Day 3 (D) and Day 7 (E) among the three groups (n = 5). (F–H) Quantification of mean fluorescence intensity (MFI) of IL-6 (F), IL-1β (G), and DSPP (H) at the pulp exposure sites at Day 7 (n = 4). (I) Representative micro-CT images of treated teeth from control, LPS, and LPS + IDR-1002 groups at Day 28. Solid triangles indicate the pulp exposure site. (J) Quantitative analysis of mineralized tissue formation, expressed as the ratio of MV/TV, at Day 28 (n = 5). ns, not significant (p > 0.05); *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001.

3.12.2. IDR-1002 modulates inflammatory cytokine expression and enhances odontogenic marker expression in vivo

Immunofluorescence staining showed increased IL-6 and IL-1β expression in the LPS group, whereas IDR-1002 treatment reduced the expression of both cytokines (Fig. 8C, F, and G). Conversely, DSPP expression was higher in the IDR-1002 group than in the LPS group (Fig. 8C and H).

3.12.3. IDR-1002 enhances reparative dentin formation

Micro-CT analysis at day 28 revealed more extensive mineralized tissue formation beneath the pulp exposure site in the LPS + IDR-1002 group than in the LPS group (Fig. 8I). Quantitative analysis confirmed a significantly higher MV/TV ratio following IDR-1002 treatment (Fig. 8J).

4. Discussion

The present study investigated the therapeutic potential of the innate defense regulator peptide IDR-1002 under inflammatory conditions relevant to vital pulp therapy. The findings demonstrate that IDR-1002 exhibits antibacterial, anti-inflammatory, antioxidative, and pro-regenerative activities. In addition, incorporation of IDR-1002 into an MMP-responsive hydrogel enabled inflammation-associated release of biologically active peptide following MMP-9-mediated degradation. These observations support the concept that successful vital pulp therapy depends not only on bacterial control and physical sealing but also on regulation of the inflammatory microenvironment that influences cellular behavior and tissue repair [4,7,13,14].

Current vital pulp therapy strategies are primarily based on preservation of residual pulp vitality through disinfection and placement of bioactive capping materials [[5], [6], [7], [8], [9]]. Calcium silicate–based materials, including MTA and bioceramic cements, have substantially improved clinical outcomes because of their sealing ability, alkalinity, and bioactivity [[7], [8], [9]]. Material–cell interactions may contribute to the biological responses associated with pulp healing and repair [47]. Nevertheless, treatment outcomes remain variable, particularly in teeth presenting with moderate or severe inflammation [[10], [11], [12]]. Increasing evidence suggests that residual inflammatory activity may persist within the pulp tissue even after apparent clinical hemostasis, thereby compromising tissue repair and long-term pulp vitality [8,11,12]. These limitations highlight the need for therapeutic approaches capable of simultaneously regulating bacterial activity, inflammatory signaling, and regenerative processes within the pulp microenvironment.

The present study builds upon our previous investigations involving MMP-responsive biomaterials and host defense peptide-based regenerative strategies for endodontic applications [24,25]. In our earlier work, an MMP-9-responsive PEG-based hydrogel was developed to enable inflammation-associated degradation and environmentally triggered release of IDR-1002 under simulated pulpitis conditions [25]. The current study substantially extends those findings by demonstrating that IDR-1002 not only exerts antibacterial activity but also modulates inflammatory responses, oxidative stress, stem cell behavior, and mineralized tissue formation under inflammatory conditions. These observations further support the concept that disease-responsive biomaterials may function as biologically interactive therapeutic systems rather than serving merely as passive drug carriers.

HDPs have attracted increasing interest in regenerative medicine because of their combined antimicrobial and immunomodulatory properties [15,16]. Unlike conventional antibiotics, many HDPs directly participate in regulation of innate immune responses and tissue repair pathways [15,48]. IDR peptides were specifically designed to reduce cytotoxicity while preserving host-regulatory activity [17,18,49]. Previous studies have demonstrated that IDR-1002 can regulate neutrophil activity, suppress excessive inflammatory responses, and promote tissue healing [17,18]. However, its influence on the inflamed pulp microenvironment and mineralized tissue formation during vital pulp therapy has remained insufficiently characterized.

One important aspect of the present study was the use of multiple bacterial models representing distinct stages and characteristics of pulpal infection. Multispecies biofilms were included because dental caries is now recognized as a polymicrobial ecological disease rather than an infection driven by a single pathogen [[1], [2], [3], [4]]. Interactions among bacterial species contribute to enhanced biofilm resilience, metabolic cooperation, and antimicrobial tolerance. Earlier investigations from our group further demonstrated that extracellular polymeric substance architecture and biofilm organization substantially influence antimicrobial susceptibility and recovery dynamics within oral biofilms [34,35]. Consequently, multispecies biofilm models provide greater biological relevance than simplified mono-species systems. S. mutans was selected because of its established role in cariogenic biofilm initiation and acid production during caries progression [2,3,50]. In contrast, E. faecalis was included because of its association with persistent endodontic infection and deep dentinal tubule colonization. Although E. faecalis is not considered a dominant pathogen in primary caries, its ability to penetrate dentinal tubules and survive harsh environmental conditions makes it a clinically relevant model organism for persistent dentin-associated infection [51].

The antibacterial findings demonstrated that IDR-1002 reduced bacterial viability in multispecies biofilms, S. mutans biofilms, and E. faecalis biofilms within dentinal tubules. The peptide also inhibited planktonic bacterial growth and reduced multispecies biofilm biomass. These observations are consistent with the antimicrobial and antibiofilm properties previously reported for HDPs [15,16,46]. Importantly, IDR-1002 penetrated dentinal tubules and reduced intratubular bacterial viability, which may be clinically relevant because residual intratubular bacteria are considered an important contributor to persistent pulpal inflammation following caries excavation [8]. The relatively small molecular size and cationic characteristics of IDR-1002 may facilitate diffusion within the hydrated dentin matrix, thereby supporting antibacterial activity within deeper dentin regions.

Several limitations of the antibacterial evaluation should be acknowledged. Although the biofilm models used in this study provide greater biological relevance than mono-species systems, they do not fully reproduce the complexity of the oral environment. In addition, antibacterial activity was evaluated primarily through bacterial viability, growth, and biofilm biomass assays without comparison to clinically used antimicrobial agents. Future studies incorporating CFU-based analyses, clinically relevant controls, and long-term biofilm models would further strengthen assessment of the antibacterial performance of IDR-1002.

The inflammatory response within pulp tissue plays a dual role in both defense and repair [3,4]. Excessive or unresolved inflammation, however, disrupts tissue homeostasis and impairs regenerative processes. Different sources of LPS were used in the in vitro and in vivo experiments because the objectives of the two models were distinct. E. coli LPS was selected for hDPSC studies because it is one of the most widely used inflammatory stimuli in dental pulp cell culture models, whereas P. gingivalis LPS was employed in the rat pulpitis model because it originates from an oral pathogen and is therefore more representative of oral inflammatory disease [52]. Although direct quantitative comparisons between these models should be interpreted cautiously, IDR-1002 consistently reduced inflammatory responses in both systems, supporting its anti-inflammatory activity across complementary experimental models.

In the present study, LPS and IDR-1002 were administered simultaneously to evaluate whether IDR-1002 could modulate the initiation of inflammatory responses. Therefore, the current model primarily reflects the capacity of IDR-1002 to influence the establishment of inflammation rather than its ability to reverse an already established inflammatory state. Further studies employing post-treatment protocols will be necessary to determine whether IDR-1002 can also attenuate established inflammatory responses under more clinically relevant conditions.

It should also be recognized that the inflammatory models used in the present study represent simplified experimental systems. Although LPS stimulation reproduces important aspects of inflammatory activation, clinical pulpitis is driven by a considerably more complex microenvironment involving polymicrobial biofilms, bacterial metabolites, damage-associated molecular patterns, infiltrating immune cells, and dynamic interactions among multiple residents and recruited cell populations [3,4,27,28]. Consequently, the biological effects of IDR-1002 observed in the current study should be interpreted within the context of these controlled experimental conditions. Future studies employing more physiologically relevant models, including multispecies biofilm-derived stimuli, macrophage–hDPSC co-culture systems, and other immune cell–containing platforms, will be valuable for further evaluating the therapeutic potential of IDR-1002 in inflamed pulp tissue.

In the present study, IDR-1002 attenuated LPS-induced expression of IL-6, IL-1β, and TNF-α in hDPSCs at both transcriptional and protein levels without substantially compromising cell viability. Since LPS stimulation itself reduced cell viability, it is possible that partial restoration of cell viability by IDR-1002 indirectly contributed to the reduction in cytokine expression. Alternatively, IDR-1002 may directly regulate inflammatory signaling pathways independent of its cytoprotective effects. Exploratory transcriptomic analysis provided preliminary support for this latter possibility, as RNA-seq identified enrichment of pathways associated with receptor regulation, receptor ligand activity, and extracellular matrix organization following IDR-1002 treatment. Some of these pathways may be relevant to cellular responses involved in inflammation and tissue repair. Although the precise signaling mechanisms underlying these effects were not investigated, previous studies have reported that IDR-1002 can modulate innate immune responses through regulation of NF-κB- and MAPK-associated pathways [17,18]. Persistent activation of these pathways contributes to sustained expression of proinflammatory cytokines and promotes tissue-destructive responses within inflamed pulp tissue [4,28]. Therefore, the reduction in inflammatory cytokine expression observed in the present study may reflect regulation of inflammatory signaling rather than complete suppression of immune activity. This distinction is biologically important because successful tissue repair requires transition from a destructive inflammatory state toward a regulated reparative microenvironment rather than elimination of inflammatory responses altogether [13,14].

The inflammatory response observed in the present study was accompanied by increased intracellular ROS generation. Oxidative stress is increasingly recognized as an important contributor to pulpal degeneration and stem cell dysfunction under inflammatory conditions [26]. Excessive ROS accumulation can amplify inflammatory signaling while impairing cellular migration, extracellular matrix synthesis, and odontogenic differentiation [53,54]. In the present study, IDR-1002 reduced LPS-induced ROS accumulation and improved hDPSC wound closure, suggesting that modulation of cellular redox homeostasis may contribute to its overall protective effects.

The mechanism responsible for the reduction in ROS remains unclear. Previous studies suggest that IDR-1002 may enhance endogenous antioxidant defenses through activation of Nrf2-associated pathways and upregulation of antioxidant enzymes, including HO-1 and NQO1 [18,55,56]. Therefore, the observed reduction in ROS may reflect regulation of cellular antioxidant responses rather than direct free-radical scavenging. However, these mechanisms were not investigated in the present study and require further validation.

The promotion of mineralized tissue formation further highlights the regenerative potential of IDR-1002. Successful vital pulp therapy requires not only suppression of inflammation but also stimulation of reparative dentin formation [8,9]. Under inflammatory conditions, odontogenic differentiation is often impaired because inflammatory signaling disrupts extracellular matrix production and mineralization-associated pathways [57]. In the present study, IDR-1002 enhanced mineral deposition and increased expression of the odontogenic markers DSPP and DMP1. Both markers are closely associated with odontoblast differentiation and dentin matrix mineralization. These findings suggest that modulation of the inflammatory microenvironment by IDR-1002 may create conditions that are more favorable for reparative mineralization.

Exploratory transcriptomic analysis identified enrichment of biological processes related to extracellular matrix organization, GAG binding, receptor regulation, and receptor ligand activity following IDR-1002 treatment. Given the limited number of biological replicates, these findings should be interpreted as exploratory and hypothesis-generating. Nevertheless, the identified biological processes are relevant to tissue remodeling and cellular signaling and may provide useful directions for future mechanistic investigations.

An additional focus of this study was the development of an MMP-responsive delivery strategy. Stimuli-responsive biomaterials have attracted increasing interest because they allow therapeutic activity to be linked to pathological cues within diseased tissues [[22], [23], [24], [25]]. Matrix metalloproteinases, particularly MMP-9, are strongly associated with pulpal inflammation and tissue degradation [[27], [28], [29], [30], [31]]. The MMP-cleavable peptide linker therefore provided a biologically relevant mechanism for inflammation-associated release of IDR-1002. The hydrogel platform employed in this study was based on a previously characterized MMP-responsive PEG system [25]. Consequently, the primary objective of the present work was not comprehensive physicochemical characterization of the hydrogel but rather evaluation of the biological activity of IDR-1002 following MMP-mediated release.

The present findings demonstrate that biologically active IDR-1002 can be released following MMP-9-triggered hydrogel degradation. Since MMP activity was inhibited prior to cell treatment, the hydrogel-derived products reflected peptide release following enzymatic triggering rather than continuous enzymatic degradation during cell culture. The observed biological activities therefore indicate that IDR-1002 retained functional activity after release from the hydrogel system. This inflammation-responsive release strategy aligns with the emerging concept of precision-based vital pulp therapy, in which therapeutic activity may be dynamically linked to the local pathological microenvironment [23,24].

Across multiple assays, PHD consistently exhibited greater biological activity than PHL, including anti-inflammatory, antioxidative, wound-closure-related, and mineralization-related effects. These findings are in agreement with our previous study demonstrating that MMP-9-mediated hydrogel degradation markedly enhanced IDR-1002 release compared with passive diffusion and generated stronger antibacterial effects under simulated inflammatory conditions [25]. The superior performance of PHD therefore likely reflects the increased availability of bioactive IDR-1002 released following enzymatic degradation of the hydrogel network. In addition, because elevated MMP activity is a characteristic feature of inflamed pulp tissue, PHD may better represent the products generated from the hydrogel under pathophysiologically relevant conditions. Nevertheless, the concentrations of released IDR-1002 and other degradation-derived components were not determined in the present study. Accordingly, the mechanisms underlying the differential biological activities of PHD and PHL remain to be clarified through future quantitative release and compositional analyses.

Cyclo(-RGDfC) functionalization improved hDPSC adhesion and spreading on PEG hydrogels, consistent with the known role of integrin-binding motifs in promoting cell–material interactions [[40], [41], [42], [43]]. PEG-based hydrogels are intrinsically bioinert and generally lack the adhesive motifs required for integrin-mediated cell attachment [40,41]. Because the hydrogel is intended to function as a pulp-contacting biomaterial, optimization of cell adhesion remains an important design consideration for future translation.

The rat pulpitis model reproduced several pathological features associated with inflamed pulp tissue encountered clinically during vital pulp therapy [26,58,59]. Within this inflammatory environment, IDR-1002 reduced inflammatory infiltration, suppressed IL-6 and IL-1β expression, improved tissue organization, and was associated with increased DSPP expression and mineralized tissue formation. The in vivo findings were generally consistent with the in vitro observations and further supported the therapeutic potential of IDR-1002 under inflammatory conditions.

Interpretation of the in vivo mineralization findings nevertheless requires caution. Because iRoot BP Plus is highly radiopaque, material-related interference in micro-CT analysis cannot be completely excluded despite careful definition of the ROI. Furthermore, day-28 histological sections did not demonstrate formation of a continuous reparative dentin bridge. Therefore, the extent of newly formed mineralized tissue should be interpreted in conjunction with the histological and immunofluorescence findings. Additional histological approaches for evaluating matrix maturation and mineralization would further strengthen assessment of the reparative response.

Several limitations should be acknowledged. First, although hDPSCs represent an important cellular component of pulp repair, the dental pulp microenvironment contains multiple immune cell populations that were not directly investigated in the present study [[60], [61], [62]]. Future studies incorporating immune cell–containing co-culture systems may provide additional mechanistic and translational insight. Second, the molecular mechanisms underlying the antibacterial, anti-inflammatory, antioxidative, and pro-regenerative effects of IDR-1002 were not directly examined. Therefore, the observed effects should be interpreted primarily as functional outcomes rather than definitive evidence of specific signaling pathways. Third, the transcriptomic analysis was exploratory and performed using only two biological replicates per group. Consequently, the enrichment results should be interpreted cautiously and regarded as hypothesis-generating. Finally, although the MMP-responsive hydrogel enabled release of biologically active IDR-1002 in vitro, the hydrogel platform itself was not evaluated in vivo, and the specificity of the MMP-cleavable linker toward MMP-9 was not assessed. Further studies are needed to evaluate the safety, responsiveness, and therapeutic potential of this delivery strategy under clinically relevant conditions.

5. Conclusion

Within the limitations of this study, the innate defense regulator peptide IDR-1002 demonstrated antibacterial, anti-inflammatory, antioxidative, and pro-regenerative activities in experimental models relevant to vital pulp therapy. IDR-1002 reduced bacterial viability, attenuated inflammatory cytokine expression and intracellular oxidative stress, and promoted wound closure and mineralization-related responses in hDPSCs under inflammatory conditions. Incorporation into an MMP-responsive hydrogel enabled inflammation-associated release of biologically active IDR-1002 following MMP-9-mediated degradation, while cyclo(-RGDfC) functionalization improved the cell-interactive properties of the hydrogel platform. In a rat pulpitis model, IDR-1002 reduced pulpal inflammation and was associated with improved tissue organization, increased DSPP expression, and enhanced mineralized tissue formation. These findings support further investigation of host-regulatory peptides and inflammation-responsive delivery systems as biologically based approaches for vital pulp therapy.

Ethics approval and consent to participate

This study was approved by the Clinical Research Ethics Committee of the University of British Columbia (approval numbers H23-00624 and H12-02430). Human-derived samples were collected following informed consent procedures approved by the ethics committee. All animal procedures were approved by the Institutional Animal Care and Use Committee of Sun Yat-Sen University (SYSU-IACUC-2025-001326) and were conducted in accordance with the institutional and national guidelines for laboratory animal welfare.

Availability of data and materials

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

CRediT authorship contribution statement

He Liu: Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing – original draft. Chanchan Chen: Data curation, Investigation, Software, Visualization, Writing – review & editing. Mengjie Li: Data curation, Investigation, Software, Visualization, Writing – review & editing. Lari Häkkinen: Methodology, Validation, Writing – review & editing. Ahmed Hieawy: Methodology, Writing – review & editing. Xi Wei: Methodology, Writing – review & editing. Markus Haapasalo: Methodology, Writing – review & editing. Ya Shen: Conceptualization, Funding acquisition, Methodology, Project administration, Resources, Supervision, Validation, Writing – review & editing.

Declaration of competing interest

Ya Shen is an associate editor for Bioactive Materials and was not involved in the editorial review or the decision to publish this article. He Liu is an editorial board member for Bioactive Materials and was not involved in the editorial review or the decision to publish this article. All authors declare that there are no competing interests.

Appendix A. Supplementary data

The following is the Supplementary data to this article: