5-ALA photodynamic metabolite-powered zero-waste ferroptosis amplifier for enhanced hypertrophic scar therapy

Chen, Yuan; Wang, Shan; Mao, Congxiu; Lu, Qinyi; Zhu, Xingyu; Fan, Dongqi; Liu, Yiping; Chen, Xu; Zhan, Jinglei; Yang, Zixin; Ji, Ping; He, Qingqing; Chen, Tao

doi:10.1038/s41467-025-63438-7

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Published: 18 September 2025

5-ALA photodynamic metabolite-powered zero-waste ferroptosis amplifier for enhanced hypertrophic scar therapy

Yuan Chen¹^na1,
Shan Wang¹^na1,
Congxiu Mao¹,
Qinyi Lu¹,
Xingyu Zhu¹,
Dongqi Fan¹,
Yiping Liu¹,
Xu Chen¹,
Jinglei Zhan¹,
Zixin Yang¹,
Ping Ji¹,
Qingqing He¹ &
…
Tao Chen ORCID: orcid.org/0000-0002-5658-5525¹

Nature Communications volume 16, Article number: 8321 (2025) Cite this article

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Abstract

Hypertrophic scars are a stubborn form of dermal fibrosis that impairs quality of life. Although 5-ALA-mediated photodynamic therapy holds promise, its efficacy is undermined by poor transdermal delivery and rapid metabolism into non-photosensitive heme. Here, we introduce a “zero-waste” strategy that repurposes 5-ALA-derived heme to synergistically amplify ferroptosis. This is achieved by co-encapsulating 5-ALA and baicalin within human H-ferritin, subsequently incorporated into polyvinylpyrrolidone microneedles. The resulting system enables targeted delivery to hypertrophic scar fibroblasts with pH-responsive, programmable drug release. Upon administration, 5-ALA generates protoporphyrin IX to initiate photodynamic therapy. Baicalin is then released to induce ferroptosis and synergize with the reactive oxygen species and heme accumulated during photodynamic therapy, thereby overstimulating the HO-1–heme metabolic axis. This cascade promotes the release of Fe²⁺ and CO, further amplifying ferroptotic responses. Moreover, the ferroptotic stress triggers mitophagy and mitochondrial Fe²⁺ efflux. By harnessing 5-ALA metabolic byproducts, this strategy achieved markedly prolonged anti-scar efficacy in the female rabbit ear HS tissues, surpassing that of conventional therapies.

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Introduction

Hypertrophic scar (HS), characterized by excessive proliferation of fibroblasts and over-deposition of extracellular matrix, has been regarded as a complex skin disorder, imposing substantial social and financial burdens^1,2,3. Studies have reported that the incidence of HS following linear surgical wounds is around 35%, with the rate rising to as high as 72% in burn patients^4,5. Local corticosteroid injections, particularly triamcinolone acetonide (TA) and compound betamethasone (CB), remain the first-line treatments for HS^6,7,8. However, their variable efficacy and potential side effects, such as skin atrophy and pigment deposition, limit their use. As an innovative alternative, photodynamic therapy (PDT) is gaining attention for its high efficacy and precise controllability. The Food and Drug Administration (FDA) has approved 5-aminolevulinic acid (5-ALA) as a photosensitizer precursor, which is metabolized to protoporphyrin IX (PpIX), generating abundant reactive oxygen species (ROS) for hypertrophic scar fibroblasts (HSFs) killing upon light exposure⁹. However, the further metabolism of PpIX into heme not only results in the loss of the photosensitizer but also poses potential toxicity to normal tissues^10,11. For decades, 5-ALA-derived heme has been regarded as a metabolic waste, with numerous studies focused on mitigating its accumulation^12,13,14. From the perspective of natural material recycling, metabolic byproducts may be repurposed into functional resources under suitable conditions. Inspired by this, we are exploring the possibility of turning these useless heme into a potential “auxiliary energy source” to amplify the therapeutic efficacy of PDT.

Numerous studies have demonstrated that heme can be catabolized into Fe²⁺ and CO through the action of heme oxygenase-1 (HO-1), thereby inducing iron overload and ferroptosis^15,16,17,18. Coincidentally, it has been observed that HSFs are naturally addicted to iron, suggesting ferroptosis as a potential alternative treatment¹⁹. Inspired by this, 5-ALA-derived heme may become a potential source for ferroptosis, amplifying the therapeutic effect for HS. However, due to the limited abundance of HO-1 within cells, catalyzing the excessive accumulation of heme to facilitate the process of ferroptosis presents a significant challenge. Therefore, efforts to excessively activate the HO-1-heme metabolic axis and optimize the utilization of 5-ALA metabolic waste are critical for establishing an efficient ferroptosis bioreactor post-PDT.

Baicalin is a flavonoid extracted from the root of Scutellaria baicalensis. As one of the oldest medicinal plants recorded in Chinese ancient pharmacopeia, baicalin has been widely used to treat inflammation, infection and cancer for centuries^20,21,22. Recent studies have highlighted baicalin’s ability to enhance HO-1 expression, suggesting that co-delivering baicalin with 5-ALA may initiate a complex cascade reaction^23,24. Furthermore, baicalin has been shown to independently trigger ferroptosis in cancer treatment^25,26,27,28. When combined with the enhanced HO-1-heme metabolic axis, this may create a continuous ferroptosis bioreactor within HSFs. Compared to other ferroptosis inducers such as erastin, baicalin is more affordable and exhibits lower toxicity to normal tissues. However, its poor solubility and lack of specific targeting severely restrict its clinical application^29,30. Recently, various nanocarriers have been employed to improve the poor bioavailability of hydrophobic drugs and enhance their therapeutic efficiency^31,32. Among these, human H-ferritin (HFn) has garnered significant interest due to its excellent water solubility, high drug loading capacity and intrinsic targeting ability towards transferrin receptor 1 (TFR1), which is often overexpressed in tumor cells^33,34,35. Interestingly, HSFs also overexpress TFR1, suggesting that the HFn-based delivery strategy may hold promise for the targeted therapy against HS¹⁹.

Herein, a zero-waste “ferroptosis amplifier” capable of targeting HSFs was engineered by co-encapsulating 5-ALA and baicalin into an HFn nanocage (FAB NPs). To enhance transdermal delivery, these FAB NPs were integrated with polyvinylpyrrolidone (PVP) microneedles (FAB@MN), providing a painless and convenient mode of administration (Fig. 1A). Once penetrating the epidermal barrier, FAB@MNs target HSFs via the TFR1-HFn interaction, thereby enhancing intracellular drug delivery and retention (Fig. 1B). Then, a synergistic therapy for HS can be implemented by PDT and ferroptosis based on multistage cascade metabolites (Fig. 1C). Specifically, 5-ALA is converted into PpIX to initiate PDT, followed by the release of baicalin to induce ferroptosis. Meanwhile, baicalin synergizes with the ROS and heme accumulated during PDT, activating the HO-1-heme metabolic axis to release Fe²⁺ and CO, further amplifying ferroptosis. This was not merely an amplification of a single process but a multi-layered feedback loop. As iron-catalyzed Fenton reactions continue, the generated ROS further upregulate HO-1, thus driving heme degradation and propelling ferroptosis. Unlike traditional approaches that focus solely on the immediate effects of PDT, this strategy repurposes 5-ALA metabolic waste as an “auxiliary energy source” to trigger a sustained ferroptosis cascade. Therefore, each brief PDT session sparks a sustained ferroptosis effect, enabling FAB@MN to significantly inhibit HSFs after a single laser exposure (Fig. 1D). Despite the potential for further optimization, this approach offers a compelling strategy for harnessing 5-ALA-derived byproducts to enhance the synergistic efficacy of PDT and ferroptosis, with promising implications for the treatment of HS and other related disorders.

**Fig. 1: Schematic overview of FAB@MN for HS treatment.**

Results and discussion

Fabrication and characteristics of FAB@MN

Herein, we reported a one-pot method to co-encapsulate 5-ALA and baicalin within the hollow core of the HFn nanocage, which were then incorporated into PVP MN patches (Fig. 2A). Specifically, 5-ALA and baicalin were mixed with HFn solutions and heated to 60 °C. It has been demonstrated that hyperthermia can expand the channels of HFn, enabling the encapsulation of small molecules without compromising the protein structure^33,36. Transmission electron microscopy (TEM), dynamic light scattering (DLS), and Zeta potential analysis were performed to characterize HFn and the newly synthesized FAB NPs. As shown in Fig. 2B, HFn exhibited a typical ring-like structure with a hollow core, while FAB NPs appeared as solid spheres, likely due to the filling of 5-ALA and baicalin. Notably, dual drug loading did not alter the high monodispersity and good colloidal stability of HFn, while markedly enhancing the water solubility of baicalin, thus amplifying its potential for biomedical applications (Fig. 2B, C and Fig. S1). According to the DLS data, the average diameters of HFn and FAB NPs were 10.11 nm and 11.68 nm, respectively (Fig. 2D). The slight increase in the size of FAB NPs is likely due to the encapsulated drugs. And the more negative Zeta potential of FAB NPs may be attributed to the inherent negative charge of baicalin molecules (Fig. 2D)³⁷.

**Fig. 2: The synthesis and characterization of FAB@MN.**

Ultraviolet-visible (UV-Vis) spectroscopy further confirmed the successful encapsulation of 5-ALA and baicalin within HFn, as indicated by their distinct characteristic absorption peaks (Fig. 2E). The high-performance liquid chromatography (HPLC) analysis determined that the encapsulation efficiency of 5-ALA was 57.42 ± 2.46%, with a drug loading capacity of 19.43 ± 0.67%. For baicalin, the encapsulation efficiency was 37.99 ± 0.60%, and the drug loading capacity was 12.86 ± 0.19% (Fig. S2). To confirm that the drug loading process did not affect the structure of HFn, comprehensive analyses including native polyacrylamide gel electrophoresis (Native-PAGE), circular dichroism (CD) spectra and size-exclusion chromatography (SEC) were conducted. As shown in Fig. 2F, FAB NPs appeared as a ≈ 440 kDa protein in Native-PAGE, similar to that of HFn. The CD spectra and SEC elution profiles of FA NPs (HFn encapsulating 5-ALA), FB NPs (HFn encapsulating baicalin) and FAB NPs also closely matched those of HFn, confirming the secondary structure and oligomerization states of HFn remained unchanged after drug encapsulation (Fig. 2G(i, ii)). Additionally, the stability of FAB NPs in cell culture media was assessed, revealing no significant change in size over 48 h under physiological conditions (Fig. S3).

To facilitate the in vivo biomedical applications, we developed a percutaneous delivery system for FAB NPs utilizing PVP MNs (Fig. 2H). Specifically, a 20% PVP solution containing FAB NPs was cast into a poly-dimethylsiloxane (PDMS) mold to dry overnight. The optical images demonstrated that the fabricated MN patch consisted of 225 needles spread over a 14 × 14 mm² area, with the patch turning slightly yellow after loading with FAB NPs (Fig. 2I). Scanning electron microscopy (SEM) further revealed that each MN was pyramid-shaped, with 340 μm in diameter at the base and 800 μm in height (Fig. 2K). Additionally, the mechanical hardness of the fabricated FAB@MN was evaluated, showing a force of 0.11 N/needle, which was sufficient for skin penetration (Fig. 2J). Intriguingly, the mechanical strength of FAB@MN was found to be higher than that of the blank PVP MN, possibly due to the hydrogen bonding and electrostatic interactions between the proteins and polymers. The penetration ability of FAB@MN was further demonstrated on rabbit ear HS models, where regular perforations were observed in the skin following insertion (Fig. 2L). And the fluorescence staining images revealed these MNs successfully penetrated the epidermal layer, capable of delivering Rhodamine B (RhB)-labeled FAB NPs to the deeper dermal layer (Fig. 2M(i, ii)).

Targeting ability and programmable release behavior of FAB NPs

Although various nanodrugs have been explored for eliminating HSFs, the enhancement of their targeting capabilities has often been neglected. Recently, the HFn protein-based drug delivery system has gained widespread use in cancer treatments, emerging as a hotspot in the field of nanomedicine^33,38. Studies have demonstrated that HFn not only possesses excellent drug-loading capacity but also exhibits pH-responsive disassembly and high targeting specificity for TFR1^39,40,41. In this study, HFn was employed to encapsulate 5-ALA and baicalin (FAB NPs), aiming to optimize the therapeutic effects of free drugs while minimizing damage to normal tissues. To further explore the therapeutic potential of these FAB NPs for HS, their targeting ability and release behavior were thoroughly characterized.

Initially, double-blind molecular docking was conducted using HFn (PDB ID: 7CK9) as the receptor to evaluate the structural stability of 5-ALA and baicalin encapsulation. As shown in Fig. 3A, both molecules exhibited stable binding on the HFn surface, forming fourteen intermolecular contacts comprising five hydrogen bonds, seven van der Waals forces, and two π–alkyl interactions. The binding energies for 5-ALA and baicalin with HFn were calculated as −4.4 and −9.4 kcal mol⁻¹ respectively, indicating that the resulting complex structures were stable, thereby laying a solid foundation for FAB NPs to retain the TFR1-targeting capability of native HFn. Previous studies have successfully employed HFn to target the overexpressed TFR1 in tumor cells, yielding promising therapeutic outcomes^33,41. According to our results from western blotting and quantitative reverse transcription polymerase chain reaction (qRT-PCR), the TFR1 level in HSFs was significantly higher than that in normal fibroblasts (Fig. 3B(i–iii) and Fig. S4). The immunohistochemical staining also demonstrated the increased TFR1 levels in scarring tissues, which potentially offered more binding sites for FAB NPs (Fig. S5). To verify the targeting ability of FAB NPs towards HSFs, we constructed RhB-labeled FAB NPs and co-incubated them with normal fibroblasts and HSFs, respectively. As shown in Fig. 3B(iv) and Fig. S6, the fluorescence intensity in HSFs was 7.43-fold higher than in normal fibroblasts, demonstrating the superior targeting efficacy of FAB NPs toward HSFs. Flow cytometry analysis further confirmed these findings, with results consistent with the observed fluorescence patterns (Fig. S7).

**Fig. 3: The targeting, penetration and drug release kinetics of FAB NPs.**

As many drugs proven effective in 2D cell cultures lose their potency in dense scar tissues in vivo, we further investigated the deep penetration abilities of FAB NPs in 3D scarring spheroid models using confocal laser scanning microscopy (CLSM) (Fig. 3C(i)). As shown in Fig. 3C(ii), the spheroids treated with free 5-ALA and baicalin displayed weak fluorescence, with drugs predominantly confined to the periphery. In contrast, spheroids incubated with FAB NPs exhibited widespread fluorescence throughout the entire structure. The fluorescence intensity of the treated spheroids was further quantified by 3D surface plots, revealing distinct signals penetrating deep into the spheroid core following incubation with FAB NPs (Fig. 3C(iii)). Considering the excellent targeting and penetration abilities of FAB NPs in both 2D and 3D scar models, we proceeded to assess their specific cytotoxicity towards HSFs. Normal fibroblasts and HSFs were exposed to varying concentrations of FAB NPs, with the Cell Counting Kit-8 (CCK8) assay revealing significantly greater cytotoxicity towards HSFs, highlighting their marked selectivity for HS (Fig. S8). Based on this, we further developed a microneedle system for the localized delivery of FAB NPs, enhancing treatment precision while minimizing potential harm to surrounding tissues.

The subcellular location and release behavior of FAB NPs were also explored. As shown in Fig. 3D(i) and Fig. S9, a large portion of FAB NPs colocalized with Lyso-Tracker (a typical marker of lysosome) after internalization. Recent studies have indicated that HFn remains stable in neutral environments, but its subunit interactions weaken and disassemble under acidic conditions^33,42. Leveraging this property, FAB NPs were expected to selectively release 5-ALA and baicalin in acidic lysosomes, thus minimizing premature leakage during administration. As depicted, a complete release of 5-ALA and baicalin was observed after 24 h of incubation at pH 5.0, significantly faster than that at pH 7.4 (Fig. 3D(ii)). Moreover, notable discrepancies were observed in the release rates of 5-ALA and baicalin, particularly within the initial 4 h, with approximately 81.41% of 5-ALA and 53.87% of baicalin released. To explore the mechanisms behind these differential release behaviors, we conducted molecular dynamics simulations and computed the free energy landscape (FEL) and free binding energy for each drug-protein interaction. In cases of weak or unstable protein-ligand interactions, the FEL typically exhibits multiple minima clusters. As expected, the 5-ALA-HFn complex had four small energy clusters, whereas the baicalin-HFn complex exhibited only a single concentrated energy cluster, indicating relatively stronger binding stability (Fig. 3D(iii)). Consistently, the binding free energy of 5-ALA-HFn was calculated as −10.68 kcal mol⁻¹, higher than that of baicalin-HFn at −42.05 kcal mol⁻¹. A lower binding free energy often indicates a stronger binding affinity, which may explain the slower release of baicalin compared to 5-ALA from the FAB NPs (Fig. S10). The diagram in Fig. 3E summarizes the precise targeting and unique drug release properties of FAB NPs in treating HS. Initially, FAB NPs selectively bind to the significantly overexpressed TFR1 on the surface of HSFs, achieving targeted drug localization and minimizing side effects on surrounding healthy tissues. Once internalized, FAB NPs degraded within the lysosomal acidic environment, initiating a programmable sequential release of the encapsulated drugs. Specifically, the rapid release of 5-ALA quickly initiates PDT, providing early therapeutic effects, while the slow release of baicalin offers sustained support. This sequential release pattern allows for a programmable approach to treat HS, significantly improving the precision and efficiency of the therapy.

Enhanced anti-scarring effects through FAB@MN-mediated 5-ALA metabolic waste reutilization

5-ALA serves as a precursor of photosensitizer PpIX, demonstrating potent photodynamic activity against HSFs^9,43. However, a significant challenge arises as PpIX is quickly metabolized into non-photosensitive heme^13,14. To address this challenge, we developed a waste-to-energy strategy to fully reutilize 5-ALA metabolic waste heme to enhance its anti-scar efficacy. In this strategy, FAB@MN releases 5-ALA to biosynthesize PpIX for PDT, during which substantial heme accumulates. Subsequently, baicalin significantly boosts the expression of HO-1 through its synergistic interaction with the ROS and heme accumulated from PDT. This upregulation facilitates the decomposition of heme into therapeutically beneficial Fe²⁺ and CO, thereby inducing oxidative damage to HSFs. Moreover, this cascade establishes a multi-layered feedback loop, where iron-catalyzed Fenton reactions enhance ROS production, thus further promoting HO-1 overexpression. Through this strategy, we aim to transform the “end” metabolic waste heme into a bioavailable intracellular fuel to suppress HSF activity (Fig. 3I).

As the rate-limiting enzyme in heme catabolism, HO-1 efficiently converts heme into Fe²⁺, CO, and biliverdin^44,45. Studies have shown that excessive HO-1-mediated heme metabolism can exacerbate cellular oxidative damage, offering potential for cancer treatment^46,47,48. In this study, we observed differential HO-1 expression levels across the FA@MN, FB@MN and FAB@MN groups, which correlate closely with their cellular effects. As shown, HO-1 expression was elevated in both the FA@MN and FB@MN groups compared to the control, but significantly lower than in the FAB@MN group (Fig. 4A, B). The slight upregulation of HO-1 expression observed in the FA@MN group may be attributed to a negative feedback response triggered by elevated levels of ROS and heme following PDT^49,50,51. And the increase in the FB@MN group suggested that baicalin was a potent enhancer of HO-1. To better understand the interactions between baicalin and HO-1 protein at the atomic level, molecular docking studies were subsequently conducted. As shown, baicalin formed hydrogen bond interactions with multiple residues of HO-1, showing a strong binding affinity of −12.5 kcal mol⁻¹ (Fig. 4C). These findings suggest that baicalin may directly bind to the HO-1 protein, thereby modulating its activity or expression to activate the HO-1–heme metabolic axis. Particularly following 5-ALA-mediated PDT, the increased ROS and heme further amplified the baicalin’s effects, initiating a powerful synergy to upregulate HO-1 in the FAB@MN group.

**Fig. 4: Biosynthesis of therapeutic metabolites in HSFs following treatment with FAB@MN.**

We then analyzed the key metabolites within the 5-ALA-PpIX-heme-Fe²⁺/CO/biliverdin axis across different groups. Utilizing enzyme-linked immunosorbent assay (ELISA) and fluorescence imaging techniques, we quantified the PpIX and heme levels. As shown in Fig. S11, both the FA@MN and FAB@MN groups exhibited a similar increase in PpIX. However, significant differences were observed in heme, with the FA@MN group showing a notable increase compared to the control, while the FAB@MN group exhibited minimal changes, indicating enhanced heme degradation by FAB@MN (Fig. 4D). Tracking the heme decomposition products, we observed no significant biliverdin fluorescence across the groups, likely due to its rapid degradation by abundant biliverdin reductase enzymes (Fig. S12)^18,52. However, the FA@MN group exhibited enhanced fluorescence for Fe²⁺ and CO relative to the control, confirming the excess heme and ROS generated during PDT upregulated HO-1, thus promoting heme degradation (Fig. 4E). During 5-ALA treatment, the conversion of PpIX to heme typically increased iron uptake, explaining the higher free Fe²⁺ levels in the FA@MN group, despite substantial Fe²⁺ binding by heme^18,53,54. Notably, the FAB@MN group displayed the strongest fluorescence for Fe²⁺ and CO, which was largely reversed by the HO-1 inhibitor, emphasizing the critical role of overexpressed HO-1 in heme reutilization. Interestingly, no detectable CO was observed in the FB@MN group, likely due to the limited heme available in cells. And the elevated Fe²⁺ levels in this group could be attributed to baicalin’s independent ferroptosis mechanism as reported in previous researches^25,26.

Recently, Fang et al. made a groundbreaking discovery by leveraging downstream metabolites of 5-ALA-derived heme, such as Fe²⁺ and CO, to amplify oxidative stress in cancers¹⁸. Inspired by this, we explored whether the FAB@MN could effectively promote oxidative cytotoxicity against HSFs through the enhanced 5-ALA-PpIX-heme-HO-1 metabolic axis. Using DCFH-DA staining, we evaluated the oxidative stress levels of HSFs across different groups. The results revealed that FA@MN, FB@MN and FAB@MN all elevated intracellular ROS levels, with FAB@MN inducing the most significant increase (Fig. 4F and S13). Although baicalin can be used as an antioxidant, numerous studies have highlighted its dual role in inducing ROS and ferroptosis, with this activity varying based on different cellular environments^55,56,57. Notably, FAB@MN elicited a synergistic effect that exceeded the additive impact of FA@MN and FB@MN, resulting in a pronounced elevation of intracellular ROS. And this increase was significantly mitigated by the HO-1 inhibitor, further supporting the pivotal role of the HO-1-heme metabolic axis in driving ROS storm in HSFs. As the iron-catalyzed Fenton reaction progresses, the accumulated ROS act as an activator to further upregulate HO-1 expression, thus establishing a self-perpetuating feedback loop (Fig. 4I(i))⁵¹. By repurposing 5-ALA–derived heme as a therapeutic energy source, FAB@MN offers a promising strategy to enhance anti-scarring effects following PDT.

As indicated by the CCK-8 assay, the relative cell viability in the FA@MN and FB@MN groups decreased to 72.2% and 61.9% respectively, while only 13.8% of HSFs survived in the FAB@MN group after a single irradiation, significantly outperforming the combined effects of FA@MN and FB@MN (Fig. 4H). However, inhibition or knockdown of HO-1 notably reduced FAB@MN’s therapeutic efficacy, increasing cell survival to 36.6% and 42.9% respectively (Fig. 4H and Fig. S14). Flow cytometry analysis also revealed that the HO-1 inhibitor significantly lowered the apoptosis rate in the FAB@MN group, emphasizing FAB@MN’s role in enhancing HSFs killing by upregulating the HO-1-heme metabolic axis (Fig. 4G and Fig. S15). Building on these findings, it can be concluded that FAB@MN not only generates PpIX to initiate PDT but also upregulates HO-1 to decompose heme for oxidative damage (Fig. 4I). This chemical reaction storm has been demonstrated to enhance the anti-scarring effects of PDT, effectively converting waste metabolites into treasure.

FAB@MN triggers continuous ferroptosis in HSFs

Ferroptosis is a regulated form of cell death characterized by the accumulation of Fe²⁺ and lipid peroxidation (LPO)^58,59. Recent studies have shown that enhanced heme degradation and the subsequent release of Fe²⁺ serve as potent inducers of ferroptosis^16,60. Concurrently, the role of CO in exacerbating mitochondrial damage and driving ferroptosis has become increasingly evident^18,61,62. Building on these findings, we propose that FAB@MN-mediated reutilization of 5-ALA-derived heme can effectively trigger ferroptosis in HSFs (Fig. 5A). Additionally, studies have indicated that baicalin not only promoted HO-1 expression but may also independently induce ferroptosis, with this dual action further reinforcing the continuous ferroptosis effects of FAB@MN^25,26,27,28.

**Fig. 5: FAB@MN-induced continuous ferroptosis in HSFs.**

To elucidate the role of ferroptosis in the effects of FAB@MN, we introduced the necroptosis inhibitor necrostatin-1 (Nec-1), apoptosis inhibitor Z-VAD-FMK, and ferroptosis inhibitor deferoxamine (DFO) for cell viability assays (Fig. S16). The results showed that Nec-1 and Z-VAD-FMK provided minimal rescue effects, likely by suppressing PDT-induced damage. In contrast, DFO significantly increased the cell viability from 14.9% to 76.2%, suggesting ferroptosis played a dominant role in FAB@MN-induced cell death independent of PDT. According to our strategy, FAB@MN transformed 5-ALA metabolic waste heme into a potent storm of Fe²⁺ and CO. This cascade, along with baicalin’s inherent ferroptosis potential, initiated robust and sustained ferroptosis in HSFs. To demonstrate it, we investigated several ferroptosis biomarkers across different groups. As shown in Fig. 5B, FAB@MN-treated HSFs exhibited the highest labile iron pool (LIP), which was 6.07-fold higher than that of the control group. While the level of antioxidants such as GSH dropped to 35.70% of the control, significantly lower than those in other groups (Fig. 5C). Furthermore, using the C11-BODIPY581/591 probe to assess intracellular LPO, we observed heightened green fluorescence in FAB@MN-treated HSFs compared to the FB@MN group, indicating increased LPO production via heme biotransformation following PDT (Fig. 5E and Fig. S17). Intracellular malondialdehyde (MDA), the end product of lethal LPO, also increased by 1075.53% in FAB@MN-treated HSFs, far exceeding other groups (Fig. 5D). Notably, DFO treatment significantly reversed these ferroptosis biomarkers, reinforcing the role of ferroptosis in FAB@MN-induced cell death. Furthermore, FAB@MN-treated HSFs showed the most pronounced downregulation of GPX4 and xCT, with expression levels significantly lower than those observed in the FB@MN group (Fig. 5F, H). Previous research has shown that baicalin can downregulate GPX4 and xCT, triggering LPO accumulation and ferroptosis^25,63. Additionally, excessive activation of the HO-1-heme metabolic axis may further exacerbate the downregulation of GPX4 and xCT^64,65,66. To validate this, we performed HO-1 knockdown, resulting in a significant restoration of GPX4 and xCT, thus confirming the pivotal role of the 5-ALA-PpIX-HO-1-heme-Fe²⁺ metabolic axis in FAB@MN-mediated ferroptosis (Fig. S18).

As previously reported, mitochondrial dysfunction is closely involved in ferroptosis. Therefore, we investigated the mitochondrial structure and mitochondrial membrane potential (MMP) of HSFs treated with different materials. Initially, the JC-1 and TMRM probe was utilized as the fluorescent indicator for MMP detection. As shown, FAB@MN-treated HSFs exhibited the brightest JC-1 green fluorescence and the weakest red fluorescence, alongside a significant decrease in TMRM intensity, suggesting the most pronounced MMP depolarization (Fig. 5I, L and Fig. S19). Subsequently, mitochondrial morphology analysis revealed that FAB@MN induced abnormal mitochondrial structures in HSFs, including rod-like, spherical and fragmented shapes, in stark contrast to the elongated and interconnected mitochondria observed in the control group (Fig. 5J). TEM images confirmed severe mitochondrial shrinkage and reduced cristae in FAB@MN-treated HSFs, consistent with mitochondrial changes typically associated with ferroptosis¹⁹. And the quantitative analysis via ImageJ software showed an increase in mitochondrial number but a reduction in branching length after FAB@MN treatment (Fig. 5M and Fig. S20). Notably, all observed mitochondrial damage observed in the FAB@MN group could be rescued by DFO, further confirming that the mitochondrial damage in HSFs was associated with ferroptosis.

FAB@MN-induced crosstalk between ferroptosis and mitophagy in HSFs

FAB@MN acted as a ferroptosis bioreactor by leveraging 5-ALA metabolic waste heme and the inherent ferroptosis property of baicalin. To further explore its underlying mechanisms, transcriptome sequencing was conducted. As shown in Fig. 6A, the volcano plot revealed a total of 2413 differentially expressed genes (DEGs) following the treatment with FAB@MN, with 1364 up-regulated and 1049 down-regulated. Gene ontology (GO) enrichment analysis revealed significant enrichment of DEGs in processes such as mitochondrial membrane permeability, stress response to metal ions, positive regulation of fatty acid oxidation, and cellular response to oxidative stress (Fig. 6B). Gene set enrichment analysis (GSEA) based on the Kyoto Encyclopedia of Genes and Genomes (KEGG) database highlighted the activation of pathways related to HIF1, mitophagy and ferroptosis, with key genes like HO-1, xCT, p62, and LC3B implicated (Fig. 6C, D). Additionally, protein-protein interaction (PPI) network analysis of DEGs enriched in both ferroptosis and mitophagy revealed extensive interactions, indicating a possible association between FAB@MN-induced ferroptosis and mitophagy (Fig. 6E). Previous research have revealed the intricate crosstalk between ferroptosis and mitophagy^67,68. In the early stages of ferroptosis, mitophagy is typically activated to remove damaged mitochondria and alleviate oxidative stress. However, excessive mitophagy can release iron from mitochondria, further amplifying ferroptosis and perpetuating a vicious cycle^69,70. Based on this, we hypothesized that FAB@MN not only induced ferroptosis but also triggered mitophagy, with potential direct or indirect interactions between these two processes.

**Fig. 6: The mechanism of FAB@MN-induced ferroptosis in HSFs.**

As shown in Fig. 6F(i), TEM analysis revealed FAB@MN significantly enhanced the formation of mitophagosomes in HSFs. And the fluorescence imaging also confirmed a marked increase in colocalization between mitochondria (stained with Mito-Tracker) and lysosomes (stained with Lyso-Tracker), further supporting the enhancement of mitophagy activity (Fig. 6F(ii, iii)). To elucidate the molecular mechanism of FAB@MN-induced mitophagy, the expression levels of several mitophagy-related proteins including LC3B, p62, PINK1 and Parkin were subsequently measured by immunoblotting assays (Fig. 6G). As expected, FAB@MN markedly upregulated the protein ratio of LC3B-II and LC3B-I, while decreasing p62 levels, indicating the formation of autophagosomes and the occurrence of autophagy. Furthermore, the abundance of PINK1 and Parkin proteins also significantly increased, suggesting that FAB@MN activated mitophagy via the classical PINK1/ Parkin pathway. Under normal conditions, PINK1 is rapidly degraded. However, when mitochondria are damaged, it accumulates and acts as a beacon to recruit Parkin to the surface of mitochondria. Parkin, endowed with E3 ubiquitin ligase activity, subsequently tags these damaged mitochondria, recruiting substantial p62 to the mitochondrial outer membrane. This promotes LC3B-I lipidation, ultimately leading to autophagosomal engulfment of the damaged mitochondria^71,72,73.

To investigate the potential interplay between mitophagy and ferroptosis mediated by FAB@MN, we assessed the influence of the iron chelator DFO on mitophagy regulation. As expected, DFO effectively reduced the accumulation of Parkin and PINK1 while promoting the upregulation of p62, demonstrating that FAB@MN-induced iron overload played a pivotal role in abnormal mitophagy activation within HSFs (Fig. S21). We also explored the involvement of mitophagy in FAB@MN-mediated ferroptosis by silencing PINK1 (Fig. S22). As shown, PINK1 knockdown partially restored cell viability in the FAB@MN-treated group, boosting survival from 16.7% to 44.81% (Fig. 6H). Further analysis revealed that PINK1 knockdown not only reduced ROS levels induced by FAB@MN but also significantly decreased the fluorescence intensities of Fe²⁺ and LPO (Fig. 6I–K and Fig. S23). The role of mitophagy in ferroptosis remains controversial, often viewed as a double-edged sword. In some cases, mitophagy can remove damaged mitochondria and maintain cellular function⁶⁸. However, under specific conditions, excessive mitophagy may release free Fe²⁺ to promote ferroptosis⁷⁴. In this study, FAB@MN triggered iron overload, which subsequently mediated aberrant mitophagy activation via the PINK1/Parkin pathway, leading to elevated Fe²⁺ and LPO levels in HSFs and amplifying ferroptosis (Fig. 6L).

In vivo anti-scarring effects of FAB@MN

Despite the availability of various treatment options, managing HS remains a significant challenge, underscoring the urgent need for a universally effective and minimally invasive therapeutic approach^7,75,76. Motivated by the promising in vitro results of FAB@MN, we further investigated its anti-scarring effects in vivo. Specifically, we established rabbit HS models and applied treatments once a week for a month (Fig. 7A). The results revealed that a single light exposure induced the highest levels of ROS in FAB@MN group, supporting the synergy between PDT and ferroptosis (Fig. 7B, C). After four-week treatments, scars in the FAB@MN group became smooth and indistinguishable from the surrounding healthy skin, with therapeutic effect significantly surpassing those of FA@MN, FB@MN, and even commercial corticosteroids like TA and CB (Fig. 7D and Fig. S24). Consistent with previous studies, although TA and CB injections led to some lightening of the scars, residual pigmentation persisted following treatments^6,8. To assess efficacy more comprehensively, we measured scar color, thickness and hardness in each group (Fig. S25A). As shown in Fig. S25B, the redness value of FAB@MN-treated scars (2.00 ± 0.10) was closest to that of normal skin (1.97 ± 0.31). Ultrasound imaging further confirmed FAB@MN’s superior ability to reduce scar thickness, with a significant reduction from 3.70 ± 0.54 mm to 0.46 ± 0.09 mm (Fig. 7E and Fig. S24, S27A). Moreover, FAB@MN treatment resulted in a marked decrease in scar hardness, with a 53.9% reduction (17.53 ± 2.11 HOO) compared to untreated scars (38.03 ± 2.97 HOO) (Fig. S25C). Notably, both FA@MN and FB@MN exhibited limited therapeutic effects in vivo, suggesting short-term PDT or baicalin-induced ferroptosis alone was ineffective. While FAB@MN leveraged a series of 5-ALA metabolic waste to establish a sustained bioreactor integrating PDT and ferroptosis, thereby providing an effective strategy for scar treatment and recurrence prevention.

**Fig. 7: The therapeutic efficacy of FAB@MN in rabbit ear HS models.**

Histological analyses were further conducted to assess the therapeutic outcomes of various treatments. Hematoxylin-eosin (H&E) staining showed the HS treated with FAB@MN returned to the smooth appearance of normal skin, while the other groups exhibited varying degrees of protuberance, with densely packed and disorganized fibers (Fig. 7F and Fig. S26A). Numerous studies have shown that abnormally proliferating HSFs secrete excessive collagen, particularly resulting in the accumulation of type I collagen and disrupting the collagen balance^9,77. Therefore, direct targeting and destroying HSFs could potentially restore the collagen structure and alleviate fibrosis^9,19,78. As expected, Masson staining demonstrated that FAB@MN significantly reduced collagen deposition and promoted more organized collagen fiber alignment at the HS sites (Fig. 7G and Figs. S26B, S27B). Sirius Red staining revealed a significant reduction in yellow-stained type I collagen in the FAB@MN group, suggesting substantial remodeling of the abnormal collagen matrix (Fig. 7G). Besides, the GPX4 level in HS tissues was also evaluated to assess the ferroptosis effects via immunohistochemical staining. As shown, FAB@MN notably reduced the area of GPX4-positive cells and exhibited a more effective therapeutic outcome than FB@MN, likely due to the ferroptosis cascade initiated by 5-ALA photodynamic waste (Fig. 7G, and Fig. S27C).

The biosafety of FAB@MN in vivo was verified by the H&E staining of vital organs (heart, liver, spleen, lung, and kidney), as well as hematological and biochemical analyses. No significant organ damage was observed in FAB@MN-treated group, and all blood parameters remained within normal limits after one month of treatment, confirming its negligible systemic toxicity in vivo (Figs. S28, S29). As a targeted ferroptosis nanomedicine for HS, FAB@MN not only enhances the therapeutic outcomes but also minimizes damage to healthy tissues (Fig. 7H). Furthermore, by utilizing byproducts from PDT, we have optimized the use of 5-ALA, ensuring efficient waste conversion and enhancing the synergistic effects between PDT and ferroptosis. Even compared to first-line corticosteroids, FAB@MN exhibits superior therapeutic efficacy, offering a promising alternative for HS management and presenting a potential strategy applicable to other pathological scars, such as keloids.

Methods

Ethical statement

This study complies with all applicable ethical regulations. Both human tissue collection and animal experiments were approved by the Ethics Committee of the Affiliated Stomatological Hospital of Chongqing Medical University (Approval No. CQHS-REC-2024-058). Human tissues were obtained with written informed consent from all participants and processed in accordance with the Declaration of Helsinki. Animal procedures were conducted in accordance with the Guidelines for the Care and Use of Laboratory Animals issued by the National Institutes of Health (NIH Publication No. 86–23, revised 1985) and the ARRIVE guidelines.

Cell culture and animals

Nine HS tissues and the matched normal skin tissues were provided by patients undergoing surgical treatment, and each patient has voluntarily signed an informed consent form. All participants in this study did not have any known systemic diseases and were not receiving any treatments that could potentially affect the study results. In brief, tissues were first cut into 3 mm-thick slices, and then digested overnight with 0.25% neutral protease II. Subsequently, the epidermal layer was peeled off, and further digestion was carried out for 4–6 h at 37 °C with 0.2% Collagenase II. After digestion, cells were filtered and collected using a cell strainer with a 75 μm pore size, then resuspended in Dulbecco’s Modified Eagle Medium (DMEM) containing 10% FBS and cultured at 37 °C in an atmosphere of 5% CO₂. The method of isolating fibroblasts from normal skin was similar to the aforementioned process for obtaining HSFs with minor modifications.

Eighteen female Oryctolagus cuniculus (New Zealand White rabbits, 8–10 weeks old, ~2.5–3.0 kg) were obtained from the Experimental Animal Center of Chongqing Medical University (Chongqing, China). Animals were fed standard laboratory chow (Catalog No. 1016708478371979264, Beijing Keao Xieli Feed Co., Ltd., China) containing 17% protein, 3% fat, 10–15% fiber, and 11% moisture. They were housed individually under controlled conditions (25 ± 2 °C, 55 ± 2% relative humidity, 12-h light/dark cycle). Group allocation was performed in a blinded manner by a technician using a random number table to ensure unbiased distribution and intergroup comparability. Animals were monitored daily for general health, including activity, posture, intake of food and water, body weight, and wound healing. Signs of pain or distress were closely monitored. Humane euthanasia was performed under veterinary supervision upon evidence of sustained weight loss, anorexia, or pain-related behaviors. At the end of the study, all animals were humanely euthanized via CO₂ inhalation using a gradual-fill method (20–30% of chamber volume per minute) in a sealed chamber. Death was confirmed by the cessation of respiration and loss of corneal reflex.

Preparation of FAB@MN

The synthesis of FAB@MN followed a detailed protocol. Initially, 5 mg of 5-ALA was dissolved in PBS, while 5 mg of baicalin was dissolved in ethanol. Then, 10 mg of HFn was dissolved in Tris-HCl buffer (20 mM, pH 7.5). The solutions of 5-ALA and baicalin were gradually added to the HFn solution under gentle stirring to prevent ethanol-induced denaturation of the protein. After mixing, the blend was subsequently incubated in a 60 °C water bath for 4 h, followed by centrifugation at 16,099 g for 15 min. The supernatant was collected and dialyzed four times in PBS using a dialysis bag (10 kDa), with the dialysis fluid replaced every 4 h to remove unencapsulated baicalin and 5-ALA. Subsequently, the obtained FAB NPs were mixed with a 20% PVP solution and poured into a PDMS mold, then placed under a vacuum of 600 mmHg for 1 min to ensure the solution completely filled the MN cavities. Finally, it was dried overnight in the dark before being demolded.

Characterizations of FAB NPs and FAB@MN

The protein structure changes of HFn were characterized with size-exclusion chromatography (SEC, Agilent 1260 Infinity II, USA) and circular dichroism (CD, JASCO J-1500, Japan). Particle size and Zeta potential of the materials were measured using a Zetasizer Nano ZS90 (Malvern, UK). The ultraviolet absorption spectra of the materials were determined using a multimode microplate reader (PerkinElmer EnSpire, USA). Nanoparticle morphology was visualized using a JEM-1400 Plus transmission electron microscope (TEM, Tokyo, Japan), with grid staining performed using 1% (w/v) uranyl acetate for several minutes before observation. Molecular docking simulation was performed with Auto Dock Vina software (Version 1.2.0), and molecular dynamics simulation was performed by the GROMACS (2022) program. The fabricated MN patches were then examined by scanning electron microscopy (SEM, FEI Hillsboro, USA), and their mechanical properties were assessed using a universal material testing machine (INSTRON, USA). To evaluate the skin penetration ability of FAB@MN (with RhB-labeled FAB NPs), we vertically inserted it into rabbit ear HS for 5 min. The treated skin sample was then embedded in optimal cutting temperature (OCT), and sliced into 10 μm sections using a cryotome (CM1860, Leica). Subsequently, these skin sections were placed on adhesive microscope slides and observed using a multi-function microscope.

Targeting ability and cellular uptake of FAB NPs

To investigate the targeting ability of FAB NPs towards HSFs, RhB-labeled FAB NPs were added into HSFs for 4 h. Then, cells were fixed in 4% cold formaldehyde, and the nuclei and cytoskeleton were stained with DAPI and Actin-Tracker-Red respectively. The fluorescence emitted from the treated cells was evaluated using a CLSM (Leica TCS SP8, Germany) and flow cytometry (BD Biosciences, USA). To further assess the deep penetration capability of FAB NPs in a 3D cell model, we initially planted 5 × 10⁴ cells/well in ultra-low attachment round-bottom 48-well plates (Corning, USA) to cultivate 3D cell spheroids. After the formation of 3D scarring spheroids, the fluorescent-labeled NPs were co-incubated for 4 h, and the fluorescence images were captured and analyzed using a CLSM.

In vitro drug release and subcellular colocalization of FAB NPs

We employed High-Performance Liquid Chromatography (HPLC, Agilent, USA) for the precise quantification of 5-ALA and baicalin concentrations. The encapsulation efficiency (%) and loading capacity (%) of 5-ALA and baicalin were calculated using the following formulas: encapsulation efficiency (%) = amount of encapsulated drugs/amount of drugs added; loading capacity (%) = amount of encapsulated drugs /amount of FAB NPs. To examine the release profiles of FAB NPs, 5 mL of FAB NPs solution were put into a dialysis bag (10 kDa) that was directly immersed into 20 mL PBS at 37 °C with pH values of 7.4 and 5.0. At predetermined time intervals, the concentrations of the released drugs were tested. Stability evaluation was conducted by incubating FAB NPs with cell culture medium containing 10% FBS for 48 h, and assessed their stability using a Zetasizer Nano ZS90 (Malvern, UK). For the subcellular colocalization imaging experiments, HSFs were seeded into 48-well plates at a density of 1 × 10⁴ cells/well and incubated with RhB-labeled FAB NPs for 4 h. After removing the culture medium, the cells were washed twice with cold PBS. Lysosomes and nuclei were stained with Lyso-Tracker and Hoechst 33342 for 30 and 10 min respectively. After staining, the intracellular fluorescence signals were captured with CLSM.

In vitro analysis of 5-ALA metabolic products in HSFs

To evaluate the metabolic products of 5-ALA generated in HSFs, the cells were co-cultured with different materials in the dark for 4 h, then irradiated with a 633 nm laser for 10 min, and subsequently incubated at 37 °C for 24 h. For heme assessment, the cells were gently scraped and subjected to ultrasonication. The cell suspension was then centrifuged at 11,180 g for 10 min to collect the supernatant, and the heme content was analyzed using an ELISA kit. Simultaneously, the cells were stained with COP-1 (Ruixi Biotechnology, China) for CO detection and FerroOrange (Beiren Chemical Technology, Beijing) for Fe²⁺ detection. These metabolic products were then examined using a fluorescence microscope, allowing for the measurement of fluorescence signals for CO, Fe²⁺, and biliverdin (autofluorescence). Molecular docking of baicalin with HO-1 was performed using Auto Dock Vina software (Version 1.2.0).

In vitro cytotoxicity assay

In vitro cytotoxicity was assessed using the Cell Counting Kit-8 (CCK-8). In brief, HSFs were seeded at a density of 5 × 10³ cells/well in a 96-well plate and cultured for 24 h. Subsequently, these cells were exposed to various experimental materials for 4 h, followed by laser irradiation at 633 nm (40 mW/cm², LWRPD630, China) for 10 min. After an additional 24-h incubation period, a diluted CCK-8 solution was administered to each well and incubated at 37 °C for 4 h. The absorbance at 450 nm was then measured using a microplate reader. Similarly, the treated HSFs underwent Annexin V-FITC/PI staining following the supplied protocol and were then analyzed with flow cytometry.

Ferroptosis evaluation of FAB@MN

HSFs were seeded at a density of 5 × 10⁴ cells/well in 24-well plates and cultured overnight. Subsequently, these cells were exposed to various experimental materials for 4 h, followed by laser irradiation for 10 min. After an additional 24-h incubation, the HSFs were stained with C11-BODIPY 581/591 for LPO analysis for 30 min, and analyzed using a fluorescence microscope and flow cytometry. In addition, the LIP, LPO, MDA and GSH amounts were monitored using an LIP, LPO, MDA and GSH assay kit according to the instructions. To assess the mitochondrial structure and MMP of HSFs, JC-1, TMRM and Mito-Tracker probes were incubated with cells for 30 min respectively, and the intracellular fluorescence was evaluated with CLSM or flow cytometry. For a detailed examination of the mitochondrial ultrastructure, the treated cells were fixed with 2.5% glutaraldehyde and observed using biological transmission electron microscopy (Bio-TEM, Tokyo, Japan). Mitophagy was evaluated using a dual staining method with Lyso-Tracker Green and Mito-Tracker Deep Red. Specifically, the treated HSFs were first stained with Mito-Tracker Deep Red at 37 °C for 30 min, then incubated with Lyso-Tracker Green for another 30 min, and finally stained with Hoechst 33342 for 10 min to label the cell nuclei.

Western blot analysis

Cells were collected using a cell scraper and transferred into lysis buffer containing RIPA: PMSF: phosphatase inhibitors: protease inhibitors at a ratio of 100:1:2:2. The mixture was then sonicated and centrifuged at 14,000 g for 15 min at 4 °C, and the protein concentration was measured using a BCA assay kit. Depending on the molecular weight of the proteins, appropriate Sodium Dodecyl Sulfate–Polyacrylamide Gel Electrophoresis (SDS-PAGE) gels were selected for protein separation, and the proteins were transferred to a polyvinylidene fluoride (PVDF) membrane. The membrane was incubated overnight with specific antibodies, followed by incubation with horseradish peroxidase (HRP)-conjugated secondary antibodies (1:2000, Zhongshan Golden Bridge, China). At last, the membranes were analyzed using an image reader (LAS-4000; Fujifilm, Japan). All uncropped scans of blot presented in the main text have been included in the Source Data file, and those related to the Supplementary Figs. have been provided in Supplementary Information.

In vivo anti-scarring efficacy of FAB@MN

The HS model was established using a previously detailed method. Under anesthesia, a 1 cm diameter full-thickness wound was created on the ventral side near the proximal end of the rabbit’s ear, with careful removal of the perichondrium to avoid damaging the cartilage beneath. Four weeks post-surgery, a firm and dense HS had formed at the wound sites, and these scars were randomly divided into six groups: Control group, FA@MN group, FB@MN group, FAB@MN group, TA-TRANSTON^® group and CB-DIPROSPAN^® group. Four hours post MN insertion, the treated regions were exposed to laser irradiation for 10 min, and this process was repeated weekly for a total of four weeks. The scar morphology within each group was captured and recorded using a digital camera. Upon completion of the treatment, ultrasound imaging of the scars was conducted using the Aixplorer ultrasound system (Super-Sonic Imagine, France), enabling direct measurement of scar thickness. Specifically, ultrasound transmission gel was uniformly applied over the scarred region, and the scars were analyzed with an ultrasonic probe. Scar hardness and color were measured using a Shore durometer (HT-65100, Lantai Instruments Co., Ltd, China) and a 3nh spectrophotometer (three Enshi technology Co., Ltd, Guangdong, China). Following these procedures, the rabbits were sacrificed and HS tissues were collected for histological analysis through H&E, Masson and Sirius Red staining. Additionally, immunohistochemical staining with GPX4 antibody was performed. For ROS assays, 10 µL of a 10 mM DCFH-DA solution was precisely injected into the rabbit ear scar tissue before treatment. Upon completing the treatment, detailed imaging of the scar areas was performed using the animal in vivo imaging system (IVIS). Meanwhile, the scar tissue samples were harvested for fluorescence analysis.

Statistical analysis

All analyses and comparisons were performed using SPSS 20.0 software (IBM, USA). Results were presented as mean ± SD. The two-sided t-test was employed for comparisons between two groups, while one-way ANOVA was used for analyzing the variance among multiple groups. p-value < 0.05 indicated a significant difference between groups.

Reporting summary

Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability

All data generated this study are provided in the main text and Supplementary Information. The transcriptomic data generated in this study have been deposited in the NCBI Gene Expression Omnibus (GEO) under accession code GSE300586: https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE300586. Source data are provided with this paper.

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Acknowledgments

This work was supported by the National Science Fund for Excellent Young Scholars (32322044, C.T.), National Natural Science Foundation of China (32071362, C.T.), Key International (Regional) Joint Research Program funded by National Natural Science Foundation of China (NSFC) (82220108019, J.P.), the Science Fund for Distinguished Young Scholars of Chongqing (CSTB2022NSCQ-JQX0012, C.T.), the CQMU Program for Youth Innovation in FutureMedicine (W0077, C.T.).

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These authors contributed equally: Yuan Chen, Shan Wang.

Authors and Affiliations

The Affiliated Stomatological Hospital of Chongqing Medical University, Chongqing Key Laboratory of Oral Diseases, Chongqing Municipal Key laboratory of Oral Biomedical Engineering of Higher Education, Chongqing Municipal Health Commission Key Laboratory of Oral Biomedical Engineering, Chongqing, China
Yuan Chen, Shan Wang, Congxiu Mao, Qinyi Lu, Xingyu Zhu, Dongqi Fan, Yiping Liu, Xu Chen, Jinglei Zhan, Zixin Yang, Ping Ji, Qingqing He & Tao Chen

Authors

Yuan Chen
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Shan Wang
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Congxiu Mao
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Qinyi Lu
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Xingyu Zhu
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Dongqi Fan
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Yiping Liu
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Xu Chen
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Jinglei Zhan
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Zixin Yang
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Ping Ji
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Qingqing He
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Tao Chen
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Contributions

Y.C. conceived the idea and designed the experiments; Y.C., S.W., C.X.M., Q.Y.L., and D.Q.F. prepared and characterized the materials; Y.C., S.W., Y.P.L., X.C., J.L.Z., and Z.X.Y performed in vitro and in vivo experiments; Y.C., S.W., and X.Y.Z. contributed to analysis and discussion of the results; Y.C. and S.W. wrote the paper; Q.Q.H., P.J., and T.C. revised the manuscript; T.C. supervised the overall study.

Corresponding authors

Correspondence to Ping Ji, Qingqing He or Tao Chen.

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Nature Communications thanks Salvador Gonzalez, Cheng Zhan, who co-reviewed with Guoshu Bi, and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. A peer review file is available.

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Chen, Y., Wang, S., Mao, C. et al. 5-ALA photodynamic metabolite-powered zero-waste ferroptosis amplifier for enhanced hypertrophic scar therapy. Nat Commun 16, 8321 (2025). https://doi.org/10.1038/s41467-025-63438-7

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Received: 29 May 2024
Accepted: 04 August 2025
Published: 18 September 2025
Version of record: 18 September 2025
DOI: https://doi.org/10.1038/s41467-025-63438-7