Astaxanthin Alleviates Bleomycin‐Induced Pulmonary Fibrosis in Mice by Suppressing the Fibroblast‐to‐Myofibroblast Transition and Collagen Deposition

ABSTRACT

Idiopathic pulmonary fibrosis (IPF) is a progressive and fibrotic lung disease characterized by epithelial injury, inflammatory cell infiltration, fibroblast activation and extracellular matrix deposition. Currently, the treatments for IPF have significant adverse effects. Astaxanthin (AST) is a natural xanthophyll carotenoid with unique antioxidant and anti‐inflammatory properties. This study investigated the therapeutic potential of AST against pulmonary fibrosis using both in vivo (bleomycin‐induced mouse model) and in vitro (TGF‐β1‐induced HFL1 fibroblasts) approaches. Results showed that AST significantly improved lung function and attenuated fibrosis in mice, as evidenced by improved respiratory function, reduced histological damage, and decreased expression of fibrosis markers (Col1, Col3, α‐SMA, FN). Consistent with in vivo findings, AST suppressed the TGF‐β1‐induced differentiation of fibroblasts into myofibroblasts in HFL1 cells as determined by western blot, immunofluorescence, and quantitative RT‐PCR. Furthermore, transcriptome analysis revealed multiple key genes (e.g., LUM and INHBB) and potential pathways (including TGF‐β signaling pathway, extracellular matrix (ECM) remodeling pathway) involved in the regulatory effects of AST on pulmonary fibrosis. These findings demonstrate that AST alleviates BLM‐induced pulmonary fibrosis by inhibiting the fibroblast‐to‐myofibroblast transition. This study supports the development of AST as a functional food for IPF management, providing a new strategy for alleviating pulmonary fibrosis.

Astaxanthin, a natural carotenoid, alleviates bleomycin‐induced pulmonary fibrosis in mice by inhibiting TGF‐β1‑driven fibroblast‑to‑myofibroblast transition and collagen deposition, supporting its potential as a functional food for managing idiopathic pulmonary fibrosis.

1. Introduction

Idiopathic pulmonary fibrosis (IPF) is a chronic, progressive lung disease characterized by worsening respiratory symptoms and physiological impairment [1, 2]. Significant advances in the understanding of the pathobiology of IPF have emerged in recent years [1]. It involves alveolar epithelial injury and hyperplasia, the recruitment and activation of innate immune cells, fibroblast differentiation and proliferation, and excessive extracellular matrix deposition with fibrotic lesion formation [2, 3, 4, 5]. However, the mechanisms underlying IPF pathogenesis remain incompletely understood. The two clinically available drugs, pirfenidone (PFD) and nintedanib, exhibit limited therapeutic efficacy, accompanied by significant side effects [6, 7, 8]. Currently, lung transplantation remains the only curative intervention for IPF patients, due to the irreversibility of IPF. Therefore, the development of novel therapeutic agents with enhanced safety and efficacy is imperative for the treatment of pulmonary fibrosis.

Astaxanthin (AST; 3,3′‐dihydroxy‐β, β′‐carotene‐4,4′‐dione) is a natural lipid‐soluble and red‐orange xanthophyll carotenoid, primarily found in marine organisms [9, 10]. AST is widely known for its strong antioxidant capacity, and other biological properties, such as anti‐inflammatory, antiapoptotic, antiaging, and anticancer properties [9, 11, 12]. Due to its unique bioactivities and safety, AST has been approved as a dietary supplement since 1999 [12, 13]. Previous studies indicated that AST can play a protective role in age‐related diseases, type 2 diabetes, cardiovascular diseases, lung diseases, organ fibrosis and other conditions [9, 11, 14].

Several studies also suggested that AST attenuates pulmonary fibrosis [15, 16, 17]. Wang et al. have observed that AST ameliorated lung fibrosis in vivo, and inhibited cell growth and promoted cell apoptosis in transforming growth factor‐β1(TGF‐β1) induced A549 and MRC‐5 cells [15]. In addition, one previous study demonstrated that AST prevented pulmonary fibrosis by promoting myofibroblast apoptosis dependent on Drp1‐mediated mitochondrial fission [17]. Meanwhile, AST also has been shown to suppress the apoptosis in alveolar epithelial cells in vivo and in vitro [16]. Thus, AST may be a potent compound in ameliorating pulmonary fibrosis mainly by inducing the apoptosis of myofibroblasts and inhibiting EMT in lung tissues [9].

The migration, proliferation, and activation of fibroblasts are involved in the development of pulmonary fibrosis. Particularly, fibroblast activation, also known as myofibroblast differentiation, is considered to be the main pathological mechanism for fibrosis [18, 19]. However, whether AST could alleviate pulmonary fibrosis by inhibiting myofibroblast differentiation and extracellular matrix deposition should be further verified. In this study, we aimed to investigate the effects of AST on bleomycin (BLM)‐induced pulmonary fibrosis and TGF‐β1 induced myofibroblast differentiation in HFL1. To further explore the underlying involved mechanisms, transcriptome analysis was also applied to AST treatment on TGF‐β1 induced HFL1 cells. Our study provides the critical additional findings to demonstrate that AST might be a promising antifibrotic agent for treating pulmonary fibrosis.

2. Materials and Methods

2.1. Reagents and Antibodies

AST (purity ≥ 97% by HPLC) was sourced from Solarbio Co., Ltd. (Beijing, China). Pirfenidone (PFD) was from Aladdin Biochemical Technology Co., Ltd. (Shanghai, China). Complete Ham's F‐12K medium supplemented with 10% fetal bovine serum was purchased from Meiwang Biological Technology Co., Ltd. (Shanghai, China). Penicillin‐streptomycin mixture (100 ×) was from Servicebio Technology Co., Ltd (Wuhan, China). Human TGF‐β1 protein (100‐21‐2UG) was obtained from Thermo Fisher Scientific (Waltham, USA). Cell Counting Kit‐8 (M4839) was from AbMole Bioscience Inc (Shanghai, China). BCA Protein Assay Kit (P0012), Trizol (R0016), Cell Lysis Buffer for western or IP (P0013), RIPA Lysis Buffer (Strong) (P0013B) and BeyoECL Plus (P0018S) were from Beyotime Co., Ltd (Shanghai, China). Eastep Super Total RNA Extraction Kit (LS1040) was from Promega (Beijing, China). PrimeScript RT Master Mix (Perfect Real Time) (RR036A) and TB Green Premix Ex Taq II (RR820A) were purchased from Takara (Beijing, China). Primary antibodies Col1 (14695‐1‐AP), Col3 (22734‐1‐AP), FN (66042‐1‐Ig), β‐actin (66009‐1‐Ig), and α‐tubulin (66031‐1‐Ig) were purchased from Proteintech group Co., Ltd. (Wuhan, China). Primary antibody α‐SMA was obtained from BOSTER Biological Technology Co., Ltd. (Wuhan, China). Polyvinylidene difluoride membranes were from Merck KGaA (Darmstadt, Germany).

2.2. Animals and Treatments

Wild‐type C57BL/6J male mice (6–8 weeks old; 20–22 g) were obtained from Hunan SJA Laboratory Animal Co. Ltd. (Changsha, China). All experimental procedures were conducted with permission from the National Institutes of Health Guidelines for the Care and Use of Laboratory Animals and were approved by the Ethics Committee of Hainan University (HPIACUC2025006). All animal procedures complied with the China Animal Welfare Legislation, with efforts made to minimize the number of animals used and their discomfort. All animals were fed under standard conditions with free access to food and water. After acclimatization for 7 days, mice were randomly divided into four groups: (1) the control group, (2) the BLM group, (3) BLM + AST group, and (4) BLM + PFD group. The pulmonary fibrosis model of mice was established via intratracheal instillation of BLM as previously described [3], and the control group received an equal volume of normal saline. Starting from the next day, AST was administered to the mice by oral gavage at a dose of 3 mg/kg. AST was freshly prepared before each administration. Briefly, AST powder was first dissolved in a vehicle consisting of 5% DMSO and 10% Kolliphor HS 15 (Sigma‐Aldrich, USA), followed by dilution with 85% sterile saline to obtain the final working solution. The mice in the PFD group were administered PFD at a dose of 300 mg/kg via oral gavage. The treatments were administered once daily for 21 consecutive days. Then, the pulmonary function test of mice was carried out by DSI Buxco FinePointe Pulmonary Function Test (DSI, USA). Finally, the mice were anaesthetized with Zoletil50 and the lung tissues were harvested. A part of lung tissue was fixed in 4% paraformaldehyde for histological assessment, and the rest was snap‐frozen in liquid nitrogen and stored at −80 °C for PCR and western blotting.

2.3. Lung Histopathology

The lung tissues were fixed in 4% paraformaldehyde, dehydrated, and then embedded in paraffin. Thin sections were cut at a thickness of 5 µm and subjected to hematoxylin‐eosin (HE) staining and Masson's trichrome staining to evaluate histopathological damage and tissue fibrosis, respectively. Ashcroft score was applied to assess the severity of fibrosis [20]. In addition, collagen deposition in Masson's trichrome‐stained sections was quantified using ImageJ software.

2.4. Cell Culture and Treatment

The human embryonic lung fibroblast cell line HFL1 (GNHu28) was obtained from the Cell Bank of the Chinese Academy of Sciences. HFL1 cells were cultured in complete Ham's F‐12K medium supplemented with 10% fetal bovine serum and 1% Penicillin‐streptomycin solution at 37 °C in a humidified incubator with 5% CO₂. For experimental treatments, HFL1 cells were seeded in six‐well plates at a density of 1 × 10⁵ cells per well and serum‐starved for 24 h. The cells were then treated with 5 ng/mL TGF‐β1 for establishing the fibrosis model and three different concentrations of AST (10, 20, and 40 µM, diluted from a 100 mM DMSO stock), simultaneously. After 48 h of incubation, the cells were harvested for subsequent RNA or protein extraction.

2.5. Western Blotting

Total proteins were first extracted from HFL1 cells or mouse lung tissues and then quantified with BCA protein assay kits, according to the manufacturer's instructions. Equal amounts of protein (5–20 µg) were separated by 8% SDS‐PAGE and then transferred to polyvinylidene difluoride membranes. The membranes were blocked with 5% skim milk for 1 h at room temperature, and then incubated with a primary antibody overnight at 4°C. After washing with TBST, the membrane was incubated with a secondary antibody for 1 h at room temperature. Protein‐level abundances were visualized using an ECL chemiluminescence kit. The densities of bands were analyzed using Image J. Protein levels were standardized by comparison with α‐tubulin or β‐actin.

2.6. CCK‐8 Assay

Cell survival rates were determined by the CCK‐8 assay. HFL1 cells were seeded uniformly into 96‐well plates at a density of 5 × 10³ cells per well and cultured overnight in Ham's F‐12K medium containing 1% FBS. Then, cells were treated with various concentrations of AST (1, 5, 10, 20, 40, and 80 µM) in complete medium and incubated for 48 h. Cell‐free wells without AST were used as the blank group (blank), and wells containing cells without AST served as the control group (control). Three replicate wells were used for each experimental condition. 10 µL of CCK‐8 reagent was added to each well, followed by additional incubation for 2 h. The absorbance was measured at 450 nm using a microplate reader. The experiment was performed independently in triplicate.

2.7. Quantitative Real Time PCR

Total RNA was isolated from mouse lung tissues or HFL1 cells using Eastep Super Total RNA Extraction Kit and reverse transcribed into cDNA using the PrimeScript RT Master Mix kit according to the manufacturer's protocol. The resulting cDNA was then used as a template for quantitative real time PCR (qRT‐PCR) using the TB Green Premix Ex Taq II kit. Primer sequences for RT‐qPCR are listed in Table 1, were synthesized by Sangon Biotech (Shanghai) Co., Ltd. The relative expression levels of the target genes were quantified using the 2^–ΔΔCt method and normalized to the GAPDH or β‐actin mRNA level.

TABLE 1.

Primer sequences for qRT‐PCR.

Primers	Species	F primer sequence (5’–3’)	R primer sequence (5’–3’)
Col1	Mouse	CTTTGCTTCCCAGATGTCCT	CGGTGTCCCTTCATTCCAG
	Human	AAGGTGTTGTGCGATGACG	TGGTCGGTGGGTGACTCTG
Col3	Mouse	GGCAGTGATGGGCAACCT	TCCCTTCGCACCGTTCTT
	Human	GAGCTGGCTACTTCTCGC	TCTATCCGCATAGGACTGAC
α‐SMA	Mouse	GGGAGTAATGGTTGGAATG	CTCAAACATAATCTGGGTCA
	Human	GGGGTGATGGTGGGAATG	GCAGGGTGGGATGCTCTT
FN1	Mouse	CATTGTTACCAACTGGGACG	CCAGAGGCATACAGGGAC
	Human	ATGGAGGAAGCCGAGGTT	AGCGGTTTGCGATGGTAC
Col5	Mouse	GGACTCGGCGGAACATTG	GGGAGTTGAGGGAACCAAA
	Human	TGGCAAGTGGCACAGAATT	TCACCCTCAAACACCTCCTC
GAPDH	Mouse	TGTTTCCTCGTCCCGTAG	CAATCTCCACTTTGCCACT
β‐Actin	Human	GGAAATCGTGCGTGACATT	CAGGCAGCTCGTAGCTCTT

2.8. Immunofluorescence

Cells with various treatments were seeded into confocal dishes, fixed with 4% paraformaldehyde for 15 min, and then permeabilized with 0.5% Triton X‐100 for 15 min. Subsequently, non‐specific antigen‐binding sites were blocked by 10% goat serum for 30 min. Cells were incubated with primary antibodies against Col1, Col3, FN, and α‐SMA overnight at 4 °C. After washing, cells were then incubated with a secondary antibody for 1 h at room temperature. Finally, the nuclei were counterstained with DAPI for 15 min. Cells were kept from light before observed with a fluorescence microscope.

2.9. RNA‐Seq Analysis

RNA‐Seq analysis of HFL1 cells with treatments was carried out by Beijing Novogene Co., Ltd. (China). The integrity and quantity of RNA were assessed using an Agilent 2100 Bioanalyzer (Agilent Technologies, CA, USA). mRNA was enriched from total RNA using oligo(dT) magnetic beads for subsequent library preparation and sequencing. Illumina sequencing was performed, and clean reads were aligned to the reference genome. Differentially expressed genes (DEGs) were identified using the DESeq2 R package, and screened according to the criteria of p < 0.05 and |log2FC| > 1. Functional enrichment analysis of DEGs was implemented by the clusterProfiler R package. Additionally, Gene‐set enrichment analysis (GSEA) was conducted through GSEA analysis tool (http://www.broadinstitute.org/gsea/index.jsp) as an alternative approach using pre‐ranked gene lists.

2.10. Statistical Analysis

Statistical analyses were performed using SPSS software (version 23.0; IBM Corp., Armonk, N. Y., USA). All data are presented as mean ± standard deviation of at least three independent experiments. Multiple comparisons were performed using one‐way ANOVA followed by the least significant difference (LSD) post‐hoc analysis, with differences considered statistically significant at p < 0.05. Graphs were plotted using GraphPad Prism 7.0 software (GraphPad Software Inc., San Diego, CA, USA).

3. Results

3.1. AST Ameliorated BLM‐Induced Impairment of Pulmonary Respiratory Function in Mice

To evaluate the antifibrotic effects of AST in vivo, the effects of AST on BLM‐induced pulmonary fibrosis in C57BL/6J mice were examined. Following 21 consecutive days of AST administration at a dose of 3 mg/kg once daily via oral gavage, the respiratory function of mice was determined by a pulmonary function testing system. As shown in Figure 1, compared with the saline group, these pulmonary function parameters from the BLM group, including PIF (peak inspiratory flow), IC (inspiratory capacity), FEV50 (forced expiratory volume in 50 ms), FEV100 (forced expiratory volume in 100 ms), PEF (peak expiratory flow), CChord (lung compliance), FVC (forced vital capacity), and VC (forced vital capacity) reduced significantly. In contrast, the AST group showed significantly augment in those parameters compared with the BLM group. Meanwhile, RI (airway resistance) was significantly elevated in the BLM group compared to the saline group, whereas it was significantly reduced in the AST group. Additionally, the positive control PFD group exhibited a similar trend of changes to the AST group. These results indicated that treatment with AST (3 mg/kg) ameliorated BLM‐induced impairments of pulmonary respiratory function returned to the level of the normal control group.

FIGURE 1.

Effects of AST on pulmonary respiratory function in BLM‐induced mouse pulmonary fibrosis model. Respiratory function parameters, including RI, PIF, IC, FEV50, FEV100, PEF, Cchord, FVC, and VC were assessed using the pulmonary function test. Data are presented as mean ± SD. n = 6 Biologically independent mice per group. Multiple comparisons were performed using one‐way ANOVA followed by the least significant difference (LSD) post‐hoc analysis. *p < 0.05 and **p < 0.01.

3.2. AST Suppressed BLM‐Induced Destruction of Lung Tissue Architecture and Collagen Deposition in Mice

To further investigate the severity of fibrosis in lung tissues, HE staining and Masson's trichrome staining were carried out. As shown in Figure 2, HE staining revealed that compared with the saline control group, mice in the BLM group exhibited disrupted alveolar structures and extensive inflammatory infiltration in lung tissues. Treatment with AST ameliorated these histopathological alterations, and restored lung tissue architecture similar to that of the normal control group. Similarly, the PFD group also displayed pulmonary histology comparable to the normal control group. Meanwhile, Ashcroft score indicated that the severity of fibrosis was induced by BLM but significantly reduced by AST and PFD, respectively. Additionally, Masson's trichrome staining results showed that substantial collagen deposition was observed in the BLM group compared to the control group, whereas administration of AST reduced collagen fiber accumulation significantly. Collagen deposition was minimal in both the AST group and the PFD group, comparable to that in the control group. In addition, the results of fibrosis score and collagen volume fraction provided further support. These data suggest that AST ameliorated BLM‐induced lung tissue structural damage and collagen deposition.

FIGURE 2.

HE and Masson trichrome staining showed the effects of AST on lung tissue morphology and collagen deposition in BLM‐induced mouse pulmonary fibrosis. (A) Histopathological evaluation of lung tissues through HE and Masson staining (scale bar: 100 µm). (B) Quantification of Ashcroft scores (HE staining) and collagen content (Masson trichrome staining). Data are presented as mean ± SD. n = 6 Biologically independent mice per group. Statistical analysis was performed using one‑way ANOVA with LSD post‑hoc analysis. *p < 0.05 and **p < 0.01.

3.3. AST Reduced the Expression Levels of Fibrotic Proteins in the Lungs of BLM‐Treated Mice

The expression levels of fibrosis‐related proteins in lung tissues were also determined by qRT‐PCR and western blotting. The results indicated that the mRNA and protein levels of Col1, Col3, α‐SMA, and FN were obviously upregulated in BLM‐treated mice compared to the control group (Figure 3A,B). After treatment with AST, the mRNA and protein expressions of these fibrotic proteins were significantly reduced (Figure 3A,B). The mRNA level of Collagen V (Col5) also displayed a comparable trend: it was increased in the BLM group but decreased after AST treatment relative to the control group (Figure 3A,B). Moreover, the expression levels of these fibrosis proteins were also reduced in the PFD group compared with the BLM group (Figure 3A,B). These results indicated that AST reduced the expression levels of fibrotic proteins in the lungs of BLM‐treated mice. The effect was comparable to that observed in the positive control PFD group.

FIGURE 3.

AST treatment ameliorated BLM‐induced pulmonary fibrosis in mice. (A) Western blot analysis of fibrotic protein expression (Col1, Col3, α‐SMA, and FN) in the lung tissue. (B) qRT‐PCR analysis of fibrotic gene expression (Col1, Col3, Col5, α‐SMA, and FN) in the lung tissue. Data are presented as mean ± SD. n = 6 Biologically independent mice per group. Comparisons among multiple groups were performed using one‑way ANOVA followed by LSD post‑hoc analysis. *p < 0.05 and **p < 0.01.

3.4. AST Inhibited TGF‐β1‐Mediated Fibrogenesis In Vitro

To determine the appropriate concentrations of AST in HFL1 cells, the cell viability after incubation with different concentrations of AST was detected by CCK‐8 assay. As shown in Figure 4A, no significant cytotoxicity was observed when HFL1 cells were exposed to the dose up to 80 µM of AST for 48 h. Therefore, AST concentrations of 10, 20, and 40 µM were utilized in the following experiments in HFL1. We further examined the therapeutic effects of AST on differentiation of fibroblasts using TGF‐β1 (5 ng/mL)‐induced HFL1 cell as the fibrosis model in vitro. As illustrated in Figure 4B, protein expression levels of Col1, Col3, α‐SMA, and FN were significantly elevated in the model group compared with the control group, which was dose‐dependently reversed by AST. The PFD group showed significantly decreased protein expression levels of Col1, Col3, α‐SMA, and FN compared to the model group.

FIGURE 4.

Effects of different concentrations of AST on fibrotic protein expression in TGF‐β1‐induced HFL1 cells. (A) Cell proliferation/cytotoxicity was assessed by CCK‑8 assay. (B) Western blot analysis of fibrotic protein expression in HFL1 cells treated with three different concentrations of AST (10, 20, and 40 µM). Except for the control group, all other groups were induced with 5 ng/mL TGF‐β1. Data are presented as mean ± SD from three independent experiments. Statistical analysis was performed using one‑way ANOVA with LSD post‑hoc analysis. *p < 0.05 and **p < 0.01.

The effect of AST (40 µM) on TGF‐β1‐induced fibrosis in HFL1 cells was also evaluated by immunofluorescence analysis, as shown in Figure 5A. The results showed that the fluorescence intensity of Col1, Col3, α‐SMA, and FN in the model group exhibited significantly upregulation compared to the control group. In contrast, the high‐dose of AST treatment led to the significant reduction of Col1, Col3, α‐SMA, and FN overexpression induced by TGF‐β1.

FIGURE 5.

Effects of a high dose of AST (40 µM) on fibrotic protein expression in TGF‑β1‑induced HFL1 cells. (A) Immunofluorescence staining of fibrotic proteins (Col1, Col3, α‐SMA, and FN) (n = 6; Data are presented as mean ± SD). (B) qRT‐PCR analysis of fibrotic gene expression (Col1, Col3, Col5, α‐SMA, and FN). Data are presented as mean ± SD from three independent experiments. Statistical analysis was performed using one‑way ANOVA with LSD post‑hoc analysis. *p < 0.05 and **p < 0.01.

The mRNA expression levels of fibrotic markers Col1, Col3, Col5, α‐SMA, and FN were also detected by qRT‐PCR (Figure 5B). The results also demonstrated that, relative to the normal group, these fibrotic markers were all upregulated in the model group. The significant decrease was observed in the mRNA expression of these proteins after the treatment with AST (40 µM). Based on these results, AST inhibited the differentiation of fibroblasts and ECM accumulation in vitro.

3.5. Transcriptomic Analysis Revealed the Potential Molecular Pathways Involved in the Inhibition of Pulmonary Fibrosis by AST

To comprehensively explore the molecular pathways underlying the potential antifibrotic effects of AST, RNA‐seq sequencing and analysis was performed on HFL1 cells, including the control, BLM‐treated, and AST treatment groups. Principal component analysis (PCA) revealed that a distinct separation of gene expression profiles among the three groups and the high biological reproducibility of the groups (Figure 6A). As shown in Figure 6B, the volcano plots indicated that 524 upregulated and 694 downregulated DEGs were found in the model group versus the control group, whereas 84 upregulated and 63 downregulated DEGs were found in the AST group versus the model group. Subsequently, a Venn analysis was conducted to identify the overlapping genes between these two DEG sets. The results revealed that, 42 genes underwent significant changes in BLM group and were significantly regulated by AST among these DEGs (Figure 6C). Based on the previous literature, we also screened for several potential fibrosis‐related genes and visualized their expression patterns using a heatmap (Figure 6D), including Col3a1 (collagen type III alpha 1 chain), ELN (elastin), MMP1 (matrix metallopeptidase 1), LUM (lumican), CLDN1 (Claudin‐1), DTX3L (deltex E3 ubiquitin ligase 3L), INHBB (inhibin subunit beta B), BMP6 (bone morphogenetic protein 6), PRSS35 (serine protease 35), AGTR1 (angiotensin II receptor type 1), and IL15RA (interleukin 15 receptor subunit alpha). only MMP1, CLDN1, DTX3L, and IL15RA were significantly downregulated in the model group and upregulated after AST treatment, whereas the remaining genes exhibited the opposite pattern (Figure 6D). Further, GO, KEGG, and GSEA enrichment analyses were performed on these DEGs. GO enrichment analysis indicated that they were enriched in extracellular matrix. The most enriched pathways included TGF‐β signaling pathway, PI3K‐Akt signaling pathway, AGE‐RAGE signaling pathway, and Cytokine‐cytokine receptor interaction. GSEA of KEGG pathways confirmed the enrichment of several upregulated pathways in the model group compared to the control, including the TGF‐β signaling pathway, ECM‐receptor interaction, glycosaminoglycan (GAG) biosynthesis, AGE‐RAGE signaling pathway, and focal adhesion (Figure 6E). This enrichment was downregulated following AST treatment (Figure 6E). Additionally, the GSEA results indicated that the PI3K‐Akt signaling pathway (p = 0.095) was not significantly enriched (Figure 6E).

FIGURE 6.

Transcriptome analysis revealed the effects of AST on fibrosis‐related gene expression and key signaling pathways in TGF‐β1‐induced HFL1 cells. (A) Principal component analysis (PCA) of the control, model (TGF‑β1), and AST (40 µM) groups. (B) Volcano plot of differentially expressed genes (DEGs) between the model versus control groups and the model versus AST groups. (C) Venn diagram of DEGs among the three groups. (D) Heat map showing selected fibrosis‑related DEGs. (E) Gene‐set enrichment analysis (GSEA). Control is the control group, with C1, C2, and C3 being three biological replicates. The model group is induced by TGF‐β1, with M1, M2, and M3 being three biological replicates. The concentration of AST used was 40 µM, and A1, A2, and A3 are three biological replicates. Except for the control group, all other groups were induced with 5 ng/mL TGF‐β1.

4. Discussion

IPF is a chronic and progressive interstitial lung disease characterized by the excessive deposition of ECM. Although PFD and nintedanib are clinically approved for the treatment of pulmonary fibrosis, the therapeutic efficacy is limited, associated with severe adverse effects, such as gastrointestinal disturbances, rash, and photosensitivity [6, 7, 8]. Therefore, there is an urgent need to discover novel, safe and effective therapeutic options for IPF. Compounds derived from natural dietary sources generally possess a more favorable safety profile. AST is a natural, lipid‐soluble xanthophyll carotenoid derived from marine organisms and has been approved as a dietary supplement since 1999 [12, 13]. Studies in recent decades have demonstrated that AST has significant antioxidant, anti‐inflammatory, antiproliferative, and antiapoptotic properties [11, 12]. In the present study, we demonstrated that daily administration of AST (3 mg/kg) significantly ameliorates BLM‐induced pulmonary fibrosis in mice. The BLM‐induced impairment of lung respiratory function, disruption of lung tissue architecture, and excessive collagen deposition were alleviated by AST treatment. Furthermore, AST also downregulated both the mRNA and protein of fibrotic markers, including α‐SMA, FN, Col1, and Col3. Notably, the antifibrotic efficacy of AST (3 mg/kg) was comparable to that of the clinically approved drug PFD (300 mg/kg) in BLM‐induced mice. Additionally, to evaluate the inhibitory effect of AST on fibroblast differentiation, TGF‐β1‐induced HFL1 cells were utilized as an in vitro model of pulmonary fibrosis. AST at concentrations up to 80 µM exhibited no significant cytotoxicity in HFL1 cells. Treatment with AST at concentrations of 20 and 40 µM significantly suppressed the TGF‐β1‐induced overexpression of fibrotic markers (α‐SMA, FN, Col1, and Col3). It is noteworthy that α‐SMA is a hallmark of myofibroblasts, the key effector cells in IPF pathogenesis [5, 21]. Furthermore, both medium‐ and high‐dose of AST (20 and 40 µM, respectively) demonstrated a more potent antifibrotic effect compared with 10 µM PFD in vitro. Collectively, these results indicate that AST exhibits potent antipulmonary fibrosis activity in both in vivo and in vitro models.

Previous studies also suggested that AST plays a protective role in various diseases including pulmonary fibrosis [9, 11, 14, 15, 16, 17]. However, most of them concentrated on the apoptosis of alveolar epithelial cells or myofibroblasts involved in pulmonary fibrosis [15, 16, 17]. The effect of AST on the fibroblast‐to‐myofibroblast differentiation, a key pathological mechanism in pulmonary fibrosis, has not been fully elucidated. Therefore, our study provides solid evidence that AST alleviates pulmonary fibrosis through inhibiting myofibroblast differentiation and ECM deposition both in vivo and in vitro. This finding is consistent with previous studies and further elucidates the inhibitory effect of AST on pulmonary fibrosis.

AST belongs to the C40 xanthophyll family, which also includes β‑carotene, lycopene, and lutein. Previous comparative studies have revealed that among these four common dietary carotenoids, AST exhibits the strongest antioxidant capacity, with a singlet oxygen quenching ability approximately 10‑fold higher than that of either β‑carotene or lutein [22]. A comparison of reported effective doses in rat models of BLM‑induced pulmonary fibrosis reveals notable differences among carotenoids [23]. Lycopene was effective at 5 mg/kg in Sprague‑Dawley rats [24], crocin at 25 mg/kg in Wistar rats [25], whereas AST exhibited significant anti‑fibrotic effects at a lower dose of 2 mg/kg in Sprague‑Dawley rats [15]. After adjusting for body surface area to estimate human equivalent doses, AST demonstrates a more potent anti‑fibrotic efficacy compared to lycopene and crocin. Although direct comparisons across studies should be interpreted with caution due to variations in animal strains, protocols, and formulations, these data collectively support that AST possesses superior anti‑fibrotic activity at a lower dosage among the tested carotenoids. Notably, among these four typical carotenoids, AST exhibited the lowest retention rate (9.49%) after 24 h of in vitro gut fermentation, indicating its highest susceptibility to microbial degradation [26]. Besides, in cancer models, AST has shown anti‑proliferative effects in breast and colon cancer cells, often outperforming β‑carotene and lycopene [22]. Given its well‑documented efficacy in preclinical pulmonary fibrosis models, AST remains a promising candidate for further therapeutic development.

Based on transcriptomic analyses conducted in HFL1 cells, this study has also identified potential key genes and molecular pathways that may mediate the antipulmonary fibrosis effects of AST. Col3a1 and ELN may represent potential effector proteins in fibrosis [5]. LUM, an ECM proteoglycan known to regulate collagen fibrillogenesis, has been reported to contribute to hepatic and cardiac fibrosis by enhancing collagen deposition and stability [27, 28, 29]. It has been established that tubule‐derived INHBB promotes renal fibrosis through the paracrine activation of interstitial fibroblasts [30]. AGTR1, a major effector of the renin‐angiotensin system, has been reported to promote fibrosis through activation of downstream profibrotic signaling pathways such as TGF‐β/Smad and ROS generation, leading to extracellular matrix deposition and tissue remodeling [31, 32]. PRSS35 is a serine protease that is upregulated in renal fibrosis. It contributes to the pathogenesis of fibrosis by degrading col1, thereby influencing extracellular matrix dynamics [33]. In contrast, DTX3L has been demonstrated to suppress skeletal muscle fibrosis by promoting ubiquitin‐mediated degradation of the profibrotic transcription factor Runx2, thereby reducing collagen deposition and ECM remodeling [34]. IL15RA plays a protective and antifibrotic role in hepatic fibrosis. Conversely, its deficiency exacerbates fibrosis [35]. The expression profiles of these genes were consistent with the previously reported results.

In addition, MMP‐1 is primarily responsible for degrading interstitial collagens types I, II, and III. Previous studies have reported that MMP‐1 is upregulated in lung tissue, peripheral blood, and bronchoalveolar lavage fluid obtained from patients with IPF [36, 37]. However, in vitro evidence suggests that MMP‐1 may exert protective effects in IPF. This apparent discrepancy may be explained by the fact that MMP‐1 is predominantly localized in reactive alveolar epithelium rather than fibroblasts [38, 39]. Furthermore, MMP‐1 has been demonstrated to play a protective role in other fibrotic diseases, which is consistent with our findings [38, 39, 40]. However, BMP6 and CLDN1 displayed an expression pattern opposite to that reported in published studies [41, 42]. This requires further clarification. Subsequently, enrichment analysis revealed significant enrichment of the TGF‐β signaling pathway, ECM‐receptor interaction, PI3K‐Akt signaling pathway, GAG biosynthesis, AGE‐RAGE signaling pathway, cytokine‐cytokine receptor interaction, and focal adhesion. These pathways have been consistently reported to be upregulated during pulmonary fibrosis progression, as they play critical roles in fibroblast‐to‐myofibroblast transition, cellular proliferation, migration, and apoptosis [43]. These results provide important clues for further elucidating the key genes and pathways involved in the mechanism by which AST alleviates pulmonary fibrosis through inhibiting the fibroblast differentiation.

Among these, the TGF‑β signaling pathway is widely recognized as a master regulator of pulmonary fibrosis, driving the activation and differentiation of fibroblasts into collagen‑producing myofibroblasts. In the canonical pathway, binding of TGF‑β to its type II receptor (TGFBR2) leads to recruitment and phosphorylation of type I receptor (TGFBR1), which subsequently phosphorylates receptor‑regulated Smads, primarily Smad2 and Smad3. Phosphorylated Smad2/3 then form a heteromeric complex with co‑Smad4, translocate to the nucleus, and activate transcription of pro‑fibrotic target genes, including α‑SMA, Col1a1, Col3a1, and FN. This transcriptional program promotes fibroblast‑to‑myofibroblast differentiation, excessive ECM deposition, and ultimately tissue fibrosis [44, 45]. Previous studies have demonstrated that AST may exert its anti‑fibrotic effects, at least in part, by modulating this pathway. In hepatic stellate cells, AST was reported to prevent TGF‑β1‑induced pro‑fibrogenic gene expression by inhibiting Smad3 activation [46]. In renal fibrosis, AST attenuated fibroblast activation through modulation of Smad2, Akt, and STAT3 pathways [47]. A comprehensive review by Li et al. [11] further summarized that AST inhibits the development of fibrosis in multiple organs, with the TGF‑β1/Smad axis being one of the most consistently implicated signaling cascades. These established literature supports the hypothesis that AST may alleviate pulmonary fibrosis by interfering with the TGF‑β/Smad‑mediated fibroblast‑to‑myofibroblast differentiation and collagen deposition.

The present study did not directly measure the bioavailability or pharmacokinetic profile of AST. However, our selected dosage and route of administration were supported by previous rodent pharmacokinetic studies, which demonstrated that orally administered AST undergoes systemic absorption, accumulates in lung tissue, and achieves concentrations above its EC₅₀ for antioxidant activity [48]. Future studies should incorporate direct pharmacokinetic analyses and clarify the precise association between AST systemic exposure and its antifibrotic efficacy. Overall, our study suggests that AST can ameliorate BLM‐induced pulmonary fibrosis by inhibiting fibroblast‐to‐myofibroblast differentiation, and reducing collagen deposition. Moreover, the dosage required for AST to exert its therapeutic effects in BLM‐induced mice was significantly lower than that of PFD. As an edible natural compound, AST demonstrates potent antioxidant and anti‐inflammatory activities. Therefore, this study indicates that AST may offer a safer and more effective therapeutic alternative for pulmonary fibrosis. It also provides a theoretical foundation for the use of natural food‐derived compounds in the attenuation of fibrotic lung diseases. It should be noted, however, that the present findings were obtained exclusively in male mice, given the well‑recognized sexual dimorphism of pulmonary fibrosis [49]. In addition, since carotenoids may elicit distinct cellular responses in females, the protective effect of AST in female mice remains to be determined. In future studies, the specific targets through which AST inhibits fibroblast‐to‐myofibroblast differentiation will be further elucidated, and the therapeutic efficacy of AST will be systematically assessed in both male and female mice.

Author Contributions

J.M. data curation, formal analysis, investigation, methodology, X.Z. data curation, formal analysis, investigation, methodology, Y.Z. software, formal analysis, Y.Z. validation, methodology, H.W. methodology, formal analysis, Y.Y. writing original draft, conceptualization, methodology, funding acquisition, X.X. conceptualization, supervision, writing/reviewing and editing, resources, and funding acquisition. All authors reviewed the manuscript.

Funding

This work was supported by Hainan Provincial Natural Science Foundation of China (no. 822MS053) and Collaborative Innovation Center of Life and Health, Hainan University (no. XTCX2022JKB08).

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

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