In zebrafish, Mollugin at safe doses (≤200 µg/mL) was associated with improved tail regeneration, accompanied by changes in apoptosis-related gene expression, inflammatory markers, antioxidant enzyme activity, and macrophage-associated markers. These findings suggest that Mollugin may modulate the local injury response during zebrafish tail regeneration.
Research Article
Mollugin Supports Zebrafish Tail Regeneration in Association with Altered Inflammatory and Oxidative Responses
* These authors contributed equally
In This Article
Summary
Abstract
Mollugin is a natural naphthoquinone compound with reported anti-inflammatory and antioxidant activities, yet its role in tissue regeneration remains incompletely understood. In this study, a zebrafish tail amputation model was used to investigate the effects of Mollugin on wound repair and associated immune and redox responses. Zebrafish embryos were treated with Mollugin at concentrations of 22 µg/mL, 66 µg/mL, and 200 µg/mL following tail injury. Mollugin treatment was associated with improved tail-area recovery in a dose-dependent manner, accompanied by reduced expression of apoptosis-related genes and suppression of pro-inflammatory mediators, including IL-1β, IL-6, TNF-α, TLR4, and NF-κB1. In parallel, Mollugin restored antioxidant enzyme activities of superoxide dismutase and catalase. Fluorescence imaging in Tg(mpeg1:GFP) zebrafish showed a time-dependent effect of Mollugin on macrophage accumulation, with no obvious reduction at 24 h, but decreased macrophage-positive area at 48 h in the 66 and 200 µg/mL groups compared with the injured model group. qRT-PCR analysis further showed reduced expression of the M1-associated marker iNOS and increased expression of the M2-associated marker arginase-1. Collectively, these findings suggest that Mollugin may support zebrafish tail regeneration in association with changes in inflammatory, oxidative stress, apoptosis-related, and macrophage-associated markers. Further studies are needed to define the underlying cellular and molecular mechanisms.
Introduction
Tissue regeneration is a critical physiological process by which organisms repair damage, involving complex regulation of cell proliferation, differentiation, and the maintenance of microenvironmental homeostasis1. Among vertebrate models, zebrafish have emerged as a pivotal system for studying wound repair mechanisms due to their remarkable regenerative capacity. In particular, tail fin regeneration in zebrafish encompasses the spatiotemporal coordination of multiple biological events, including inflammation regulation, oxidative response balance, and apoptosis control2,3. In recent years, the pharmacological activities of natural products have become a focal point in regenerative medicine research. Naphthoquinone compounds, in particular, have been widely studied because of their diverse biological effects4. Mollugin is a naphthoquinone compound derived from Rubiaceae plants and has shown anti-inflammatory, antioxidant, and cell cycle regulatory activities in previous studies5,6. However, its mechanism of action in tissue regeneration remains unclear, and its potential role in modulating the inflammatory microenvironment and immune cell polarization warrants systematic investigation.
After injury, inflammation is rapidly activated and directly influences tissue repair. Moderate inflammatory signaling is essential for clearing necrotic tissue and initiating the regeneration program, whereas an excessive inflammatory cascade can exacerbate oxidative stress and lead to tissue fibrosis7. Cytokines, including IL-1β, IL-6, and TNF-α, increase markedly after injury and are accompanied by activation of TLR4/NF-κB signaling, leading to amplified inflammatory responses and tissue remodeling8,9. Intercellular adhesion molecule-1 (ICAM-1) and CC chemokine ligand 2 (CCL-2) are also elevated after injury and are associated with increased recruitment of neutrophils and macrophages to the damaged tissues10. Notably, oxidative imbalance further aggravates tissue damage, and reduced superoxide dismutase (SOD) and catalase (CAT) activities indicate impaired antioxidant capacity2,11.
Macrophages serve as key regulators of the inflammatory microenvironment, and their functional polarization significantly impacts the regenerative process12. M1 macrophages are associated with inducible nitric oxide synthase (iNOS) expression and inflammatory activity, while M2 macrophages express arginase-1 (Arg-1) and are linked to tissue repair13,14. Macrophage polarization is influenced by local cytokines, oxidative stress, and apoptosis-related signals. Within this context, the apoptotic regulatory network composed of B-cell lymphoma-2 (Bcl-2) family proteins and caspase-3 not only affects cell survival at the injury site but also indirectly modulates the chemotactic behavior of immune cells through the release of "find-me" signals15. Therefore, exploring intervention strategies that simultaneously modulate inflammatory responses, oxidative stress, and macrophage polarization may help improve tissue regeneration.
Natural compounds have demonstrated unique advantages in modulating these biological processes. Recent studies have confirmed that naphthoquinone substances, owing to their conjugated structures, possess the potential to scavenge free radicals; their benzoquinone core can affect redox-sensitive signaling pathways via electron transfer mechanisms16,17. Mollugin has been reported to attenuate inflammatory responses in DSS-induced colitis, with reduced pro-inflammatory cytokine production and decreased TLR4 expression in colon tissues18. Mollugin has also been shown to inhibit TAK1-NF-κB/MAPK signaling and activate the Keap1-Nrf2 pathway in macrophages6; however, its impact on macrophage polarization and the apoptosis–anti-apoptosis network in an in vivo wound model has not yet been elucidated. Moreover, the dose-dependent biological effects of Mollugin and its regulatory role during the regeneration process remain to be systematically clarified.
The present study investigated the effects of Mollugin in a zebrafish tail fin regeneration model. Specifically, we examined tail regrowth, apoptosis-related gene expression, inflammatory signaling, antioxidant enzyme activity, and macrophage-associated responses after Mollugin treatment. Our findings suggest that Mollugin treatment is associated with improved tail regeneration and changes in multiple injury-response markers in this zebrafish model.
Protocol
All animal procedures followed the National Research Council Guide for the Care and Use of Laboratory Animals and were approved by the Ethics Committee of Fuzhou First General Hospital (approval no. 202408002).
Zebrafish breeding
All experiments were conducted using zebrafish (Danio rerio) maintained under standard laboratory conditions. Wild-type AB zebrafish were used for the toxicity assay and tail-amputation regeneration assay. Tg(mpeg1:GFP) zebrafish were used specifically for macrophage imaging. Tg(mpeg1:GFP) is a macrophage reporter line in which GFP expression labels macrophage-lineage cells under the control of the macrophage-expressed mpeg1 promoter. All zebrafish lines were maintained at 28.5 °C under a 14-h/10-h light/dark cycle. Only morphologically normal embryos were used for experiments. For all experiments, embryos were incubated in Danieau’s buffer, which was prepared using reagent-grade chemicals including NaCl, KCl, Ca(NO₃)₂·4H₂O, MgSO₄·7H₂O, and HEPES adjusted to pH 7.2.
Drug preparation and safety determination
Mollugin (40 mg) was dissolved in 1 mL DMSO to generate a 40 mg/mL stock. The stock was first diluted 1:10 in Danieau’s buffer to 4 mg/mL, followed by further dilution to 22, 66, and 200 µg/mL for treatments. To determine the maximum safe concentration, 48-hour post-fertilization (hpf) AB strain embryos were dechorionated and disinfected with 0.003% sodium hypochlorite before being distributed into 6-well plates (n=20, triplicate wells per group). Embryos were incubated with Mollugin (22 µg/mL, 66 µg/mL, 200 µg/mL, 240 µg/mL, 280 µg/mL, 320 µg/mL, or 360 µg/mL) in E3 medium for 48 h at 28.5 °C. Survival was monitored under a stereomicroscope, and the highest concentration with ≥90% embryo viability was established as the safe upper limit for subsequent experiments.
Tail amputation and regeneration model construction
For regeneration studies, 48 hpf wild-type AB embryos were anesthetized with 0.03% tricaine prepared in embryo medium and positioned laterally under a stereomicroscope. Before tail amputation, embryos were maintained in Danieau’s buffer. A standardized transverse tail amputation was performed under stereomicroscopic guidance. The amputation line was placed immediately posterior to the notochord tip, with the posterior end of the notochord used as the anatomical landmark. The tail fin fold distal to this landmark was removed using a microscalpel to ensure a consistent injury level among specimens. Immediately following amputation (0 h), images were acquired with a stereo fluorescence microscope. After amputation, embryos were transferred to the corresponding E3-based treatment medium for the regeneration assay. Mollugin working solutions were prepared by diluting the Mollugin stock solution into E3 medium to final concentrations of 22, 66, and 200 µg/mL. Mollugin was administered by bath exposure during the regeneration period. To match vehicle exposure, the uninjured control group and injured model group were maintained in E3 medium containing 0.5% DMSO, corresponding to the final DMSO concentration in the 200 µg/mL Mollugin group. Embryos were maintained in 6-well plates at 28.5 °C for 2 days, with daily refreshment of the corresponding E3-based treatment medium. Regeneration was documented by imaging at 24 and 48 h post-amputation under both white light and fluorescence settings to assess tail regrowth and macrophage distribution. Image measurements were carried out in ImageJ.
For regeneration quantification, the recovered tail area was measured as an absolute area in µm2 and was not normalized to embryo size or original tail size. To minimize variability, all embryos were amputated at the same anatomical landmark immediately posterior to the notochord tip, and the same landmark was used to define the measurement region across groups. Regions of interest were manually delineated in ImageJ based on the tail outline, and the analysis was performed by an investigator blinded to the treatment groups. For macrophage-positive area quantification, GFP fluorescence images of Tg(mpeg1:GFP) embryos were analyzed in ImageJ. The tail injury/regeneration region was manually defined as the region of interest using the same anatomical landmark applied for tail-area measurement. Images were converted to 8-bit grayscale, and background fluorescence was subtracted using the same background-correction settings for all images within the same experiment. A fixed fluorescence threshold was then applied uniformly to all images within each experiment to identify GFP-positive macrophage-associated signals. The macrophage-positive area was calculated as the threshold-positive GFP area within the defined ROI. Measurements were based on fluorescence-positive area rather than fluorescence intensity or manual cell counting. All image analyses were performed by an investigator blinded to the treatment groups.
RNA Extraction and quantitative real-time PCR (qRT-PCR)
Following completion of the regeneration assay, embryos from each experimental condition were collected and pooled (at least 20 embryos per group) for RNA analysis. For each condition, one biological replicate consisted of a pooled sample containing at least 20 embryos from the same experimental group. Three independent pooled biological replicates were prepared for each condition and used for statistical analysis. Total RNA was isolated after mechanical homogenization in a phenol/guanidine-based RNA extraction reagent. RNA quality and concentration were assessed spectrophotometrically by measuring A260/A280 ratios. cDNA was synthesized using a commercial reverse transcription kit. PCR amplification was performed with SYBR Green reagents. The qRT-PCR cycling conditions were as follows: initial denaturation at 95 °C for 30 s, followed by 40 cycles of denaturation at 95 °C for 5 s and annealing/extension at 60 °C for 30 s. Expression levels of genes related to apoptosis (caspase-3 and Bcl-2), inflammation (TNF-α, IL-1β, IL-6, NF-κB1, TLR4, CCL2, and ICAM1), and macrophage phenotype (iNOS and arginase-1) were measured. β-Actin served as the reference gene. Fold changes in gene expression were determined using the 2−ΔΔCt approach. Primer information is listed in Table 1.
Activity assays for SOD and CAT
SOD and CAT levels were used to evaluate antioxidant status after tail injury. At 48 h post-amputation, zebrafish from each group were collected and homogenized in cold PBS. After centrifugation, the cleared extracts were used for SOD and CAT measurement with commercial activity assay kits. SOD activity was measured using a WST-1-based assay kit according to the manufacturer’s instructions. Samples were loaded onto 96-well assay plates together with standards, and absorbance was read on a microplate reader. Enzyme activities were obtained from standard curves. All assays were performed in triplicate.
Statistical analysis
Data are shown as mean ± SEM from at least three independent experiments. For the toxicity assay, each independent experiment consisted of one embryo batch, with 20 embryos per group and triplicate wells. For regeneration imaging and macrophage imaging in Tg(mpeg1:GFP) zebrafish, individual embryos were used as the measurement units, and 10 embryos were analyzed per group. For qRT-PCR analysis, one biological replicate consisted of a pooled sample containing at least 20 embryos from the same experimental group, and three independent pooled biological replicates were analyzed per condition. For SOD and CAT activity analysis, five independent pooled biological replicates were analyzed per condition. Group differences were analyzed by one-way ANOVA with Tukey’s post hoc test. Statistical analyses were performed in GraphPad Prism. Differences with p < 0.05 were regarded as significant.
Results
Toxicity and safe dose determination
To investigate the effects of Mollugin, we first performed toxicity tests. The results indicated that zebrafish embryo mortality remained low at concentrations up to 200 µg/mL, whereas increased mortality was observed at concentrations above 200 µg/mL (Table 2). Therefore, the highest safe dose was determined to be 200 µg/mL, while the low and medium doses were set at 22 and 66 µg/mL, respectively.
Tail amputation model and early tail-area recovery assessment
Subsequently, a tail amputation model was established in 48 hpf wild-type AB zebrafish, and successful model construction was confirmed by microscopic observation (Figure 1). Following this, the safe doses of Mollugin were applied to the model group. At 24 h post-amputation, treatment with 200 µg/mL Mollugin was associated with increased recovered tail area compared with the injured model group (P < 0.01). At 48 h post-amputation, both 66 and 200 µg/mL Mollugin were associated with significantly increased recovered tail area compared with the injured model group (P < 0.01). Because tail-area recovery remained incomplete at 48 h post-amputation, these data indicate improved early tail-area recovery within the observation window rather than a definitive increase in final regenerative capacity (Figure 2). qRT-PCR showed increased caspase-3 expression and reduced Bcl-2 expression in the injured model group. Mollugin treatment was associated with reduced caspase-3 expression and increased Bcl-2 expression, suggesting changes in apoptosis-related gene expression after treatment (Figure 3). ICAM-1 and CCL2 levels were markedly elevated in the model group (P < 0.001). Mollugin treatment significantly reduced both markers, indicating decreased inflammatory cell recruitment at the injury site (Figure 3).
Mollugin is associated with reduced inflammatory marker expression at 48 h post-amputation
In the model group, inflammatory markers including IL-1β, IL-6, TNF-α, TLR4, and NF-κB1 increased significantly increased at 48 h post-amputation. Mollugin treatment lowered these markers, and the highest dose produced the strongest effect (Figure 4).
Mollugin restores antioxidant enzyme activity
The antioxidant effects of Mollugin were examined by measuring the levels of CAT and SOD. SOD and CAT were reduced in the model group, indicating impaired antioxidant capacity. Mollugin increased both enzymes in a concentration-dependent manner, with the highest dose showing the strongest recovery (Figure 5). These findings indicate that the oxidative stress-induced cellular damage and tissue inflammation, resulting from the redox imbalance caused by tail amputation, can be dose-dependently alleviated by Mollugin.
Mollugin modulates macrophage aggregation and distribution
To assess macrophage involvement in wound repair, we examined their aggregation using fluorescence microscopy in Tg(mpeg1:GFP) zebrafish. Macrophage accumulation increased markedly at the tail injury site in the model group, with macrophage-positive area being considerably higher than those in the normal group. After 24 h of treatment, macrophage accumulation remained similar among all Mollugin doses. However, at 48 h, treatment with 66 and 200 µg/mL Mollugin decreased macrophage-positive area at the tail injury site, although still higher than in the normal group—indicative of an ongoing tissue repair process. Compared with the injured model group, Mollugin treatment reduced the macrophage-positive area within the injury region at 48 h post-amputation, particularly at 66 µg/mL and 200 µg/mL. These results suggest that Mollugin may modulate macrophage-associated responses during the later phase of tail regeneration (Figure 6).
Mollugin modulates macrophage phenotype-related marker expression
Macrophage phenotype-related markers were evaluated by measuring iNOS, a commonly used M1-associated marker, and Arg-1, a commonly used M2-associated marker. iNOS expression increased in the injured model group and was significantly reduced following Mollugin treatment, whereas Arg-1 expression was further increased after Mollugin treatment (Figure 7). These changes suggest that Mollugin may influence macrophage phenotype-related responses during tail regeneration. However, because the current evidence is based on marker-expression analysis rather than direct validation of macrophage subsets, further studies are required to determine whether Mollugin directly affects M1/M2 macrophage polarization. Notably, the effects of high-dose Mollugin on these polarization markers were more pronounced than those observed at lower doses.
DATA AVAILABILITY:
All raw data supporting the findings of this study have been uploaded as Supplementary File 1, Supplementary File 2, Supplementary File 3, Supplementary File 4, Supplementary File 5, and Supplementary File 6. These files include the raw data for zebrafish embryo survival, tail-area recovery measurements, qRT-PCR analysis, SOD and CAT activity assays, and macrophage-positive area quantification.
Figure 1: Establishment of the zebrafish tail amputation regeneration model. Representative images of wild-type AB zebrafish embryos at 48 hpf before and immediately after tail amputation. A standardized transverse incision was performed under stereomicroscopy to generate a consistent injury model for regeneration studies. The amputation site indicates the starting point for subsequent tail-area recovery measurements. Please click here to view a larger version of this figure.
Figure 2: Mollugin supports early tail-area recovery after zebrafish tail amputation. (A) Representative images of zebrafish tails at 24 and 48 h post-amputation following treatment with Mollugin (22 µg/mL, 66 µg/mL, and 200 µg/mL). (B) Quantification of the recovered tail area corresponding to (A). The recovered tail area was measured in µm2 using ImageJ, corresponding to (A). Tail regrowth was measured using ImageJ. The recovered tail area was measured as an absolute area and was not normalized to embryo size or original tail size. Each data point represents an individual embryo. Values are shown as mean ± SEM (n = 10 embryos). Group differences were analyzed using one-way ANOVA with Tukey’s test. ns,, not significant; **p < 0.01; ***p < 0.001. Please click here to view a larger version of this figure.
Figure 3: Mollugin modulates apoptosis-related and chemotactic gene expression after tail amputation. (A–D) qRT-PCR analysis of caspase-3 (A), Bcl-2 (B), ICAM-1 (C), and CCL2 (D) expression in zebrafish embryos at 48 h post-amputation with or without Mollugin treatment (22 µg/mL, 66 µg/mL, and 200 µg/mL). Data are presented as relative mRNA expression normalized to β-actin and calculated using the 2−ΔΔCt method. Each data point represents one pooled biological replicate. Values are shown as mean ± SEM (n = 3 pooled biological replicates per group; each replicate contained 20 embryos). One-way ANOVA with Tukey’s test was used for group comparisons. ns, not significant; **p < 0.01; ***p < 0.001. Please click here to view a larger version of this figure.
Figure 4: Mollugin suppresses inflammatory cytokine expression in the zebrafish tail amputation model. (A–E) Relative expression of IL-1β (A), IL-6 (B), TNF-α (C), TLR4 (D), and NF-κB1 (E) in zebrafish embryos at 48 h post-amputation following Mollugin treatment (22 µg/mL, 66 µg/mL, and 200 µg/mL). Data are presented as relative mRNA expression normalized to β-actin and calculated using the 2−ΔΔCt method. Each data point represents one pooled biological replicate. Data are shown as mean ± SEM (n = 3 pooled biological replicates per group; each replicate contained 20 embryos). One-way ANOVA with Tukey’s test was used for statistical analysis. ns, p > 0.05; *p < 0.05; **p < 0.01; ***p < 0.001. Please click here to view a larger version of this figure.
Figure 5: Mollugin restores antioxidant enzyme activity after tail amputation. (A,B) Activity assays of superoxide dismutase (SOD) (A) and catalase (CAT) (B) activities in zebrafish embryos at 48 h post-amputation with or without Mollugin treatment. Data represent antioxidant enzyme activities measured from pooled embryo extracts. Each data point represents one pooled biological replicate. Values are shown as mean ± SEM (n = 5 pooled biological replicates per group; each replicate contained 20 embryos). One-way ANOVA with Tukey’s test was used. **p < 0.01; ***p < 0.001. Please click here to view a larger version of this figure.
Figure 6: Mollugin reduces macrophage-positive area at the injury site. (A) Representative images of GFP-positive macrophages in Tg(mpeg1:GFP) zebrafish embryos at 24 and 48 h post-amputation following Mollugin treatment (22 µg/mL, 66 µg/mL, and 200 µg/mL). The outlined region indicates the tail injury/regeneration area used for macrophage fluorescence quantification. (B) Quantification of macrophage-positive area within the outlined injury/regeneration region. Each data point represents an individual embryo. Values are shown as mean ± SEM (n = 10). Group comparisons were performed using one-way ANOVA with Tukey’s test. ns, not significant; p > 0.05; *p < 0.05; **p < 0.01; ***p < 0.001. Please click here to view a larger version of this figure.
Figure 7: Mollugin modulates macrophage phenotype-related marker expression. (A,B) qRT-PCR analysis of iNOS (A) and Arg-1 (B) mRNA expression in zebrafish embryos at 48 h post-amputation with or without Mollugin treatment. Data are presented as relative mRNA expression normalized to β-actin and calculated using the 2−ΔΔCt method. Each data point represents one pooled biological replicate. Expression levels are shown as mean ± SEM (n = 3 pooled biological replicates per group; each replicate contained 20 embryos). One-way ANOVA with Tukey’s test was used. ***p < 0.001. Please click here to view a larger version of this figure.
| Genes | Primer sequence (5 '-3') |
| TNF-α-F | TATCAGACAACCGTGGCACC |
| TNF-α-R | GCTTCAGCACTTTTCCGTGG |
| IL-6-F | ATGACGGCATTTGAAGGGGT |
| IL-6-R | TCAGGACGCTGTAGATTCGC |
| IL-1β-F | TGCGGGCAATATGAAGTCA |
| IL-1β-R | TTCGCCATGAGCATGTCC |
| Caspase3-F | CGGCACTGATACATCTGT |
| Caspase3-R | TCCTTTCACGACCATCTG |
| Bcl2-F | CTGCTGAGGAAGATGACA |
| Bcl2-R | CAGGCATTCAGAGTTGTTC |
| NFκB1-F | CTCCAAACGCCTCCAATCTG |
| NFκB1-R | AGATGGGCTTCTGGAGGTTC |
| TLR4-F | ACAGATCACCTGGACAGCAAG |
| TLR4-R | TGCTTGAAAGTCCCGCATGT |
| CCL-2-F | GATTTGTCCCAGAGTCCCGA |
| CCL-2-R | CTCAGTCGGGTTAGTGCAGA |
| ICAM-1-F | CCACAGTCACCTATGGCAAC |
| ICAM-1-R | AGTGTCTCCTGGCTCTGGTT |
| iNOS-F | GATAACCACTGCTCTGCTGC |
| iNOS-R | GAGCCATCCTTGTAGTTGCG |
| Arg-1-F | GGGGCTGTGCAAAAGATCAAG |
| Arg-1-R | GTGTGTTGATGTCTGCGTGTG |
| β-actin-F | GTATTGCTGACCGTATGC |
| β-actin-R | CTGCCTCATCGTATTCCT |
Table 1: qRT-PCR primer sequences. Primer sequences used for qRT-PCR analysis of apoptosis-related genes, inflammatory mediators, macrophage phenotype-related markers, and the reference gene β-actin are listed.
| Mollugin | Replicate 1 deaths/20 embryos | Replicate 2 deaths/20 embryos | Replicate 3 deaths/20 embryos |
| 0 | 0/20 | 0/20 | 0/20 |
| 22 μg/mL | 0/20 | 0/20 | 0/20 |
| 66 μg/mL | 0/20 | 0/20 | 0/20 |
| 200 μg/mL | 0/20 | 0/20 | 0/20 |
| 240 μg/mL | 10/20 | 10/20 | 11/20 |
| 280 μg/mL | 20/20 | 20/20 | 20/20 |
| 320 μg/mL | 10/20 | 10/20 | 11/20 |
| 360 μg/mL | 20/20 | 20/20 | 20/20 |
| 360 μg/mL | 20/20 | 20/20 | 20/20 |
Table 2: Zebrafish embryo survival at different Mollugin doses. Survival of zebrafish embryos exposed to different concentrations of Mollugin was assessed to determine the safe concentration range for subsequent tail regeneration experiments.
Supplementary File 1: Raw data for recovered tail area measurements in zebrafish embryos at 24 h and 48 h post-amputation across control, model, and Mollugin-treated groups.Please click here to download this file.
Supplementary File 2: Raw qRT-PCR data for apoptosis-related and chemotactic gene expression markers (Caspase-3, Bcl-2, ICAM-1, and CCL2) in control, model, and Mollugin-treated zebrafish groups at 48 h post-amputation. Please click here to download this file.
Supplementary File 3: Raw qRT-PCR data for inflammatory marker expression (IL-1β, IL-6, TNF-α, TLR4, and NF-κB1) in control, model, and Mollugin-treated zebrafish groups at 48 h post-amputation. Please click here to download this file.
Supplementary File 4: Raw antioxidant enzyme activity data for superoxide dismutase (SOD) and catalase (CAT) in control, model, and Mollugin-treated zebrafish groups at 48 h post-amputation. Please click here to download this file.
Supplementary File 5: Raw macrophage-positive area measurements in Tg(mpeg1:GFP) zebrafish embryos at 24 h and 48 h post-amputation across control, model, and Mollugin-treated groups. Please click here to download this file.
Supplementary File 6: Raw qRT-PCR data for IL-1β and IL-6 expression in control, model, and Mollugin-treated zebrafish groups at 48 h post-amputation. Please click here to download this file.
Discussion
In this work, we examined the effects of Mollugin on zebrafish tail regeneration. Within the tested dose range, Mollugin treatment was associated with improved tail-area recovery after injury, along with changes in apoptosis-related gene expression, inflammatory markers, antioxidant enzyme activity, and macrophage-associated markers.
Zebrafish, as a prominent model for regeneration studies, exhibit tail regeneration that involves the coordinated action of multiple signaling pathways19. After injury, caspase-3 levels increased while Bcl-2 expression decreased, indicating enhanced apoptosis. This excessive activation of apoptosis not only depletes the pool of regenerative cells but also exacerbates secondary inflammatory responses, leading to further tissue damage20,21. Treatment with Mollugin restored the balance of apoptotic markers, suggesting its potential to inhibit excessive caspase-3 activity or enhance the protective effects of Bcl-2, thereby preserving cellular resources critical for regeneration.
Inflammation is important in tissue repair. While moderate inflammation aids in pathogen clearance and necrotic tissue removal, excessive or prolonged inflammation results in the release of pro-inflammatory cytokines and upstream regulators. These factors amplify local inflammatory signals, promote apoptosis, and exacerbate oxidative stress, creating a vicious cycle9,22,23. Additionally, cell adhesion molecules such as ICAM-1 and chemokines like CCL-2 play pivotal roles in leukocyte recruitment and inflammatory cell aggregation24. Mollugin lowered these inflammatory markers and reduced local inflammation, which may support tissue repair.
Oxidative stress further aggravates tissue damage after injury. ROS overproduction not only directly damages cellular membranes, proteins, and DNA but also activates inflammatory signaling and induces apoptosis, further aggravating tissue injury25. Key antioxidant enzyme activity is diminished under injury conditions, leading to redox imbalance26. Our results demonstrate that Mollugin dose-dependently restored SOD and CAT activity, effectively mitigating ROS accumulation and oxidative damage. This suggests that Mollugin not only protects cells from direct oxidative injury but also indirectly attenuates ROS-induced inflammatory and apoptotic signaling.
Immune regulation, especially macrophage polarization, strongly influences tissue repair. Macrophages are essential for zebrafish fin regeneration, and the present findings should not be interpreted as indicating that macrophage depletion promotes tissue regrowth. Previous studies using macrophage depletion or ablation models demonstrated that macrophages are required for regenerative outgrowth, tissue patterning, blastema formation, and TNFα-dependent regeneration27,28. Importantly, a previous study showed that TNFα-expressing M1-like macrophages are recruited rapidly after fin amputation and reach a peak around 6 h post-amputation in larval zebrafish28. In contrast, the present study did not delete or deplete macrophages, and our macrophage imaging and marker-expression analyses were performed at 24 h and 48 h post-amputation. Therefore, the observed reduction in macrophage-positive area and inflammatory/M1-associated markers at 48 h should not be interpreted as suppression of the early pro-regenerative TNFα⁺ macrophage response. Mollugin treatment did not obviously reduce macrophage accumulation at 24 h, and the macrophage-positive area at 48 h remained above the uninjured control level. Thus, the observed change more likely reflects modulation of sustained macrophage-associated inflammatory dynamics during the later phase examined, rather than loss of macrophages from the injury site.
Macrophages are also functionally heterogeneous during tissue repair and regeneration. Early inflammatory macrophage responses contribute to debris clearance and activation of regenerative programs, whereas subsequent inflammation-resolving or pro-repair macrophage states participate in tissue remodeling29,30. Consistent with this concept, Mollugin treatment reduced the expression of the M1-associated marker iNOS and increased the expression of the M2-associated marker Arg-1, together with reduced inflammatory cytokine expression. These results suggest that Mollugin may influence macrophage phenotype-related responses during tail regeneration. However, because the current evidence is based on qRT-PCR and Tg(mpeg1:GFP) reporter imaging, further studies are needed to determine whether Mollugin directly regulates macrophage polarization or macrophage-dependent regenerative processes.
In summary, the present findings support an association between Mollugin treatment and improved tail-area recovery in the zebrafish tail amputation model. Rather than indicating a single defined mechanism, the observed changes in apoptosis-related genes, inflammatory mediators, antioxidant enzyme activities, and markers of the macrophage phenotype suggest that Mollugin may reshape multiple components of the local injury response. These results provide a basis for further mechanistic studies to determine how Mollugin influences the cellular events that coordinate zebrafish tail regeneration.
This study also has several limitations. First, because the present study assessed tail-area recovery only up to 48 h post-amputation, the data indicate improved early tail-area recovery rather than a definitive increase in final regenerative capacity. Complete restoration of tail morphology requires a longer observation period, and untreated amputated tails may continue to regenerate beyond 48 h. Therefore, future longer-term time-course experiments are needed to determine whether Mollugin accelerates the early phase of regeneration or alters the final extent of regeneration. Second, some experimental design and reporting issues should be considered when interpreting the results. Although the revised methods clarify that the uninjured control group and injured model group were maintained in medium containing 0.5% DMSO to match vehicle exposure, the potential influence of vehicle conditions cannot be completely excluded.
In addition, the tail amputation procedure was standardized as much as possible, but minor variation in the amputation level may still have influenced recovered tail-area measurements. Third, the mechanistic interpretation of this study is limited by the assays performed. Apoptosis-related changes were evaluated mainly by qRT-PCR analysis of caspase-3 and Bcl-2 expression, without direct cell-death assays such as acridine orange staining, TUNEL staining, or cleaved caspase-3 immunofluorescence. Macrophage-associated responses were assessed using Tg(mpeg1:GFP) reporter imaging and qRT-PCR analysis of iNOS and Arg-1, which cannot directly validate macrophage subsets or establish causality. Future studies should include direct cell-death staining, protein-level validation, macrophage subset analysis, and functional perturbation experiments. Finally, this study did not directly assess neutrophil recruitment using neutrophil reporter lines such as mpx:GFP or lys:GFP. Because neutrophils are among the earliest immune cells recruited after tissue injury and infection, future studies should evaluate whether Mollugin affects early neutrophil recruitment. In addition, qRT-PCR was performed using pooled whole-embryo samples, which may dilute tissue-specific molecular changes in the regenerating tail region. Region-restricted sampling from the anus to the tail, together with additional regeneration-related markers such as cyp26b1 and sp7, will help clarify the tissue-specific effects of Mollugin during zebrafish tail regeneration31. Therefore, the current results should be interpreted as marker-level associations rather than definitive evidence of a causal mechanism.
Overall, our findings indicate that Mollugin treatment is associated with improved tail-area recovery after zebrafish tail amputation and with coordinated changes in local injury-response markers. These results provide preliminary evidence that Mollugin may modulate the regenerative response in this zebrafish model and support further investigation of its role in regeneration-related contexts.
Disclosures
The authors declare no competing interests.
Acknowledgements
The present study was Sponsored by Health Commission of Fujian Province of China (Grant NO. 2022QNA085).
Materials
| Name | Company | Catalog Number | Comments |
|---|---|---|---|
| 6-well tissue culture plates | Corning | 3516 | |
| Catalase (CAT) assay kit | Nanjing Jiancheng Bioengineering Institute | A007-1-1 | |
| Chloroform | Sigma-Aldrich | 288306 | |
| CMax Plus microplate reader | Molecular Devices | iD3s | |
| Danieau’s buffer – Ca(NO3)2·4H2O | Guangzhou Pharmaceutical Holdings Ltd., China | N/A | Final concentration in Danieau’s buffer: 0.6 mM. |
| Danieau’s buffer – HEPES | Guangzhou Pharmaceutical Holdings Ltd., China | N/A | Final concentration in Danieau’s buffer: 5 mM; pH adjusted to 7.2. |
| Danieau’s buffer – KCl | Guangzhou Pharmaceutical Holdings Ltd., China | N/A | Final concentration in Danieau’s buffer: 0.7 mM. |
| Danieau’s buffer – MgSO4·7H2O | Guangzhou Pharmaceutical Holdings Ltd., China | N/A | Final concentration in Danieau’s buffer: 0.4 mM. |
| Danieau’s buffer – NaCl | Guangzhou Pharmaceutical Holdings Ltd., China | N/A | Final concentration in Danieau’s buffer: 58 mM. |
| Dimethyl sulfoxide (DMSO) | Sigma-Aldrich | D2650 | |
| E3 embryo medium | Laboratory-prepared (standard recipe) | N/A | Standard recipe. |
| Gene-specific qPCR primers (tnfα, il1β, il6, nfkb1, tlr4, ccl2, icam1, inos, arginase-1, caspase-3, bcl2, β-actin) | Sangon Biotech | N/A | |
| GraphPad Prism | GraphPad Software, San Diego, CA, USA | N/A | Version 8.0. |
| ImageJ software | National Institutes of Health (NIH) | N/A | Version 1.52n. |
| Mollugin (≥98%, HPLC) | Sigma-Aldrich | SMB00431 | Active compound from Rubia cordifolia, dissolved in DMSO to 40 mg/mL and diluted to 22, 66 and 200 µg/mL in E3 medium. |
| Motorized tissue homogenizer | IKA, T10 basic ULTRA-TURRAX (or equivalent) | N/A | |
| NovoScript 1st Strand cDNA Synthesis SuperMix | Novoprotein | abs60077 | |
| Phosphate-buffered saline (PBS) | Gibco, Thermo Fisher Scientific | 10010023 | pH 7.4 |
| Real-time PCR system (Archimed X4) | Sansure Biotech Inc. | Archimed X4 | |
| Refrigerated microcentrifuge | SCILOGEX | CF1524R | |
| RNA-free tips | Sangon Biotech | F603222-0010 | |
| RNAiso Plus | Takara Bio Inc. | 9109 | Phenol/guanidine-based RNA extraction reagent. |
| RNase-free microcentrifuge tubes, 1.5 mL | Axygen / Eppendorf | N/A | |
| SMZ800N fluorescence stereomicroscope | Nikon | SMZ800N | Fluorescence stereo microscope used to image Tg(mpeg1:GFP) larvae and quantify macrophage aggregation. |
| Sodium hypochlorite solution | McLean, USA | N/A | 0.003% working solution. |
| SYBR Green qPCR Mix (2× SYBR qPCR Mix) | Novoprotein | E096 | |
| SZX16 stereomicroscope | Olympus | SZX16 | Used for morphological observations, toxicity assessment and tail fin amputation of larvae. |
| Total Superoxide Dismutase (T-SOD) assay kit | Nanjing Jiancheng Bioengineering Institute | A001-3 | |
| Transgenic zebrafish Tg (mpeg1:GFP) | Anburui, Fujian, China (or in-house colony) | N/A | Macrophage reporter line. |
| Tricaine methanesulfonate (MS-222) | Sigma-Aldrich | E10521 | 0.03% working solution. |
| UltraPure DNase/RNase-Free distilled water | Invitrogen | 10977023 | |
| Wild-type zebrafish, CZ98 strain (Danio rerio) | Anburui, Fujian, China | N/A | Fertilized embryos and larvae were obtained from adult breeding stock. |
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