Paeoniflorin (purity: 98.04%) was acquired from MedChemExpress (Monmouth Junction, NJ, United States). Tin protoporphyrin IX (SnPP) and dimethyl sulfoxide (DMSO) were obtained from Sigma-Aldrich (St. Louis, MO, United States). Primary antibodies were used as following: C-caspase3 (ab32351), Bax (ab32503), Bcl-2 (ab182858), and Cytochrome C (ab133504) were obtained from Abcam (Cambridge, United Kingdom); Keap1 (sc-514914) and poly (ADP-ribose) polymerase (PARP) (sc-7150) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, United States). Primary antibodies against Nrf2 (16396-1-AP), HO-1 (27282-1-AP), Lamin B (12987-1-AP), NF-κB (66535-1-Ig) and β-actin (66009-1-Ig) were acquired from Proteintech Group (Chicago, IL, United States), and 4′, 6-diamidino-2-phenylindole (DAPI) was received from Beyotime (Shanghai, China). ML385 was purchased from ChemeGen (Shanghai, China). All experimental consumables for cell culture are from Gibco (Grand Island, NY, United States).

HUVECs were acquired from ATCC (Manassas, VA, United States) and cultured in DMEM/F12 (Gibco, Invitrogen, Grand Island, NY) with 10% heat-inactivated FBS, and 1% penicillin and streptomycin were added to a 5% CO₂ incubator at 37°C. To elucidate the role of PF in HUVEC survival, cells were exposed to differing PF (0, 5, 10, 20, and 50 μM) concentrations over 24 h. To induce oxidative stress and apoptosis in vitro, cells were cultured with a range of concentrations of TBHP (0, 100, 200, 500, and 1000 μM), and cell toxicity was measured. After determining the optimal TBHP concentration, HUVECs were exposed to different concentrations of PF (0, 5, 10, and 20 μM) and TBHP (500 μM), and worked together for 24 h to detect the PF-mediated apoptosis and deregulation protection. To delineate the function of HO-1 in this process, HUVECs were pretreated with an HO-1 inhibitor SnPP (20 μM) for 2 h before PF exposure. The data presented here represent replicas of three separate experiments.

HUVEC viability were evaluated with the Cell Counting Kit-8 (CCK-8) assay (MedChemExpress LLC; Monmouth Junction, NJ, United States), following operational guidelines. In brief, second-generation HUVECs were cultured in 96-well plates (3 × 10³ cell/well) and exposed to a combination of differing PF (0, 5, 10, and 20 μM) concentrations and TBHP (500 μM), before incubation in DMEM/F12 medium at 37°C. After 24 h, the cells were PBS-rinsed and exposed to 10 μl CCK8 in DMEM/F12 serum-free medium for an additional 2 h. Finally, absorbance was detected at 450 nm with a microwell plate reader (Thermo Fisher, Waltham, MA, United States).

Isolation of cytosol and mitochondria was performed according to the instructions for the Mitochondria/Cytosol Isolation Kit (Applygen Technologies, Beijing, China) (Zhang et al., 2014). Fifty million HUVECs were resuspended with ice-cold mito-cyto isolation buffer (Applygen Technologies Beijing, China) and homogenized. The homogenate was centrifuged at 800 × g for 10 min at 4°C. The supernatants were combined in a new tube and centrifuged at 12,000 × g for 10 min at 4°C. The supernatant contained the cytosolic fraction and the pellet contained intact mitochondria.

Western blot was conducted, as described before (Lin et al., 2018). In brief, HUVECs treated with differing PF (0, 5, 10, and 20 μM) concentrations and TBHP (500 μM) were lysed in 1 mM PMSF (phenylmethanesulfonyl fluoride) radioimmunoprecipitation analysis buffer to extract protein and were subsequently subjected to protein quantification with the BCA Protein Assay Kit (cat# P0012S, Beyotime, Shanghai, China). The equivalent amount of proteins (30 µg per sample) were separated by gel electrophoresis and transferred to the PVDF membrane (BioRad, United States). Next, the membranes were blocked in 5% skim milk at room temperature for 2 h, exposed to primary antibodies at 4°C for 18 h: cleaved caspase-3 (C-caspase3) (1:1,000), Bax (1:1,000), Bcl-2 (1:1,000), β-actin (1:1,000), Cytochrome C (1:1,000), Nrf2 (1:1,000), HO-1 (1:1,000), or Lamin B (1:800), followed by washing and exposure to corresponding HRP-conjugated secondary antibodies for 2 h at room temperature. The strips were imaged using a chemical XRS + imaging system (Bio-Rad, United States). The final protein quantification was done with Image LabV 3.0 (Bio-Rad, United States).

Apoptotic cells were detected with the in situ Cell Death Detection Kit (Roche, South San Francisco, CA, United States), following operational guidelines. In brief, HUVECs were treated with different concentrations of PF (0–20 μM) for 24 h with or without 500 μM TBHP for 2 h. The treated HUVECs were PBS-rinsed thrice and fixed in 4% paraformaldehyde for 20 min at room temperature, followed by incubation with 3% hydrogen peroxide and freshly made 0.1% triton X-100 for 10 min. At every step, the cells were PBS-rinsed thrice. We used the TUNEL reagent to detect the cells, and the amount of apoptotic cells were calculated from three arbitrary fields, using a fluorescence microscope (Olympus Inc., Tokyo, Japan).

The chamber glass slide was first prepared with endothelial cell matrix gel solution (BD Biocoat, NJ, United States) and kept in a humidified environment at 37°C for 1 h to facilitate matrix solidification. HUVECs were treated with different PF concentrations (0, 5, 10, and 20 μM) for 24 h with or without 500 μM TBHP for 2 h. Pretreated HUVECs were next harvested with trypsin/ethylenediaminetetraacetic acid, plated onto the already made matrigel-coated glass slides, and maintained at 37°C for 6 h. Tube generation was detected with a phase contrast microscope (Olympus Inc., Tokyo, Japan) and counted using randomly selected areas within each well.

HUVEC migration was evaluated using an 8 μm polycarbonate membrane Boyden chamber inserted in a transwell system (CoStar, Cambridge, MA, United States). HUVECs were treated with different concentrations of PF (0–20 μM) for 24 h with or without 500 μM TBHP for 2 h, as mentioned previously. Next, HUVECs were separated, centrifuged, and resuspended before 4 × 10⁴ cells, from each treatment, were plated into the transwell system in 400 μl of non-FBS DMEM/F12 medium. Furthermore, 600 μl of medium with 1% FBS was introduced to the bottom chamber. After a 12 h incubation in 5% CO₂ humidified environment, the membranes were PBS-washed thrice, and the cells were fixed with 4% paraformaldehyde. Additionally, any cells on top of the transwell system were removed with a cotton swab, and it was stained with crystal violet. Migratory cells that traveled from the top to the lower surface of the insert were calculated in three arbitrary fields, using an inverted microscope (Olympus Inc., Tokyo, Japan).

Double-stranded siRNA was designed and synthesized for silencing human Nrf2 gene (RiboBio, Guangzhou, China). The sequence is listed below: sense strand 5′-CGT​CAT​TGA​TGA​TGA​GGC​T-3′. HUVECs were incorporated with 50 nM siRNA and Lipofectamine 2000 reagent (Thermo Fisher, UT, United States) for 36 h, following operational guidelines. Subsequently, the cells were exposed to different concentrations of PF (0–20 μM) for 24 h with or without 500 μM TBHP for 2 h, as described before. Finally, relevant protein levels were assessed with Western blot.

The production of intracellular ROS was determined by oxidation sensitive dye (DCFDA). HUVECs were inoculated at a density of 1 × 10⁴ cell/ml and treated with a specified concentration of PF (0–20 μM) for 24 h, following by 500 μM TBHP for 2 h. The cells were rinsed with PBS and immediately treated with 10 μM DCFDA. Intracellular ROS generation was observed by fluorescence microscope (Olympus Inc., Tokyo, Japan).

HUVECs were treated with different concentrations of PF (0, 5, 10, and 20 μM) for 24 h with or without 500 μM TBHP for 2 h. Then treated HUVECs were washed with PBS three times and placed on ice for 15 min. SOD (S0086, Beyotime, China), CAT (S0082, Beyotime, China), and GPX (S0058, Beyotime, China) assay kits were used to detect the expression levels. We used the ATP Assay Kit (S0026, Beyotime, China) to assess the mitochondrial energy production in HUVECs.

HUVECs were treated with different concentrations of PF (0, 5, 10, and 20 μM) for 24 h with or without 500 μM TBHP for 2 h. The treated HUVECs were used with JC-1 staining to determine the changes of mitochondrial membrane potential (MMP). The level of mitochondrial ROS production and mitochondrial distribution was detected by MitoSox staining and MitoTracker (C1035-250μg, Beyotime, China), according to the vendors’ instructions. Fluorescence microscope (Olympus, Tokyo, Japan) was applied to capture the images.

Seventy-two adult male Sprague–Dawley rats (250–300 g) were purchased from Wenzhou Medical University (license no. SYXK [ZJ] 2020-0014) and maintained at 23 ± 2°C with 50 ± 5% humidity, and a 12 h light/dark cycle. Additionally, all the animal experimental protocols were agreed upon by the Wenzhou Medical University’s Animal Research Committee (wydw 2021-0256). The rats were randomly assigned to four experimental categories, namely, control (n = 18), PF (n = 18), ML385 (n = 18), and PF+ML385 (n = 18).

All rats were anaesthetized with 2% (w/v) sodium pentobarbital (10 mg/kg) via oral administration. Random-pattern skin flaps were created from the central dorsum, using the modified McFarlane flap technique (Lin et al., 2018). In short, an electric razor and hair removal cream were used to eliminate back hair. Next, a caudal skin flap (3 cm × 9 cm) was excised from the subcutaneous deep fascia of the rat’s back. Following this, the bilateral exposed iliac arteries were cut and sectioned. Next, the flap was placed on the donor bed and immediately sutured using 4-0 non-absorbable sutures. The flap on the back can be divided into three equal and independent areas: zone I (near the tail of the flap), zone II (middle region), and zone III (distal region). The PF group animals were administered 10 mg/kg PF via daily oral administration for 7 days after the establishment of the random-pattern skin flaps (Dong et al., 2020). Animals in the control group were given the same amount of saline (p.o 10 mg/kg, daily). The ML385 group received 30 mg/kg ML385 daily dose. Last, the PF+ML385 animals received 10 mg/kg PF and 30 mg/kg ML385 daily dose, similar to the other groups. After seven consecutive days of treatment, the rats were sacrificed with excess anesthesia, and tissue samples (1 cm × 1 cm) were collected from the zone II flaps for further study.

Macroscopic properties, such as appearance, color, and hair condition of the flap, were observed and recorded on day 7 after the operation. On the 7th day, the percentage of the flap survival area was assessed using ImageJ software (ver. 6.2; Media Cybemetics). The survival area (in %) was obtained as listed below: % survival = survival area ÷ total area × 100%.

To examine vascularization under the flap, rats were anesthetized and made to lie prone, while vascularization was recorded with a laser dopplerometer on postoperative day 7 (POD 7). The data obtained were analyzed with moor LDI review (ver.6.2; Moor Instruments), and perfusion units (PU) were used to assess the blood flow intensity. Scans were performed thrice, and the mean of the results was used to search for statistical significance.

Six tissue specimens (1 cm × 1 cm) were harvested per group from area II to conduct hematoxylin and eosin (H&E) staining. These specimens were fixed in 4% (v/v) paraformaldehyde for 1 day, embedded in paraffin wax before being cut into 4 µm thick sections, and stained with H&E. The sections were observed under an optical microscope (Leica, Wetzlar, Germany). Microvessels were measured on six arbitrary fields per section, and the microvessel quantity per unit area (/mm²) was counted to assess the level of microcirculation.

Rat skin tissues obtained from area II of the flaps were fixed in 4% paraformaldehyde. Following deparaffinization of the tissue sections in xylene, the sections were rehydrated via graded ethanol bath. Next, the samples were PBS-washed thrice, blocked with 3% hydrogen peroxide, and repaired in 10.2 mM sodium citrate buffer before an overnight exposure with following primary antibodies: CD34 (1:100), Nrf2 (1:100), VEGF (1:100), HO-1 (1:100), NF-κB (1:100), Bax (1:100), and C-caspase3 (1:100). Subsequently, tissue samples underwent exposure to HRP-conjugated secondary antibody, followed by staining with the DAB kit, and counterstained with hematoxylin, before visualization under a light microscopy (200× magnification). Quantified absorption values were estimated via Image-Pro Plus to detect CD34, Nrf2, HO-1, Bax, NF-κB, VEGF, and C-caspase3 expression levels. Six randomly chosen fields from six sections were analyzed for immunohistochemistry (IHC) statistical calculation.

For cell immunofluorescence, HUVECs were grown in a 6-well plate for 24 h exposed to PF (20 μM) for 24 h, subsequently, the cells were again exposed 500 μM TBHP for 2 h. Next, the cells were PBS-rinsed, fixed with 4% paraformaldehyde for 15 min, permeabilized with 0.1% TritonX-100 in PBS for 10 min, blocked with 5% bovine serum albumin (BSA) for 1 h at room temperature, with subsequent overnight exposure at 4°C to primary antibodies against Nrf2 (1:200; Abcam), followed by exposure to Alexa Fluor^®488- or Alexa Fluor^®594 goat anti-rabbit IgG (H+L) secondary antibodies (1:300; Jackson Immunoresearch, PA, United States) for 1 h at 37°C before staining with DAPI (Beyotime, China) for 5 min, and finally observed under a Nikon ECLIPSE Ti microscope (Nikon, Japan). Six arbitrary fields were analyzed for each slide. Fluorescence quantification was done with Image-Pro Plus (Media Cybernetics, Rockville, MD, United States).

For tissue immunofluorescence, the tissue specimen was deparaffinized, rehydrated, and PBS-washed in the initial steps of immunofluorescence preparation. Meanwhile, the endogenous peroxidase was quenched with 3% hydrogen peroxide, and tissue antigen was repaired using 10.2 mM sodium citrate buffer. The slides were then incubated with primary antibody against a-SMA overnight at 4°C. After washing with PBS, the slices were incubated with Texas red-conjugated anti-IgG secondary antibody and stained by DAPI. Visualization was done with a fluorescence microscope (Zeiss, LSM800, Germany).

Total RNA was extracted from the skin flaps using TRIzol reagent according to the manufacturer’s specifications. Quantification was performed using reverse transcription (RT) and PCR. RT reaction containing 0.5 μg of RNA, 2 μl of 5× TransScript all-in-one SuperMix for qPCR, and 0.5 μl of gDNA remover in a total volume of 10 μl. The reactions were performed in a GeneAmp^®9700 PCR system (Applied Biosystems, United States) for 15 min at 42°C and 5°s at 85°C. The RT reaction mix was diluted in nuclease-free water and stored at −20°C. The real-time PCR was performed using a LightCycler^®480 II real-time PCR instrument (Roche, Switzerland) in 10 μl PCR mixture containing 1 μl of cDNA, 5 μl of 2× PerfectStartTM Green qPCR SuperMix, 0.2 μl of forward primer, 0.2 μl of reverse primer, and 3.6 μl of nuclease-free water. The samples were incubated in a 384-well optical plate (Roche, Switzerland) at 94°C for 30°s, followed by 45 cycles of 94°C for 5°s and 60°C for 30°s. Each sample was assayed in triplicate. At the end of PCR, melting curve analysis was performed to validate specific generation of expected PCR products. The primer sequences were designed in the laboratory and synthesized by TsingKe Biotech based on the mRNA sequences obtained from the NCBI database as follows: Nrf2, 5′- GCA​TTT​CGC​TGA​ACA​CAA-3′ (forward), and 5′-CTC​TTC​CAT​TTC​CGA​GTC​A-3′ (reverse); H O -1, 5′-AGA​GTT​TCT​TCG​CCA​GAG​G-3′ (forward), and 5′-GAG​TGT​GAG​GAC​CCA​TCG-3′ (reverse); Bax, 5′-CGG​CTG​CTT​GTC​TGG​AT-3′ (forward), and 5′- TGG​TGA​GTG​AGG​CAG​TGA​G -3′ (reverse); Bcl-2, 5′-CGG​GAG​AAC​AGG​GTA​TGA-3′ (forward), and 5′- AGG​CTG​GAA​GGA​GAA​GAT​G-3′ (reverse); glyceraldehyde 3-phosphate dehydrogenase (GAPDH) expression was used as an internal control using the 2^−ΔΔCt method to evaluate the expression level of the target gene.

Nrf2 (PDB ID: 4ZY3) was subjected to docking studies (Khan et al., 2017). The protein was downloaded from PDB (https://www.rcsb.org/) before being prepared for docking. The lowest energy conformations for docking were determined via default parameters after being minimized using PYMOL 2.3.4. The protein–ligand docking analysis was carried out using AutoDock Vina 1.1.2, which can provide ligand binding flexibility with the binding pocket residues. Finally, the images were generated using UCSF PyMoL.

Data are represented by the means ± standard deviation. All statistical analyses were performed by one-way analysis of variance (ANOVA) with Tukey’s multiple comparisons test to assess for differences among the groups. p value < 0.05 was regarded as statistically significant.

To estimate whether PF exerts cytotoxic effects on HUVECs, HUVECs were treated with various doses of PF (Figure 1B). Damaged cells were not observed below 20 μM PF in HUVECs. Based on this, 20 μM PF was determined as the highest nontoxic concentration for subsequent experiments. To assess the optimal concentration of TBHP as a suitable exogenous oxidative stimulus source, the HUVECs were then treated with TBHP concentration gradient (0–1,000 μM), and subsequently, the cell activity was detected. With the increasing of TBHP concentration, the cell morphology atrophy gradually increased, and apoptotic bodies were visible. Compared with unprocessed group, the cell viability was significantly decreased to 54.6 ± 4.5% at 500 μM TBHP (Figure 1C). Subsequently, PF demonstrated a protective effect on TBHP-treated cell damage in a dose-dependent manner (64.9 ± 1.8%, 76.8 ± 3.4%, 89.3 ± 1.9%, respectively, at 5, 10, and 20 μM PF, comparing with that of TBHP-treated group), indicating that PF had exerted cytoprotective effect on TBHP-mediated cytotoxicity (Figures 1D,E). Additionally, TUNEL were introduced for the assessment of PF-induced protection of TBHP-stimulated HUVECs. As expected, the amount of TUNEL+ cells was upregulated with exposure to TBHP but alleviated with PF in a concentration-dependent manner (35.8 ± 1.7%, 28.3 ± 0.8%, 11.8 ± 3.7%, respectively, at 5, 10, and 20 μM PF, comparing with that of TBHP-treated group (51.5 ± 3.4%) (Figures 2A,B). To assess whether the cytoprotective effect of PF was attributable to antioxidant activity, the expression of antiapoptotic proteins such as PARP and Bcl-2 was measured, and proapoptotic proteins such as Bax and cleaved caspase-3 were calculated in HUVECs treated with or without TBHP. TBHP significantly downregulated PARP and Bcl-2, which was shown in Figure 2C. However, PF restored these protein levels in a dose-dependent manner compared with the TBHP-induced group, especially at 20 µM. PF also significantly increased the levels of PARP and Bcl-2 compared with the TBHP-administered group, suggesting that PF could potentially promote the expression of antiapoptotic proteins in HUVECs treated with TBHP (Figures 2D,F). As envisioned, compared with the unadministered group, TBHP had induced massive expression of Bax and cleaved caspase-3; furthermore, these effects were reversed in the PF-treated group, indicating that PF inhibited the expression of Bax and the cleavage of caspase-3 in TBHP-induced oxidative stress (Figures 2E,G). In conclusion, these results suggested that PF had protected HUVECs from apoptosis in response to TBHP-mediated oxidative stress.

Then, the effect of PF on TBHP-induced mitochondrial depolarization and mtROS generation was investigated. To assess the expression of related proteins in mitochondrial dysfunction, the expression of cytochrome C was measured. The Western blotting results had revealed that cytoplasmic cytochrome C was significantly increased by PF in a concentration-dependent manner, while the level of mitochondrial cytochrome C had decreased (Figures 3A,B). Next, the mitochondrial functional damage and mtROS production in TBHP-treated HUVECs were explored. The high expression of mtROS and mitochondrial dysfunction severely affected the HUVECs functional state. MitoSox red staining revealed that compared with the untreated group, the TBHP-treated group resulted in 172.7 ± 5.2% of mtROS. Then, compared with untreated group, the effect at 20 µM of PF co-treated with TBHP (500 µM) had nearly restored the mtROS generation (Figure 3D). Consistent with the MitoSox data, it was also found that fluorescence images of MitoTracker and JC-1 had largely restored the mitochondrial membrane potential decreased by TBHP. Immunofluorescence staining also revealed that treatment with TBHP significantly increased mtROS (MitoSOXRed) generation in the mitochondria (MitoTracker); however, PF had completely suppressed TBHP-mediated mtROS generation (Figures 3C,E). In addition, TBHP-treated HUVECs, co-cultured with PF, also restored the activities of SOD, CAT, and GPx (Figures 3F–H), along with ATP levels (Figure 3I). In conclusion, these results have suggested that PF downregulated the intracellular ROS generation in TBHP-induced HUVECs by stabilizing the MMP and subsequently inhibiting oxidative stress–mediated apoptosis.

To determine the therapeutic benefit of PF on cellular function, neovascularization evaluation was first performed via the tube formation assay, and the number of endothelial cell tubules in each group was counted. Based on a dose-dependent manner, it was found that TBHP markedly reduced HUVEC neovascularization, while PF pretreatment prevented TBHP effects (Figures 4A,D). Next, to elucidate the role of PF on HUVEC migration, transwell examination was performed. It was proved that markedly fewer cells had underwent migration after treated with TBHP, while PF reversed these effects (Figures 4B,E). Finally, to assess the effect of PF on HUVEC adhesion, a fibronectin adhesion assay was performed (Figures 4C,F). Based on the analysis, TBHP stimulation markedly lowered the quantity of adherent cells, and this effect was reversed by PF treatment.

Nrf2 nuclear translocation directly regulates HO-1 promoter activity, which protects cells against oxidative stress and a variety of toxins (Ndisang, 2017). It was explored in the study whether PF could upregulate HO-1 under the oxidative stress condition. The results have shown that compared with the TBHP-processed group, PF had significantly increased HO-1 expression in a concentration-dependent manner (Figure 5A). The role of HO-1 in PF-mediated cytoprotection was demonstrated using SnPP, a known HO-1 inhibitor. The existence of SnPP vastly eliminated the inhibition of DCFDA intensity by PF mediation in TBHP-induced oxidative stress, which markedly promoted the intensity regardless of the presence of PF, indicating that PF inhibited ROS production by activating HO-1 expression (Figures 5B,C). The results have shown that HO-1 acted as the key detoxifying enzyme against TBHP-induced oxidative stress conditions. The Western blotting results also showed that TBHP, combined with HO-1 inhibitor SnPP, attenuated the protective effect on HUVECs induced by PF (Figures 5D,E). In conclusion, these experimental data have suggested that PF played a cytoprotective role through the activation of Nrf2/HO-1 axis.

Since Nrf2 is an important transcription factor for maintaining the balance of redox responses (Khitan et al., 2014), the beneficial ways in which PF may be involved in HUVECs exposed to TBHP-mediated oxidative stress were investigated. First, nucleoprotein components were prepared to assess the expression of Nrf2 in different stimulation states through Western blotting. PF and TBHP co-treatments significantly upregulated the expression of Nrf2 and its nuclear translocation. However, Nrf2 was only slightly increased under the stimulation of TBHP. In particular, pretreatment with PF significantly increased the nuclear Nrf2 expression induced by TBHP, allowing much Nrf2 to translocate to the nucleus (Figures 6A,B). Immunostaining further confirmed the nuclear translocation of Nrf2 (Figure 6C). In addition, Western blotting showed that PF had significantly downregulated Keap1 expression with or without TBHP stimulation (Figure 6D), indicating that PF-mediated degradation of Keap1 would cause Nrf2 to migrate to the nucleus. And Nrf2 silencing abolished the PF-induced cytoprotection. After transfection with Nrf2 small interfering ribonucleic acid, Western blot analysis showed that small interfering RNA against Nrf2 had markedly diminished nuclear Nrf2 levels and reduced cytoplasmic HO-1 levels in TBHP and PF co-exposed HUVECs (Figures 6G,H). Furthermore, Nrf2 siRNA reversed PF-induced alteration of apoptotic proteins (Figures 6E,F). Furthermore, whether there was any affinity between PF and Nrf2 or upstream proteins in the Nrf2 pathway was assessed via computational molecular docking analysis (Luo et al., 2020). For this analysis, the PF chemical structure was used, as shown in Figure 1A. After the examination of all generated models, PF was found to have clearly interacted with and docked in the Nrf2 binding site (Figure 6I), with macro- and local-level views of these interactions shown using a ribbon model. A space-filling model was additionally used to illustrate this interaction. High-affinity (−9.2 kcal/mol) hydrogen binding events between PF and the Ser508 and Ser555 residues of Nrf2 were observed, indicating that PF might have functioned to partially inhibit the development of the flap survival through its ability to interact with Nrf2 in a manner promoting its nuclear translocation. These results indicated that PF treatment had significantly stimulated the nuclear translocation of Nrf2, leading to the activation of antioxidant defense mechanisms.

After operation, the distal skin flap appeared pale and swollen (Figure 7A). Tissue necrosis was first evident in zone III, where it became dry, crumpled, and dark. Necrosis gradually spread from distal areas to the proximal segment on POD 7. Based on our analysis and compared to PF+ML385-treated animals, a larger percentage of the random-pattern skin flaps survived in the PF-treated animals (34.90 ± 5.85%, 74.60 ± 6.69%, 28.06 ± 5.06% and 52.84 ± 7.13%, respectively, corresponding to the control group, PF group, ML385 group, and PF+ML385 group; Figure 7B). To assess whether PF affected microvascular network on the rat’s back, laser doppler was used to visualize the blood flow. It was revealed that compared to the control group, the PF-treated animals had evident blood flow signal in the flaps. Moreover, the enhanced blood flow was impaired in animals treated with PF+ML385 (421.36 ± 17.48PU, 527.04 ± 28.52PU, 383.24 ± 9.89PU, and 417.43 ± 10.42PU, respectively, corresponding to the control group, PF group, ML385 group, and PF+ML385 group; Figures 7C,D). Next, multiple angiogenesis markers have been used to evaluate the level of neovascularization in the PF-treated flaps. Hematoxylin–eosin staining results have demonstrated that PF-treated animals had remarkably more vessel density than those in the control group. Conversely, a combined treatment of PF+ML385 had dramatically reduced vessel density comparing to PF treatment alone (174.17 ± 13.64/mm², 303.17 ± 30.62/mm², 154.17 ± 13.26/mm², and 190.00 ± 14.21/mm², respectively, corresponding to the control group, PF group, ML385 group, and PF+ML385 group; Figures 7E,F). Similarly, the quantity of CD34-positive vessels in PF-treated animals was significantly higher than that of the controls, and the opposite was observed in animals co-treated with PF+ML385 (197.83 ± 8.57/mm², 306.50 ± 11.06/mm², 185.83 ± 7.00/mm², and 218.83 ± 13.52/mm², respectively, corresponding to the control group, PF group, ML385 group, and PF+ML385 group; Figures 7G,H). It was also explored whether the positive effect of PF on flap survival might be obtained via enhancing angiogenesis. IHC was employed for the detection of VEGF expression. Compared to the control group and the PF+ML385 group, VEGF expression was considerably enhanced in the flap II area of the PF group, while VEGF expression in the ML385 group was significantly lowered, as depicted in Supplementary Figures S1A,B. The results were consistent with those obtained from immunoblotting (Supplementary Figures S1C,D). Consistent with these results, immunofluorescence was employed for the detection of a-SMA–positive microvessels, which were more significant in the PF-treated animals than in the control animals, while the PF+ML385 co-treatment vastly reduced the a-SMA positive microvessel quantity comparing to PF treatment alone (Figures 7I,J). In all, these data have suggested that PF promoted angiogenesis and played a crucial role in improving the skin flap viability.

To explore whether PF modulates the Nrf2 pathway in vivo, the levels of Nrf2 pathway-related markers with IHC were evaluated. IHC and integrated absorbance analysis indicated that the Nrf2 and HO-1 expression in the PF group were considerably increased compared to the control group and the PF+ML385 group. The Nrf2 and HO-1 expression were decreased in the ML385 group; however, no considerable variations were observed between the control group and the PF+ML385 group, as depicted in Figures 8A,B. Additionally, the level of NF-κB, C-caspase3, and Bax in the PF group analyzed by IHC showed significantly lower expression than that of those in the control group. And these protective effects of PF-induced were decreased by PF co-administration with ML385. The data of real-time quantitative PCR assays have demonstrated that transcription of Nrf2, HO-1, and Bcl-2 was reduced after being treated with ML385 (Figure 7C). The results also showed that the level of Bax expression was increased in the ML385 group compared to the control group and the PF+ML385 group, while no considerable variations were observed between the control group and the PF+ML385 group, as depicted in Figure 8C. To sum up, the IHC and real-time quantitative PCR assay results showed that PF promoted the skin flap viability via Nrf2/HO-1 signaling pathway in vivo.