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. 2022 Oct 12;31(13):1741–1751. doi: 10.1007/s10068-022-01163-3

Protein from Hylocereus polyrhizus protects MRC-5 cells against hydrogen peroxide (H2O2)-induced damage

Haomiao Ding 1, Yuzhe Zhang 1, Yue Zha 1, Sijia Zhou 1, Chaoqing Cao 1, Huajian Zhu 1, Feng Xu 2, Xiuqiang Zhang 2,✉, Caisheng Wang 1,✉
PMCID: PMC9596649  PMID: 36312996

Abstract

The cytoprotective and potential molecular mechanisms of Hylocereus polyrhizus protein (RFPP) were investigated on the hydrogen peroxide (H2O2)-triggered damage in normal human embryonic lung (MRC-5) cells. An MTT assay was conducted to assess the MRC-5 cell viability after exposure to H2O2 or RFPP. Cell cycle distribution and apoptosis were explored via flow cytometry. The contents of related proteins were assessed via western blot. MRC-5 cells exhibited markedly decreased cellular viability after treatment with H2O2; however, treatment with RFPP suppressed this decrease. Additionally, RFPP interference dampened H2O2-triggered intracellular apoptosis levels and increased H2O2-triggered intracellular S phase. In these processes, the contents of phosphorylated (p)-AKT along with p-mTOR proteins were downregulated in 120 µM H2O2-treated cells compared with vehicle-treated cells. Nevertheless, in MRC-5 cells inoculated with RFPP, the levels expression of these proteins were reversed. To conclude, RFPP protected MRC-5 cells from H2O2-triggered damage via activation of the PI3K/AKT/mTOR cascade.

Keywords: Hylocereus polyrhizus, Protein, MRC-5 cells, Damage, PI3K/AKT/ mTOR cascade

Introduction

Redox homeostasis is essential for normal body functioning (Chen et al., 2019), which is strictly regulated by reducing molecules, antioxidant enzymes, two-phase detoxification enzymes, and other molecules on cell receptor (Ye and Meng, 2021). Mounting evidence suggests that many diseases, including obstructive pulmonary disease, cataracts, arthritis, neurodegenerative diseases and cancer, are related to oxidative stress damage (Zeng et al., 2022; Zhu et al., 2022). Therefore, studying the antioxidant effects of the active ingredients in natural products is necessary for preventing, as well as treating various diseases.

The oblong-shaped, red-fleshed pitaya (Hylocereus polyrhizus), often known as dragon fruit in English or Buah Naga in Malay, has scaly features on its outer peels. (Roriz et al., 2022). It is from the Hylocereus genus and belongs to the Cactaceae family. (Zulkifli et al., 2020). Pitaya is a fruit crop that originated in Central and Northern South America and is now grown in Malaysia, Thailand, Vietnam, Australia, Taiwan, and other parts of the globe (Tsai et al., 2019). Pitaya has attracted much interest for its excellent nutritional benefits, in addition to its attractive appearance. The fruit’s flesh is sweet and delicious, with several small and gritty black seeds. Pitaya fruit seeds contain oil that is comparable to that found in grapes and linseeds, as well as berries, for instance blackberries and red raspberries. The seed oil from the pitaya fruit has been effectively extracted and is said to have potential uses in food, health, as well as cosmetics (Tako et al., 2020). Pitaya protein is a type of plant 2S albumin, however, the biological activities of the red-fleshed pitaya protein (RFPP), a byproduct from defatted pitaya seeds after extraction, remains unexplored.

Plant 2S albumin is a plant seed protein that is widely present in mono-dicotyledonous plants (Khan et al., 2016). Plant albumin is a mixed protein with a sedimentation coefficient of 2 and a molecular weight ranging from 10–20 kDa and 10–20 subunits (Freire et al., 2015). The amino acid component contains a large amount of sulfur-containing amino acids, which can be dissolved in water, dilute salt, dilute acid or dilute alkali solution, and can be heated and cured. Studies have found that albumin from plant seeds, such as peanuts, mustard, castor beans, sesame seeds, peach seeds and cashew seeds, has allergenic properties (Cristina et al., 2021). Recent studies have revealed other valuable biological activities from plant; thus, plant seed albumin has become an important topic in life science research (Soboleva et al., 2018; Tarhini et al., 2020; Zhu et al., 2018).

The phosphatidylinositol 3-kinase (PI3K)/protein kinase B (AKT)/mammalian target of rapamycin (mTOR) cascade is a signal transduction pathway closely involved in cell differentiation, survival, proliferation, angiogenesis, and metabolism (Slomovitz and Coleman, 2012). Investigations have documented that the PI3K signaling cascade affects the activation states of various downstream effector molecules and plays roles in inhibiting apoptosis and promoting cell proliferation (Rajendran et al., 2020). Bcl-2 family proteins regulate apoptosis through mitochondrial permeability. However, few published studies have reported on RFPP-associated regulation of the PI3K/AKT/mTOR and Bcl-2/Bax/Caspase-3 cascade. Here, we examined the protective effects of purified RFPP against peroxidative injury in human embryonic lung (MRC-5) cells to offer a scientific basis for further development and use of RFPP.

Materials and methods

Materials

DEAE-52 cellulose, Sephadex G-75 columns, along with FBS were obtained from GE Healthcare Life Sciences (Logan, UT, USA). H2O2 was obtained from Shanghai Aladdin Science and Technology Co., Ltd. (Shanghai, China). DMSO, MTT, (protein extraction kits, BCA) protein assay kits, reactive oxygen species (ROS) assay kit, as well as TBST were provided by Beijing Solarbio Science and Technology Co., Ltd. (Beijing, China). Penicillin G, Dulbecco’s modified Eagle’s medium (DMEM), along with streptomycin were supplied by Gibco, Thermo Fisher Scientific, Inc. (Waltham, MA, USA). FITC annexin V apoptosis detection kits coupled with PI/ RNase staining buffer were supplied by BD Biosciences (San Diego, CA, USA). The test kits for SOD, GSH-Px, MDA along with nitric oxide (NO) were provided by Nanjing Jiancheng Bioengineering Research Institute Co., Ltd. (Nanjing, China). GAPDH rabbit monoclonal antibody (mAb; cat. no. 5174), PI3K antibody (cat. no. 4257), Akt rabbit mAb (cat. no. 9272), phospho-Akt (Ser473) rabbit mAb (cat. no. 2764), goat anti-rabbit IgG-HRP (cat. no.7074), mTOR antibody (cat. no. 2972), along with phospho-mTOR antibody (cat. no. 2971), Bax antibody (cat. no. 9292), Bcl-2 antibody (cat. no. 7223), Caspase-3 antibody (cat.no. 9662) were supplied by Cell Signaling Technology, Inc. (Boston, MA, USA). All other reagents were of analytical grade.

Extraction of RFPP from H. polyrhizus

RFPP (purity > 99.0% and molecular weight 19.8 kDa) was obtained from red-fleshed pitaya extracts as previously described (Chen et al., 2017). Hylocereus polyrhizus was purchased from the Cambodian red meat variety of Ningbo Green Garden Agricultural Development Co., Ltd., Zhejiang Province, China, and was selected from mature fresh red pitaya in July or August of that year. Seeds were extracted from pulp and rinsed with running tap water in order to get rid of mucilage before being dried in the lab. Dried seeds were pounded into a powder and stored in airtight bottles. Deffatening of the seeds was then done via maceration with n-hexane as the solvent. Before proceeding with following protein extraction processes, the defatted seeds were allowed overnight in a fume hood to allow remaining n-hexane to evaporate.

Thirty grams of defatted seeds were pretreated with citric acid-sodium citrate buffer solution (10 mmol/L, pH 7.08; 1:30 w/v) at 74 °C and extracted for 30 min. The filtrate was collected, then span at 10,000 g for 20 min. Finally, the crude protein was extracted via lyophilizing the deproteinized supernatant. The crude extracts were refined using DEAE-52 cellulose columns, yielding two H. polyrhizus proteins that were lyophilized for future research. The first RFPP extract had a molecular weight of 13.2 kDa.

Cell lines and cultures

MRC-5 cells were supplied by the Shanghai Institute of Cell Biology, Chinese Academy of Sciences. MRC-5 cells were inoculated in DMEM enriched with 10% FBS, penicillin (100 IU/mL) along with streptomycin (100 µg/mL) under 37 °C and 5% CO2 conditions. The cells were inoculated on a culture plate, and the experiment was carried out after the cells reached the logarithmic growth phase.

Establishment of the MRC-5 cell injury model

5 × 103 MRC-5 cells per well were inoculated into a 96-well culture plate under 37 °C along with 5% CO2 conditions. When the cells reached 80% confluence, the medium was subsequently replaced with 0, 20, 40, 60, 80, 100, 120, 140, 160, 180, or 200 µM H2O2 diluted with DMEM for 0, 0.5, 1, 1.5, 2, 2.5, 3, or 3.5 h, with five parallel wells per group. The effect of H2O2 on MRC-5 cell toxicity was determined using MTT tetrazolium dye assay kits (Guo et al., 2020). After H2O2 treatment, 10 µL of 5 mg/mL MTT was introduced to every well and cultured for 4 h. The culture medium was then removed, and 100 µL DMSO was introduced to every well to dissolve the formed blue formazan crystals. Finally, the absorption value of each well was determined at 490 nm by a microplate reader (SpectraMax 190, San Jose, CA, USA). When cell viability reached around 50%, the associated H2O2 concentrations and treatment periods were considered as the optimal conditions for oxidative damage. The mean standard deviation (SD) of three experiments with five wells per treatment group is reported for all experimental data.

Effect of RFPP on cell viability in H2O2-triggered MRC-5 cells

5 × 103 MRC-5 cells per well inoculated onto 96-well plates at in 100 µL of complete culture medium for 24 h. The cells were inoculated with 120 µM H2O2 for 3.5 h, then exposed to 0, 20, 40, or 80 µM RFPP for 24 h. After treatment, MTT solution was introduced to every well and incubated at 37 °C for 4 h. The produced formazan crystals were dispersed in DMSO. The MTT assays were read on a microplate reader at an absorbance of 490 nm, and the cell viability owing to RFPP treatment was calculated as the percentage of viable cells relative to the control cells treated with DMEM alone.

Measurement of antioxidant parameters

1 × 105 MRC-5 cells per well were inoculated in 6-well plates under 5% CO2 conditions for 24 h. Afterwards, we inoculated the cells with 120 M H2O2 for 3.5 h before being incubated in different dosages of RFPP for 24 h. After that, we homogenized the cells in RIPA lysis solution before being span at 12,000 g for 15 min at 4 °C. The antioxidant parameters were assessed using the supernatant. Commercial assay kits were utilized to assess SOD along with GSH-Px activity, NO release, as well as MDA levels as per the manufacturer’s recommendations. Measurement of intracellular ROS was performed using 2, 7-dichlorofluorescin-diacetate (DCFH-DA). Cells were incubated with 5 µM DCFH-DA diluted in DMEM for 20 min at 37 °C in the dark. For flow cytometry, cells were collected for analysis.

Apoptosis analysis using Hoechst 33258 staining

The nuclear morphologic alterations in apoptotic cells were explored via Hoechst 33258 staining (Zhang et al., 2019). 1 × 105 MRC-5 cells/well were inoculated in six-well plates with a coverslip for 24 h. After that, the cells were inoculated with 120 µM H2O2 for 3.5 h before being inoculated with 20, 40, or 80 µM RFPP for another 24 h. The cells on the coverslip were stained with 500 µL Hoechst 33258 per well, as directed by the manufacturer, and observed under a fluorescence microscope (Leica DM4000, Leica, Germany).

Flow cytometry analysis of apoptosis of MRC-5 cells

Flow cytometry (BD FACSVerse, Franklin Lakes, New Jersey, USA) was utilized to verify RFPP-triggered apoptosis with the annexin V-FITC/PI apoptosis detection kit. we inoculated 1 × 105 MRC-5 cells/well into 6-well plates for 24 h and the inoculated them with 120 µM H2O2 for 3.5 h, followed by inoculation with 20–80 µM RFPP in complete DMEM enriched with 10% FBS for 24 h. Cells were rinsed twice in cold PBS and re-suspended in 1⋅ binding buffer at 1 × 106 cells per mL, then moved to 5 mL culture tubes in 100-L aliquots containing 1 × 105 cells each. The cells were vortexed gently and incubated for 15 min at 25 °C in the dark after addition of 5 µL FITC annexin V and 5 µL PI. Triplicate assays were carried out.

Cell cycle analysis

Flow cytometry (BD) was utilized to confirm RFPP-triggered apoptosis. Flow cytometry was adopted to measure the DNA content of the cells throughout the cell cycle (Xu et al., 2020a). The nuclear DNA-docking agent PI was utilized. The fluorescence intensity of the distinct cell cycle stages was adopted to identify them. After 24 h of RFPP treatment, MRC-5 cells were collected, rinsed twice in PBS, spun at 500 g for 10 min, and fixed overnight in 70% ethanol at 20 °C. Flow cytometry was adopted to assess cells that had been stained with 0.5 mL PI/RNase staining solution at 25 °C for 15 min.

Western blot analysis

Isolation of total proteins from the MRC-5 cells was done using a protein extraction kit. Thereafter, quantitation of proteins was done with the BCA assay kit. Subsequently, fractionation of 20 µg aliquots of cytoplasmic or nuclear extracts was done on the SDS-PAGE-gels, and blotted onto nitrocellulose membranes. Afterwards, blocking of membranes was done using 5% skimmed milk powder for 2 h at 25 °C, then overnight inoculated with the following primary antibodies at indicated dilutions: anti-Bax (1:1000), anti-Bcl-2 (1:1000), anti-Caspase-3 (1:1000), anti-PI3K (1:1000), anti-AKT (1:1000), anti-p-AKT (1:2000), anti-mTOR (1:1000), or anti-p-mTOR (d1:2000) or the anti-GAPDH (1:2000) control at 4 °C. After that, we rinsed the membranes three in TBST for 15 min, then inoculated with HRP-linked secondary antibodies (1:2000) for 2 h at 25 °C (Ding et al., 2021). Visualization of antibody binding was done via electrochemiluminescence (BeyoECL Star; Beyotime, Hangzhou, China) and imaged using a ChemiDoc imaging platform (BIO-RAD, Hercules, CA, USA).

Statistical analysis

Mean values are given as the average of three replicates ± SD. Comparisons were assessed via Student’s t-test or one-way analysis of variance followed by Tukey’s test implemented in the SPSS 21.0 software (SPSS Inc., Chicago, IL, USA). p < 0.05 signified statistical significance.

Results and discussion

Effect of RFPP on cell viability in H2O2-triggered MRC-5 cells

The MTT assay was used to examine the concentration along with time-dependent viability losses in MRC-5 cells inoculated with H2O2. MRC-5 cell viability dramatically diminished as the H2O2 concentration increased from 20 to 200 µM (p < 0.05; Fig. 1A). After pretreatment with H2O2 for 3.5 h, the cell viabilities were 49.39 ± 2.10% in 120 µM, 45.15 ± 4.12% in 140 µM, 35.14 ± 0.54% in 160 µM, 27.73 ± 0.82% in 180 µM and 24.02 ± 1.17% in 200 µM. Thus, H2O2 greatly damaged the MRC-5 cell viability. In vitro oxidative damage has been frequently triggered using H2O2, an essential active oxygen molecule with highly stable characteristics (Wang et al., 2015). Previous research has demonstrated that when cell viability was about 50%, the associated H2O2 concentration and treatment periods provided the optimal circumstances for oxidative damage (Wang et al., 2019a). Thus, we used H2O2 to establish an MRC-5 cell model of oxidative damage. Stimulating MRC-5 cells using 20–200 µM H2O2 greatly decreased the viability of MRC-5 cells. The viability of MRC-5 cells decreased to 51.25 ± 2.67% after treatment with 120 µM H2O2 for 3.5 h. Hence, we used a 3.5-h exposure time with 120 µM H2O2 in our follow-up experiments.

Fig. 1.

Fig. 1

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The viability of MRC-5 cells. (A) Impact of different H2O2 concentration (0, 20, 40, 60, 80, 100, 120, 140, 160, 180, or 200 µM) diluted with DMEM for 0, 0.5, 1, 1.5, 2, 2.5, 3, or 3.5 h on MRC-5 cell viability. (B) Impact of different RFPP concentration (0, 20, 40, or 80 µM) for 24 h on the viability of 120 µM H2O2-treated MRC-5 cells after 3.5 h. Cell viability was measured using MTT assay. The experimental results are given as the mean ± SD, n = 5

The most obvious indicator of the extent of cell damage triggered by the external environment is cell viability. Viability of the cells treated with H2O2, a stable active oxygen molecule often used in models of oxidative damage, decreased remarkably compared with that of the vehicle (DMEM only)-treated cells (p < 0.01; Fig. 1B), illustrating that the model of oxidative damage was effectively developed. RFPP treatment (20–80 µM) markedly increased cell viability dose-dependently in H2O2-treated MRC-5 cells in contrast with that of the H2O2-only group (p < 0.05). These data imply that RFPP protected against H2O2-mediated oxidative damage in the MRC-5 cells.

Effects of RFPP on SOD and GSH-Px activities, NO release and MDA levels in H2O2-triggered MRC-5 cells

To clarify whether antioxidant properties cause the protective impact of RFPP on H2O2-triggered MRC-5 cells, we used commercial kits to determine the SOD along with GSH-Px activities, NO release and MDA contents. The SOD coupled with GSH-Px contents were markedly decreased in MRC-5 cells, while the NO release along with MDA contents were remarkably increased in the 120 µM H2O2-inoculated cells in contrast with the controls (p < 0.05; Fig. 2A, B). On the other hand, intervention with RFPP (20–60 µM) improved the SOD along with GSH-Px activities and attenuated the NO release, as well as MDA contents in H2O2-triggered MRC-5 cells (p < 0.05; Fig. 2C, D). Flow cytometry indicated that RFPP decreased ROS levels in H2O2-triggered MRC-5 cells (Fig. 2E). Our results imply that the protective impact of RFPP on MRC-5 cell oxidative damage triggered by H2O2 was due to improved cellular antioxidant systems.

Fig. 2.

Fig. 2

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Impacts of RFPP on antioxidant enzyme activities, NO release and MDA levels in H2O2-triggered MRC-5 cells. (A) SOD activities; (B) GSH-Px activities; (C) NO release; (D) MDA levels. (E) The cells treated with RFPP were stained with DCFH-DA to measure intracellular ROS production by flow cytometry. Values are given as the means ± SD, n = 3

Cellular antioxidant systems, particularly SOD along with GSH-Px, can improve the capacity of cells to handle H2O2-triggered oxidative damage. MDA is the end product of lipid peroxidation in vivo by free radicals, and its levels indirectly indicate the extent of free radical damage to cells (Choi et al., 2022; Marimoutou et al., 2015). Additionally, NO release can partially reflect the degree of oxidative damage to cells (Weng et al., 2017). Thus, the most common indications of oxidative damage are intracellular NO release and MDA levels. The H2O2-treated MRC-5 cells exhibited significant increases in intracellular NO release and MDA levels. However, 20–80 µM RFPP treatment effectively prevented these H2O2-mediated events. This suggests that RFPP can protect MRC-5 cells from oxidative damage by inhibiting NO release and MDA levels. Previous research established that H2O2 triggered oxidative stress in cells primarily via its attack on the antioxidant system. Antioxidant enzyme defenses are critical in protecting cells from suffering oxidative damage. Increasing evidence suggests that over-expression of SOD along with GSH-Px has cytoprotective influences in human hepatocellular carcinomas (HepG2), human umbilical vein endothelial (HUVEC), and retinal pigment epithelial (RPE) cells (Hu et al., 2019; Iloki-Assanga et al., 2015; Ma et al., 2016). These data illustrated that H2O2-triggered oxidative damage lowered SOD along with GSH-Px activity, but RFPP administration effectively restored this effect. Our findings illustrate that RFPP protected cells from oxidative damage by increasing antioxidant enzyme activity or reducing cell ROS levels in H2O2-inoculated MRC-5 cells.

Effect of RFPP on apoptosis-triggered morphological changes in H2O2-triggered MRC-5 cells

Hoechst 33258 is a benzimidazole dye that stain DNA by attaching to the minor groove of adenine, as well as thymine-rich regions. When attached to double-stranded DNA, it exhibits blue fluorescence and is used to label nuclei in cell cycle research and to differentiate nuclear morphology in dead cells. Normal cell nuclei glow diffusely and uniformly in blue, while apoptotic cells fluoresce strongly in blue (Fig. 3). Hoechst staining exhibited a substantial increase in the number of H2O2-triggered apoptotic cells in contrast with the control group, but RFPP administration decreased the amount of apoptotic cells. The normal nuclear disintegration, chromosomal condensation, and cell shrinkage (Fig. 3A). As shown in Fig. 3B, the H2O2-treated cells were reduced in size. We used Hoechst 33258 to stain the MRC-5 cell nuclei and found that RFPP protected H2O2-treated MRC-5 cells by suppressing apoptosis (Fig. 3C–F).

Fig. 3.

Fig. 3

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Effects of RFPP on the apoptotic morphological changes triggered by H2O2. (A) Control group; (B) H2O2 group; (C) 20 µM RFPP; (D) 40 µM RFPP; (E) 80 µM RFPP; (F) Analysis of statistical results of Hoechst 33258 detection

RFPP ameliorated H2O2-triggered apoptosis in MRC-5 cells

To assess apoptosis in MRC-5 cells, we explored the incidence of phosphatidylserine externalization on the cell surface via annexin V-FITC/PI staining. This staining with annexin V is often employed in combination with a vital dye, such as PI, to distinguish between the early (annexin V+, PI−) and late phases of apoptosis (annexin V+, PI+). Flow cytometry analysis exhibited a substantial reduction in the number of double-positive cells (upper right quadrant; 4.08%) (Fig. 4A) in contrast with H2O2-inoculated cells (56.2%) (Fig. 4B). The three RFPP dosages tested remarkably reduced this amount of apoptosis, ranging from 45.6 to 10.18%, in contrast with H2O2 treatment alone (Fig. 4C–E). The treatment with RFPP protected the MRC-5 cells from H2O2-triggered apoptosis (Fig. 4F).

Fig. 4.

Fig. 4

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Impacts of RFPP on the cell apoptosis of H2O2 triggered MRC-5. (A) Control group; (B) H2O2 group; (C) 20 µM RFPP; (D) 40 µM RFPP; (E) 80 µM RFPP; F Quantitative analysis of the percentage of apoptotic cells; Q1 indicates dead cells; Q2 indicates cells in end-stage apoptosis; Q3 indicates cells undergoing apoptosis; Q4 indicates viable cells not undergoing apoptosis

To establish if the growth triggered by RFPP was related with protective impacts, we measured the proportion of apoptotic cells via flow cytometry technique with dual labeling with annexin V-FITC/PI. The shift of the inner to the outside leaflet of the plasma membrane is one of the first visible alterations in cells undergoing apoptosis (Mou et al., 2019). This modification of the cell membrane is now viewed as a vital early characteristic of apoptosis (Ma et al., 2016). This translocation exposes phosphatidylserine to the external cellular environment, which may be determined via incubating cells with fluorochrome-linked phospholipid-docking proteins, for instance phycoerythrin-labeled annexin V. Annexin V-FITC is employed to assess apoptosis in its early stages. While viable cells with undamaged membranes are impermeable to PI, the membranes of dead, as well as injured cells undergoing apoptosis are permeable. Thus, PI may infiltrate the cell membrane and stain cells in the intermediate and terminal phases of apoptosis (Wang et al., 2019b). Thus, the combination of annexin V-FITC and PI permits the differentiation of cells at various phases of apoptosis. This suggests that RFPP protected the MRC-5 cells from H2O2-triggered apoptosis.

Effect of RFPP on H2O2-triggered cell cycle arrest

To further investigate the mechanism by which RFPP protects MRC-5 cells from H2O2 damage, we investigated how RFPP affects cell cycle progression. We monitored the cell cycle phase distribution of MRC-5 cells after treatment with RFPP for 24 h. In H2O2-treated cells, treatment with RFPP remarkably decreased the cells in G0/G1 phase (from 52.99 to 40.16%; p < 0.01) and concomitantly increased the cells in the S phase (from 38.41 to 42.22%) and G2/M phase (from 8.6 to 17.62%; p < 0.05; Fig. 5C–E). MRC-5 cells were increased in the S phase in a concentration-dependent manner after 24 h of exposure to RFPP compares to the H2O2-treated group (Fig. 5F). This alteration in cell cycle progression likely caused the increases in cell proliferation and viability.

Fig. 5.

Fig. 5

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Effects of RFPP on the cell cycle arrest of H2O2-triggered MRC-5 cells. (A) Control group; (B) H2O2 group; (C) 20 µM RFPP; (D) 40 µM RFPP; (E) 80 µM RFPP; (F) Statistical analysis of the percentage S phase of MRC-5 cells

Exposure to 120 µM H2O2 remarkably increased the fraction of apoptotic cells in MRC-5 cells, which was reverted concentration-dependently by RFPP. Senescence is inextricably linked to reduced cell proliferation, since senescent cells are incapable of dividing. When the cell cycle is halted, cells irreversibly lose their ability to proliferate, resulting in their senesce. Along with morphological changes, we reported that RFPP supplementation greatly boosted cell viability, revealing RFPP’s capacity to enhance cell proliferative capability. The length of cell cycle arrest dictates the rate at which proliferative ability is lost (Jalal et al., 2019). To further elucidate this impact, we used flow cytometry to assess the cell cycle distribution. We established that 120 µM H2O2 caused the G0/G1 cell cycle arrest and that RFPP treatment remarkably increased the G2/M phase. Thus, by interfering with the cell cycle, giving RFPP to H2O2-treated cells may enhance cell proliferation.

Effects of RFPP on relative protein contents of PI3K/AKT/mTOR and Bcl-2/Bax/Caspase-3 in H2O2-triggered MRC-5 cells

Because RFPP appears to protect MRC-5 cells from oxidative damage, we conducted further experiments to determine the potential molecular mechanisms of RFPP. H2O2-stimulated MRC-5 cells showed markedly decreased relative protein contents of p-Akt, Bcl-2 and p-mTOR in contrast with those of the control cells (p < 0.05; Fig. 6A and B). Conversely, MRC-5 cells treated with RFPP (20–80 µM) showed enhanced relative protein expressions of p-Akt/AKT, Bcl-2/Bax p-mTOR/mTOR (Fig. 6D–F) and reduced relative protein expression of Caspase-3 (Fig. 6G). Therefore, RFPP protected MRC-5 cells against oxidative damage by activating the PI3K/AKT/mTOR and Bcl-2/Bax/Caspase-3 pathway.

Fig. 6.

Fig. 6

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Effect of RFPP on the contents PI3K/AKT/mTOR and Bcl-2/Bax/Caspase-3 signaling cascade-linked proteins in H2O2-treated MRC-5 cells. (A) PI3K, AKT, p-AKT, mTOR and p-mTOR expressions were measured by western blot; (B) Bcl-2, Bax and Caspase-3 expressions were measured by western blot; (C) PI3K; (D) p-AKT/AKT; (E) p-mTOR/mTOR; (F) Bcl-2/Bax; (G) Caspase-3

Previous investigations showed that the PI3K/Akt cascade positively activated Akt and mTOR in cells (Xu et al., 2020b). Activated p-Akt inhibits ROS production; promotes proliferation, migration and angiogenesis; and inhibits proapoptotic Bcl-2 family members, for instance Bad, Bax and Caspase-3 (Jiang et al., 2020). Additionally, activated Akt helps promote mTOR phosphorylation, thus stimulating endothelial cells to attenuate NO release levels (Chu and Zhang, 2018). These data illustrate that the PI3K/Akt/mTOR and Bcl-2/Bax/Caspase-3 signaling cascades mediate sequences of beneficial physiological activities. We established that H2O2 remarkably dampened phosphorylation of Akt and mTOR, while interventions with different RFPP concentrations remarkably dose-dependently upregulated p-Akt, Bcl-2, p-mTOR expressions and downregulated Bax and Caspase-3 expressions. Thus, RFPP successfully reversed H2O2-triggered cell injury via activating the PI3K/Akt/mTOR and Bcl-2/Bax/Caspase-3 cascade.

Acknowledgements

This study was supported by Ningbo Major Public Welfare Science and Technology Project (2019C10005), Zhejiang Wanli University Scientific Research and Innivation Team (No. SC1032110880210), Zhejiang Provincial Top Discipline of Biological Engineering (No. KF2022007).

Declarations

Conflict of interest

The authors declare that they have no conflict of interest.

Footnotes

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Contributor Information

Haomiao Ding, Email: 18368406641@163.com.

Yuzhe Zhang, Email: 1119739465@qq.com.

Yue Zha, Email: 1633942053@qq.com.

Sijia Zhou, Email: 3237189220@qq.com.

Chaoqing Cao, Email: caochaoqing@zwu.edu.cn.

Huajian Zhu, Email: 1179640717@qq.com.

Feng Xu, Email: 1621510719@qq.com.

Xiuqiang Zhang, Email: 1601091039@nbu.edu.cn.

Caisheng Wang, Email: wangcaisheng@zwu.edu.com.

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