All chemicals and solvents used were of the highest analytical grade available. Cell culture supplies and media, including fetal bovine serum (FBS), phosphate-buffered saline (PBS), and penicillin-streptomycin were purchased from Thermo Fisher Scientific, Inc. (Waltham, MA, USA). The Cell Counting Kit-8 (CCK-8; cat. no. CK04-11) was from Dojindo Molecular Laboratories, Inc. (Kumamoto, Japan). Protease inhibitor cocktail (cat. no. P8340) and D-GalN (cat. no. G0500) were purchased from Sigma-Aldrich; EMD Millipore (Billerica, MA, USA). Anti-proliferating cell nuclear antigen (PCNA; cat. no. ab15497), anti-Tomm20 (cat. no. ab56783) and anti-Lamin B1 (cat. no. ab16048) antibodies were purchased from Abcam (Cambridge, UK). Anti-poly (ADP) ribose polymerase (PARP; cat. no. 9532S) antibody was purchased from Cell Signaling Technology, Inc. (Beverly, MA, USA). Anti-B-cell lymphoma 2 (BCL2; cat. no. SC-7382), anti-Bcl-2-associated X protein (BAX; cat. no. SC-526), anti-DRAM (cat. no. SC-81713), anti-SOD1 (cat. no. SC-17767), anti-SOD2 (cat. no. SC-130345), anti-gluta-thione peroxidase (GPx; cat. no. SC-133160), anti-Catalase (cat. no. SC-271358), anti-Keap1 (cat. no. SC-365626), anti-HO-1 (cat. no. SC-136960), anti-p53 (cat. no. SC-126), anti-GAPDH (cat. no. SC-20357), and anti-Nrf2 (cat. no. SC-81342) antibodies were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA, USA). Anti-microtubule-associated protein 1A/1B-light chain 3 (LC3)I/II (cat. no. PM036), anti-phospho-rlyated-p62 (cat. no. PM074), and anti-p62 (cat. no. PM045) antibodies were purchased from MBL International Corporation (Woburn, MA, USA). hPH (Laennec inj.) was manufactured by Green Cross WellBeing Corporation (Seongnam, Korea) through the hydrolysis of human placenta with HCl and pepsin.

Sprague-Dawley rats (SD) rats (male, 6–8 weeks old, weighing ~200–250 g each) were purchased from Raonbio, Inc. (Yongin, Korea). These rats were provided with adequate food and water ad libitum and were housed in clean cages for 1 week. The laboratory temperature was 24±1°C and relative humidity was 40–80%. All animal experiments were performed according to the Guide for the Care and Use of Laboratory Animals as published by the US National Institutes of Health. The present study was reviewed and approved by the Animal Welfare and Research Ethics Committee at Chung-Ang University (Seoul, Korea; 2017-00003). Acute liver injury was induced by intraperitoneal injection of LPS (15 µg/kg) together with D-GalN (700 mg/kg) dissolved in normal saline, which can increase the sensitivity of hepatocytes. Blood was collected from the inferior vena cava 24 h following injection of D-GalN/LPS. The SD rats were then dissected, and liver tissues were removed immediately for histological detection. Normal PBS was used in control rats. hPH (1.2, 2.4, and 3.6 ml/kg) was injected subcutaneously into each mouse 24, 48, and 72 h prior to D-GalN/LPS injection. As a negative control, only D-GalN/LPS was injected.

The collected blood samples were stored overnight at 4°C. The serum was isolated the subsequent day following centrifugation at 15,928 × g for 10 min at 4°C. The ALT and AST were detected using a Hitachi 7600 Series automatic biochemical analyzer (Hitachi, Ltd., Tokyo, Japan).

Based on a previous study, blood was collected for measuring TNF-α (cat. no. 438207) and IL-6 (cat. no. 437107) at 24 h post-D-GalN/LPS injection. The serum was separated by centrifugation at 15,928 × g at 4°C for 10 min. The cytokines were measured using mouse ELISA kits (BioLegend, Inc., San Diego, CA, USA) according to the manufacturer's protocol.

The liver tissues were immersed in normal 10% neutral buffered formalin and fixed for 48 h, dehydrated in a series of graded ethanol, embedded in paraffin wax, and cut into 5-µm sections. The paraffin-embedded sections were stained with hematoxylin and eosin (H&E) for pathological analysis under a light microscope. Histological changes were evaluated using a point-counting method for the severity of hepatic injury using an ordinal scale, as previously described (27). Briefly, the H&E-stained sections were evaluated at ×400 magnification using the point-counting method for the severity of hepatic injury with an ordinal scale as follows: grade 0, minimal or no evidence of injury; grade 1, mild injury consisting of cytoplasmic vacuolation and focal nuclear pyknosis; grade 2, moderate to severe injury with extensive nuclear pyknosis, cytoplasmic hypereosinophilia, and loss of intercellular borders; and grade 3, severe necrosis with disintegration of hepatic cords, hemorrhage, and neutrophil infiltration.

TUNEL was performed to analyze DNA fragmentation indicative of cellular apoptosis using the in situ cell death detection kit (cat. no. ab206386, Abcam), according to the manufacturer's protocol. The paraffin wax-embedded tissue sections were treated with proteinase K, and endogenous peroxidase activity was blocked with hydrogen peroxide. The sections were incubated at 37°C with the terminal TdT nucleotide mixture for 1 h. The reactions were then terminated, and the slides were rinsed with PBS. Nuclear labeling was developed with horseradish peroxidase and diaminobenzidine. The number of apoptotic cells in each slide from at least five different fields was analyzed using Image-Pro Plus software (version 6.0 for Windows; Media Cybernetics, Inc., Rockville, MD, USA) with the assistance of a microscope (DM750; Leica Microsystems GmbH, Wetzlar, Germany). The number of positive cells was averaged for statistical analysis.

The paraffin sections were de-paraffinized and rehydrated. Endogenous peroxidase was inactivated by incubation in 0.3% hydrogen peroxide in absolute methanol for 30 min. The sections were incubated in 5% skim milk for 30 min at room temperature. Antigen retrieval was performed by microwave (700 W) treatment in 10 mM citrate buffer (pH 6.0) for 15 min. The sections were incubated overnight at 4°C with anti-PCNA primary antibody (Abcam) at a dilution of 1:500. Following washing with PBS, the sections were incubated at room temperature for 30 min in secondary antibody (Goat Anti-Rabbit; cat. no. ab205718; Abcam; 1:1,000). A brown color was developed with 3 diami-nobenzidine for 2–4 min, and the sections were then washed in distilled water. The number of positively stained brown nuclei (denoting PCNA) in each slide from at least five different fields was analyzed using Image-Pro Plus software (version 6.0 for Windows) with the assistance of a microscope (DM750, Leica Microsystems GmbH). The number of positive cells was averaged for statistical analysis.

Human HepG2 cells, the HepG2 (ATCC HB 8065) cells were purchased from the Korea Cell Line Bank (Seoul, Korea), and were cultured in Dulbecco's modified Eagle's medium (DMEM) with 10% (v/v) heat-inactivated FBS and 1% penicillin and incubated in a humidified incubator with 5% CO₂, 95% air at 37°C. A dose of 5% hPH was used in the time-course investigation of the induction of antioxidant enzymes by hPH. To investigate the protective effects of hPH and its active mechanism on HepG2 cells challenged by D-GaIN (Sigma; EMD Millipore), the cells were pretreated with hPH for 2 h, stimulated with 50 mM D-GaIN at 37°C, and harvested for the indicated experiments.

The HepG2 cells, treated as described above, were further treated with 50 mM D-GalN and incubated for another 24 h, following which the cell viability was determined using CCK-8 (Dojindo Molecular Technologies, Inc.). To determine cell viability, the HepG2 cells were seeded in 96-well plates at a density of 1×10⁴ cells/ml and grown for 24 h in a 37°C incubator. When the cells attained 70–80% confluence, they were either left untreated (control group) or treated with D-GalN (50 mM), 5% hPH, or 5% hPH + D-GalN (50 mM). Following incubation for 24 h, cell morphology changes were observed under Olympus inverted microscopes (Olympus Corporation, Tokyo, Japan). The supernatants were discarded, following which 10% CCK-8 solution in fresh DMEM (100 µl) were added to each well. The cells were then incubated at 37°C for 1 h, and the absorbance was measured at 450 nm directly in the wells using a SpectraMax i3x Multi-Mode detection platform (Molecular Devices LLC, Sunnyvale, CA, USA).

The LDH in the culture medium was assessed with 0.2 mM NADH and 0.4 mM pyruvic acid in up to 200 µl PBS at pH 7.4. The LDH concentration in the sample was proportional to the rate of NADH oxidation measured by absorbance at 334 nm (OD/min) using the SpectraMax i3x Multi-Mode detection platform. The LDH concentration in the culture media was calculated using a commercial standard LDH assay kit purchased from Abnova (cat. no. KA0785, Taipei, Taiwan).

Morphological changes in the apoptotic cells were assessed by fluorescent microscopy following DAPI staining. Briefly, the HepG2 cells were seeded at a density of 1×10⁵ cells/ml in 6-well plates and grown for 24 h in a 37°C incubator. Following washing once with PBS, the cells were either left untreated (control group) or treated with D-GalN (50 mM), 5% hPH, or 5% hPH + D-GalN (50 mM). Following incubation for 24 h, the cells were fixed in 4% formaldehyde for 1 h, and permeabilized with 0.1% Triton X-100 (Biosesang, Inc., Seongnam, Korea) for 5 min. Subsequently, DNA-specific fluorochrome DAPI (Invitrogen; Thermo Fisher Scientific, Inc.) was applied to each well, following which samples were incubated for 10 min in the dark at room temperature. Finally, the cells were washed three times with PBS and examined using a confocal microscope (Carl Zeiss AG, Oberkochen, Germany).

For apoptotic cell analysis, the HepG2 (1×10⁴/well) cultures in 96-well black plates were exposed to the indicated treatments for 12 h. The assay was performed according to the manufacturer's protocol (BD Biosciences, San Jose, CA, USA). Briefly, the cells were washed twice with PBS and again with binding buffer. Annexin V-fluorescein isothiocyanate and PI were added to each sample, and the mixture was incubated at room temperature for 15 min. The absorbance was read directly in the wells at 494/518 nm for FITC Annexin V and 535/617 nm for PI using the SpectraMax i3x Multi-Mode detection platform. The populations staining positive for Annexin V and Pl were compared with the untreated group.

The HepG2 cells were grown to ~70% confluence; 1×10⁵ cells were collected in a Petri dish and either left untreated (control group) or treated with D-GalN (50 mM), 5% hPH, or 5% hPH + D-GalN (50 mM); and then incubated for 24 h. The cells were then washed with PBS and stained with 10 µg/ml PI for 5 min, and fluorescent images were observed under a DP70 fluorescence microscope with DP Controller software (Olympus Corporation; ver. 3.2.276.2).

Cytosolic and mitochondrial ROS were measured on a 96-well plate reader as previously described (28). Briefly, the HepG2 cells were seeded at a concentration of 1×10⁴ cells per well in black 96-well flat bottom plates (Thermo Fisher Scientific, Inc.) and allowed to adhere overnight. Following seeding, the cells were washed and incubated with serum-starved medium, D-GalN (50 nM), 5% hPH, or 5% hPH + D-GalN (50 mM) for 4 h, followed by incubation with 2′,7′-dichlorofluorescin diacetate (DCFH-DA; 10 µM, ex/em: 518/605 nm; Invitrogen; Thermo Fisher Scientific, Inc.) or MitoSOX™ (5 µM, ex/em: 510/580 nm; Invitrogen; Thermo Fisher Scientific, Inc.) for 20 min. The oxidative products were measured with the SpectraMax i3x Multi-Mode detection platform.

The ΔΨm of intact cells was measured as previously described, (29) with modifications. Briefly, the cells were washed with PBS and TMRE (200 nM, ex/em: 549/582 nm; Invitrogen; Thermo Fisher Scientific, Inc.) was added to the cell suspension. The cells were incubated at 37°C for 30 min in the dark. The ΔΨm was measured using the SpectraMax i3x Multi-Mode detection platform. The percentage and mean fluorescence intensity level of the mass were calculated for each sample.

Mitochondrial mass was measured according to fluorescence levels following staining with MitoTracker Green FM (100 nM, ex/em: 490/525 nm; Invitrogen; Thermo Fisher Scientific, Inc.) at 37°C for 30 min. Subsequently, the cells were washed once in PBS and promptly evaluated using the SpectraMax i3x Multi-Mode detection platform. The percentage and mean fluorescence intensity level of the mass were calculated for each sample.

The cells (1×10⁶ cells/well) were prepared on sterilized glass coverslips (BD Biosciences) in triplicate and then fixed in 4% paraformaldehyde in PBS for 10 min, permeabilized with 0.25% Triton X-100 in PBS for 10 min and incubated with primary antibodies against Tomm20 (Abcam; 1:500), Nrf2 (Santa Cruz Biotechnology, Inc.; 1:500), and LC3 (Abcam; 1:500) for 12 h at 4°C incubator. The cells were washed to remove excess primary antibodies and incubated with the appropriate fluorescently labeled secondary antibodies (1:1,000) for 1 h at room temperature. Following mounting of slides, fluorescence images were acquired using a confocal microscope (LSM700, Carl Zeiss AG).

Western blot analysis

The cells were lysed in cell lysis buffer [62.5 mM Tris-HCl (pH 6.8), 2% sodium dodecyl sulfate (SDS), 5% β-mercaptoethanol, 2 mM phenylmethylsulfonyl fluoride, protease inhibitors (Complete™; Roche Diagnostics GmbH, Mannheim, Germany), 1 mM Na₃VO₄, 50 mM NaF and 10 mM EDTA]. The protein content of the lysates was determined using BCA Protein Assay reagent (cat. no. 23225; Pierce; Thermo Fisher Scientific, Inc.). A 20 µg sample of protein per lane was separated by 12% SDS-polyacrylamide gel electrophoresis and blotted onto polyvinylidene fluoride membranes (EMD Millipore) which were then saturated with 5% powdered milk in Tris-buffered saline containing 0.5% Tween-20. The blots were then incubated with the appropriate primary antibodies at a dilution of 1:1,000 or 1:2,000 for 12 h in a 4°C incubator, and further incubated with horseradish peroxidase-conjugated secondary antibody (1:10,000) for 2 h at 37°C. The bound antibodies were detected using enhanced chemiluminescence (Amersham; GE Healthcare Life Sciences, Chalfont, UK). Images of the blotted membranes were captured using the LAS-1000 lumino-image analyzer (Fujifilm, Tokyo, Japan). The protein levels were compared to those of the appropriate loading control (GAPDH or non-phosphorylated proteins).

Total RNA was extracted from the dorsal skin using TRIzol (Invitrogen; Thermo Fisher Scientific, Inc.). First-strand cDNA synthesis from the total RNA template was performed with PrimeScript RT master mix (Takara Bio, Inc., Tokyo, Japan). The resulting cDNA was subjected to real-time PCR (complementary cDNA, 2 µl; primer, 3 µl; 2X premix SYBR, 5 µl), using qPCR 2X PreMIX SYBR (Enzynomics, Seoul, Korea) and a CFX-96 thermocycler (Bio-Rad Laboratories, Inc., Hercules, CA, USA). The PCR conditions used to amplify all genes were as follows: 10 min at 95°C and 40 cycles of 95°C for 10 sec, 60°C for 15 sec, and 72°C for 20 sec. The expression data were calculated from the quantification cycle (Cq) value using the ΔCq method of quantification (30). GAPDH was used for normalization. The oligonucleotides that were used for qPCR were as follows: Human ATG8, forward 5′-CGC ACC TTC GAA CAA AGA GT-3′ and reverse, 5′-GAC CAT GCT GTG TCC GTT C-3′; human beclin 1 (BECN1), forward 5′-AAC CTC AGC CGA AGA CTG AA-3′ and reverse, 5′-CCT CTA GTG CCA GCT CCT TT-3′; human cathepsin D (CTSD), forward 5′-CAA GTT CGA TGG CAT CCT GG-3′ and reverse 5′-CGG GTG ACA TTC AGG TAG GA-3′; human lysosomal-associated membrane protein 1 (LAMP1), forward 5′-CTT TCA AGG TGG AAG GTGz GC-3′ and reverse 5′-GAT AGT CTG GTA GCC TGC GT-3′; human activating transcription factor (ATF)6, forward 5′-GTG TCA GAG AAC CAG AGG CT-3′ and reverse 5′-GGT GCC TCC TTT GAT TTG CA-3′; human ATF4, forward 5′-ACA CTG CTT ACG TTG CCA TG-3′ and reverse 5′-AGA CCC ACA GAG AAC ACC TG-3′; human C/EBP homologous protein (CHOP), forward 5′-CAT TGC CTT TCT CCT TCG GGG-3′ and reverse 5′-CCA GAG AAG CAG GGT CAA GA-3′.

All quantitative data are presented as the mean ± standard error of the mean for three independent experiments. Statistical analyses were performed using the statistical package for SPSS software version 17.0 (IBM Corp., Armonk, NY, USA). Differences between two groups were evaluated using a paired Student's t-test. For multiple comparisons, one-way analysis of variance was used followed by Tukey's multiple comparisons test. P<0.05 was considered to indicate a statistically significant difference.

Acute liver injury leads to high mortality rates in rats. Therefore, the present study assessed the effects of hPH on D-GalN (700 mg/gg)/LPS (15 µg/kg)-induced mortality. To investigate whether hPH can protect against liver injury in rats due to D-GalN/LPS, the survival rate of D-GalN/LPS + hPH-treated rats was monitored for 24 h (n=12). All saline-injected rats survived as the normal control group (n=12). No significant differences in gross liver examination, body weight, or liver weight were found between groups (Fig. 1A-C). As shown in Fig. 1D, a number of rats died 7 h following D-GalN/LPS injection, and the survival rate was 33.3% at 24 h. However, it was found that hPH pretreatment effectively increased the survival rate of rats with liver injury induced by D-GalN/LPS. Pretreatment with hPH (1.2 ml/kg) increased the survival rate to 83.3%, and all hPH (2.4 and 3.6 ml/kg)-treated rats survived. These results suggested that pretreatment with hPH effectively protected against D-GalN/LPS-induced mortality.

The serum levels of AST, ALT, LDH, and inflammatory cytokines are well-established markers of hepatic injury (31). In order to investigate the effects of hPH on D-GalN/LPS-induced liver damage, hematological tests were performed. Serum (n=4–6) was collected to detect AST, ALT, LDH, IL-6, and TNF-α. As shown in Fig. 1E-G, LPS/D-GalN markedly increased serum levels of AST, ALT, and LDH, whereas hPH reduced these elevations. As hepatocyte injury resulting from acute liver injury is accompanied by an elevation in inflammatory cyto-kines (32), the present study examined the effect of hPH on hepatotoxicity by measuring serum levels of IL-6 and TNF-α (Fig. 1H and I). The D-GalN/LPS-treated rats exhibited significant increases in levels of IL-6 and TNF-α compared with normal rats. In addition, hPH pretreatment significantly inhibited the release of these pro-inflammatory cytokines compared with the LPS/D-GalN group. These results suggested that hPH effectively protected against LPS/D-GalN-induced liver degeneration and inflammatory responses.

The present study also investigated histological changes following liver injury to confirm the protective effects of hPH. The rats treated with D-GalN/LPS showed severe histopathological degeneration evidenced by severe intrahepatic hemorrhaging, apoptosis, necrosis, ballooned hepatocytes, and hepatocytes expanded by fat vacuoles, whereas hPH (1.2, 2.4 and 3.6 ml/kg) treatment ameliorated these changes (Fig. 2A). It has been suggested that D-galN/LPS-induced imbalance between apoptosis and proliferation in hepatocytes is responsible for the impairment of liver function. Therefore, hPH-mediated modulations of proliferation (PCNA) and apoptosis (TUNEL) were also evaluated by immunohistochemical analysis (Fig. 2B and C). A high expression of PCNA and low numbers of TUNEL-positive cells were observed in the control group. However, D-GalN/LPS markedly reduced PCNA immunore-activity and significantly increased the number of apoptotic cells. By contrast, pre-administration of hPH resulted in an increase of PCNA-positive cells and reduced the number of apoptotic cells in all three dose groups. Graphs showing results of the staining experiments are shown in Fig. 2D-F. These observations indicated that hPH pretreatment ameliorated hepatic injury by reducing D-GalN/LPS-induced hepatocyte damage in rats.

Hepatocyte apoptosis is important in the pathogenesis of liver disease, including ALF (33). D-GalN induces hepatocyte cell death in vivo (34) and in vitro (35); it offers a suitable experimental model based on its capacity to reduce the intracellular pool of uridine monophosphate in hepatocytes, inhibiting the synthesis of RNA and proteins, and leading to cell apoptosis. Therefore, the present study investigated whether D-GalN-induced hepatotoxicity occurred in HepG2 cells. Following D-GalN stimulation, cell viability decreased markedly in a dose-dependent manner. The results revealed a reduction of 50% at 50 mM D-GalN and of 82% at 100 mM D-GalN in the HepG2 cells (Fig. 3A). The cytotoxic effects of hPH on HepG2 cells have not been examined previously. The present study used a CCK-8 assay to evaluate the dose-dependent cytotoxic effects of hPH on HepG2 cells. The results showed that stimulation with hPH had no significant effect on cell viability at various concentrations (1.25–5% in serum-free DMEM) over 24 h. Therefore, hPH was used at a concentration of 5% in subsequent experiments (Fig. 3B). In addition to the D-GalN-induced decrease in cell viability being attenuated by pretreatment with hPH, the concentration of LDH released from the hepatocytes was also reduced in the D-GalN + 5% hPH-stimulated cells, compared with the D-GalN only-treated cells (Fig. 3C and D). These results indicated that hPH protected against D-GalN-induced hepato-toxicity in HepG2 cells. Changes in cellular morphology and cell viability were also evaluated to examine the protective effects of hPH in D-GalN-stimulated HepG2 cells. As shown in Fig. 3E, the D-GalN-stimulated HepG2 cells exhibited a marked reduction in the number of hepatocytes. However, hPH pretreatment effectively improved the D-GalN-mediated cell damages.

To examine the inhibitory mechanisms of hPH in D-GalN-induced cytotoxicity, the HepG2 cells were assessed by Annexin V and PI staining, followed by microplate analysis (Fig. 3F and G) or fluorescence microscopy (Fig. 3H). D-GalN stimulation alone facilitated the induction of apoptosis, including levels of Annexin V⁺ (841%)/PI⁺ (948%), compared with the controls. By contrast, pretreatment with hPH significantly inhibited D-GalN-induced apoptosis (Annexin V⁺, 801%; PI⁺, 280%). D-GalN-induced DNA fragmentation was also effectively attenuated by hPH pretreatment (Fig. 3H). The present study further examined whether hPH regulated PARP-dependent cell death. D-GalN induced an increase of Bax (pro-apoptotic protein) and a decrease of Bcl-2 (anti-apoptotic protein) (Fig. 3I and J). Furthermore, the cleavage of PARP was inhibited in the HepG2 cells by pretreatment with hPH (Fig. 3I and J). These findings suggested that hPH treatment significantly improved hepatocyte degeneration through the attenuation of apoptosis in HepG2 cells.

The rapid generation of ROS is associated with hyperpolarization of the mitochondrial membrane potential and apoptosis in D-GalN-treated hepa-tocytes (36). In addition, D-GalN-dependent cell necrosis is associated with mitochondrial membrane depolarization (37). To investigate whether the D-GalN-mediated increase in abnormal mitochondria was rescued by hPH treatment, the present study first evaluated mitochondrial morphology using Tomm20 (mitochondrial outer membrane) staining (Fig. 4A). Mitochondrial morphology was altered at 24 h in the D-GalN-stimulated HepG2 cells. However, the increased mitochondrial damage was attenuated significantly by hPH. In addition, mitochondrial mass decreased, by ~55%, in the D-GalN-stimulated HepG2 cells, and this was rescued by hPH pretreatment (Fig. 4B). ΔΨm was then evaluated using microplate analysis. The D-GalN-stimulated HepG2 cells exhibited decreased ΔΨm; however, this decrease was signifi-cantly attenuated in the hPH-pretreated cells and unstimulated cells (Fig. 4C). These findings suggested that hPH improved D-GalN-mediated mitochondrial dysfunction by improving ΔΨm and inhibiting quantitative mitochondria loss.

Subsequently, the present study determined whether hPH was involved in the negative regulation of oxidative stress. The HepG2 cells were stimulated with D-GalN in the presence or absence of hPH, and ROS generation was determined using DCFH-DA or MitoSOX (mitochondrial-specific superoxide indicator) (38) as a probe for microplate analyses (Fig. 4D and E). D-GalN alone led to the potent intracellular generation of ROS, whereas the D-GalN-induced generation of ROS in HepG2 cells was attenuated significantly by hPH pretreatment (Fig. 4D). Mitochondrial ROS increased significantly following treatment with D-GalN (Fig. 4E), whereas D-GalN-induced mitochondrial ROS generation was decreased by pretreatment with hPH. These results suggested that hPH protected against D-GalN-mediated apoptotic cell death by regulating the generation of cytosolic and mitochondrial ROS.

As the induction of antioxidant enzymes is a universal stress response against liver degeneration and has been widely shown to have an anti-apoptotic effect, the expression levels of antioxidative enzymes SOD1, SOD2, GPx, and catalase were detected by western blot analysis (Fig. 5A). hPH induced antioxidative enzyme expression in the HepG2 cells. These enzymes were elevated 1 h following hPH treatment and were maintained at this level until 8 h. Furthermore, to investigate the antioxidant mechanisms of hPH, Nrf2 pathway-related gene expression on HepG2 cells was evaluated (Fig. 5B-D). The induction of Nrf2 upregulates antioxidant proteins, including SOD, catalase, and HO-1, reducing ROS (15). hPH treatment resulted in a significant decrease of Keap1 at 4–8 h. However, hPH increased the protein levels of cytoplasmic p-p62 and HO-1. Additionally, p62 was upregulated 4–8 h following hpH treatment (Fig. 5B). Furthermore, the expression of Nrf2 was decreased in the cytosol and increased in the nucleus (active Nrf2) following hPH treatment, indicating that translocation to the nucleus was affected by hPH treatment (Fig. 5C and D). Together, these findings indicated that the induction of anti-oxidative enzymes by hPH treatment was modulated by the Keap1-p62-Nrf2 pathway.

Autophagy, mitochondria, and oxidative stress are closely intertwined in redox signalling (39). Additionally, D-GalN-induced cell damage regulates the autophagy process in early stages. To investigate whether the regulation of autophagy is influenced by D-GalN stimulation and the effects of hPH, the present study measured the level of autophagy regulation by staining cells with anti-LC3 I/II antibody (Fig. 6A). LC3 I/II is an established autophagy marker due to its involvement in autophagosome membrane formation (40). In the present study, compared with the cells treated with vehicle control or D-GalN alone, the cells treated with hPH exhibited signifi-cantly fewer LC3 puncta per cell (Fig. 6A). The changes in autophagy-related genes were then quantified through assessment of RNA and protein levels in the hPH-treated HepG2 cells. As shown in Fig. 6B and C, hPH treatment induced a decrease in the conversion of LC3-I to LC3-II (lipidated form), the latter of which is a reliable marker of autopha-gosome generation. Furthermore, the expression levels of DRAM, CHOP, and p53 (proteins associated with autophagic cell death) indicated that a minimized autophagy process was induced in the hPH-treated cells (Fig. 6B) (20,41,42). Similarly, the transcription levels of autophagy-related genes (ATG8, CTSD, BECN1, and LAMP1) and of ER stress-related genes (ATF4, ATF6, and CHOP) were lower in the hPH-treated cells compared with those in cells treated with D-GalN only (Fig. 6C). These results suggested that hPH treatment minimized the escalation of D-GalN-induced autophagy processes in HepG2 cells.