The AML-12 cells (cat. no. CL-0602) were provided by Wuhan Pricella Biotechnology Co., Ltd. (Elabscience Bionovation Inc.). Assay kits for alanine aminotransferase (ALT; cat. no. C009-2-1), aspartate aminotransferase (AST; cat. no. C010-2-1), GSH (cat. no. A006-2-1), malondialdehyde (MDA; cat. no. A003-1-2) and superoxide dismutase (SOD; cat. no. A001-3-2) were obtained from Nanjing Jiancheng Institute of Bioengineering. The anti-p38γ (cat. no. 20184-1-AP), anti-IL-6 (cat. no. 21865-1-AP), anti-IL-1β (cat. no. 16806-1-AP), anti-albumin (cat. no. 16475-1-AP), Akt (cat. no. 10176-2-AP), anti-phosphorylated (p-)Akt (cat. no. 66444-1-Ig), anti-TNF-α (cat. no. 17590-1-AP), anti-peroxisome proliferator-activated receptor (PPAR)-α (cat. no. 66826-1-Ig), anti-sterol regulatory element-binding protein (SREBP)-1 (cat. no. 14088-1-AP) and anti-Fasn (cat. no. 10624-2-AP) antibodies were purchased from Proteintech Group, Inc. Anti-NADPH oxidase 4 (NOX4) (cat. no. A11274) and anti-inducible nitric oxide synthase (iNOS) (cat. no. A14031) were purchased from ABclonal Biotech Co., Ltd. Anti-β-actin (cat. no. AF7018), anti-PI3K (cat. no. AF3241) and anti-p-PI3K (cat. no. AF6241) were purchased from Affinity Biosciences, Ltd. ELISA kits for IL-1β (cat. no. MLB00C-1), IL-6 (cat. no. M6000B-1) and TNF-α (cat. no. MTA00B-1) were purchased R&D Systems, Inc. The detection of IL-1β, IL-6 and TNF-α was carried out in accordance with the manufacturer's instructions.

All animal care and experimental protocols were approved by the Anhui Medical University Experimental Animal Ethics Committee (approval no. LLSC20241364). The protocols for the use and care of medical laboratory animals were followed in all operations. For the animal experiments, 8-week-old male C57BL/6J mice weighing 20-22 g were obtained from Jiangsu GemPharmatech Co., Ltd. Mice were kept in a specific pathogen-free facility at the Anhui Medical University Experimental Animal Center. The number of animals used in each group was 6, with a total of 24 C57BL/6J mice used in the experiment. All 24 mice were euthanized at the scheduled time point and no mice were found dead during the experiment. The housing conditions were set as follows: The breeding room temperature was maintained at 22±2°C, the relative humidity was 50±10% and the light/dark cycle was 12/12 h. Mice had free access to standard laboratory chow and sterile drinking water throughout the experiment.

Before the animal model was established, mice were allowed to acclimatize for 1 week, during which they were fed an adaptable diet and had access to sufficient quantities of water. Following randomization, the animals were weighed and split into four groups: Normal, APAP, APAP + AAV9-Empty and APAP + AAV9-shRNA-p38γ. For the intervention groups, mice were slowly injected via the tail vein with either AAV9-Empty or AAV9-shRNA-p38γ. The AAV9-shRNA-p38γ used in the present study was provided by Hanbio Biotechnology Co., Ltd. The sequences were as follows: 5'-GGCCAAGAACUACAUGGAATT-3' and 5'-UUCCAUGUAGUUCUUGGCCTT-3'. An intraperitoneal dose of APAP (250 mg/kg; Beijing Solarbio Science & Technology Co., Ltd.) was used to induce liver injury (39,40) 14 days after the AAV injection. Liver injury was induced in the APAP, APAP + AAV9-Empty and APAP + AAV9-shRNA-p38γ groups via an intraperitoneal injection of APAP. At 24 h after APAP injection, blood was collected via retro-orbital bleeding using sterile heparinized microhematocrit capillary tubes in the SPF animal room. Mice were anesthetized with isoflurane inhalation (induction 3-4%, maintenance 1.5-2%) prior to collection. In total, 0.5-1.0 ml of blood was obtained from each mouse and transferred into plain serum separation tubes. The samples were allowed to clot at room temperature for 30 min, and serum was separated by centrifugation at 2,000 × g for 10 min at 4°C. Serum was stored at −80°C until analysis. The mice were euthanized 24 h after APAP administration using CO₂ at 30-70% vol/min displacement in compliance with the AVMA guidelines: 2020 Edition (41). The absence of a heartbeat and respiratory movement, along with a lack of response to physical stimulation, were used to confirm animal death. The health and behavior of the mice were monitored twice a day during the acclimatization and experimental periods, including the observation of mental state, activity, food intake, water intake and hair condition. The humane endpoints followed in the present study included: Severe weight loss (>20% of the body weight), inability to eat or drink independently, severe lethargy, abnormal posture, difficulty in breathing and severe liver injury-related symptoms. No animals reached these humane endpoints during the experiment and all mice were euthanized as scheduled.

The livers were collected and divided into tiny pieces, then fixed in 4% paraformaldehyde at room temperature for 48 h for histology. The remainder of the tissue was frozen in liquid nitrogen and kept at −80°C for further examination. To minimize bias, group assignment was performed randomly using a computer-generated random number sequence. Furthermore, while the investigators administering the treatments were necessarily aware of group identities, all subsequent data acquisition and analysis were performed under blinded conditions. The personnel conducting histopathological scoring, biochemical assays, western blot quantification and statistical analysis were unaware of the group codes until after all data collection was complete.

The AML-12 cells were cultured in DMEM/F-12 media (Gibco; Thermo Fisher Scientific, Inc.; cat. no. 11320033) supplemented with 10% fetal bovine serum (Gibco; Thermo Fisher Scientific, Inc.; cat. no. 10099141), 1% penicillin-streptomycin solution (10,000 units/ml penicillin and 10,000 μg/ml streptomycin; Gibco; Thermo Fisher Scientific, Inc.). The cells were maintained in a 37°C incubator with 5% CO₂. Cells were treated with 10 mM APAP for 24 h to establish an in vitro model of liver injury (42,43). Cells between passages 5 and 15 were used for all experiments. For treatment, cells were seeded and allowed to reach 70-80% confluency before the addition of APAP or transfection.

For IHC, 4 μm sections were cut from the paraffin-fixed liver tissues. Following deparaffinization and rehydration, heat-induced epitope retrieval was performed by incubating the slides in sodium citrate buffer (10 mM, pH 6.0) at 95°C for 20 min. The sections were then allowed to cool to room temperature. After washing with PBS, the sections were incubated with 3% hydrogen peroxide for 10 min at room temperature to quench endogenous peroxidase activity, washed three times with PBS for 5 min each, then blocked with 2% BSA (Sigma-Aldrich; Merck KGaA; cat. no. A8806) for 30 min at room temperature. Subsequently, after overnight incubation at 4°C with anti-TNF-α, anti-IL-1β or anti-IL-6 (all 1:150), the sections were washed three times with PBS for 5 min each and then incubated for 1 h at room temperature with a biotinylated IgG secondary antibody (Proteintech Group, Inc.; cat. no. 16795-1-AP; 1:200). Next, the sections were washed with PBS, then incubated with Vectastain ABC reagent for 1 h at room temperature and diaminobenzidine (Shanghai Aladdin Biochemical Technology Co., Ltd.) substrate solution for 5 min at room temperature. Finally, the slides were counterstained with hematoxylin at room temperature for 3 min and examined under a light microscope.

The procedure for immunofluorescence microscopy was performed as described previously (33,44). Briefly, cells were plated on coverslips coated with 10 μg/ml fibronectin. After fixing the cells for 10 min at room temperature with 3.7% paraformaldehyde, the cells were washed three times with PBS. Cells were permeabilized with 0.5% Triton X-100 at 37°C for 5 min. Subsequently, the cells were blocked with goat serum (Beyotime Biotechnology; cat. no. C0265) at room temperature for 30 min, then incubated with anti-p38γ and anti-albumin primary antibodies (both 1:100) in a wet chamber at 4°C overnight. This was followed by incubation with Alexa Fluor 488-conjugated anti-rabbit/mouse (cat. no. P11047; 1:500; Invitrogen; Thermo Fisher Scientific, Inc.) or Alexa Fluor 594-conjugated anti-rabbit (cat. no. S32356; 1:500; Invitrogen; Thermo Fisher Scientific, Inc.) secondary antibodies for 1 h at 37°C. Cells were washed with PBS, then cell nuclei were counterstained with DAPI (1 μg/ml; Beyotime Biotechnology) for 5 min at room temperature. The cells were mounted with Prolong Gold anti-fade reagent and imaged using a YB710FL fluorescence microscope.

The radioimmunoprecipitation assay lysis buffer (Beyotime Biotechnology; cat. no. P0013B) supplemented with 1 mM phenylmethanesulfonyl fluoride was used to extract total protein from the liver tissues and cultured cells, then protein concentration was measured using a BCA protein assay kit (Beyotime Biotechnology). Equal quantities of protein (30 μg) were loaded per lane on a 10% SDS gel, resolved using SDS-PAGE, transferred to PVDF membranes at 400 V for 30 min and then blocked for 2 h at room temperature in 5% non-fat milk in Tris-buffered saline with 0.05% Tween-20 (TBST). Subsequently, the membranes were incubated with primary antibodies at 4°C overnight. The following day, the membranes were washed TBST and incubated with horseradish peroxidase-conjugated secondary antibodies (cat. no. 7074S; Cell Signaling Technology, Inc.) diluted in blocking buffer (1:5,000) for 1 h at room temperature. An enhanced chemiluminescence kit (SuperSignal west pico trial kit; Pierce; Thermo Fisher Scientific, Inc.) was used to visualize signals. The signals were captured using a ChemiDoc MP Imaging System (Bio-Rad Laboratories, Inc.). β-actin was used as the loading control. Densitometry analysis was performed using ImageJ software (version 1.53k; National Institutes of Health). The primary antibodies were diluted as follows: Anti-PI3K (1:1,000), anti-p-PI3K (1:1,000), anti-Akt (1:1,000), anti-p-Akt (1:1,000), anti-p38γ (1:1,000), anti-IL-6 (1:1,000), anti-IL-1β (1:2,000) anti-TNF-α (1:1,000), anti-NOX4 (1:2,000), anti-iNOS (1:2,000), anti-PPAR-α (1:1,000), anti-SREBP-1 (1:1,000), anti-Fasn (1:1,000) and anti-β-actin (1:1,000).

The putative wild-type (WT) or mutant (MUT) 3'-UTR of p38γ containing the miR-125 binding site was synthesized and cloned into the pmirGLO Dual-Luciferase vector (Promega Corporation; cat. no. E1330). AML-12 cells were co-transfected with the constructed reporter vectors (luc-p38γ-WT or luc-p38γ-MUT) and either miR-125 mimics or negative control (NC) mimics using Lipofectamine™ 3000 (Invitrogen; Thermo Fisher Scientific, Inc.). After 48 h, luciferase activity was measured using the Dual-luciferase Reporter Assay Kit (Promega Corporation) according to the manufacturer's protocol. Firefly luciferase activity was normalized to Renilla luciferase activity for each sample.

The levels of serum (0.5-1.0 ml per mouse) from mice ALT, AST, MDA, GSH and SOD were measured using commercial kits according to the manufacturer's instructions.

Liver specimens were fixed in 4% paraformaldehyde, embedded in paraffin and cut into 50 μm-thick sections. Samples were stained with hematoxylin and eosin (H&E) as follows: Initially, samples were incubated with hematoxylin for 5 min and then washed with water, differentiated with hydrochloric acid solution for ~3 sec, washed with tap water, counterstained with ammonia solution for ~3 sec and washed with tap water again. Under the microscope, nuclei appeared pale blue to blue, with a pale blue or nearly colorless background. Sections were dehydrated sequentially in 85 and 95% ethanol solutions for 4 min each, then immersed in eosin solution for 1-2 min. Dehydration was performed using absolute ethanol, followed by clearing with n-butanol and xylene. Sections were then transferred to a square container containing clean xylene and mounted with neutral resin.

For the Masson's trichrome staining procedure, paraffin-embedded sections were dewaxed, immersed in water for 2 min, then sequentially immersed in xylene, absolute ethanol, 95% ethanol, 80% ethanol and 70% ethanol for varying durations, followed by washing with distilled water. Sections were then immersed in ferric hematoxylin stain for 3-8 min, then washed under running water. Subsequently, sections were stained with Alizarin Red-Acid Fuchsin for 10 min. Next, the sections were placed in phosphomolybdic acid for differentiation for several seconds, then washed with water, followed by staining in aniline blue solution for 5 min and washing with 0.2% glacial acetic acid. Sections were sequentially immersed in 95% ethanol, anhydrous ethanol and xylene for varying durations. After removing sections from xylene, they were air-dried briefly and mounted in neutral resin. Sirius red staining for collagen was performed according to the manufacturer's protocol (Beyotime Biotechnology; cat. no. C1006).

All stained sections were examined under a light microscope (Nikon Eclipse Ci-L) by two independent pathologists blinded to the experimental grouping. All steps were performed at room temperature.

To determine the intracellular ROS levels, a fluorescence-based probe sensitive to oxidation, 2',7'-dichlorodihydrofluorescein diacetate (DCFH-DA; Beyotime Biotechnology) was used. After treatment, cells were washed twice with PBS and then incubated for 20 min at 37°C with 10 μmol/l DCFH-DA. Non-specific esterase within the cells deacetylate DCFH-DA and ROS are then oxidized to produce the fluorescent chemical 2',7'-dichlorofluorescein (DCF). The cells were then washed three times with pure DMEM. Fluorescence microscopy (Yuescope; cat. no. YB710FL) was used to detect the ROS levels with excitation/emission settings of 485/535 nm. To validate the assay, untreated cells were used to establish the baseline fluorescence (negative control) and cells treated with hydrogen peroxide (200 μM for 30 min) were used as a positive control to confirm the responsiveness of the DCFH-DA probe.

Total RNA was extracted from tissues or cells using TRIzol^® reagent (Invitrogen; Thermo Fisher Scientific, Inc.; cat. no. 15596026) according to the manufacturer's protocol. RNA concentration and purity were assessed using a NanoDrop™ spectrophotometer (Thermo Fisher Scientific, Inc.). A total of 1 μg total RNA was reverse-transcribed into cDNA using the Transcriptor First-Strand cDNA Synthesis Kit (Roche Diagnostics GmbH; cat. No 04379012001) with oligo(dT) primers according to the manufacturer's protocol. qPCR was performed using FastStart Universal SYBR Green Master (Rox) (Roche Diagnostics GmbH; cat. no. 04913850001) on a QuantStudio 3 Real-Time PCR System (Applied Biosystems, Thermo Fisher Scientific, Inc.). The thermocycling conditions were as follows: 95°C for 30 min, followed by 40 cycles of 95°C for 15 sec and 60°C for 30 min. All reactions were run in triplicate. The relative mRNA expression levels were calculated using the 2^-ΔΔCq method (45) and normalized to GAPDH. The sequences of the primers are listed in Table I.

To downregulate or upregulate the expression of miR-125 and p38γ, miR-125 mimic, miR-125 inhibitor, p38γ small interfering (si)-RNA, p38γ plasmid (pEGFP-C1-p38γ) or corresponding control constructs were used. A total of 50 nM nucleic acid constructs (miR-125 mimic, miR-125 inhibitor or p38γ siRNA) and 2 μg plasmid DNA (pEGFP-C1-p38γ) or corresponding control constructs were transfected using Lipofectamine™ 3000 (Invitrogen; Thermo Fisher Scientific, Inc.) according to the manufacturer's protocol. Cells were transfected for 6 h, after which, the culture medium was changed and APAP was added. AML-12 cells were cultured at 37°C in a humidified incubator supplied with 5% CO₂ for 24 h and then collected for RT-qPCR or western blot analysis. All transfected constructs were purchased from Shanghai GenePharma, Co., Ltd. The sequences of the miR-125 mimic, miR-125 inhibitor, p38γ-siRNA and the corresponding NCs are listed in Table II.

The supernatants of AML-12 cells in the different treatment groups were collected and the expression levels of TNF-α, IL-1β and IL-6 were assessed by ELISA kits according to the manufacturer's instructions.

MTT reagent was purchased from Sigma-Aldrich (Merck KGaA). Before the experiment, MTT reagent was prepared into a 5 mg/ml working solution with serum-free medium, filtered and sterilized, and stored at 4°C in the dark for later use. AML-12 cells in the logarithmic growth phase were seeded into 96-well culture plates at a density of 1×10⁴ cells per well with a volume of 100 μl per well and cultured in a 37°C, 5% CO₂ incubator until the cells adhered to the wall. Then, different concentrations of APAP treatment solution were added (the control group was added with the same volume of solvent) and cultured for 0, 6, 12, 24 and 48 h respectively. After the incubation, 20 μl of MTT working solution was added to each well and the incubation was continued for 4 h to form purple formazan crystals. After the incubation, the supernatant in the wells was carefully aspirated and discarded, 150 μl of DMSO was added to each well and the mixture was shaken at low speed on a shaker for 10 min to ensure complete dissolution of formazan crystals. A microplate reader was used to measure the absorbance of each well at a wavelength of 570 nm with a reference wavelength of 630 nm. The cell viability of the control group was set as 100% and the relative cell viability of each treatment group was calculated. Each treatment group was set with 3 replicate wells and the experiment was independently repeated 3 times.

The male C57BL/6J mice (8 weeks old, body weight 20-22 g) were obtained from Jiangsu GemPharmatech Co., Ltd. Mice were housed under specific pathogen-free conditions at 22±2°C, 50±10% relative humidity and a 12 h light/dark cycle, with free access to standard chow and water. Before the hepatocytes were isolated, mice were allowed to acclimatize for 1 week. Then, isoflurane (induction 3-4%, maintenance 1.5-2%) was used to anesthetize the mice, followed by perfusion with a washing buffer [Hank's balanced salt solution (HBSS; Beyotime Biotechnology; cat. no. C0218) with 0.1% EDTA (Sigma-Aldrich; Merck KGaA; cat. no. E6758)] via the inferior vena cava. Next, a digestive solution (HBSS with 0.5 mg/ml collagenase D; Roche Diagnostics GmbH; cat. no. 11088866001) was used for perfusion. Once the perfusion was complete, the mice were euthanized using CO₂ and the livers were extracted and primary hepatocytes were isolated by carefully slicing the liver lobes. The cells underwent filtration, centrifugation at 50 × g for 5 min at 4°C, washing twice with cold William's E medium (Gibco; Thermo Fisher Scientific, Inc.; cat. no. A1217601) by centrifugation at 50 × g for 5 min at 4°C, and were resuspended in William's E medium. They were then seeded at a suitable density for further experiments.

Total RNA was extracted from AML-12 cells, both untreated and treated with 10 mM APAP, using TRIzol reagent, with three replicates per group. NanoDrop spectrophotometry was used to evaluate the concentrations, quality and integrity of the extracted RNA. Afterward, 3 μg of RNA (RNA integrity number, 10.0; a score of 10.0 indicates intact, high-quality RNA with no degradation) was extracted from each sample, which was sequenced and analyzed by Shanghai Personal Biotechnology Co., Ltd. Briefly, the construction of the sequencing library was completed using the TruSeq RNA Sample Preparation Kit (cat. no. RS-122-2001; Illumina, Inc.). Library quantification was achieved through a two-step method: Preliminary quantification was performed using RT-qPCR, followed by precise detection using the Agilent High Sensitivity DNA Kit on the Agilent 2100 Bioanalyzer System (Agilent Technologies, Inc.). After quantification, the library was sequenced on the Illumina Novaseq 6000 sequencing platform (300 cycles; cat. no. 20012860; Illumina, Inc.), generating paired-end reads of 150 bp in length. The final library was loaded at a concentration of 150 pM, determined by RT-qPCR and verified using the Agilent 2100 Bioanalyzer with the High Sensitivity DNA Kit. The raw sequencing data was filtered using FastQC software (version 0.11.9; https://www.bioinformatics.babraham.ac.uk/projects/fastqc/) to remove low-quality sequences and then the qualified reads were aligned to the human reference genome (version hg38) using HISAT2 alignment software (version 2.2.1; https://daehwankimlab.github.io/hisat2/). HTSeq v0.9.1 (https://htseq.readthedocs.io/en/release_0.9.1/) was used to calculate read counts, which were then normalized and expressed as fragments per kilobase of transcript per million mapped reads. Genes showing a fold change >2 and a normalized P<0.05 were considered differentially expressed. A heatmap of differentially expressed genes (DEGs) was created using R (version 4.2.2; https://www.R-project.org/). Gene Ontology (GO) (http://www.geneontology.org/) was employed to analyze biological processes and the Kyoto Encyclopedia of Genes and Genomes (KEGG) was used for pathway analysis. Unlike the previous analyses that focused on identifying DEGs by filtering with fold change >2 and normalized P<0.05, and visualizing DEGs via a heatmap, GO_BP enrichment, KEGG analysis and Gene Set Enrichment Analysis were performed on DEGs specifically to annotate their biological functions, enrich associated biological processes and explore the involved signaling pathways. Circle plots were used to illustrate the DEGs associated with specific BP terms.

Data are presented as the mean ± SD of three repeats. One-way ANOVA with Bonferroni correction was used to compare multiple groups. Otherwise, a two-tailed unpaired t-test was used. All data were analyzed using SPSS (version 21.0; IBM Corp.). P<0.05 was considered to indicate a statistically significant difference.

To assess p38γ expression during the progression of liver injury, a liver injury model was established using APAP. H&E staining was used to determine the degree of liver injury (Fig. 1A). Additional analysis showed that ALT and AST activity was increased compared with the normal group (Fig. 1B). IHC analysis showed markedly increased levels of TNF-α, IL-6 and IL-1β (Fig. 1A). These results were further confirmed by western blot analysis (Fig. 1C). Further western blot analysis showed a significant decrease in the expression level of PPAR-α, whereas the SREBP-1, Fasn, iNOS and NOX4 expression levels were increased compared with the normal group (Fig. 1D and E). Next, APAP was used to construct a liver injury cell model in vitro. To explore the pharmacological effects of APAP, an MTT assay was conducted after 0, 6, 12, 24 and 48 h of incubation. The results showed that a reduction in cell viability from 99.33 to 51.67% was observed after incubating with APAP for 24 h (Fig. S1A). In addition, the expression of IL-6 and p38γ in AML-12 cells treated with APAP increased and peaked after 24 h of treatment (Fig. S1B). A liver injury model in AML-12 cells was established using APAP at 0, 2.5, 5, 10 and 20 mM. As shown in Fig. S1C, a reduction in cell viability from 99.67 to 52.00% was observed after incubating with 10 mM of APAP for 24 h. Additionally, the western blot results showed that APAP significantly increased the p38γ and p-Akt expression levels compared with the normal group, which peaked with APAP treatment at 10 mM (24 h) (Fig. S1D). Hence, AML-12 cells were treated with APAP at 10 mM for 24 h. The results showed that the expression levels of TNF-α, IL-6, IL-1β, SREBP-1, Fasn, iNOS and NOX4 was increased in AML-12 cells treated with APAP compared with the normal group (Fig. 1F-H).

Next, RNA-seq analysis was used to compare gene expression in control AML-12 cells and APAP-induced AML-12 cells. A total of 2,138 significantly DEGs were identified, consisting of 766 upregulated and 1,372 downregulated genes. Our previous study indicated that p38γ was upregulated in the liver injury after exposure to APAP and ethanol, and its inhibition could alleviate the cell damage and inflammatory response in the combined model (27). Moreover, another of our studies demonstrated that p38γ significantly increased during the NAFLD process and knockdown of p38γ could inhibit lipid accumulation in liver cells by suppressing the activation of the JAK/STAT signaling pathway (28), providing an important preliminary foundation for the present study. Additionally, p38γ is increasingly recognized as being related to liver diseases. For example, a study indicated that p38γ could regulate the proliferation of HCC cells (22). However, the role of p38γ as a therapeutic target for APAP-induced liver injury remains poorly understood. Based on the preliminary research of our previous studies and the core role of inflammation, oxidative stress and lipid metabolism disorders in APAP-induced liver injury, p38γ was focused on for further study among the significantly upregulated genes. Of note, the results showed that p38γ expression was upregulated in APAP-treated AML-12 cells (Fig. 2A-C). The upregulation of p38γ provides a very reasonable and novel mechanism connection between APAP-induced liver injury (associated with oxidative stress, lipid buildup and inflammatory responses) and the molecular regulatory network mediated by p38γ. The results of the GO enrichment analysis showed that the liver injury induced by APAP was linked to oxidative stress, lipid buildup and inflammatory responses (Fig. 2D). Double immunofluorescence staining showed that p38γ co-localized with hepatocyte albumin in the liver tissues (Fig. 2E), suggesting that p38γ was expressed in hepatocytes.

In addition, immunofluorescence, RT-qPCR and western blot analysis results showed that the p38γ expression level was significantly increased in the liver tissues of APAP-treated mice compared with the normal group (Fig. 2F, G and I). Similarly, p38γ expression was also upregulated in APAP-treated AML-12 cells (Fig. 2F, H and J). Collectively, these findings from both in vivo (APAP-treated mice) and in vitro (APAP-treated AML-12 cells) models confirmed that APAP successfully induced liver injury and consistently demonstrated p38γ upregulation in this APAP-induced liver injury model, suggesting that p38γ is functionally involved in the pathogenesis of APAP-induced liver injury. Thus, the specific role of p38γ in this process was further investigated.

A growing body of evidence underscores the importance of oxidative stress and lipid accumulation in liver injury induced by APAP (46-48). Thus, whether p38γ is functionally involved in regulating oxidative stress and lipid accumulation was next assessed. To this end, the p38γ-siRNA and pEGFP-C1-p38γ were used to downregulate and upregulate p38γ expression, respectively. Satisfactory transfection efficiency was obtained at 24 h post-transfection (Figs. S2A-D and S3C). Next, it was found that the MDA level was increased, while the GSH and SOD levels were decreased in APAP-induced AML-12 cells compared with normal cells. p38γ knockdown reduced MDA levels and increased the levels of GSH and SOD. By contrast, overexpression of p38γ increased MDA levels and decreased the GSH and SOD levels (Fig. 3A-C). Oil red O staining showed accumulation of cellular lipid in APAP-treated AML-12 cells compared with the normal group. Lipid accumulation was reduced by p38γ knockdown compared with p38γ NC treatment, whereas overexpression of p38γ resulted in increased lipid accumulation (Fig. 3D). The results of DCF, DHE and MitoSOX staining revealed increased ROS production within the APAP group compared with the normal group. Downregulation of p38γ inhibited the production of ROS compared with the APAP + p38γ NC group. Conversely, ROS generation was increased following p38γ overexpression compared with the APAP + pEGFP-C1 group (Fig. 3E). Next, the expression levels of SREBP-1, Fasn and PPAR-α were assessed, three proteins that are linked to lipogenesis (49). The results showed that APAP treatment increased the expression of SREBP-1 and Fasn, and decreased the expression of PPAR-α. The effect of APAP treatment on these lipid accumulation-related proteins was promoted by pEGFP-C1-p38γ (Fig. 3G) and reduced by p38γ siRNA (Fig. 3F). The expression levels of oxidative stress-related proteins (iNOS and NOX4) in AML-12 cells were assessed; their expression was increased in APAP-induced AML-12 cells. p38γ overexpression increased the impact of APAP treatment on the production of NOX4 and iNOS (Fig. 3G), whereas p38γ knockdown reduced their expression (Fig. 3F), indicating the regulatory function of p38γ in oxidative stress. Overall, these results demonstrated that p38γ increased oxidative stress and lipid accumulation aggravated by APAP in liver injury.

Inflammation is a critical cause of APAP-induced hepatic injury. Therefore, the impact of p38γ on the inflammatory response in APAP-treated AML-12 cells was next assessed. The ELISA results indicated that the expression of TNF-α, IL-6 and IL-1β were elevated by APAP. This increase in inflammatory cytokine release was significantly suppressed by p38γ knockdown and enhanced by p38γ overexpression (Fig. 4A and B). Furthermore, the results of the RT-qPCR analysis showed that, after p38γ-siRNA treatment, the expression levels of IL-1β, TNF-α and IL-6 were significantly downregulated (Fig. 4C). However, p38γ overexpression significantly increased the expression levels of IL-1β, TNF-α and IL-6 (Fig. 4D). Notably, these results were further validated through western blot analysis (Fig. 4E and F). Consequently, these findings provided evidence supporting the involvement of p38γ in the release of inflammatory cytokines during APAP-induced liver injury.

TargetScan was used to determine the upstream regulatory mechanism of p38γ. TargetScan analysis showed that a large number of miRNAs could target the 3'-UTR region of p38γ, such as miR-4319. However, among all the predicted miRNAs targeting p38γ, miR-125 had a very high 'context++ score', which is a key indicator reflecting binding affinity and conservation, and identified miR-125 as a regulator targeting the 3'-UTR of p38γ (Fig. 5A). To further investigate if the predicted binding site of miR-125 to the 3'-UTR of p38γ mRNA was responsible for the downregulation of p38γ, a WT and MUT 3'-UTR of p38γ was cloned into a luciferase reporter vector. WT-p38γ and miR-125 mimics or scrambled control were co-transfected into AML-12 cells to perform a luciferase reporter assay. In AML-12 cells transfected with miR-125 mimics, the luciferase activity was significantly reduced compared with the scrambled control cells (Fig. 5B). In addition, the miR-125 mimics and miR-125 inhibitor were used to upregulate and downregulate the miR-125 expression, respectively. Satisfactory transfection efficiency was obtained at 24 h post-transfection (Fig. S3A-D). RT-qPCR results indicated that the mRNA level of p38γ was downregulated by miR-125 mimics and upregulated by miR-125 inhibitor (Fig. 5C); western blot analysis further validated these findings (Fig. 5D). Next, AML-12 cells were co-transfected with pEGFP-C1-p38γ and miR-125 mimics. Western blot analysis demonstrated a significant decrease in TNF-α, IL-6 and IL-1β protein levels in the co-transfection compared with the pEGFP-C1-p38γ transfected group (Fig. 5E). The expression levels of miR-125 in APAP-induced liver injury mice were assessed using RT-qPCR; miR-125 expression was decreased in primary hepatocytes and liver tissues (Fig. 5F). Similarly, APAP treatment significantly suppressed miR-125 expression in AML-12 cells (Fig. 5G). Taken together, these results indicated that miR-125 directly targeted the 3'-UTR regions of p38γ to inhibit its expression.

To explore the effect of miR-125 on the etiology and progression of liver injury induced by APAP treatment, additional analysis on the impact of miR-125 on indicators associated with oxidative stress and lipid accumulation was performed. The results demonstrated that knockdown of miR-125 increased MDA levels and decreased GSH and SOD levels. Notably, the increase in MDA levels and decrease in GSH and SOD levels could be reversed by the downregulation of p38γ (Fig. 6A-C). Conversely, the MDA level was reduced and the GSH and SOD levels were elevated in AML-12 cells transfected with miR-125 mimics compared with AML-12 cells transfected with mimics NC. However, the MDA level was increased and the GSH and SOD levels were decreased in AML-12 cells transfected with miR-125 mimics and pEGFP-C1-p38γ compared with AML-12 cells transfected with miR-125 mimics (Fig. 6A-C). Furthermore, the results of Oil red O staining showed that knockdown of miR-125 increased lipid accumulation compared with AML-12 cells treated with miR-NC. When AML-12 cells were treated with miR-125 inhibitor and p38γ-siRNA, lipid accumulation was reduced compared with AML-12 cells treated with miR-125 inhibitor. In addition, lipid accumulation was repressed by miR-125 overexpression and reversed by p38γ overexpression (Fig. 6D). The findings of the DCF, DHE and MitoSOX staining indicated that the generation of ROS was increased in AML-12 cells transfected with miR-125 inhibitor and decreased in AML-12 cells transfected with miR-125 mimics. However, the increase and decrease in ROS generation were reversed by p38γ-siRNA and pEGFP-C1-p38γ, respectively (Fig. 6E). Next, the expression levels of SREBP-1, Fasn and PPAR-α we evaluated. The results suggested that APAP suppressed the expression of PPAR-α and promoted the expression of SREBP-1 and Fasn. Overexpression of miR-125 increased the expression of PPAR-α and decreased the expression of SREBP-1 and Fasn. Knockdown of miR-125 significantly increased the expression of SREBP-1 and Fasn, and inhibited the expression of PPAR-α. However, the effect of miR-125 inhibitor and miR-125 mimics treatment on lipid accumulation-related proteins (PPAR-α, SREBP-1 and Fasn) was reversed by pEGFP-C1-p38γ and p38γ-siRNA, respectively (Fig. 6F and G). The expression levels of oxidative stress-related proteins (iNOS and NOX4) in AML-12 cells were next detected. It was found that the expression of iNOS and NOX4 were increased by knockdown of miR-125, and this was reversed by p38γ knockdown. By contrast, overexpression of miR-125 reduced the expression of iNOS and NOX4, and upregulation of p38γ reversed the inhibition (Fig. 6F and G), suggesting the regulatory role of miR-125 in oxidative stress. Overall, these results indicated that miR-125 primarily targeted p38γ, inhibiting the aggravation of oxidative stress and lipid accumulation in APAP-induced liver injury.

This regulatory axis was next examined in the context of inflammation. ELISA, RT-qPCR and western blotting were used in APAP-treated AML-12 cells to investigate the impact of miR-125 on inflammatory responses. Consistent with prior observations, ELISA results confirmed elevated expression levels of TNF-α, IL-6 and IL-1β following APAP treatment. However, knockdown of miR-125 significantly increased the levels of TNF-α, IL-6 and IL-1β in AML-12 cells induced by APAP, which was reversed by transfection with p38γ siRNA (Fig. 7A). Overexpression of miR-125 suppressed the levels of TNF-α, IL-6 and IL-1β, which was reversed by transfection with pEGFP-C1-p38γ in AML-12 cells treated with APAP (Fig. 7B). Furthermore, RT-qPCR results showed that knockdown of miR-125 increased the expression levels of IL-1β, TNF-α and IL-6; however, p38γ-siRNA decreased the production of these inflammatory cytokines. By contrast, overexpression of miR-125 decreased the expression levels of IL-1β, TNF-α and IL-6, but pEGFP-C1-p38γ increased the production of these inflammation cytokines (Fig. 7C and D). Furthermore, the validity of these findings was confirmed through the utilization of western blot analysis (Fig. 7E and F). In summary, these results provide evidence that, in the setting of APAP-induced liver damage, miR-125 may directly target p38γ to inhibit the production of inflammatory cytokines.

The KEGG pathway enrichment analysis of the aforementioned RNA-seq data revealed the emergence of the 'PI3K-Akt signaling pathway' (Fig. 8A), suggesting that this pathway may play a role in APAP-induced liver damage. Based on these results, western blot analysis confirmed that APAP could promote the phosphorylation of PI3K and Akt (Fig. 8B and C). The expression levels of p-PI3K and p-Akt were markedly decreased by downregulating p38γ and overexpressing miR-125 (Fig. 8D and F). By contrast, upregulation of p38γ and knockdown of miR-125 significantly increased the expression levels of p-PI3K and p-Akt (Fig. 8E and G). In APAP-induced AML-12 cells, LY294002 (40 μM; 48 h) was used to inhibit PI3K/Akt signaling. The chemical structure of LY294002 is shown in Fig. 8H. In APAP-induced AML-12 cells, oxidative stress, lipid accumulation and inflammation-related proteins were detected. The results of the western blot analysis showed that the APAP + p38γ NC+ LY294002 group had higher expression levels of PPAR-α and lower expression levels of SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β compared to the p38γ NC group (Figs. 8I and 9C). The APAP + miR-125 NC + LY294002 group had higher expression levels of PPAR-α and lower expression levels of SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β compared with the miR-125 NC group (Figs. 8B and 9F). Conversely, LY294002 treatment did not further alter the expression levels of PPAR-α, SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β in the p38γ siRNA or miR-125 mimics group compared with the p38γ siRNA or miR-125 mimics group, indicating that the protective effects of p38γ knockdown or miR-125 overexpression are largely mediated through the PI3K/Akt signaling pathway (Figs. 8I, 9B, C and F). Conversely, compared with the pEGFP-C1/miR-125 NC group, the pEGFP-C1/miR-125 NC + LY294002 group showed decreased expression levels of SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β, and increased expression levels of PPAR-α. However, the pEGFP-C1-p38γ/miR-125 inhibitor + LY294002 group exhibited repressed expression levels of SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β, and elevated expression levels of PPAR-α compared to the pEGFP-C1-p38γ/miR-125 inhibitor group (Figs. 8J, 9A, D and E). These results suggested that miR-125 may inhibit oxidative stress, lipid accumulation and inflammatory response by targeting p38γ via the PI3K/Akt signaling pathway in APAP-induced liver injury.

AAV9-shRNA-p38γ was injected into the tail vein to knock down p38γ expression to investigate the possible role of p38γ in APAP-induced liver injury in more detail. The results showed that the p38γ knockout mice were successfully constructed (Fig. 10A and B). Immunofluorescence and western blot results indicated that AAV9-shRNA-p38γ reduced the expression level of p38γ (Fig. 10C and D). H&E staining results showed that inflammatory cell infiltration, vacuolar degeneration and the area of necrosis were significantly decreased, indicating liver injury was reduced in AAV9-shRNA-p38γ mice (Fig. 10E). IHC staining results showed that the expression of TNF-α, IL-6 and IL-1β were decreased in the AAV9-shRNA-p38γ mice (Fig. 10F). In addition, the AAV9-shRNA-p38γ group showed higher serum GSH and SOD levels compared with the AAV9-Empty group (Fig. 10I and K). By contrast, the serum levels of ALT, AST and MDA were lowered (Fig. 10G, H and J). Oil red O staining suggested that the lipid accumulation was decreased in the AAV9-shRNA-p38γ group (Fig. 10L). Crucially, primary hepatocytes were extracted from both the AAV9-shRNA-p38γ group and the AAV9-Empty group. The DFH staining findings indicated that the generation of ROS was reduced in the AAV9-shRNA-p38γ group (Fig. 10M). Additionally, the expression of PPAR-α was increased in the AAV9-shRNA-p38γ group; however, the expression of SREBP-1, Fasn, iNOS, NOX4, TNF-α, IL-6 and IL-1β was suppressed, according to the findings of the western blot analysis (Fig. 10N and O). Collectively, these findings indicate that the notable impacts of hepatic p38γ depletion are partially governed by the reduction of APAP-induced inflammation, oxidative stress and lipid accumulation. Essentially, miR-125 may regulate p38γ, which may function as an important modulator in the setting of APAP-induced liver damage.