Journal of Animal Science and Biotechnology
Research

Mechanisms of high-glucose-induced mitochondrial damage and glycolipid accumulation in largemouth bass

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Abstract

Background

The carnivorous fish, largemouth bass (Micropterus salmoides), has difficulty metabolizing dietary carbohydrates, frequently resulting in issues with energy metabolism and fatty liver disease. Nevertheless, the molecular mechanisms involved are still not fully understood.

Results

The results of high-carbohydrate (HC) diets and high-glucose (HG) treatments in largemouth bass hepatocytes showed that high-glucose causes liver damage and glycolipid accumulation. High-glucose promoted the lipogenesis process by activating AMPK/ACC/SREBP-1 pathway and reduced bile acid synthesis by downregulating cholesterol 7-hydroxylase (cyp7a1) and sterol 12-hydroxylase (cyp8b1). Concurrently, HG treatments also caused mitochondrial fission and damage by increasing the expression of dynamin-related protein 1 (Drp1), leading to impaired mitochondria accumulation and mitochondria-dependent apoptosis via the p38 MAPK/Bcl-2/Casp3 pathway. Additionally, HG treatments decreased Sirt1 expression and relocated it from the nucleus to the cytoplasm, where it interacts with autophagosomes and lysosomes, inhibiting Pink1/Parkin-mediated mitophagy. This also led to the cytoplasmic translocation of Pink1 and its co-localization with Sirt1, indicating that Sirt1 regulates high glucose-induced metabolic stress by inhibiting the Pink1/Parkin mitophagy pathway.

Conclusion

In summary, HG treatment induces mitochondrial damage and glycolipid accumulation in largemouth bass through mechanisms involving AMPK/SREBP1/ACC1-mediated lipogenesis, bile acid metabolism, Sirt-mediated mitophagy, and p38 MAPK/Bcl-2/Casp3-activated apoptosis.

Background

In recent years, aquacultural production of largemouth bass (Micropterus salmoides) in China has rapidly expanded in parallel with the development and optimization of compound feed. By 2025, Chinese aquaculture production of largemouth bass is predicted to reach 1,000 thousand tons [1]. Multiple studies have demonstrated that an excessive carbohydrate content (exceeding 10%) in the formula feed of largemouth bass can lead to liver damage [2,3,4,5,6]. Additionally, high-carbohydrate diets have been shown to induce vacuolization of liver parenchymal cells and reduce survival in largemouth bass, which may be correlated to increased liver damage [7, 8]. Therefore, low-carbohydrate feed has become the predominant choice within the largemouth bass industry [9]. However, the excessive use of fish meal in low-carbohydrate feed leads to elevated costs and environmental concerns, significantly impeding the industry's development. 

Excessive lipid and glycogen accumulation in the liver has been identified as a critical factor contributing to growth retardation and hepatic damage in fish subjected to a high-carbohydrate diet [10, 11]. Hepatic fat accumulation generally arises from an imbalance between lipogenesis and lipolysis, mediated by various transcriptional regulators and enzymatic activities involved in hepatic lipid homeostasis [12]. Xie et al. [13] found that high-carbohydrate diets promote greater fat accumulation in Nile tilapia (Oreochromis niloticus) than high-fat diets. In hybrid grouper (Epinephelus fuscoguttatus ♀ × E. lanceolatus ♂), a high-carbohydrate diet significantly downregulated adipose triglyceride lipase (atgl) expression, leading to hepatic triglyceride accumulation [14]. The expression levels of fatty acid synthase (fas) and acetyl-CoA carboxylase 1 (acc1) were significantly up-regulated, resulting in increased plasma cholesterol and total fat content in grass carp fed with high-carbohydrate diets [15]. Furthermore, it was also shown that AMP-activated protein kinase (AMPK) and sterol regulatory element-binding protein 1 (srebp1) are both involved in triglyceride accumulation in largemouth bass fed high-carbohydrate diet [16]. To fully clarify the regulatory effect of a high-carbohydrate diet on fatty metabolism, we analyzed the related genes of lipolysis/fatty acid oxidation, lipogenesis, and fatty acid transport.

Within the classic bile acid synthesis pathway FXR-mediated shp decreases bile acid biosynthesis by inhibiting cyp7a1 expression [17]. Concurrently, FXR stimulates FGF-19 in the enterocytes, which subsequently activates fibroblast growth factor receptor 4 (fgfr4). This activation triggers the c-JUN and ERK signaling pathways, leading to a reduction in new bile acid synthesis through negative feedback mechanisms [18]. However, research on MAPK signaling pathways in farmed fish remains nascent, with limited understanding of the detailed mechanisms and regulatory functions [19,20,21,22,23]. In Nile tilapia, it has been observed that the NF-κB pathway, rather than the p38 MAPK pathway, is implicated in intestinal inflammation induced by high-carbohydrate diets [24]. In largemouth bass, high-carbohydrate intake may impair spleen immune function through an inflammatory response mediated by the MAPK/FoxO pathway [25]. To elucidate the mechanisms underlying high-carbohydrate-induced lipid metabolic disorders in largemouth bass, this study would concentrate on the MAPK signaling pathway with in vivo and vitro.

In mammals, mitochondrial dysfunction constitutes a major pathogenic factor in metabolic disorders such as insulin resistance and diabetes [26]. Fish, as ectothermic organisms, exhibit unique mitochondrial adaptations that are essential for their survival in diverse aquatic environments [27]. Oxidized fish oils have been demonstrated to induce significant depletion of mitochondrial membrane potential and drastically reduce ATP production in the liver of yellow catfish (Pelteobagrus fulvidraco) [28]. Complementary transcriptomic analyses by Prisingkorn et al. [29] revealed that high-fat-high-carbohydrate diets substantially downregulate expression of key mitochondrial biogenesis genes, including PPARγ-coactivator 1α (PGC-1α), establishing a direct molecular link to mitochondrial impairment. Substantiating these findings, Shen et al. [30] documented profound mitochondrial dysfunction in high-glucose cultured tilapia hepatocytes, evidenced by diminished mitochondrial density, coupled with markedly attenuated activities of respiratory chain Complexes II and III. Investigations in blunt snout bream (Megalobrama amblycephala) further demonstrated that high-carbohydrate diets disrupt mitochondrial dynamics through dysregulated fission–fusion equilibrium, consequently suppressing oxidative phosphorylation capacity [31]. Therefore, understanding how high-glucose impacts mitochondrial function could provide insights into metabolic disorders in fish and inform dietary strategies to mitigate these effects.

Sirt1, as a central regulator of energy metabolism, forms a synergistic network with the AMPK signaling pathway to maintain mitophagic homeostasis [‌32]. Mechanistic studies reveal that Sirt1 activates AMPK through deacetylation modifications, directly promoting autophagosome formation and enhancing mitophagic flux‌ [33]. This regulatory mechanism is particularly critical for eliminating mitochondria damaged by oxidative stress [‌34]. The Pink1/Parkin pathway is a core executor of mitophagy and exhibits functional coupling with Sirt1 [35]. The Sirt1-Sirt3 regulatory axis enhances mitochondrial membrane localization stability of Pink1 and optimizes Parkin-mediated ubiquitination, thereby improving the recognition and clearance of dysfunctional mitochondria [36]. Notably, Sirt1 modulates the opening dynamics of mitochondrial permeability transition pores (mPTP), which governs cytochrome c release kinetics and acts as a pivotal checkpoint in metabolic inflammation and apoptotic cascades [37]‌.

Current research on the metabolic effects of high-carbohydrate diets in largemouth bass primarily focuses on animal models (e.g., feed substitution, growth and enzyme activity analysis), while molecular mechanism validation at the cellular level remains insufficient [38,39,40]. To address this gap, this study employed integrated in vivo and in vitro models to comprehensively evaluate the impacts of high-glucose conditions on hepatic health, lipid metabolism, bile acid homeostasis, and mitochondrial dynamics. Furthermore, the study revealed the mechanism by which the ‌Sirt1/Pink1/Parkin axis‌ improves metabolic disorders through optimized mitochondrial quality control. These findings provide a theoretical foundation for feed formulation optimization‌ ‌and targeted metabolic intervention strategies‌, ultimately supporting the sustainable development of aquaculture‌.

Methods

Feeding trial and sampling

Juvenile mixed gender largemouth bass were randomly assigned to two diets: a control diet (CON) and a high-carbohydrate diet (HC) for 8-week trial starting at 8.24 ± 0.01 g initial weight. As shown in Table 1, these two diets were made, packed, and stored according to our standard laboratory procedures [41]. Briefly, all ingredients were ground into powder and mixed thoroughly by a feed mixer (A-200 T Mixer Bench Model unit, Resell Food Equipment Ltd, Ottawa, Canada). A screw-press pelletizer was used to obtain 2.0 mm pellets from the mixture containing fish oil, soy oil, and soya lecithin and water (F-26, South China University of Technology, Guangzhou, China). Pellets were dried at 16 °C in a well-ventilated condition until moisture content dropped below 10%, and then stored at −20 °C. A total of 240 fish were randomly allocated into 6 tanks (capacity: 100 L) with 30 fish per tank. There were four replicates of each diet. All fish were cultured in an experimental system with commercial feed (Tongwei Co., Ltd., China) for two weeks to adapt to the experimental conditions. During the 8-week trial, fish were fed two times per day at 9:00 and 17:00 and maintained under recirculating aquaculture system of 25–28 °C water temperature, 9.0 mg/L dissolved oxygen, 7.9–8.2 pH, and lower than 0.2 mg/L ammonia nitrogen level. After the trial, fish were anaesthetized and euthanized with MS-222 (200 mg/L; Sigma, USA) and then weighed. The viscera and liver of each fish were also weighed and photographed for the evaluation of hepatic and visceral lipid accumulation. For future analyses, tissues (including liver, heart, brain, intestine, and head kidney; n = 4) were promptly frozen in liquid nitrogen and stored at −80 °C.

Table 1 Ingredients and nutrient compositions of the experimental diets (dry matter basis)
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Periodic acid-Schiff (PAS) staining and glycogen content detection of the liver

Liver tissue sections (1–4 µm) were fixed in 4% paraformaldehyde (Servicebio, Wuhan, China) for 24 h and embedded in paraffin. Liver glycogen staining was conducted using the periodic acid-Schiff (PAS) reaction. Images were acquired by a light NikonNi-U microscope (Nikon Corporation, Japan) with 20× magnification. The glycogen content was measured by spectrophotometry using standard commercial kits from Wuhan Abbkine Co., China (KTB1340).

Primary hepatocyte isolation and treatment

Primary hepatocytes were isolated from juvenile healthy largemouth bass (weight: 10–20 g) livers, which feed with commercial diets (Tongwei Co., Ltd., China) two times per day at 9:00 and 17:00, and cultured following established protocols [41]. Briefly, healthy largemouth bass were first bled and sacrificed by gill cutting under sterile conditions. The livers were aseptically excised and finely minced using scissors in phosphate-buffered saline (PBS) at pH 7.4. Subsequently, the minced liver tissue underwent enzymatic digestion with trypsin (25200072; Thermo Fisher, USA) at 28 °C for 40 min. The digested liver tissue was filtered through a sterile 70 μm cell strainer to remove undissociated tissue fragments and the filtrate was collected as a single-cell suspension. Then the suspension was centrifuged at 1,000 r/min for 5 min at room temperature to collected the cell pellet for subsequent cell counting. The harvested cell pellet was cultured in low-glucose medium (66113-18508; 1,000 mg/L; Vigonob, Guangzhou, China) containing 20% FBS and 1% penicillin–streptomycin. A total of 1 × 106 living cells per well were seeded into 6-well plates and culture at 28 °C in a humidified incubator with air charge of 5% CO2. Cells were treated with either low-glucose (LG; 1,000 mg/L) or high-glucose (HG; 4,500 mg/L) for 24, 48, and 72 h when they reached 70%–80% confluence. The cells were pretreated with various concentrations of SB203580 (p38 MAPK pathway inhibitor; S1076, Selleck Chemicals, Houston, Texas, USA) for 2 h, then treated with HG for 48 h.

Cell viability assays

Cell viability assays were examined by Cell Counting Kit-8 (CCK8) (FD3788; Fdbio Science, Hangzhou, China). The isolated primary hepatocytes were seeded in 96-well plates (100 μL/well) at a density of 1 × 104 cell/well. After 24 h, 48 h, and 72 h cultured in either LG or HG medium, 10 μL of CCK8 was added to each well 2 h prior to measuring the absorbance at 450 nm using a multifunctional microplate reader.

Annexin V-FITC/PI staining

Cell apoptosis was evaluated with flow cytometry (BL110A; Biosharp Life Sciences, Beijing, China). Primary hepatocytes were seeded in 6-well plates and subsequently digested with EDTA-free trypsin after LG or HG treatments. According to the manufacturer's instructions, both the supernatants and cell pellets were collected and stained with 5 μL of Annexin V-FITC and 10 μL of PI staining solution, respectively. The apoptotic cells were detected by CytoFLEX flow cytometer (Beckman Coulter, Indianapolis, IN, USA).

Assessment of mitochondrial membrane potential (MMP)

Assessment of the electrical potential across the mitochondrial membrane (ΔΨm). The MMP alterations were evaluated using the MMP assay kit containing JC-1 (C2006; Beyotime Biotechnology, Shanghai, China). Hepatocytes were initially cultured with LG and HG for 48 h, followed by washing with ice-cold PBS and staining with 1 mL of JC-1 working solution at 28 °C for 20 min. Subsequently, stained cells were examined using a Leica DM1000 fluorescence microscope (Leica Microsystems GmbH, Wetzlar, Germany).

Measurement of ROS

Generation of intracellular ROS was identified by utilizing the H2DCF-DA probe (C-2938; Invitrogen™, Waltham, MA, USA). After LG or HG treated for 48 h, cells were incubated in serum-free DMEM with 15 μmol/L H2DCF-DA for 30 min, shielded from light, and then the medium were replaced with pre-warmed PBS. ROS fluorescence was visualized under a Leica SP8 STED confocal laser scanning microscope (Leica Microsystems GmbH, Wetzlar, Germany), and the intensities were measured using Image-Pro Plus software. Additionally, the level of ROS was also detected by Cytoflex flow cytometry (Beckman Coulter, Inc., Brea, CA, USA).

Electron microscopy

For transmission electron microscopy (TEM), liver tissues and cells were fixed in 2.5% glutaraldehyde (AAPR46; Servicebio, Wuhan, China) and rinsed with PBS. The samples were dehydrated in a graded series of ethanol and embedded in pure resin overnight. In order to observe various structures within the livers and cells, a transmission electron microscope (JEM-1400 Flash, Tokyo, Japan) was used for observation and photography.

Oil Red O staining and PAS staining

For the cellular experiments, Oil Red O staining and ‌PAS staining were performed according to the manufacturer’s protocol (cat. no. C0157 and C0142 respectively; Beyotime Institute of Biotechnology, Shanghai, China). Lipid and glycogen accumulation in cells was observed using a 20× fluorescence microscope (Leica DM1000; Leica Microsystems GmbH, Wetzlar, Germany).

Analysis of mitochondrial-related indicators

Mitochondrial staining assay was determined by staining the cells with Mito Tracker Red (cat. no. M7521; Invitrogen; Thermo Fisher Scientific, Inc., Waltham, MA, USA) and Mito Tracker Green FM (cat. no. M7514; Invitrogen; Thermo Fisher) according to the manufacturer’s protocol. The fluorescence intensity was assessed in 1 × 104 cells by flow cytometry. For mitochondrial reactive oxygen species (mROS) production and mitochondrial morphology analysis, cells were labeled with 5 μmol/L MitoTracker™ Red CM-H2XRos (cat. no. M7513, Thermo Fisher) for mROS and 5 μmol/L MitoTracker™ Red FM (cat. no. M22425, Thermo Fisher) in PBS at 28 °C for 30 min, followed by super-resolution imaging with a Leica SP8 STED microscope (Leica Microsystems GmbH, Wetzlar, Germany).

Confocal microscopy

GFP-LC3 and pRK5-flag-ev were stored in our laboratory. pcDNA3.1(+)-MITO/Turbo RFP was purchased from Yunzhou Biology Company (VB211020-1156jcn). To construct the pRK5-flag-sirt1 plasmids, sirt1 mRNA was amplified using the following primer pairs: sirt1 forward, 5′-ACAAGGACGACGATGACAAGATGGCGGACGGAGAGAGCAGT-3′ and reverse, 5′-AGGTCGACTCTAGAGGATCCTTAAAGGTGTGTGGTGCTCTGAG-3′; pRK5-flag-ev was amplified using the following primer pairs: pRK5-flag-ev forward, 5′-GGATCCTCTAGAGTCGACCTGCAG′ and reverse, 5′-CTTGTCATCGTCGTCCTTGTAGTCCA-3′. Subsequently, Sirt1 and pRK5-flag-ev were ligated by homologous recombination (cat. no. C5891; Clone Smarter Technologies, Beijing, China), and identified using EcoRI enzyme digestion. siRNA targeting Sirt1 (5′-GCCAAUGAGGCCACAUCAATT-3′), and the negative control (5′-UUGAUGUGGCCUCAUUGGCTT-3′) were synthesized by Shanghai Sangon Biotech Co., Ltd. Plasmids and si-Sirt1 were transfected into the cells with jetPRIME® (101000046; Polypolus, Strasbourg, France) in a laser confocal culture dish (80100215). To stain the acidic compartments, live cells were stained with 50 nmol/L LysoTracker Red (L7528; Thermo Fisher) for 2 h at 28 °C in the dark and fixed with 4% paraformaldehyde (AAPR12). Subsequent, cells were permeabilized using 0.1% Triton X-100 (AAPR96) and blocked using 2% BSA (AAPR305) in TBST. The primary antibody dilutions were as the follows: Tom20 (1:100), Sirt1 (1:100), Flag (1:100), Pink1 (1:100), p-P38 (1:100). The membranes were washed with TBST and incubated with appropriate secondary antibodies (Alexa Fluor goat anti-rabbit 594, 1:100) for 1 h. Finally, ProLong Gold Antifade (P36941) was used to stain the nuclei and prevent fluorescence quenching. The slips were imaged using a Leica SP8 STED confocal laser scanning microscope (Leica Microsystems GmbH).

Quantitative real-time PCR (RT-PCR) and western blot

Total RNA was extracted from various tissues of largemouth bass and cells using RNAiso Plus reagent (9109, Takara Bio, Inc., Japan) and reverse transcribed to cDNA on the specification (AK2601; Takara Bio, Inc., Japan). RT-PCR was performed using the Roche Light Cycler 480II Real-Time System (Switzerland), and the gene expression levels were normalized with reference to ef-1α (GenBank accession no. KT827794) using the 2−ΔΔCq method. PCR amplification primer sequences were shown in Table 2.

Table 2 Sequences of primers used in this study
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A mixture of protease inhibitors (FD1002; Fdbio science, Hangzhou, China) was added to RIPA lysis buffer (FD009; Fdbio science, Hangzhou, China; 1:100) to extract total protein from livers and cells. All details of the primary antibodies and corresponding secondary antibodies used were stated in Table 3. Bands were visualized using an Azure 300 ultra-sensitive chemiluminescence imager (Azure Biosystems, USA). Protein levels were standardized to β-actin levels and quantified using Image-Pro Plus software.

Table 3 Antibodies used in this study
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Statistical analysis

Analysis of variance (ANOVA) or unpaired/paired t-tests of three independent repeats were performed with GraphPad Prism 8 (GraphPad Software, Inc., La Jolla, CA, USA). Data from three or four independent biological replicates were expressed as mean ± SEM. Normality was assessed by Shapiro-Wilk test, and homogeneity of variance was confirmed by Brown-Forsythe test. For comparisons between multiple groups, one-way ANOVA with Tukey’s post hoc test was applied when parametric assumptions were met; otherwise, Kruskal–Wallis test with Dunn’s correction was used. Paired/unpaired t-tests (two-tailed) were selected for two-group comparisons after verifying Gaussian distribution.

Results

High-glucose treatment induced hepatic glycolipid accumulation, liver damage and oxidative stress in both largemouth bass and primary hepatocytes

Following the 8-week experimental period, compared to the CON diet, the hepatosomatic index (HSI) and viscerosomatic index (VSI) were significantly increased in the largemouth bass fed HC diet (Fig. 1A and B). Consistent with the HSI and VSI, HC-fed largemouth bass showed obvious white liver and enlargement (Fig. 1C). PAS staining in the liver tissue of largemouth bass further corroborated that HC diet caused profound glycogen accumulation (Fig. 1D). Correspondingly, hepatic glycogen content in the HC diet was elevated by approximately 50% relative to the CON diet (Fig. 1E). Largemouth bass primary hepatocytes were exposed to a high-glucose medium to replicate in vivo conditions associated with high-carbohydrate intake, aiming to elucidate the impact of a high-carbohydrate diet on hepatic glycolipid accumulation. The results indicated that HG treatment significantly inhibited cell proliferation at 48 and 72 h (Fig. 1F). Consequently, a 48-h treatment duration was chosen for subsequent high-glucose exposure in primary hepatocytes. The results of Oil Red O staining and PAS staining proved that HG treatment promoted the accumulation of lipid droplets and glycogen in primary hepatocytes (Fig. 1G and H). Meanwhile, we also evaluated the expression of glycogenesis-related genes and the content of glycogen in primary hepatocytes. The results exhibited that HG treatment induced glycogen synthesis, as evidenced by an increase in the mRNA levels of gsy2 (Fig. 1L) and the content of glycogen (Fig. 1K). Moreover, generation of ROS was observed by fluorescence microscopy to detect whether HG treatment can result in oxidative stress. In primary hepatocytes, HG treatment increased mean fluorescence intensity (MFI) of H2DCF-DA dye, indicating elevated ROS production as compared to LG treatment (Fig. 1I). Flow cytometry analysis also demonstrated that HG treatment resulted in approximately 9% increased intracellular ROS levels in primary hepatocytes (Fig. 1J). Additionally, the gene expression level of nrf2 was up-regulated, and the gene expression levels of nqo1 was down-regulated after HG treatment (Fig. 1M). 

Fig. 1
figure 1

High-glucose treatment induced hepatic glycolipid accumulation, liver damage and oxidative stress in both largemouth bass and primary hepatocytes. A Hepatosomatic index (HSI, n = 4). B Viscerosomatic index (VSI, n = 4). C Liver morphology. D PAS staining of liver sections. E The content of glycogen in the liver. F Cell Counting Kit-8 test (n = 3). G Oil Red O staining of cells. H PAS staining of cells. I Intracellular ROS were determined using a fluorescence microscope. J The measurement of intracellular ROS. K The content of glycogen in cells (n = 4). L Relative expression of glycogenesis-related genes (ennp1, gys2, and acadm) in cells (n = 3). M Relative expression of anti-oxidant capacity-related genes (nrf2, keap1a, keap1b and nqo1) in cells (n = 3)

High-glucose treatment disturbs fatty acid metabolism, bile acid metabolism and activates MAPK signal pathway in both largemouth bass and primary hepatocytes

Given that HC diet can lead to fat accumulation in visceral tissues, we further investigated the impact of HC diet on fatty acid metabolism and bile acid metabolism. Compared with CON diet, HC diet resulted in the upregulation of lipolysis/fatty acid oxidation, lipogenesis and fatty acid transport-associated genes, as evidenced by upregulated expression of atgl, fas, acc1, srebp1, ppar-γ, apob, apob100 and fabp1. The gene expression levels of cyp7a1, cyp8b1, and shp were down-regulated, and the gene expression level of fgfr4 was up-regulated in largemouth bass fed HC diet (Fig. 2A). Western blot results indicated that HC diet effectively inhibited the phosphorylation of AMPK and promoted the phosphorylation of ACC (Fig. 2B). To further characterize the impact of high-glucose treatment on fatty acid metabolism, the phosphorylation of in vivo-validated AMPK and ACC were further analyzed in an in vitro model. Consistent with the in vivo results, HG treatment activated the phosphorylation of ACC and suppressed the phosphorylation of AMPK (Fig. 2C). Since Fgfr4 activation has been shown to initiate receptor tyrosine kinase signaling cascades, thereby activating the c-JUN and ERK signaling pathways [43], we further investigated whether HC diet might activate the MAPK signal pathways. Compared with CON diet, HC diet increased the phosphorylation of ERK, JNK and p38MAPK (Fig. 2D). As indicated in Fig. 2E, the phosphorylation of ERK, JNK and p38MAPK were up-regulated after HG treatment. Thus, both in vivo and in vitro experiments demonstrated that high-glucose treatment could promote fat synthesis by regulating the AMPK/ACC/SREBP-1 pathway and activate ERK, JNK and p38MAPK signal pathway.

Fig. 2
figure 2

High-glucose treatment disturbs fatty acid metabolism, bile acid metabolism and activates MAPK signal pathway in both largemouth bass and primary hepatocytes. A Relative expression of lipolysis/β-oxidation-related genes (cpt1, hsl, atgl and ppar-α), lipogenesis/proliferation-related genes (fas, acc1, srebp1 and ppar-γ), fatty acid transport -related genes (lpl, apob, apob100 and fabp1) and bile acid metabolism-related genes (cyp7a1, cyp8b1, fxr, rxrα, shp, hgmcr, fgfr4, fgf19, and fgf21) in the livers of largemouth bass (n = 4). B Relative expression of fatty metabolism-related proteins (p-ACC, ACC, p-AMPK, and AMPK) in the livers of largemouth bass (n = 3). C Relative expression of fatty metabolism-related proteins (p-ACC, ACC, p-AMPK, and AMPK) in cells (n = 3). D Relative expression of MAPK signal pathway (p-P38, P38, p-ERK, ERK, p-JNK, and JNK) in the livers of largemouth bass (n = 3). E Relative expression of MAPK signal pathway (p-P38, P38, p-ERK, ERK, p-JNK, and JNK) in cells (n = 3)

High-glucose treatment damages mitochondria function in both largemouth bass and primary hepatocytes

Transmission electron microscopy (TEM) was used to further visualize the livers' ultramicroscopic characteristics and structural attributes. Largemouth bass fed HC diet exhibited a reduced mitochondria number, an increased damaged mitochondria and a large glycogen accumulation (Fig. 3A). It was well-established that mitochondrial cytb transcription decreases with mitochondrial mass [44]. HC diet markedly decreased the expression of cytb in liver and elevated the expression of cytb in intestine, whereas the expression of the other tissues did not significantly change (Fig. 3B). Other than that, flow cytometry analysis revealed that HG treatment promoted cell apoptosis (Fig. 3C). Mito Tracker Red/Mito Tracker Green intensity ratio was significantly decreased in HG-treated cells, indicating an increase in the proportion of damaged mitochondria (Fig. 3D). The decline in mitochondrial membrane potential serves as an initial indicator of cell apoptosis. HG treatment resulted in a significant increase in the number of cells exhibiting depolarized mitochondria (green), indicating a decline in mitochondrial membrane potential in primary hepatocytes (Fig. 3E). To visualize the morphological changes in primary hepatocytes treated HG, the cellular ultramicroscopic structures were also view by TEM. The mitochondria of HG-treated cells exhibited significant damage, characterized by a decrease in the number of mitochondria, mitochondrial swelling with partial vacuolation (yellow arrow), an increase in the number of lipid droplets and an increase in the number of mitophagosomes (red arrow) (Fig. 3F). This was consistent with what we observed in vivo.

Fig. 3
figure 3

High-glucose treatment damages mitochondria function in both largemouth bass and primary hepatocytes. A The ultramicroscopic characteristics and structure of livers under electron microscopy. N: Nucleus; Red frame: Glycogen; Yellow arrow: Damaged mitochondria; M: Normal mitochondria; Scale bar, 2 μm. B Relative expression of cytb genes in different tissue of largemouth bass fed two different diets (n = 4). C Flow cytometry for apoptosis (n = 3). D Mito Tracker Red and Mito Tracker Green staining were measured using flow cytometry in cells (n = 3). E Mitochondrial membrane potential (MMP) analyzed by fluorescence microscope. F The ultramicroscopic characteristics and structure of cells under electron microscopy. N: Nucleus; L: Lipid droplet; M: Normal mitochondria; Red frame: Glycogen; Red arrow: Damaged mitochondria. Scale bar, 1 μm

High-glucose treatment affects mitochondrial morphology and mitochondrial dynamics in primary hepatocytes

In the in vitro experiments, mitochondrial fusion and fission were initially detected using Mito-Tracker Red staining, and morphological alterations were observed via confocal microscopy. Under normal physiological conditions, mitochondria typically exhibit large, elongated structures with distinct networks. However, our results revealed that a significant reduction in mitochondrial length after HG treatment, indicating that HG treatment facilitated mitochondrial fission and degradation (Fig. 4A). Furthermore, the results showed that HG treatment notably elevated mitochondrial reactive oxygen species (mitoROS) production (Fig. 4B). A mitochondrial marker, Tom20, was immunofluorescence stained to further assess mitochondrial morphology. The results indicated that mitochondrial length decreased and division increased after HG treatment, which further indicated that HG treatment would lead to mitochondrial fracture and promote mitochondrial fission (Fig. 4C). In addition, the expression of related proteins was determined by western blot, which indicated a decrease in Tom20 and an increase in Drp1 levels following HG treatment, suggesting that HG treatment may play an important role in mitochondrial fission (Fig. 4D).

Fig. 4
figure 4

High-glucose treatment affects mitochondrial morphology and mitochondrial dynamics in primary hepatocytes. A Morphological changes in the mitochondria of cells. Scale bar, 10 μm. B Intensity of mROS in cells. C Fluorescence photomicrograph of Tom20 examined in cells. Scale bar, 10 μm. D The expression of mitochondrial dynamics-related proteins (Tom20, Hsp60, and Drp1) in cells (n = 3)

High-glucose treatment promotes mitophagy and autophagy flow in primary hepatocytes

Mitochondrial dynamics, encompassing fusion and fission processes, alongside autophagy, function as critical quality control mechanisms for maintaining mitochondrial homeostasis. An array of proteins associated with mitophagy, notably Pink1 and Parkin, in addition to autophagic proteins such as LC3II, were investigated (Fig. 5A). In response to HG treatment, there was a marked upregulation of Pink1, Parkin, and LC3II, indicating that HG treatment may induce mitochondrial damage linked to Pink1/Parkin-mediated mitophagy. In order to further investigate the regulatory mechanism of HG treatment on autophagy, primary hepatocytes were transfected with a GFP-LC3 plasmid to quantify autophagosome formation. The results demonstrated that LG treated cells exhibited a limited number of autophagosomes, whereas a substantial increase in autophagosome formation was observed following HG treatment (Fig. 5B). Subsequently, laser confocal microscopy was employed to assess the colocalization of autophagosomes with lysosomes, mitochondria with lysosomes, and autophagosomes with mitochondria. The results demonstrated that HG treatment significantly enhanced GFP-LC3 lysosomal fusion (Fig. 5C), indicating increased autophagic flux. Furthermore, HG treatment promoted GFP-LC3 and Mito-RFP colocalization (Fig. 5D), while simultaneously inducing distinct colocalization between Mito-Tracker Green-labeled mitochondria and Lyso-Tracker Red-labeled lysosomes (Fig. 5E). Collectively, these findings confirm that HG treatment activates Pink1/Parkin-mediated mitophagy through the specified pathway in vitro.

Fig. 5
figure 5

High-glucose treatment promotes mitophagy and autophagy flow in primary hepatocytes. A Relative expression of mitophagy-related proteins (LC3II, Pink1 and Parkin) in cells (n = 3). B GFP-LC3 staining was used to assess autophagy. Scale bar, 10 μm. C The cells were stained using Lyso-Tracker Red (50 μmol/L; red) and transfected with GFP-LC3 (green) to detect colocalization of autophagosomes and lysosomes. Scale bar, 10 μm. D Mito-RFP (red) and GFP-LC3 (green) were co-transfected to detect the binding of autophagosomes to mitochondria. Scale bar, 10 μm. E The cells were stained using Lyso-Tracker Red (50 μmol/L; red) and Mito-Tracker Green (50 nmol/L; green) to detect the binding of lysosomes to mitochondria. Scale bar, 10 μm

High-glucose treatment causes mitochondrial apoptosis in primary hepatocytes

In order to elucidate the impact of high-glucose treatment on mitochondrial apoptosis. RT-PCR was used to quantify the expression of Bcl-2 family (bcl-2, bax and bad) and Caspase family (casp3, casp8, casp9, casp10 and p53). As expected, HG treatment boosted mitochondrial apoptosis, as evidenced by elevated expressions of bax, bad, casp3, and casp9, and decreased expressions of bcl-2 (Fig. 6A). P-P38 immunofluorescence was markedly enhanced both in the cytoplasm and nucleus in HG treated primary hepatocytes (Fig. 6B). Thus, we ensured that HG treatment significantly promoted the p38MAPK signal pathway. Primary hepatocytes were pretreated with different concentrations of SB203580 for 2 h and incubated with HG for another 48 h. The gene and protein levels of Casp3 were significantly downregulated by the addition of SB203580 (Fig. 6C and D). Furthermore, the gene expression levels of bcl-2, bax and casp3 were significantly altered in cells treated with HG in the presence of SB203580 (Fig. 6D). The results suggested that high-glucose treatment can induce mitochondrial apoptosis by activating the p38MAPK/bcl-2/Casp3 signaling pathway.

Fig. 6
figure 6

High-glucose treatment causes mitochondrial apoptosis in primary hepatocytes. A The expression levels of bcl-2, bax, bad, casp3 and casp9 of primary hepatocytes (n = 3). B immunofluorescence for p-P38. C Primary hepatocytes were pretreated with SB203580, expression levels of p-P38, P38 and Casp3 were analyzed using western blotting (n = 3). D Primary hepatocytes were pretreated with SB203580, expression levels of bcl-2, bax, bad, casp3 and casp9 were analyzed using RT-PCR (n = 3)

Sirt1 suppresses high-glucose-induced Pink1/Parkin-mediated mitophagy

Sirtuins‌ are a class of NAD+-dependent deacetylases that regulate metabolic processes [45]‌. In the Sirtuins family (sirt1–sirt7), HG treatment specifically downregulated sirt1‌ expression without affecting sirt2–sirt7 (Fig. 7A)‌. Phylogenetic analysis revealed that the NAD+-binding domain‌ and ‌catalytic core sequence‌ of Sirt1 are highly conserved across species, underscoring its evolutionary stability in energy metabolism regulation‌. Notably, the sirt1 gene of Micropterus salmoides exhibited closer phylogenetic proximity to carnivorous fish such as Micropterus dolomieu and Siniperca chuatsi, while forming a distinct evolutionary clade from Oncorhynchus mykiss and Ctenopharyngodon idella (Fig. 7B), suggesting a potential link between Sirt1-mediated metabolic regulation and dietary/ecological adaptations‌. Western blot‌ confirmed that HG treatment significantly reduced Sirt1 protein levels (Fig. 7C)‌. ‌Immunofluorescence staining‌ of exogenous Sirt1 revealed its predominant nuclear localization in primary hepatocytes under normal conditions, whereas HG treatment induced cytoplasmic Sirt1 puncta formation (Fig. 7D)‌. ‌Confocal microscopy‌ further demonstrated nuclear-to-cytoplasmic translocation of Sirt1 under high-glucose treatment, with cytoplasmic Sirt1 colocalizing with ‌autophagosomes‌ (GFP-LC3) and ‌lysosomes‌ (LysoTracker), implicating its role in modulating the autophagy-lysosome pathway to counteract metabolic stress (Fig. 7E)‌. ‌SiRNA-mediated Sirt1 knockdown‌ in HG treatment cells markedly upregulated key mitophagy proteins (‌Pink1‌, ‌Parkin‌, and ‌LC3II‌), indicating that Sirt1 deficiency enhances mitophagy activity (Fig. 7F)‌. Meanwhile, laser scanning confocal microscopy also revealed that high-glucose induces cytoplasmic translocation of endogenous Pink1, which colocalized with Sirt1, suggesting a regulatory role of Sirt1 in Pink1 expression (Fig. 7G).‌

Fig. 7
figure 7

Sirt1 suppresses high-glucose-induced Pink1/Parkin-mediated mitophagy. A Relative expression of Sirtuins family genes (sirt1-sirt7) in cells (n = 3). B The phylogenetic tree of Sirt1 protein base sequence was constructed by MEGA 3.1 software. C Relative expression of Sirt1 protein in cells (n = 3). D The distribution of exogenous Sirt1 was detected by immunofluorescence. Scale bar, 10 μm. E The cells were co-transfected with GFP-LC3 (green) and Flag-Sirt1 (purple), and stained with Lyso Tracker Red (red) to visualize the binding of autophagosomes, lysosomes, and the Sirt1 protein. Scale bar, 10 μm. F Effects of si-Sirt1 transfection on the protein expression levels of LC3II, Pink1, Parkin and Sirt1 (n = 3). G The binding of endogenous Sirt1 to Pink1 protein was detected by confocal laser. Scale bar, 10 μm

Discussion

The hepatosomatic index (HSI), recognized as a key indicator for evaluating liver size and overall health in fish [46, 47], showed significant elevation in grouper (Epinephelus akaara) [48] and largemouth bass fed HC diet, with similar changes observed in VSI. It was well known that carnivorous fish typically alleviate high glucose loads by enhancing glycolysis (involving hexokinase, PFK-1, and pyruvate kinase) and activating gluconeogenesis [49], this study's PAS staining, electron microscopy, and enzymatic analyses revealed excessive hepatic glycogen accumulation, which may due to dietary carbohydrate levels exceeding metabolic regulatory capacity. When glucose metabolism becomes overloaded, the AMPK/SREBP-1c axis drives hyperactivated lipogenesis, converting excess carbon sources into stored triglycerides [50]. Subsequent cellular experiments revealed a greater accumulation of lipid droplets in cells treated with high-glucose, further corroborating that a high-carbohydrate diet induced an over-accumulation of lipid in the liver. A similar phenomenon was also observed in Nile tilapia (Oreochromis niloticus) [51], gibel carp (Carassius gibelio) [52], and rainbow trout (Oncorhynchus mykiss) [53]. Additionally, prior research on largemouth bass fed an 18% carbohydrate diet demonstrated reduced serum insulin levels, downregulated hepatic insulin gene expression, and upregulated insulin resistance-related genes [41], indicating that high-glucose-induced liver injury may correlate with impaired insulin synthesis.

Lipid accumulation is well recognized to be mediated by AMPK and SREBP. The activated form of SREBP-1 is responsible for the upregulation of gene expression related to lipid biosynthesis, leading to an increase in lipid droplet formation and overall lipid content [54]. It has been shown that the high-glucose treatment of hepatocytes suppresses the expression of SREBP-1 target genes by AMPK by repressing its cleavage and translocation intranuclearly [55]. Interestingly, HC diet could increase lipid production by inhibiting AMPK, causing acc, srebp1, and fas to be increased. Additionally, FABP1 functions as a transport agent, a transporter, and a metabolic regulator of fatty acids [56]. This study found that HC diet significantly upregulated apob, apob100 and fabp1 in largemouth bass that had fatty liver, indicating that it promoted the absorption and transport of fatty acids and the synthesis of triglycerides. Notably, the HC diet exerted limited effects on key genes regulating fatty acid oxidation (cpt1, ppar-α), indicating that lipid accumulation predominantly originates from enhanced synthesis rather than suppressed breakdown.

Current findings indicated that liver damage and lipid accumulation in largemouth bass can be induced by HC diet. Similar phenomena have been reported in other fish species [57,58,59]. Liver injury is primarily attributed to the initial toxic effects of excessive lipids [60]. Mounting evidence shows that the accumulation of both cholesterol and triglycerides may result in lipotoxicity [61]. The present study revealed that the HC diet disrupts cholesterol homeostasis by downregulating critical bile acid synthesis genes (cyp7a1, cyp8b1) and the regulatory gene shp. This suppression impedes the conversion of cholesterol to bile acids, causing pathological accumulation of free cholesterol. Beyond that, abnormal accumulation of free cholesterol is believed to compromise the integrity of mitochondrial and endoplasmic reticulum membranes, thereby exacerbating promote mitochondrial oxidative damage and endoplasmic reticulum stress, and finally induce and aggravate liver injury [62]. ‌Although hepatic hmgcr (the rate-limiting enzyme for cholesterol synthesis) expression remained unchanged, inhibition of the cholesterol breakdown pathway was still a central contributor to metabolic dysregulation. Similarly, we observed that the HC diet did not alter hepatic fxr or rxrα mRNA expression, aligning with previous research [63]. This might be because FXR primarily functions in the liver-gut axis, and the HC diet has a less effect on regulating fxr expression in the liver.‌

Mitochondria serve as the central hub for glucose and lipid metabolism, functioning as both sensors and executors of metabolic signals [64]. Research indicates that their homeostasis (fusion/fission) directly responds to cellular metabolic demands, while in mammals, disruption of this equilibrium has been identified as a critical factor in high-glucose/high-fat-induced hepatic injury [65, 66]. In fish, it has been reported that high-fat diet in Pelteobagrus pelteobagrus can activate mitochondrial biogenesis, promote mitochondrial fusion and induce oxidative stress, consequently leading to increased lipid accumulation [31]. Prolonged consumption of high-carbohydrate diet has been associated with the induction of oxidative stress, which may impair mitochondrial respiratory chain activity in the liver of rainbow trout [67] and Megalobrama amblycephala [68]. In this study, HG treatment upregulated the fission protein Drp1 and downregulated the fusion protein Tom20, triggering mitochondrial fragmentation. The fission process was accompanied by a burst of mitoROS, which directly suppresses respiratory chain function. Excessive mitoROS activated the Pink1/Parkin pathway to initiate mitophagy, yet sustained activation exacerbates the cellular energy crisis. Ultimately, the AMPK/ACC/SREBP-1 axis-driven enhanced lipogenesis, cholesterol dysmetabolism, and mitochondrial fission-autophagy cascade collectively drive liver injury.

In mammals, Sirt1 serves as a central hub in nonalcoholic fatty liver disease (NAFLD)/nonalcoholic steatosis (NASH) pathogenesis by integrating the regulation of lipid and glucose metabolism with inflammatory responses [69, 70]. It has reported that Sirt1 suppresses fatty acid and triglyceride synthesis by deacetylating and inhibiting the transcription factor SREBP1c, while concurrently activating the AMPK pathway to promote fatty acid oxidation, thereby alleviating hepatocellular steatosis [71].‌ ‌Furthermore, Sirt1 mitigates hepatic inflammation and fibrosis by inhibiting NF-κB pathway activation, which reduces the release of pro-inflammatory cytokines such as TNF-α and IL-6 [72]. It also enhances antioxidant capacity to counteract oxidative damage characteristic of NASH [73]. In recent years, research has begun to uncover the significant role of Sirt1 in the metabolic regulation of fish species. Resveratrol, known as a Sirt1 activator, demonstrated the ability to alleviate oxidative stress and inflammation in LPS-injected gibel carp by boosting the Sirt1/PGC-1α and Pink1/Parkin pathways [74]. In tilapia fed high-carbohydrate diets, activation of the Sirt1/AMPK signaling pathway effectively alleviated high-glucose-induced metabolic syndrome [59]. Further investigations showed that Sirt1 and FoxO1 work together to manage glucose metabolism in the kidney and intestinal tissues of blunt snout bream under conditions of high-glucose intake [75]. Recent studies also revealed that Sirt1 attenuates hepatic lipotoxicity by suppressing the activation of the JNK signaling pathway [76]. ‌This study demonstrates that high-glucose specifically suppresses sirt1 expression (but not other sirtuins) and induces its nucleocytoplasmic translocation-a mechanism that corresponds to Xu et al.'s [77] report of nuclear Sirt1 degradation via LC3 binding and the autophagosome-lysosome pathway during aging. In mammals, Sirt1 governs early-stage NAFLD progression through metabolic and inflammatory regulation, whereas the Pink1/Parkin pathway serves as the central executor of mitochondrial quality control. Here, we reveal Sirt1 primarily functions as an energy effector: high glucose-induced nucleocytoplasmic translocation promotes its binding to autophagosomes, precipitating mitochondrial dysfunction and consequent disruption of hepatocellular energy homeostasis. Additionally, the evolutionary conservation of Sirt1 across species, combined with its lineage-specific branching patterns, underscores its central role in metabolic regulation.

Conclusions

To conclude, our results demonstrate first-time evidence that the AMPK-mediated ACC/SREBP-1 pathway contributes to lipid accumulation in largemouth bass, both in vivo and in vitro. Simultaneously, this study is also the first to establish a primary hepatocyte injury model using high-glucose, confirming its role in mitochondrial damage and glycolipid accumulation. Moreover, we uncovered a correlation between mitophagy and high-glucose treatment, and determined that high-glucose compromises Sirt1's metabolic protective function by suppressing its expression and altering its subcellular localization (nucleus-to-cytoplasm translocation), thereby activating Pink1/Parkin-mediated mitophagy.

Data Availability

All data generated or analyzed during this study are included.

Abbreviations

  • acadm :: Acyl-CoA dehydrogenase medium
  • acc1 :: Acetyl-CoA carboxylase 1
  • AMPK:: Adenosine 5 ‘-monophosphate (AMP)-activated protein kinase
  • apob :: Apolipoprotein B
  • atgl :: Adipose triglyceride lipase
  • BA:: Bile acid
  • CCK8:: Cell counting kit-8
  • CON:: Control
  • cpt1 :: Carnitine palmitoyltransferase 1
  • cyp7a1 :: Cholesterol 7-hydroxylase
  • cyp8b1 :: Sterol 12-hydroxylase
  • cytb :: Cytochrome b
  • EDTA:: Ethylenediaminetetraacetic acid
  • ef-1α :: Elongation factor 1α
  • ennp1 :: Phosphodiesterase 1
  • fabp1 :: Fatty acid binding protein 1
  • fas :: Fatty acid synthase
  • FBS:: Fetal bovine serum
  • fgfr4 :: Fibroblast growth factor receptor 4
  • FITC:: Fluorescein isothiocyanate
  • FXR:: Farnesoid X receptor
  • gsy2 :: Glycogen synthase 2
  • HC:: High-carbohydrate
  • HG:: High-glucose
  • HSI:: Hepatosomatic index
  • hsl :: Hormone-sensitive lipase
  • Keap1:: Kelch-like ECH-associated protein 1
  • LG:: Low glucose
  • lpl :: Lipoprotein lipase
  • MAPK:: Microtubule-associated protein kinase
  • mitoROS:: Mitochondrial reactive oxygen species
  • MMP:: Mitochondrial membrane potential
  • NAFLD:: Nonalcoholic fatty liver disease
  • NASH:: Nonalcoholic steatosis
  • NIH:: National institutes of health
  • nqo1 :: NAD(P)H oxidoreductase 1
  • nrf2 :: Nuclear factor erythroid 2-related factor 2
  • PAS:: Periodic acid-schiff
  • PBS:: Phosphate-buffered saline
  • PI:: Propidium iodide
  • ppar-α :: Peroxisome proliferator-activated receptor alpha
  • ppar-γ :: Peroxisome proliferator-activated receptor γ
  • qRT-PCR:: Quantitative real-time PCR
  • ROS:: Reactive oxygen species
  • shp :: Small heterodimer partner
  • srebp1 :: Sterol regulatory element-binding protein 1
  • TEM:: Transmission electron microscopy
  • VSI:: Viscerosomatic index

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Acknowledgements

Funding

This research was supported by Project of National Natural Science Foundation of China (32172982), Project of Science and Technology of Guangdong Province (2021B0202050002), Project of Science and Technology of Guangdong Province (2023A1515012627). National Natural Science Foundation of China (32303014), Guangdong Basic and Applied Basic Research Foundation (2024A1515010444). We also thank the Home for Researchers ( www.home-for-researchers.com ).

Ethics Declaration

Ethics approval and consent to participate

The National Institutes of Health (NIH) guidelines for laboratory animal care and use were strictly followed during all experimental procedures. The study protocol was approved by Experimental Animal Ethics Committee of Sun Yat-Sen University. The clinical trial number was SYSU-LS-IACUC-2024-0020.

Consent for publication

No applicable.

Competing interests

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

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