Male C57BL/6 J mice were purchased from SLAC Laboratory Animal Central (Changsha, China). As suggested by the animal welfare protocol, all efforts were made to minimize animal suffering and to use only the number of animals necessary to produce reliable scientific data. All animals were housed in a climate-controlled room (temperature, 25 ± 2 °C; relative humidity, 45–60%; lighting cycle, 12 h light/dark cycle with light provided 08:00–20:00) and had free access to food and drinking water during the duration of the experiments.

After 1 week of adaptation, C57B/6 J male mice aged at 8 weeks were randomly divided into the CON group (received control diet, n = 10), the HF group (received HF diet, n = 12), and the HUR group (received high-fat diet and supplemented with 0.4 mg/mL uridine in drinking water for the last 4 weeks, n = 12) for 10 weeks (17). The schematic diagram of the animal group and treatment is shown in Figure 1A. The consumption of feed and drinking water was measured and recorded daily to calculate the average feed intake and water intake. The drinking water was renewed every 2 days. An average of 4.27 ± 0.57 mL/d of uridine water was drunk per mouse in the HUR group (Figure 1C). The control diet consisted of 10% (kcal%) fat, 20% (kcal%) protein, and 70% (kcal%) carbohydrate (D12450B; Research Diets, Inc., New Brunswick, NJ, USA); the HF diet consisted of 60% (kcal%) fat, 20% (kcal%) protein, and 20% (kcal%) carbohydrate (D12492; Research Diets, Inc.). Uridine (purity ≥ 99.90%) was provided by Meiya Pharmaceutical Co., Ltd. (Hangzhou, China). At the end of the experiment, the mice were sacrificed after being deeply anesthetized. Blood was collected from the mice by retro-orbital bleeding, centrifuged for 10 min at 16,000 × g, frozen on dry ice, and stored at −80 °C. The liver, subcutaneous, and intra-abdominal white adipose tissues (including the epididymal, perirenal, and mesenteric white adipose tissues) were weighed. The liver was snap-frozen in liquid nitrogen and then stored at −80 °C for further analysis.

Mouse duct cells were isolated from mouse liver using a digestion solution composed of DNase-1 (100 μg/mL; Sigma-Aldrich, St. Louis, MO, USA), collagenase type XI (125 μg/mL; Sigma-Aldrich, St. Louis, MO, USA), dispase (125 μg/mL; Thermo Fisher Scientific, Shanghai, China), and 1% FBS in DMEM. Following isolation, bile ducts were embedded in Matrigel (BD Bioscience, San Diego, CA, USA) and seeded in a 24-well plate. Cultures were maintained in expansion medium, refreshed every 2–3 days, and passaged weekly at a 1:2–1:4 ratio. The expansion medium consisted of the medium previously reported (17).

Oleic acid (OA) and palmitic acid (PA) (Sigma-Aldrich, St. Louis, MO, USA) were complexed with bovine serum albumin (BSA) (Sigma-Aldrich, St. Louis, MO, USA) and sterile-filtered. The free fatty acid (FFA) mixture was dissolved in the medium at a final concentration of 600 μM (OA: PA = 2:1) to induce lipid accumulation according to our previous research (17).

Mouse liver organoids collected from 12-well plates were digested into single cells using trypsin, followed by suspension in complete culture medium and centrifugation. The Annexin V-FITC/PI apoptosis detection kit was used to detect the apoptosis of uridine and free fatty acid preparation-treated mouse liver organoids according to the manufacturer’s instructions (BD Biosciences, San Diego, CA, USA). Briefly, cells were resuspended in 195 μL of Annexin V-FITC binding buffer. Then, 5 μL of Annexin V-FITC reagent was added and gently mixed, followed by incubation on ice for 10 minutes in the dark. Subsequently, 5 μL of propidium iodide (PI) staining solution was added, and the cells were further incubated for 15 minutes at room temperature in the dark. Finally, 200 μL of binding buffer was added prior to flow cytometry analysis. The ratio of apoptotic cells was determined using a FACS instrument (BD FACSCanto II, San Diego, CA, USA). Cells exhibiting PI-negative and Annexin V-FITC-positive staining were considered to be in the early stages of apoptosis, whereas cells with positivity for both PI and Annexin V-FITC were identified as late apoptotic.

The concentrations of serum triglycerides (TG), total cholesterol (TC), and high-density lipoprotein (HDL) were determined by an Automated Biochemistry Analyzer (Synchron CX Pro, Beckman Coulter, Fullerton, CA, USA) according to the commercial kits and manufacturer’s instructions (Beijing Chemclin Biotech Co., Ltd., Beijing, China). The serum leptin was assayed using mouse leptin ELISA kits (CUSABIO Co., Ltd., Wuhan, China) according to the manufacturer’s instructions.

Liver tissue samples were isolated after mice were killed by cervical dislocation, rinsed with 0.9% NaCl, snap-frozen in liquid nitrogen, and stored at −80 °C until further analysis. The concentrations of liver TG and total protein were determined by a multimode reader (Tecan Trading AG, Switzerland) based on the commercial assay kits provided by Applygen Technologies Inc. (Beijing, China) for the tissue TG content assay kit and the trace protein quantification kit (BCA method).

Total RNA was isolated from the liver tissue using Trizol Reagent (Invitrogen, Carlsbad, CA, USA) according to the manufacturer’s instructions. RNA concentrations and A260/A280 ratios were determined on a NanoDrop ND-1000 spectrophotometer (Thermo Fisher Scientific, Wilmington, DE, USA). The total RNA was reverse-transcribed into cDNA using the PrimeScript™ RT reagent kit (Takara Biomedical Technology, Japan). Quantitative real-time RT-PCR (qRT-PCR) was performed with SYBR Green I Dye (Thermo Fisher Scientific, New York, USA) using the LightCycler 480II real-time PCR system (Roche, Basel, Switzerland). The PCR cycling conditions were as follows: 95 °C for 5 min and 98 °C for 2 min, followed by 40 cycles of 5 s at 98 °C, 5 s at 60 °C, 10 s at 95 °C, and a final step of 1 s at 65 °C. The primers for the genes are listed in Table 1. The gene expression results were expressed as the mean relative mRNA level (18).

Samples that contained 100 mg of liver tissue powder were cryo-weighed, and 80% methanol (v/v H2O) was added at a ratio of 300 μL dissolver/100 mg tissue. The samples were shaken for 20 min at room temperature and centrifuged at 14,000 × g for 15 min at 4 °C. Supernatants were filtered (0.2 μm) and stored at −20 °C until the next analysis. LC–MS/MS analyses were performed using a Thermo Vanquish UHPLC with an Accucore Vanquish C18 column (50 × 2.0 mm, 1.5 μm) (19). The mobile phases of solutions A and B were composed of 95% acetonitrile with 0.1% formic acid and 10 mM ammonium acetate and 50% acetonitrile with 0.1% formic acid and 10 mM ammonium acetate in the positive mode. The flow rate was at 300 μL/min, which consisted of 2% B for 1 min, 50% B for 16 min, 50% B for 0.5 min, 2% B for 0.5 min, and 2% B for 2 min.

For the mass spectrometric assay, a mass spectrometer detector QE HF-X (Thermo Fisher Scientific Inc., Massachusetts, CA, USA) was used to analyze the metabolite ions. Electrospray ionization (ESI) settings are as follows: spray voltage: 3.2 kV; sheath gas flow rate: 35 arb; aux gas flow rate: 10 arb; and capillary temperature: 320 °C. Estimation was performed in positive and negative modes with a scan time of 20 min. MS/MS secondary scan is a data-dependent scan. Spectral data from all peaks were accumulated in the range of 100–1,500 m/z. Molecular formulas were determined by using Xcalibur™ software (Thermo Fisher Scientific, New York, USA), and the identification of constituent substances was performed by Compound Discoverer software, which was confirmed by searching online libraries of mzVault, ChemSpider, and mzCloud.

Processed datasets were analyzed by multivariate statistical analysis using SIMCA+13.0 (Umetrics, Umea, Sweden). Partial least squares discriminant analysis (PLS-DA) was used to visualize discrimination among samples. The quality of PLS-DA models was evaluated by R²Y and Q²; the goodness of fit measure was quantified by R²Y; and the predictive ability was indicated by Q². Validation and reliability of PLS-DA models were rigorously confirmed by a permutation test (n = 200). For identification of metabolites contributing to the discrimination, the intensity differences of metabolites with a variable importance in the projection value (VIP) > 1.0, showing high relevance for explaining the differences among sample groups, were analyzed using t-tests (p < 0.05). Metabolic pathway analysis was employed using the MetaboAnalyst, and metabolites that showed significant change were mapped to the Kyoto Encyclopedia of Genes and Genomes (KEGG) pathways.

The data were expressed as mean ± standard error of mean (SEM), and the results were analyzed using one-way ANOVA, followed by the LSD test using SPSS 22.0. Differences were considered statistically significant at p < 0.05.

During the experiment, the average food and water intake of the HF group was significantly lower than that of the CON group (p < 0.05; Figures 1B, C). However, the water intake was significantly increased in the HUR group compared with the HF group (p < 0.05). The body weight of mice during 10 weeks is shown in Figure 1D, and the initial body weight of mice was approximately 22 g. The body weights of mice both in the HF group (32.7 g) and the HUR group (32.5 g) exhibited a significant increase of over 20% compared to the CON group (26.2 g) at the 6th week (p < 0.01; Figure 1E), confirming the successful induction of obesity. Compared with the HF group (40.3 g), the final body weight at the 10th week of mice in the HUR group (38.8 g) showed a significant decrease (p < 0.05; Figure 1F). Obesity induces excessive storage of adipose tissue and leads to excessive accumulation of fat in other tissues, thereby posing a risk of developing metabolic disorders (20). A previous study showed that postprandial uridine metabolism was altered by obesity (21). The supplementation of uridine during both day and night significantly reduced body weight gain in the HF diet-fed mice (15). In the present study, supplementing with uridine through drinking water throughout the day also showed a decrease in body weight in HF diet mice. It may be suggested that the effect of uridine on the body weight of HF diet mice was not affected by the time of administration.

Obesity induced by HF diet feeding in mice is typically characterized by dysregulated lipid metabolism and impaired liver function (22). The weight of the liver, subcutaneous white adipose tissues, and intra-abdominal white adipose tissues of mice in the HF group was higher than that in the CON group (p < 0.01; Figures 1G–I). As the primary site for carbohydrate and lipid biosynthesis, the liver plays a central role in regulating systemic glucose and lipid flux (23, 24). Notably, the HUR group presented a lower liver weight and intra-abdominal adipose tissue weight than the HF group (p < 0.05), although these parameters remained significantly different from those in the CON group (p < 0.05). These results indicated that uridine supplementation can ameliorate adipose tissue accumulation induced by HF feeding, but it is not sufficient to fully restore it to the same level as the CON group.

Mouse liver organoids were successfully cultured from mouse liver samples (Figure 2A). Following enzymatic digestion, bile duct fragments were observed in the supernatant. After 14 h of culture, biliary epithelial cells began to reorganize, forming spherical structures. By 72 h, complete reorganization resulted in the emergence of organoids displaying characteristic spherical morphology within Matrigel. The liver organoids could be stably cultured in expansion medium from passage 1 to passage 10, maintaining similar morphology with occasional folding and budding. After passaging, organoids demonstrated rapid proliferation and showed strong proliferative activity by 5-Ethynyl-2´-deoxyuridine (EDU) staining. The mouse liver organoids were seeded into 12-well plates with a density of 1 × 10⁵ cells/well for 24 h and incubated with FFA for 24 h and then with uridine for 48 h. The results of apoptosis analysis using flow cytometry showed that FFA treatment did not affect the apoptosis rate of liver organoid cells (Figures 2B,C). However, the addition of uridine alongside FFA treatment significantly increased the proportions of total apoptosis and late apoptosis while decreasing the proportion of early apoptosis (p < 0.05). Liver homeostasis is achieved by a tightly regulated steady state in cell turnover involving proliferation and apoptosis of hepatocytes. Increased numbers of apoptotic hepatocytes and apoptosis-associated degradation products were detected in NASH patients (25). Hepatocytes and bile duct cells are the first hepatic cell types to enter the cell cycle and proceed to mitosis. Proliferation subsides by days 5–7, while during the peak proliferation period, triglycerides accumulate in hepatocytes for 2–3 days (26). A previous study reported that 1 mM uridine significantly suppressed lipid accumulation induced by FFA exposure in the liver organoids (17). Similarly, uridine promotes the renewal of intestinal cells, and 50 mM uridine significantly promotes apoptosis of IPEC-J2 cells (27). The 1 mM and 10 mM uridine also inhibit the germination of intestinal organoids and suppress the stemness of intestinal stem cells, and this effect may be mediated through the mTOR pathway (28). The uridine exhibited higher distribution of G1/M phase of the cell cycle at 400 M compared with 0 M and reduced S-phases of the cell cycle compared with 0 and 100 M (p < 0.05) (29). It indicated that uridine is involved in the regulation of stem cell stemness and cell proliferation, regulating cell renewal.

Typical dyslipidemia of obesity includes elevated serum TG and TC (30, 31). In addition, the obesity level was consistent with the change of serum leptin, which was significantly increased after 12 weeks of HF feeding (32). As shown in Figure 3, consumption of an HF diet led to higher serum TG, TC, HDL, and leptin content than that in the CON group (p < 0.05). However, this increase was significantly decreased by the uridine supplement in obese mice (p < 0.05). Fasting-induced increases of plasma uridine occurred concurrently with high levels of lipolysis (13). Activation of uridine production in adipocytes can promote lipolysis, which suggests that uridine helps to alleviate obesity-related lipid metabolic disorder (14). Consistently, it has been reported that through the elevation of circulating uridine, Lactobacillus MR1 alleviates high-carbohydrate diet-induced hepatic lipid accumulation and oxidative stress (33). Similarly, a uridine supplement decreased hepatic lipid, serum glucose, triglyceride, and cholesterol in Nile tilapia with a high-carbohydrate diet (34).

As shown in Figure 4, the HF diet increased the hepatic TG content of mice, and activated fatty acid translocase /cd36 (Fat/cd36) gene expression but inhibited low-density lipid receptor (Ldlr) mRNA levels when compared with those in the CON group (p < 0.05). Conversely, uridine supplementation significantly decreased the liver TG content and Fat/cd36 expression and increased Ldlr expression in the liver of obese mice from the HUF group (p < 0.05). Oil red staining also showed that mice in the HF group presented greater lipid droplets in the liver when compared with those in the CON group, and uridine supplementation weakened the hepatic lipid droplet production in the HF group (Figure 4E). Importantly, uridine alleviated this typical dyslipidemia of obese mice after 4 weeks of administration, which was consistent with previous reports that uridine supplementation affected liver lipid accumulation (10, 35–37). The liver is involved in the uptake and secretion of fatty acids and TG (38). Fat/cd36 is an important regulator of tissue free fatty acid (FFA) uptake from plasma and increases in the livers of obese subjects (23). Hepatocyte-specific deletion of Cd36 protects against liver lipid accumulation induced by the HF diet in mice (39). In addition, Ldlr mRNA expression levels were decreased significantly following HF in the liver (40). Therefore, uridine supplementation may restore liver TG by regulating the expression of Fat/cd36 and Ldlr genes in obese mice fed with an HF diet.

The relative gene expression related to pyrimidine metabolism in the liver is noted (Figure 5). Consumption of an HF diet significantly decreased dihydroorotate dehydrogenase (Dhodh), uridine monophosphate synthetase (UMPS), uridine phosphorylase 2 (Upp2), ribonucleoside-diphosphate reductase subunit M2 (Rrm2), and thymidine kinase 1 (Tk1) gene expression levels when compared with those in the CON group (p < 0.05). Especially, the expression of Dhodh, Upp2, Rrm2, and Tk1 was higher in the HUR group than in the HF group (p < 0.05). Particularly, pyrimidine metabolism was abnormal in obese mice, such as low expression of UDP synthesis and conversion-related genes in the hypothalamus (41). Similarly, plasma uridine was significantly elevated after fasting for 4–24 h in healthy mice, but not in HF diet mice (13). Consistent with pyrimidine metabolism abnormalities observed in obese mice, our study revealed decreased expression of hepatic genes involved in pyrimidine metabolism and de novo synthesis, which was ameliorated by uridine supplementation, further supporting the regulatory role of uridine in metabolic disturbances induced by HFD.

A total of 443 known metabolites, including 239 in the positive ion mode and 205 in the negative ion mode, were identified in the liver of mice using LC-MS metabolomic analysis. The clustering analyses based on partial least squares discriminant analysis (PLS-DA) were used to discriminate the metabolic profiles among groups (Figure 6). Subsequently, the permutation test verified that the model was not “over-fitting.” The samples in the CON, HF, and HUR groups were separated in both positive and negative ion modes, indicating that the overall metabolic state of the mice changed after treatment. For further analysis, the candidate markers were selected by examining the volcano plot and considering a fold-change (FC) threshold of 1.5, VIP > 1, and p-value less than 0.05 (Figure 7). There were 88 differential metabolites between the CON and HF groups, 100 between the CON and HUR groups, and 37 between the HF and HUR groups. To further understand the metabolic pathways in the HUR group mice (Figure 8), metabolic pathway analyses of 37 different enriched metabolites were performed using MetaboAnalyst 5.0.¹ The HUR group metabolites revealed several metabolism pathways, including α-linolenic acid metabolism, linoleic acid metabolism, arachidonic acid metabolism, and purine metabolism, compared with the HF group.

Metabolomics measures metabolic changes in the liver, and fatty acids (α-linoleic acid and arachidonic acid) can reduce hepatic steatosis and improve liver function in HF diet-fed mice (42, 43). In addition, dietary nucleotides affected levels of linoleic and arachidonic acid in rats with liver cirrhosis (44). Similarly, uridine supplements increased the percentage of polyunsaturated fatty acids in the liver (15). In the current study, the levels of α-linolenic acid and 12(S)-HPETE were higher in the HUR group than in the HF group, and uridine restored the decrease of arachidonic acid caused by HF diet feeding. It revealed that unsaturated fatty acids play an important role in the regulation of liver lipid metabolism disorder by uridine. Testosterone is a metabolic hormone that regulates the expression of key targets of lipid and glucose metabolism and may reduce fat deposition in the liver (45). In this study, the HF diet decreased the testosterone level in the liver, and uridine restored it to normal levels. Uridine metabolism affected the levels of adenosine; the deletion of uridine phosphorylase (an enzyme that catalyzes the phosphorolysis of uridine into uracil) increased adenosine to a lesser extent (46). Notably, hypoxanthine (an adenosine metabolite) showed impaired rhythmicity due to the decrease in peak concentration at ZT8 in the HF group (47). The results showed that uridine alleviated the decrease of adenosine and hypoxanthine induced by the HF diet. It may suggest that there is an interactive effect on the metabolism of different kinds of nucleosides or nucleobases. In addition, the levels of chenodeoxyglycocholic acid and taurodeoxycholic acid were altered by nucleotide supplementation in alcohol-treated rats (48). The results from the present study also showed that uridine affected the level of taurochenodeoxycholic acid. Further mechanism research can be conducted using the mouse liver organoid model, such as functional validation of differential metabolites and study of nucleotide metabolism pathways discovered in this experiment.