
==== Front
Cell Mol Gastroenterol Hepatol
Cell Mol Gastroenterol Hepatol
Cellular and Molecular Gastroenterology and Hepatology
2352-345X
Elsevier

S2352-345X(24)00134-6
10.1016/j.jcmgh.2024.101379
101379
Original Research
Microbial-derived Urolithin A Targets GLS1 to Inhibit Glutaminolysis and Attenuate Cirrhotic Portal Hypertension
Li Rui 1∗
Liu Zhile 1∗
Huang Wensou 1∗
Guo Yongjian 1∗
Xie Chan 2
Wu Hongmei 3
Liu Jianxin 1
Hong Xiaoyang 1
Wang Xiaobin 1
Huang Jingjun 1
Cai Mingyue 1
Guo Zhaoxiong 1
Liang Licong 1
Lin Liteng linliteng@yeah.net
1∗
Zhu Kangshun zhksh010@163.com
1∗
1 Laboratory of Interventional Radiology, Department of Minimally Invasive Interventional Radiology and Interventional Cancer Center, the Second Affiliated Hospital, Guangzhou Medical University, Guangzhou, China
2 Department of Infectious Diseases, the Third Affiliated Hospital, Sun Yat-sen University, Guangzhou, China
3 Department of Pathology, Guangdong Provincial People’s Hospital (Guangdong Academy of Medical Sciences), Southern Medical University, Guangzhou, China
∗ Correspondence Address correspondence to: Prof Kangshun Zhu, Guangzhou Medical University, Department of Minimally Invasive Interventional Radiology, 250 East Changgang Road, Guangzhou 510260, China. zhksh010@163.com
∗ Dr Liteng Lin, Guangzhou Medical University, Department of Minimally Invasive Interventional Radiology, 250 East Changgang Road, Guangzhou 510260, China. linliteng@yeah.net
∗ Authors share co-first authorship.

20 7 2024
2024
20 7 2024
18 4 10137922 12 2023
16 7 2024
© 2024 The Authors
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/by-nc-nd/4.0/).
Background & Aims

Cirrhotic portal hypertension (CPH) is the leading cause of mortality in patients with cirrhosis. Over 50% of patients with CPH treated with current clinical pharmacotherapy still present variceal bleeding or sometimes death owing to insufficient reduction in portal pressure. Elevated intrahepatic vascular resistance (IHVR) plays a fundamental role in increasing portal pressure. Because of its potent effect in reducing portal pressure and maintaining normal portal inflow to preserve liver function, lowering the IHVR is acknowledged as an optimal anti-CPH strategy but without clinical drugs. We aimed to investigate the protective effect of microbial-derived Urolithin A (UroA) in IHVR and CPH.

Methods

Carbon tetrachloride or bile duct ligation surgery was administered to mice to induce liver fibrosis and CPH. 16S rRNA gene sequencing was used for microbial analysis. Transcriptomics and metabolomics analyses were employed to study the host and cell responses.

Results

UroA was remarkably deficient in patients with CPH and was negatively correlated with disease severity. UroA deficiency was also confirmed in CPH mice and was associated with a reduced abundance of UroA-producing bacterial strain (Lactobacillus murinus, L. murinus). Glutaminolysis of hepatic stellate cells (HSCs) was identified as a previously unrecognized target of UroA. UroA inhibited the activity of glutaminase1 to suppress glutaminolysis, which counteracted fibrogenesis and contraction of HSCs and ameliorated CPH by relieving IHVR. Supplementation with UroA or L. murinus effectively ameliorated CPH in mice.

conclusions

We for the first time identify the deficiency of gut microbial metabolite UroA as an important cause of CPH. We demonstrate that UroA exerts an excellent anti-CPH effect by suppressing HSC glutaminolysis to lower the IHVR, which highlighted its great potential as a novel therapeutic agent for CPH.

Graphical abstract

Keywords

Glutamine Metabolism
Gut Microbiota
Innovative Therapy
Portal Hypertension
Urolithin A
Abbreviations used in this paper

ABX antibiotics

aHSC activated hepatic stellate cell

α-KG α-ketoglutaric acid

ALT alanine aminotransferase

ANOVA analysis of variance

α-SMA α-smooth muscle actin

BCA bicinchoninic acid

BDL bile duct ligation

BPTES Bis-2-(5-phenylacetamido-1,3,4-thiadiazol-2-yl) ethyl sulfide

CCK-8 cell counting kit-8

CCl4 carbon tetrachloride

Col1α1 collagen type 1α1

CPH cirrhotic portal hypertension

DEG differentially expressed gene

ECL enhanced chemiluminescence

ECM extracellular matrix

EdU 5-Ethynyl-2’-deoxyuridine

FBS fetal bovine serum

GDH glutamate dehydrogenase

GLS glutaminase

GLS1 glutaminase1

GOT2 glutamic-oxaloacetic transaminase 2

GPT2 glutamic-pyruvic transaminase 2

HE hematoxylin and eosin

HSC hepatic stellate cell

IL-1β interleukin-1β

IHVR intrahepatic vascular resistance

LC-MS/MS liquid chromatography with tandem mass spectrometry

PBS phosphate buffered saline

PDGF platelet-derived growth factor

p-MLC2 phospho-myosin light chain 2

RT-PCR reverse transcription polymerase chain reaction

SEM standard error of mean

SPR surface plasmon resonance

TCA tricarboxylic acid cycle

TGF-β transforming growth factor-β1

TIPS transjugular intrahepatic portosystemic shunt

TNF-α tumor necrosis factor-α

TIMP-1 tissue inhibitor of metalloproteinase-1

UPLC-MS/MS ultra-performance liquid chromatography coupled to tandem mass spectrometry

UroA Urolithin A

WT wild-type
==== Body
pmc Summary

Urolithin A, one metabolite derived from gut microbiota, inhibited glutaminolysis to suppress fibrogenesis and contraction of hepatic stellate cells and ameliorated cirrhotic portal hypertension by relieving intrahepatic vascular resistance. We identified Urolithin A as novel drug candidate with great clinical value for cirrhotic portal hypertension treatment.

Cirrhosis is a late-stage liver scarring (fibrosis) caused by chronic liver injury with multiple etiologies.1 Worldwide, more than one million people die of cirrhosis annually. Cirrhotic portal hypertension (CPH), the driving force behind the major complications such as gastroesophageal variceal bleeding and bacterial infections, is a leading cause of death in patients with cirrhosis. The pathobiology of CPH is characterized by an increased portal pressure resulting from hyperdynamic splanchnic circulation and elevated intrahepatic vascular resistance (IHVR) in the cirrhotic liver.2,3 The pathological basis of an elevated IHVR mainly includes abnormal contraction of intrahepatic microvasculature and the disordered liver tissue structure caused by liver fibrosis. An elevated IHVR plays a fundamental role in increasing the portal pressure and deteriorating CPH.4 Current pharmacotherapy for CPH mainly includes nonselective beta-adrenergic blockers, which lower portal pressure by decreasing portal inflow, but with no reduction in IHVR. However, over 50% of patients with CPH treated with beta-adrenergic blockers present variceal bleeding or even death.5,6 This is chiefly attributed to the lack of effective downregulation of the IHVR, resulting in insufficient reduction in portal pressure and drug discontinuance caused by intolerable side effects such as cardioinhibitory response and asthma. In search of breakthroughs in CPH pharmacotherapy, the focus of research has shifted beyond decreasing the portal inflow to the potential reduction in IHVR. Compared with the traditional beta-adrenergic blockers aiming at decreasing portal inflow, the strategy of decreasing IHVR has obvious superiority in saving the lives of patients with CPH and improving their opportunities of receiving liver transplantation. First, decreasing the IHVR is not only highly efficient in reducing the portal pressure to minimize the risk of life-threatening variceal bleeding but can also maintain normal portal inflow to guarantee sufficient liver perfusion, thereby protecting liver function.7, 8, 9 Moreover, as the decreasing IHVR strategy focuses on modulating the liver organ itself, it effectively avoids the intolerable systemic side effects that often occur during beta-adrenergic blocker use. Currently, drugs targeting IHVR are lacking. Thus, identifying key therapeutic targets and developing novel drugs to effectively overcome the increased IHVR represents a promising strategy for CPH treatment.

The gut microbiota influences multiple hepatic diseases.10,11 Recently, metabolites produced by the gut microbiota have attracted tremendous attention because of their role as the main mediators for hepatic influence of gut microbiota.12,13 In our previous studies, the gut microbiota metabolites 1-phenyl-1,2-propanedione and 3,4-dihydroxyphenylpropionic acid were found to modulate liver injury caused by acetaminophen and ischemia/reperfusion, respectively.14,15 Our findings strongly indicated the vital impact of gut microbiota metabolites on liver diseases. However, their role in CPH remains unclear. Urolithin A (UroA) is a gut microbiota metabolite mainly derived from foods (eg, pomegranate) and is rich in ellagitannins and ellagic acid.16,17 Although UroA was discovered more than 40 years ago, its functions in health and disease have only been widely explored over the past decade. UroA exhibits superior performance in modulating mitophagy and inflammation, which contributes to beneficial effects in various diseases, including muscular dystrophy, Alzheimer’s disease, and tumors.18, 19, 20

Activated hepatic stellate cells (HSCs) induced by injury factors act as the key executors of elevated IHVR by not only producing a large amount of extracellular matrix collagen to cause liver fibrosis, but also by enhancing the cell contraction phenotype to aggravate the abnormal contraction of the intrahepatic microvasculature.21 We discovered a previously unrecognized deficiency of UroA in both patients with CPH and in CPH mice, where the abundance of UroA was negatively correlated with disease severity. Based on this, our transcriptomics and metabolomics data identified glutaminolysis of HSCs as a novel therapeutic target of UroA. Furthermore, glutaminolysis plays a crucial role in the maintenance of HSC activation.22,23 We demonstrated that UroA significantly relieved CPH by inhibiting glutaminase1 (GLS1) to suppress glutaminolysis in HSCs. Moreover, our data from primary HSCs (derived from patients and mice), LX-2 cell lines, and CPH mice induced by carbon tetrachloride (CCl4) or bile duct ligation (BDL) indicated that the suppression of glutaminolysis by UroA not only inhibited HSC activation and collagen production to improve liver tissue microstructure but also inhibited HSC contraction to restrain intrahepatic microvascular shrinkage. The above therapeutic effects of UroA collectively helped to decrease the IHVR and ameliorate CPH. Additionally, we found that UroA deficiency was associated with Lactobacillus murinus (L. murinus) disturbance and that supplementation with L. murinus increased UroA levels and significantly improved CPH.

Our study provides detailed data from patients with CPH, mice, and primary cells, indicating that HSC glutamine metabolism is a promising therapeutic target for increased IHVR, and revealed a previously unreported role of the gut bacterium L. murinus and its metabolite UroA in inhibiting HSC glutamine metabolism, decreasing IHVR, and ameliorating CPH. On a positive note, clinical trials have demonstrated that UroA is biologically safe, with no obvious side effects.24, 25, 26 Taken together, we identified UroA as novel drug candidate with great clinical value for CPH treatment.

Results

The Gut Microbiota Metabolite UroA was Deficient in Patients and Mice With CPH

The role of UroA (Figure 1A) in the pathogenesis of CPH has not yet been elucidated. To understand the potential contribution of UroA to CPH, we initially examined UroA levels in the feces and plasma of patients with CPH and in age- and sex-matched healthy subjects. As shown in Figure 1B, UroA levels in the feces and plasma of patients with CPH were significantly lower than those in healthy subjects. Notably, UroA levels in the feces and plasma were negatively correlated with portal pressure (widely applied as a key clinical parameter for determining CPH severity) (Figure 1C). Consistent with the above findings, we confirmed a similar correlation between UroA abundance and other clinical parameters of CPH severity (ascites and Child-Pugh grade of liver function) in patients with CPH (Figure 1D). Furthermore, consistent with the above clinical findings, UroA levels in both the feces and plasma were significantly decreased in mice with CCl4-induced CPH compared with those in control mice (Figure 1E). Thus, our results are the first to reveal a potential relationship between UroA and CPH pathogenesis.Figure 1 The gut microbiota metabolite UroA was deficient in patients and mice with CPH. (A) The chemical structure of UroA. (B) The levles of UroA in the humans feces and plasma were determined by LC-MS/MS (n = 20 healthy subjects; n = 30 patients with CPH). (C) The statistical correlation between UroA levels and portal pressure (Spearman rank-order correlation test) (n = 30 patients with CPH). (D) The UroA levels in different patients with CPH. Child-Pugh score (A/B+C, n = 13/17 patients with CPH), ascites (no/yes, n = 12/18 patients with CPH). Mice were administered 14 weeks CCl4 to induce liver fibrosis and CPH. (E) The concentration of UroA in the feces and plasma of mice was determined by LC-MS/MS (n = 10). (F) Mice were pre-administered with antibiotics by oral gavage. The total bacterial load in mice cecum and the levels of UroA in mice (n = 6). (G) Alpha diversity (observed species) in mice was calculated using the Chao1 index, Shannon index, and observed ASVs index (n = 10). (H) Beta diversity of mice feces (microbial community structures) was calculated using a PCoA score based on the weighted UniFrac distance matrixes (n = 10). (I) The relative abundance of bacteria at the phyla and class levels in mice feces (n = 10). (J) Heatmap of bacterial genus with significant difference in control and CPH mice. Red represents high relative expression and blue represents low relative expression (n = 10). (K) The relative abundance of Lactobacillus at the genus level in mice feces (n = 10) and the statistical correlation between UroA levels and the relative abundance of genus Lactobacillus in mice (n = 20, Spearman rank-order correlation test). (L) The relative abundance of Roseburia at the genus level in mice feces (n = 10) and the statistical correlation between UroA levels and the relative abundance of genus Roseburia in mice (n = 20, Spearman rank-order correlation test). (M) The relative abundance of Streptococcus at the genus level in mice feces (n = 10) and the statistical correlation between UroA levels and the relative abundance of genus Streptococcus in mice (n = 20, Spearman rank-order correlation test). Data were expressed as mean ± SEM; ∗P < .05.

UroA is mainly synthesized by the gut microbiota; therefore, we further investigated the potential contribution of disordered gut microbiota to the decrease in UroA in CPH mice. First, the gut microbiota was successfully reduced by treating the mice with antibiotics (ABX), and the UroA levels were extremely lower in the plasma and feces of ABX-treated control mice (Figure 1F). This result confirmed that UroA was generated by the gut microbiota in mice, which is consistent with the results of previous studies on UroA sources.17 To explore our hypothesis that the low UroA level might be caused by the CPH-induced dysfunctional gut microbiota and a reduced abundance of UroA-producing gut microbes, we performed 16S rRNA sequencing analysis of the microbiota from control and CPH mice feces. As shown in Figure 1G–H, the microbial compositions of the 2 groups were significantly different. Figure 1I shows the major bacteria at the phylum and class levels in the 2 groups. For example, we found that the abundance of Firmicutes decreased, whereas that of Bacteroidota increased significantly at the phylum level. At the class level, some probiotics also decreased markedly (eg, Bacilli). At the genus level (Figure 1J), 20 species of bacteria were enriched, with significant changes in CPH mice compared with those in the control group. We found that several probiotics, such as Faecalibaculum, Lactobacillus, and Akkermansia were depleted in groups with CCl4-induced CPH. These results clearly indicated a dysfunctional gut microbiota in CPH mice. The genera of Lactobacillus, Roseburia, Streptococcus, Fusobacterium, Slakia, and Gordonibacter have been widely demonstrated as gut microbiota involved in the metabolism of UroA generation.27 Hence, we next analyzed the abundance of these bacteria using 16S rRNA sequencing. As shown in Figure 1K–M, compared with those in the control, the relative abundances of the genera Lactobacillus, Roseburia, and Streptococcus were remarkably reduced in the feces of CPH mice. Meanwhile, the abundance of Fusobacterium, Slakia, and Gordonibacter could not be detected (below the minimal detectable concentrations) in either control or CPH mice. Based on the above findings, we evaluated the correlation between the relative abundance of the genera Lactobacillus, Roseburia, and Streptococcus and the level of UroA to confirm the key members of the gut microbes responsible for the decrease in UroA in CPH. Notably, compared with Roseburia and Streptococcus, the relative abundance of the genus Lactobacillus showed the most significant correlation with the level of UroA in mice feces. Thus, these data indicate that the decreased abundance of the genus Lactobacillus might contribute greatly to the lower levels of UroA, which induces CPH progression.

UroA Attenuated CPH in Both CCl4-induced and BDL-induced Mice

To further explore whether UroA participates in CPH progression, the therapeutic effects of 10 mg/kg UroA was initially evaluated in the CCl4-induced mice (6 weeks of CCl4 treatment, only liver fibrosis without CPH; or 14 weeks of CCl4 treatment, end-stage liver fibrosis with CPH) and BDL-induced mice. Supplementation with 10 mg/kg UroA restored UroA levels in the plasma of CPH mice (Figure 2A). As shown in Figure 2B–E and Figure 3A–E, according to the representative photographs of mice livers and hematoxylin and eosin (HE) staining, prophylactic administration of UroA significantly ameliorated liver injury in CCl4-induced mice. Meanwhile, collagen deposition, HSC activation (α-smooth muscle actin [α-SMA] as the marker), HSC contraction (phospho-myosin light chain 2 [p-MLC2] as the marker), and macrophage recruitment (F4/80 as the marker) were significantly reduced in CPH mice by UroA treatment. Consistently, the hepatic mRNA levels of fibrogenic genes (α-SMA, collagen type 1α1 [Col1α1], transforming growth factor-β1 [TGF-β1], platelet-derived growth factor [PDGF], and tissue inhibitor of metalloproteinase-1 [TIMP-1]) and inflammatory genes (F4/80, tumor necrosis factor-α [TNF-α] and interleukin-1β [IL-1β]) were decreased in UroA-treated mice. In addition, UroA significantly attenuated ascites, splenomegaly, and reduced the portal pressure in CPH mice induced by 14 weeks of CCl4 treatment, suggesting that UroA has remarkable therapeutic effects on portal hypertension. Furthermore, we also established BDL-induced CPH in mice to further confirm the effects of UroA. Consistent with the above results of CCl4-induced CPH mice, prophylactic administration of UroA significantly diminished liver fibrosis and portal hypertension in BDL-induced CPH mice (Figure 3F–J). In addition, we added an interventional treatment of UroA in CPH mice. UroA (10 mg/kg) was administered from the 8th week of CCl4 treatment or the 14th day after BDL surgery. As shown in Figure 4A–J, UroA effectively improved the liver fibrosis and portal hypertension when administered at the advanced stages of fibrosis. These findings demonstrated that UroA effectively ameliorated liver injury and portal hypertension in mice with CPH.Figure 2 UroA attenuated liver fibrosis in CCl4-induced mice. Mice were administered with CCl4 for 14 weeks and treated with 10 mg/kg UroA once a day from the first day of CCl4. (A) The plasma level of UroA in each group was detect by LC-MS/MS. (n = 8). Mice were administered with CCl4 for 6 weeks to induce liver fibrosis, and treated with 10 mg/kg UroA once a day from the first day of CCl4 treatment. (B) Representative photographs of mice livers, HE, and (C) quantification of Sirius Red, α-SMA, and F4/80 in mice liver (n = 6). (D) Hepatic hydroxyproline (n = 6). (E) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, TIMP-1, F4/80, TNF-α, and IL-1β (n = 6). Representative results from 3 independent experiments with similar results. Scale bar: 100 μm. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Figure 3 The prophylactic administration of UroA attenuated liver injury and CPH in CCl4-induced and BDL-induced mice. Liver injury and CPH was induced by CCl4 administration in mice. Ten mg/kg UroA was administered to mice by oral gavage once a day. (A) Representative photographs of mice livers, HE, and (B) quantification of Sirius Red, α-SMA, p-MLC2, and F4/80 in mice liver (n = 8). (C) Hepatic hydroxyproline (n = 8) and ascites of mice (n = 8). (D) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, TIMP-1, F4/80, TNF-α, and IL-1β (n = 8). (E) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). In BDL-induced mice, 10 mg/kg UroA was administered to mice by oral gavage once a day. (F) Representative photographs of mice livers, HE, and (G) quantification of Sirius Red, α-SMA, F4/80, and p-MLC2 in mice liver (n = 8). (H) Hepatic hydroxyproline (n = 8) and ascites of mice (n = 8). (I) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, TIMP-1, F4/80, TNF-α, and IL-1β (n = 8). (J) Spleen enlargement, spleen coefficient and portal pressure (n = 8). Representative results from 3 (A–J) independent experiments with similar results. Scale bar: 100 μm. Data were expressed as mean ± SEM; ∗P < .05.

Figure 4 The interventional administration of UroA attenuated liver injury and CPH in CCl4-induced and BDL-induced mice. Liver injury and CPH was induced by CCl4 administration in mice. Ten mg/kg UroA was administered beginning in the 8th week of CCl4 treatment. (A) Representative photographs of mice livers, HE, and (B) quantification of Sirius Red, α-SMA, and p-MLC2 in mice liver (n = 8). (C) Hepatic hydroxyproline (n = 8) and ascites of CPH mice (n = 8). (D) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 8). (E) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). In BDL-induced mice, 10 mg/kg UroA was administered from the 14th day after BDL surgery. (F) Representative photographs of mice livers, HE, and (G) quantification of Sirius Red, α-SMA, p-MLC2 in mice liver (n = 8). (H) Hepatic hydroxyproline (n = 8) and ascites of CPH mice (n = 8). (I) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 8). (J) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). Scale bar: 100 μm. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

UroA Exerted the Therapeutic Effect by Acting on HSCs

We next investigated the underlying mechanisms by which UroA prevents the progression of liver fibrosis and portal hypertension. First, as shown in Figure 5A–D, transcriptome analysis was performed to determine the different gene expression profiles of the liver tissues from UroA-treated and untreated CPH mice. The expression of different genes, including those related to proline metabolism, retinol metabolism, and vascular smooth muscle contraction, which are mostly associated with HSCs, was significantly enriched. Meanwhile, the Kyoto Encyclopedia of Genes and Genomes (KEGG) and gene ontology (GO) analysis of the transcriptome indicated that the expression levels of HSC phenotype-related hepatic genes such as those associated with cell contraction, fibroblast proliferation, and collagen synthesis, were influenced by UroA treatment in CCl4-induced CPH mice. These data suggested that UroA exerts a therapeutic effect by acting on HSCs. Upon activation by various stimuli, quiescent HSCs transdifferentiate into fibrogenic myofibroblasts known as activated HSCs (aHSCs). aHSCs not only perform intense cell contraction to shrink the hepatic microvasculature, but also produce abundant extracellular matrix (ECM) to disorder the hepatic microstructure, collectively leading to increased IHVR and the progression of CPH.28,29 Therefore, we evaluated the influence of UroA on the IHVR-related cell phenotype of HSCs using primary HSCs isolated from the patients or mice liver tissues. According to the previous report,30 as well as confirming the drug safety and effectiveness by evaluating different doses of UroA (Figure 5E–F), UroA at the concentration of 10 μM was used for the experiment in vitro. As shown in Figure 6A–L, reverse transcription polymerase chain reaction (RT-PCR) and Western blot assays of both the human HSC line LX-2, human and mice HSCs showed that UroA significantly inhibited the activation (α-SMA), collagen synthesis (Col1α1, the main component of ECM), and fibrogenic genes expression (TGF-β, PDGF, and TIMP-1) of HSCs. Cell counting kit-8 (CCK-8) and EdU assays also revealed that UroA significantly inhibited HSCs proliferation. In addition, a collagen gel contraction assay and immunofluorescence of p-MLC2 (a contraction marker) expression confirmed that UroA significantly suppressed the contraction of aHSCs.Figure 5 UroA exerted the therapeutic effect by acting on HSCs. Liver injury and CPH was induced by CCl4 administration in mice. Ten mg/kg UroA was administered to mice by oral gavage once a day for 14 weeks. (A) The differences of hepatic transcriptional levels between 2 groups were analyzed by transcriptome. The heatmap was performed to display the transcriptional level of gene expression with significant difference in mice livers. Red represents high relative expression and blue represents low relative expression (n = 5). (B) KEGG signaling pathway analysis revealed HSCs-related functional terms enriched in CPH mice. (C) The volcano plots of genes expression in UroA-treated group and CCl4 group. (D) Cellular component, molecular function, and biological process were used to reveal metabolism-related functional terms enriched by the DEGs based on Cytoscape software combined with ClueGO App. (E) LX-2 cells were treated with or without UroA for 24 hours. LX-2 cells apoptosis (n = 3). (F) The mRNA levels of α-SMA and Col1α1 with different doses of UroA in primary mice aHSCs (n = 4). Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Figure 6 UroA significantly inhibited the the fibrogenic and contraction phenotype of HSCs. LX-2 cells were treated with or without 10 μmol/L UroA for 24 hours. (A) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, TIMP-1 (n = 6). (B) Cell growth determined by CCK-8 assay (n = 5) and EdU staining. (C) Collagen gel contraction and p-MLC2 fluorescent staining. (D) The protein expression and quantification of α-SMA, Col1α1 (n = 3). Primary human HSCs were treated with or without 10 μmol/L UroA for 24 hours. (E) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (F) Cell growth determined by CCK-8 assay (n = 5) and EdU staining. (G) Collagen gel contraction and p-MLC2 fluorescent staining. (H) The protein expression and quantification of α-SMA and Col1α1 (n = 3). Primary mice HSCs were treated with or without 10 μmol/L UroA for 24 hours. (I) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (J) Cell growth (n = 5) and EdU staining. (K) Collagen gel contraction and p-MLC2 fluorescent staining. (L) The protein expression and quantification of α-SMA and Col1α1 (n = 3). Representative results from three independent experiments with similar results. Scale bar: 50 μm. Data were expressed as mean ± SEM; ∗P < .05.

UroA Suppressed the Glutamine Metabolism to Regulate the Fibrogenic and Contraction Phenotype of HSCs

To our knowledge, no studies on the regulation of HSCs function by UroA have been reported. To further explore the regulatory mechanisms of UroA, the targeted metabolomics was performed on aHSCs treated with or without UroA. Figure 7A–E showed that UroA administration remarkably altered the metabolites levels in aHSCs, among which the glutamine metabolism pathways presented prominent changes. Extracellular glutamine is transported into cells via transportase and hydrolyzed into glutamate and its downstream hydrolytic products, which participate in the tricarboxylic acid cycle (TCA) cycle and supply energy for cells (Figure 7F).31 We found that glutamine metabolism was significantly suppressed, and the abundance of glutaminolysis metabolites (represented by glutamic acid and α-ketoglutaric acid [α-KG]) were considerably reduced in LX-2 cells after UroA treatment (Figure 8A). These observations confirmed that UroA might regulate the HSCs function by influencing the glutaminolysis-associated pathways. In recent years, growing evidence has suggested that glutaminolysis reprogramming plays a crucial role in supporting the fibrogenic phenotypes of HSCs, such as activation, survival, and ECM synthesis.22,32 To further verify whether the regulatory effect of UroA on HSC phenotype was mediated by glutaminolysis, we evaluated the influence of UroA treatment on LX-2 cells cultured in a glutamine-deprived medium. We found that UroA exerted no further inhibition on HSC proliferation, activation (with α-SMA as the marker), collagen synthesis (Col1α1), and profibrogenic genes expression (TGF-β, PDGF, TIMP-1), as well as cell contraction (p-MLC2 and collagen gel) after glutamine deprivation (Figure 8B–G), indicating that the regulatory efficacy of UroA on HSC phenotype was significantly blunted in the absence of glutamine. Taken together, these results demonstrated that UroA regulates HSC function, at least in part, via glutamine metabolism.Figure 7 UroA suppressed the glutamine metabolism of HSCs. LX-2 cells were treated with or without 10 μmol/L UroA, and (A) the significantly different metabolites were shown between control and UroA group by heatmap. Red represents high relative abundance, whereas blue represents low relative abundance (n = 3). (B) Cytoscape software combined with ClueGO App was used to reveal signaling pathways enriched by the significantly different metabolites. (C) Z score of metabolites of the glutamine metabolism. (D) KEGG signaling pathway analysis of metabolomics reveal glutamate metabolism-related functional terms enriched in cells. (E) The metabolites of the glutaminolysis in LX-2 cells (n = 3). (F) Metabolic pathways of glutamine. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Figure 8 UroA suppressed the glutamine metabolism to regulate the fibrogenic and contraction phenotype of HSCs. LX-2 cells were treated with or without 10 μmol/L UroA. (A) Glutamic acid and α-Ketoglutaric acid levels in LX-2 cells (n = 3). LX-2 cells were cultured in medium deprived of glutamine and treated with or without UroA, control cells were cultured in complete medium. (B) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (C) Cell growth (n = 5) and (D) EdU staining. (E) p-MLC2 fluorescent staining. (F) The protein expression of α-SMA and Col1α1. (G) Collagen gel contraction. Primary HSCs from healthy mice were isolated. Quiescent HSCs were cultured for 0 days, and activated HSCs were cultured for 7 days and treated with or without 10 μmol/L UroA. (H) The relative ATP content (n = 5). Liver injury and CPH was induced by CCl4 administration in mice. Ten mg/kg UroA was administered to mice by oral gavage once a day for 14 weeks. (I) The mRNA levels of TNF-α and IL-1β in liver macrophage of CPH mice (n = 8). Liver injury and CPH was induced by 14 weeks CCl4 administration in mice. Ten mg/kg UroA was administered to mice by oral gavage once a day. (J) Plasma ALT level (n = 8) and the relative ATP content in primary hepatocytes (n = 5). Representative results from 3 independent experiments with similar results. Scale bar: 50 μm. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Under various injury stimuli, healthy quiescent HSCs trans-differentiate into activated fibrogenic myofibroblasts (aHSCs). aHSCs depend mainly on glutamine metabolism to meet their significantly elevated ATP energy requirements to exert pro-fibrogenic and pro-IHVR behaviors.33 Hence, we also investigated whether the effect of UroA was mediated by interference with ATP production. As shown in Figure 8H, we measured the ATP levels in quiescent and activated HSCs treated with or without UroA. UroA showed no significant influence on the ATP level of quiescent HSCs, which indicated that 10 μM UroA did not interfere the energy metabolism of physiologically normal HSCs. In contrast, UroA mildly reduced ATP levels in aHSCs, which was consistent with a previous report that glutamine deprivation impairs ATP production in aHSCs.22 Given that ATP production plays an important role in supporting the pro-fibrogenic behaviors of aHSCs during CPH development, we speculated that the downregulation of ATP by UroA in aHSCs, rather than in normal quiescent HSCs, might contributed to the therapeutic action of UroA. Consequent to the reduced recruitment of macrophages (F4/80), the main executors of liver inflammation, in the CPH mice treated with UroA, we also explored whether UroA affected the pro-inflammatory response of macrophages in the pathological course of CPH. We isolated liver macrophages from CPH mice following UroA treatment to detect the expression of pro-inflammatory factors. As shown in Figure 8I, the TNF-α and IL-1β levels did not decrease significantly in the liver macrophages of UroA-treated CPH mice. This indicates that the decrease in macrophage recruitment and proinflammatory factors in the livers of CPH mice treated with 10 mg/kg UroA cannot be attributed to the direct inhibitory effect of UroA on macrophages. Thus, UroA may indirectly inhibit macrophage recruitment by inhibiting HSC activation and its inflammatory chemotaxis. Additionally, we also detected the ALT levels in plasma of CPH mice treated with or without 10 mg/kgUroA and the ATP levels in the primary hepatocytes isolated from CPH mice treated with or without 10 mg/kg UroA. As shown in Figure 8J, we found no significant difference of ALT levels between CPH mice and UroA-treated CPH mice. Moreover, the ATP levels of hepatocytes also showed no significant difference after UroA treatment, which was in line with our above findings that UroA improved CPH mainly by acting on aHSCs.

UroA Inhibited GLS1 Activity to Suppress Glutaminolysis Metabolism

According to the target metabolomics results (liquid chromatography with tandem mass spectrometry [LC-MS/MS]) aforementioned, UroA treatment showed no influence on the glutamine uptake of HSCs but significantly decreased the abundance of downstream glutamine metabolites such as glutamic acid and α-KG. Based on this, we focused on the potential effects of UroA on glutaminolysis to regulate glutamine metabolic reprogramming in HSCs. We examined the activity of the key enzymes responsible for glutaminolysis, including glutaminase (GLS), glutamate dehydrogenase (GDH), glutamic-oxaloacetic transaminase 2 (GOT2), and glutamic-pyruvic transaminase 2 (GPT2). As shown in Figure 9A–B, UroA had no significant influence on the activity of GDH, GOT2, or GPT2, whereas it significantly inhibited the activity of GLS in primary aHSCs isolated from both patients and mice, as well as in LX-2 cells. Consistent with this, UroA significantly inhibited GLS activity in primary HSCs isolated from CPH mice (Figure 9C). GLS is a vital glutaminolysis enzyme that converts glutamine into glutamate and downstream α-KG to promote ATP production and to provide key metabolic intermediates for the biosynthesis of nucleic acids, amino acids, and lipids. GLS has 2 distinct forms, including GLS1 and GLS2. GLS1 has been well-demonstrated to be the key rate-limiting enzyme for glutamine utilization and regulates liver glutamine metabolism.34,35 Notably, as shown in Figure 9D–E, UroA treatment showed not significantly suppress the protein level of GLS1. This suggests that UroA may bind to GLS1 and affect the enzymatic activity of GLS1 rather than influence its protein expression.Figure 9 UroA targets GLS1 to inhibit glutaminolysis of HSCs. LX-2 cells were treated with or without 10 μmol/L UroA for 24 hours. (A) GLS activity of primary HSCs and LX-2 cells treated with or without UroA (n = 5). (B) The GDH, GOT2, and GPT2 activity in LX-2 cells (n = 3). (C) GLS activity of primary mice HSCs (n = 8). (D–E) The protein levels and quantification of GLS1 (n = 3). (F) SPR analysis of UroA (2.5, 5, 10, and 20 μmol/L) and BPTES (10 μmol/L) binding to the recombinant human GLS1 protein. (G) Molecular docking analysis and binding sites between UroA and GLS1. (H) Molecular docking analysis and binding sites between BPTES and GLS1. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Next, the molecular mechanism underlying the inhibitory effect of UroA on GLS1 activity in HSCs was further explored. First, surface plasmon resonance (SPR) analysis was performed to determine the potential binding activity between UroA and GLS1. As shown in Figure 9F, UroA showed a direct binding signal to GLS1 in a dose-dependent manner, with a KD value of 2.66e-10M (the GLS1 inhibitor Bis-2-(5-phenylacetamido-1,3,4-thiadiazol-2-yl) ethyl sulfide [BPTES] was used as the positive control). Furthermore, we used molecular docking to confirm the potential binding sites between UroA and the 3D protein structure of the GLS1 (PDB ID: 3VOZ). As shown in Figure 9G–H, UroA was buried within the activity pocket of GLS1 (docking score of −5.074), partly overlapping with the binding sites of BPTES. Based on docking simulations, Asp-327 (D327) and Try-394 (Y394) amino acid residues were most likely to form hydrogen bonds with UroA. To validate our findings, we constructed plasmids overexpressing wild-type (WT) recombinant human GLS1 and plasmids overexpressing human GLS1 with D327 or Y394 mutations (rhGLS1D327A or rhGLS1Y394A). We used the above plasmids to transfect LX-2 cells, then the LX-2 cells were treated with or without UroA. As shown in Figure 10A−E, the GLS1 WT plasmids transfection significantly enhanced the GLS1 activity and promoted the activation (α-SMA) and fibrogenic phenotype (Col1α1, TGF-β, PDGF, TIMP-1) of aHSCs compared with those in the normal control group (NC group; transfected with blank pcDNA3.1 plasmid). However, the pharmaceutical inhibitory effect of UroA on the activation and fibrogenic phenotype of HSCs was significantly abolished by the D327 and/or Y394 mutations. To further demonstrate whether UroA interacts with the amino acid residues of GLS1 and diminishes its activity, 2 protein mutations (rhGLS1D327A and rhGLS1Y394A) were prepared, and SPR analysis was performed to determine the binding affinity of UroA with mutated GLS1. UroA exhibited a much lower binding affinity to the rhGLS1D327A (with a KD value of 3.93e-5M) and rhGLS1Y394A (with a KD value of 1.1e-5M) mutations than to the GLS1, thus demonstrating that UroA interacted with the amino acid residues of GLS1 to diminish GLS1 activity (Figure 10F). Taken together, UroA significantly inhibits GLS1 activity, probably by interacting with GLS1 at the D327 and Y394 sites, which downregulates glutaminolysis to exert its pharmaceutical effects against the activation and fibrogenic phenotype of aHSCs.Figure 10 UroA inhibited GLS1 activity to inhibited the the fibrogenic and contraction phenotype of HSCs. GLS1, rhGLSD327A, and rhGLSY394A overexpression plasmids were transfected into LX-2 cells, and cells were treated with 10 μmol/L UroA for 24 hours. (A) The mRNA levels of GLS1 in LX-2 cells (n = 4). (B) GLS activity of LX-2 cells (n = 3). (C–D) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 4). (E) The protein expression of α-SMA and Col1α1. (F) SPR analysis of UroA (2.5, 5, 10, 20 μmol/L) binding to the rhGLS1D327A and rhGLS1Y394A protein. Representative results from 3 independent experiments with similar results. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

UroA partly Depended on GLS1 Inhibition to Counteract Fibrogenesis and Contraction of aHSCs and Thus Ameliorated CPH

Next, we explored whether the beneficial effect of UroA on the HSCs phenotype was mediated by GLS1 inhibition. Figure 11A–E showed that the GLS1 inhibitors BPTES markedly inhibited GLS1 activity, cell proliferation, fibrogenic mRNA levels, activation, and collagen synthesis in HSCs. Similary, the contraction and growth of aHSCs were suppressed by BPTES. However, the administration of UroA had no additional effect on the phenotypic changes in HSCs treated with BPTES. This finding supports the notion that the inhibitory effects of UroA on the fibrogenic and contraction phenotypes of HSCs are at least partly dependent on the inhibition of GLS1. These in vitro results clarified that UroA inhibited the IHVR-related phenotype (fibrogenesis and contraction) of aHSCs by regulating GLS1 and glutaminolysis. This finding was also confirmed in the CPH mice. As shown in Figure 11F–K, the inhibition of glutaminolysis after BPTES treatment significantly alleviated liver fibrosis and CPH in mice induced by 14 weeks of CCl4 treatment. Liver injury, HSC activation (α-SMA), HSC contraction (p-MLC2), collagen deposition (Sirius Red), and the hepatic mRNA levels in fibrogenic HSCs (α-SMA, Col1α1, TGF-β, PDGF, TIMP-1) were significantly reduced in BPTES-treated CPH mice. BPTES alleviated ascites and splenomegaly, and remarkably reduced the portal pressure in CPH mice, confirming that glutaminolysis plays a vital role in CPH progression. Notably, UroA administration did not further relieve the liver fibrosis or CPH-related indicators in BPTES-treated CPH mice. In addition, the consistent therapeutic effect of another GLS1 inhibitors CB-839 in vitro was also verified in Figure 12A–G.Figure 11 The protective effect of UroA was partly dependent on GLS1 inhibition. LX-2 cells were treated with 10 μmol/L BPTES and/or 10 μmol/L UroA for 24 hours. (A) GLS activity (n = 5) and the mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (B) EdU staining and p-MLC2 fluorescent staining, scale bar: 50 μm. (C) Cell growth (n = 5). (D) The protein expression and quantification of α-SMA and Col1α1 (n = 3). (E) Collagen gel contraction. Representative results from 3 (A–E) independent experiments with similar results. Liver injury and CPH was induced by CCl4 administration in mice. Then mice were treated with 10 mg/kg BPTES and/or 10 mg/kg UroA. (F) Representative photographs of mice livers, HE, Sirius Red, and (G) quantification of Sirius Red, α-SMA, p-MLC2 in mice liver (n = 8), scale bar: 100 μm. (H) Hepatic hydroxyproline and ascites of CPH mice (n = 8). (I) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 8). (J–K) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). Representative results from 2 (F–K) independent experiments with similar results. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

Figure 12 The protective effect of UroA was partly dependent on inhibiting glutaminolysis of HSCs. LX-2 cells were treated with 1 μmol/L CB-839 and/or 10 μmol/L UroA for 24 hours. (A) GLS activity (n = 5) and (B) the mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (C) Cell growth determined by CCK-8 assay (n = 5). (D) EdU staining and (E) p-MLC2 fluorescent staining. (F) The protein expression and quantification of α-SMA and Col1α1 (n = 3). (G) Collagen gel contraction. LX-2 cells were treated with 10 μmol/L UroA and/or 4 mmol/L DKG for 24 hours. (H) The mRNA levels of α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 6). (I) Cell growth (n = 5). (J) EdU staining and (K) p-MLC2 fluorescent staining. (L) The protein expression of α-SMA and Col1α1. (M) Collagen gel contraction. Scale bar: 50 μm. Representative results from 3 independent experiments with similar results. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

α-KG, a key metabolic product of glutaminolysis, can replenish the TCA cycle for energy production and anabolism. α-KG serves as the preferred energy substrate of aHSCs, and supplementing α-KG can enhance fibrogenic phenotype in myofibroblasts.31 Hence, we also added dimethyl α-ketoglutarate (DKG, a cell-permeable substitute of the impermeable α-KG) to the culture medium to investigate whether α-KG could blunt the regulatory effect of UroA on aHSCs. As shown in Figure 12H–M, supplementation with DKG significantly counteracted the inhibition of UroA treatment on the proliferation (as evidenced by CCK8 and EdU assay results), fibrogenesis (validated by α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 levels) and contraction (as demonstrated by collagen gel contraction assay and p-MLC2 immunofluorescence) of HSCs. These data showed that the protective effects of UroA against CPH progression may be, at least in part, dependent on the inhibition of GLS1 and glutaminolysis.

Gut Bacterium L. murinus Protected Against CPH in CCl4-induced Mice by Producing Metabolite UroA

Among the significantly reduced gut microbiota (Figure 1), Lactobacillus was significantly reduced in the CPH mice. In the Lactobacillus genus, L. murinus, a previously reported probiotic,36,37 was the most abundant and showed the most obvious changes, correlated with the levels of UroA (Figure 13A). To further identify the relationship between L. murinus and UroA, CCl4-induced mice were given L. murinus alone or in combination with the GLS inhibitor BPTES to determine whether the therapeutic effect of L. murinus depended on UroA. L. murinus treatment enhanced fecal levels of UroA in the CCl4-induced mice (Figure 13B). As shown in Figure 13C–G, liver injury, collagen deposition (Sirius Red), HSC activation (α-SMA as marker), HSC contraction (p-MLC2 as marker), and the hepatic mRNA levels of fibrogenic indicators (α-SMA, Col1α1, TGF-β1, PDGF, and TIMP-1) were significantly reduced in L. murinus-treated CPH mice. Liver fibrosis and CPH (ascites, splenomegaly, and portal pressure) were also alleviated by L. murinus treatment. These results indicated the great potential of UroA-producing L. murinus as a promising probiotic for CPH treatment. However, compared with L. murinus-treated CPH mice, additional supplementation with the GLS1 inhibitor BPTES did not further relieve liver fibrosis and CPH-related indicators in L. murinus-treated CPH mice.Figure 13 Gut bacterium L. murinus protected against CPH in CCl4-induced mice by producing metabolite UroA. (A) The relative abundance of L. murinus at the species level in mice feces (n = 10), the statistical correlation between the UroA levels and the relative abundance of L. murinus (n = 20, Spearman rank-order correlation test). CPH mice were treated with L. murinus (5×10ˆ7 CFU) by oral gavage once a day for 14 weeks. (B) The UroA levels in mice feces (n = 8). Liver injury and CPH was induced by CCl4 administration in mice. CPH mice were treated with L. murinus (5×10ˆ7 CFU) and/or BPTES once a day for 14 weeks. (C) Representative photographs of mice livers, HE, Sirius Red, and (D) quantification of Sirius Red, α-SMA, and p-MLC2 in mice liver (n = 8). (E) Hepatic hydroxyproline and ascites of mice (n = 8). (F) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 8). (G) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). CPH mice were treated with 10 mg/kg UroA and/or L. murinus (5×10ˆ7 CFU) for 14 weeks. (H) Representative photographs of mice livers, HE, Sirius Red. (I) The UroA levels in mice feces and plasma (n = 8). (J) Quantification of Sirius Red, α-SMA, and p-MLC2 in mice liver (n = 8). (K) Hepatic hydroxyproline and ascites of CPH mice (n = 8). (L) The mRNA levels of hepatic α-SMA, Col1α1, TGF-β, PDGF, and TIMP-1 (n = 8). (M) Spleen enlargement, spleen coefficient, and portal pressure (n = 8). Representative results from 2 independent experiments with similar results. Scale bar: 100 μm. Data were expressed as mean ± SEM; ∗P < .05; n.s. indicates nonsignificant.

To further confirm the role of UroA in the response to L. murinus, CPH mice were treated with 10 mg/kg UroA combined with L. murinus. As shown in Figure 13H–M, UroA abundance in the feces of CPH mice showed no significant difference between the group receiving UroA alone and the group receiving co-administration of UroA and L. murinus, indicating that the additional supplementation of UroA was fully adequate and could cover the UroA produced by L. murinus. And the additional supplement of L. murinus in CPH mice treated with UroA did not further relieve the key indicators of liver fibrosis and CPH, including liver injury, HSC activation and contraction, collagen deposition, ascites, splenomegaly, and portal pressure. These data indicated that the protective effects of L. murinus against CPH progression may be, at least in part, dependent on production of the UroA.

Discussion

CPH constitutes a common final pathway of liver cirrhosis with diverse etiologies, and is a key driver of mortality in chronic liver diseases. Currently, clinical pharmacotherapy for CPH mainly includes nonselective beta-adrenergic blockers; however, less than 40% of patients achieve satisfactory hemodynamic response, and up to 15% of patients encounter drug discontinuation due to intolerable side effects. Hence, there is an urgent need to investigate the pathophysiology of CPH and to search for novel breakthroughs in CPH pharmacotherapy. In this study, we discovered for the first time that both patients and mice with CPH were deficient in a gut microbiota metabolite UroA. We demonstrated a novel role for UroA in ameliorating CPH by suppressing glutamine metabolism to inhibit the fibrogenic and contraction phenotypes of HSCs. Furthermore, UroA significantly inhibited GLS1 activity by interacting with GLS1 at the D327 and Y394 sites and downregulated glutaminolysis to exert its pharmaceutical effects against the activation and fibrogenic phenotype of aHSCs. In addition, we found that UroA deficiency was associated with L. murinus disturbance and that supplementation with L. murinus increased the UroA levels and significantly improved CPH.

Disturbances in the gut microbiota, both in experimental and human settings, are closely linked to the development of CPH. Gut microbiota metabolites have long been known as key mediators of the pathophysiological effects of the gut microbiota. However, the role of the gut microbiota metabolites in CPH remains unclear. UroA is a naturally occurring bacterial metabolite with multiple health benefits such as enhancing mitophagy, regulating immunity, and resisting aging. The present study revealed a previously unrecognized link between UroA deficiency and CPH progression. We observed significantly reduced levels of UroA in the plasma and feces of both patients and mice with CPH. More importantly, in terms of clinical significance, reduced UroA levels correlated with severity parameters (portal pressure, liver function, and ascites) in patients with CPH. On this basis, our data from CPH mice, as well as from primary HSCs and LX-2 cells, further demonstrate a new role of UroA in effectively ameliorating CPH by acting on HSCs. UroA inhibits HSC activation and proliferation, which remarkably reduces ECM production and improves the hepatic microstructure. In contrast, UroA inhibits the HSC contraction to weaken shrinkage of the hepatic microvasculature. These 2 pharmacological effects collectively endow UroA with an excellent ability to reduce IHVR, thus restraining CPH progression. Traditional nonselective beta-adrenergic blockers lower portal pressure mainly by decreasing portal inflow, which has been reported to show no survival benefit in some patients with CPH with spontaneous bacterial peritonitis and even increase the risk of liver failure.6,38 With different pharmacological mechanisms from those of nonselective β-adrenergic blockers, UroA lowers portal pressure mainly by intervening in IHVR, which may possess unique advantages because it not only ameliorates CPH but may also avoid decreasing liver perfusion of portal blood and thereby improve liver function. Our study broadens the novel role of the gut microbiota metabolite UroA as a promising drug candidate for CPH by acting on HSCs to decrease the IHVR.

UroA is primarily generated by the gut microbiota by metabolizing its precursors, including ellagitannins and ellagic acid. UroA is much more absorbable than these precursors and is mainly responsible for the health-beneficial effects of ellagitannins and ellagic acid that occur in foods such as berries and nuts.39 However, owing to the complicated components and functional changes in the disturbed gut microbiota, the biosynthesis of UroA shows considerable variation among different diseases, and the definite species of gut microbiota for UroA production remains unknown. We observed that the abundance of tannase-metaboling bacteria, including genus Lactobacillus, Roseburia, and Streptococcus, which have been reported to be correlated with the fecal content of UroA in both animals and humans, was markedly decreased in CPH mice. Among these intestinal bacteria, the Lactobacillus abundance accounted for the highest proportion and showed the most significant correlation with reduced UroA levels in the feces of CPH mice. Moreover, supplementation with L. murinus increased the fecal level of UroA and improved the symptoms of CPH in CCl4-induced mice; furthermore, the protective effects of L. murinus against CPH progression might be partly dependent on its metabolite UroA. Notably, the current clinical application of gut microbiota intervention in CPH treatment is still restricted to non-specific probiotics agents without definite effects on improving CPH prognosis. Taken together, our study lays the foundation for further development of L. murinus-based agents, contributing to the clinical application of specific intervention strategies for the gut microbiota in CPH therapy.

Apart from the activation and ECM-producing phenotype formation of HSCs, HSC contraction is attracting tremendous attention owing to its remarkable role in shrinking the hepatic microvasculature to increase the IHVR and thus aggravate CPH. In the present study, we provided a new perspective on CPH pathogenicity, involving HSC glutamine metabolism. We confirmed the findings reported in previous studies that the glutamine metabolism can support the activation, ECM-biosynthesis and pro-inflammatory effects of HSCs to promote liver fibrosis.32 Moreover, we also observed a previously unrecognized effect of enhanced glutamine metabolism on intensifying the HSC contraction phenotype to aggravate CPH. These findings suggest that glutamine metabolism in HSCs may be a novel therapeutic target in CPH. After confirming that UroA supplementation can significantly ameliorate CPH by suppressing the glutamine metabolism to inhibit the fibrogenic and contraction phenotypes of HSCs, we investigated the molecular mechanism by which UroA regulates the HSC glutamine metabolism. Using molecular docking and SPR assays, we demonstrated that UroA could directly bind to the D327 and Y394 sites of GLS1 in aHSCs, which showed no influence on GLS1 protein expression but significantly inhibited both GLS1 activity and glutamine metabolism. Our results demonstrated that UroA inhibited GLS1 activity and elucidated the possible mechanism of UroA binding to the GLS1 protein pocket, revealing a novel role of UroA as a natural inhibitor of the GLS1.

Our study demonstrated that the gut microbiota metabolite UroA inhibited GLS1 to downregulate the glutaminolytic metabolism of HSCs, which effectively ameliorated CPH. In terms of clinical transformation, supplementation with UroA or direct intervention of GLS1 appears to be a promising strategy for the treatment of CPH. First, UroA showed an effect similar to that of the classical GLS1 inhibitor BPTES in current study. Based on our findings that UroA-mediated GLS1 inhibition effectively improved the liver injury and CPH, the clinical application of UroA supplementation needs to be further verified in terms of pharmacokinetics, biosafety, adverse drug reactions, and clinical efficacy. Second, because UroA is mainly derived from the metabolism of ellagitannins-rich foods (eg, pomegranate and nuts) by the gut microbiota, dietary therapies that appropriately supplement these foods may be beneficial and directly applied to patients with CPH. Third, in addition to the previous reports on the role of L. murinus as a probiotic beneficial for intestinal ischemia/reperfusion injury and necrotizing enterocolitis, our findings further expanded the health-beneficial effects of L. murinus by increasing UroA levels in mice feces and improving CPH. Therefore, L. murinus-based agents are expected to promote the clinical application of gut microbiota in CPH therapy. In addition to the UroA supplementation strategy, direct intervention of GLS1 deserves priority as a promising strategy for CPH treatment. Currently, an oral GLS1 inhibitor (IPN60090) with good drug tolerance and potent inhibition of tumor glutaminase has been tested in phase I clinical trials, showing a 50% disease control rate in patients with lung cancer.40 However, when aiming to regulate the HSCs within the fibrotic CPH liver, the currently available GLS1 inhibitors may have potential limitations, including poor solubility, unfavorable pharmacokinetic profile, and low bioavailability. Specific and efficient GLS1 inhibition strategies, such as the nanomedicine-based delivery of GLS1 inhibitors targeting HSCs in vivo, shows a bright prospect in pushing forward novel clinical CPH therapies.

Materials and Methods

Experimental Animals

Male C57BL/6 mice, aged 8 to 10 weeks (weight, 23-26 g) were purchased from GemPharmatech Co., Ltd. Mice were housed in a standard 12-hour light/dark conditions with a temperature-controlled environment (22 ± 2 °C). All mice received food and water ad libitum. All animal procedures were approved by the Ethics Committee on the Care and Use of Laboratory Animals in the Second Affiliated Hospital of Guangzhou Medical University (Guangzhou, China).

For CCl4 treatment, mice were administered CCl4 (diluted at 1:4 in corn oil, 5 mL/kg) via intraperitoneal injection, twice a week (6 or 14 weeks) to induce liver fibrosis and CPH. The control mice were intraperitoneally injected with corn oil only. For prophylactic administration of UroA, CCl4-induced mice were treated with UroA (10 mg/kg; Macklin) or its vehicle (1% DMSO in phosphate buffered saline [PBS]) by oral gavage daily from the first day of CCl4 injection. For the therapeutic administration of UroA, 10 mg/kg UroA was administered beginning in the the 8th week of CCl4 treatment. For BDL, mice were subjected to a midline laparotomy under anesthesia to separate the bile duct, which was tied proximal to the liver hilus using a 7-0 silk ligature, and the second ligature was tied around its distal end. The bile duct was resected between these 2 ligatures. The bile duct was isolated similarly but not ligated in the sham group. Mice were subjected to BDL for 4 weeks to establish CPH. For prophylactic administration of UroA, BDL-induced mice were treated with UroA (10 mg/kg) or its vehicle (1% DMSO in PBS) by oral gavage daily from the first day of BDL surgery. For the therapeutic administration of UroA, 10 mg/kg UroA was administered beginning in the 14th day after BDL surgery.41,42

BPTES (10 mg/kg; Macklin) or its vehicle (1% DMSO in PBS) was intraperitoneally administered once a day from the first day of CCl4 injection. For ABX treatment,15 the mice were administered neomycin sulfate (200 mg/kg), metronidazole (200 mg/kg), ampicillin (200 mg/kg), and vancomycin (100 mg/kg) via oral gavage once daily for 1 week.

Human Samples

The samples from patients with CPH were obtained from The Second Affiliated Hospital of Guangzhou Medical University. The Medical Ethics Committee of the Second Affiliated Hospital of Guangzhou Medical University approved the use of human samples (2022-YJS-ks-26). Informed consent was acquired from all enrolled patients. Detailed clinical information for enrolled patients with CPH is listed in Table 1.Table 1 General Characteristics of Patients

Parameters	Value	
Patients, n	30	
Gender (female/male)	9/21	
Age, years	59 (39-77)	
Etiology (alcoholic/chronic hepatitis)	4/26	
Child score	7 (5-13)	
Child-Pugh score (A/B/C)	13/14/3	
Ascites (no/yes)	12/18	
Blood platelet count, 10ˆ9/L	101.59 ± 8.97	
White blood cell count, 10ˆ9/L	4.64 ± 0.51	
Red blood cell count, 10ˆ12/L	3.18 ± 0.17	
ALT, U/L	26.16 ± 3.03	
AST, U/L	32.41 ± 2.80	
Antibiotics	No	
Lactulose	No	
PPI	No	
Hepatic encephalopathy	No	
Note: Data are presented as number, median (range), or mean ± standard deviation.

ALT, Alanine aminotransferase; AST, aspartate aminotransferase; PPI, proton pump inhibitor.

Histology Staining

Mice liver specimens were fixed in 4% paraformaldehyde and embedded in paraffin blocks after dehydration. Next, liver sections (4-μm thick) were processed using a standard protocol of HE and Sirius Red staining. For immunohistochemical staining, liver sections were processed using a standard immunostaining protocol. Mice liver sections were separated, rehydrated, and incubated with anti-α-SMA antibody (1:200; BOSTER), anti-F4/80 (1:500; Cell Signaling Technology), or anti-p-MLC2 antibody (1:100; Cell Signaling Technology). Secondary horseradish peroxidase-conjugated anti-mouse and anti-rabbit antibodies (Zhongshan Golden Bridge Biotechnology Co., Ltd) were used at 1:500 dilution. The sections were imaged using an Olympus IX53 microscope (Olympus Co., Ltd), and the areas of positive staining were measured and quantified using Image J software.

Hepatic Hydroxyproline Content

The hepatic hydroxyproline content of the mice was measured using a commercial assay kit according to the manufacturer’s instructions. Samples were tested by a microplate reader at 550 nm (Nanjing Jiancheng Bioengineering Institute).

Portal Pressure Measurement

All patients with CPH in our study received intravascular interventional therapy with transjugular intrahepatic portosystemic shunt (TIPS) in our department, and portal pressure was directly measured in the portal vein during TIPS. TIPS is a minimally invasive procedure that has been widely approved for CPH treatment because of its potent effect in reducing portal pressure to inhibit gastroesophageal variceal bleeding.9 After jugular vein puncture, intrahepatic portal vein puncture through the hepatic vein (jugular vein-hepatic vein path for puncture equipment) is an indispensable step in TIPS to establish the final operation path (jugular vein-hepatic vein-portal vein). Thus, during TIPS, portal pressure could be directly measured in the portal vein using a catheter linked to a pressure sensor, which accurately reflects CPH severity, and the therapeutic effect after the portosystemic shunt was established during TIPS.

The portal pressure values of the CPH mice were measured using a BL-420 Biological Function Experimental System (TECHMAN). After mice were anesthetized with 1% pelltobarbitalum natricum (intraperitoneally administered, 50 mg/kg) and the instruments were calibrated, a 26G catheter attached to a pressure transducer was inserted into the portal vein of mice, and the portal pressure was recorded and analyzed.

Evaluation of Ascites

Ascites were evaluated as described in the literature.43 Briefly, the ascites fluid of mice were drained using absorbent paper strips and then weighed.

Isolation and Culture of Cells

Primary human HSCs were obtained from non-tumorous liver tissue after surgical resection of liver cancer, and were isolated by Liver Biotechnology Co., Ltd. Primary HSCs were isolated from the mice livers as described in our previous studies, where the primary HSCs at 0 days after isolation were used as quiescent HSCs, and those cultured in uncoated dishes for 7 days after isolation for spontaneous activation were used as activated HSCs.44,45 LX-2 cell line was provided by ProCell Life Science & Technology Co., Ltd.

The primary cells and LX-2 cells were cultured in Dulbecco’s modifier Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin in a 5% CO2 incubator at 37°C. UroA (10 μmol/L), alanyl-glutamine (GlutaMAX; Procell Life Science & Technology Co. Ltd), 10 μmol/L BPTES, 1 μmol/L CB-839 (Telaglenastat; MedChemExpress), 4 mmol/L DKG (Sigma Aldrich) were used to treat cells in subsequent experiments. To verify glutamine deprivation, LX-2 cells were cultured in a medium deprived of glutamine, and control cells were cultured in complete medium (4500 mg/L glucose and 4 mmol/L alanyl-glutamine).

Caspase-3 Activity Analysis

Cell apoptosis was measured using a caspase3 activity assay kit, according to the manufacturer’s instructions (Bolarbio). The cells were collected, lysed (15 minutes, 4 °C), and centrifuged at 15,000 × g for 10 minutes. The supernatants were collected according to the manufacturer’ s protocol.

Cell Proliferation and Cytotoxicity Assays

CCK-8 (APExBIO) was used to measure cell growth. Cells were seeded in 96-well plates at a density of 1 × 103 cells/well. The absorbance at 450 nm was recorded using a Synergy H1 Multi-Mode Microplate Reader (BioTek Instruments Inc) at the appointed time.

5-Ethynyl-2’-deoxyuridine

The 5-Ethynyl-2’-deoxyuridine (EdU)-488 assay kit (Beyotime) was used to detect cell proliferation. According to the manufacturer’s protocol, cells were incubated with 10 μmol/L EdU for 2 hours and were fixed in 4% paraformaldehyde and permeabilized using 0.3% Triton X-100. The samples were then incubated in the click reaction solution away from light for 30 minutes. Samples were imaged using an IX53 microscope (Olympus).

Glutaminase Activity Assay

Total protein in the cells was extracted and quantified using a commercial bicinchoninic acid (BCA) protein assay kit, and GLS activity was detected according to the manufacturer’s instructions (Micro Glutaminase Assay Kit, Solarbio).

Cell Immunofluorescence Staining

Cells were fixed in 4% paraformaldehyde and permeabilized with 0.1% Triton X-100. Then cells were blocked with 5% BSA for 1 hour at 25 °C and incubated with anti-p-MLC2 antibody (1:100). An Alexa Fluor 594-conjugated antibody (1:200; Invitrogen) was used as the secondary antibody. Images were captured by Olympus IX53 microscope.

Collagen Contraction Assays

Collagen contraction assays were used to measure the contractile activity of the HSCs. Cells were seeded in 24-well plates containing 1 mg/mL collagen type I (Corning). After collagen polymerization, the gel was gently dissociated from the edges of the well, media were added, and images were captured after 24 hours.46

Quantitative Real-time RT-PCR

Total RNA was extracted from tissues or cells using TRIzol Reagent (Invitrogen) according to the manufacturer’s instructions. RNA was reverse-transcribed using a 1-step cDNA kit (Toyobo). Real-time quantitative PCR of samples was performed using SYBR Green mastermix (Toyobo) with an ABI 7500 Sequence Detector. Threshold cycle numbers were normalized to the 18S or GAPDH values. The primers used in this study are listed in Table 2.Table 2 The Primer Sequences of Real-time Polymerase Chain Reaction

Genes	Forward primer (5′-3′)	Reverse primer (5′-3′)	
Mouse	
GAPDH	AGGTCGGTGTGAACGGATTTG	GGGGTCGTTGATGGCAACA	
 18S	AGTCCCTGCCCTTTGTACACA	CGATCCGAGGGCCTCACTA	
 α-SMA	GTTCAGTGGTGCCTCTGTCA	ACTGGGACGACATGGAAAAG	
 Col1α1	TAGGCCATTGTGTATGCAGC	ACATGTTCAGCTTTGTGGACC	
 TGF-β	GTGGAAATCAACGGGATCAG	ACTTCCAACCCAGGTCCTTC	
 PDGF	CCTTCCTCTCTGCTGCTACC	GAAGATCATCAAAGGAGCGG	
 TIMP-1	AGGTGGTCTCGTTGATTTCT	GTAAGGCCTGTAGCTGTGCC	
 F4/80	GTCTGTGGTGTCAGTGCAGG	GGATGTACAGATGGGGGATG	
 TNF-α	AGGGTCTGGGCCATAGAACT	CCACCACGCTCTTCTGTCTAC	
 IL-1β	TGCCACCTTTTGACAGTGATG	ATGTGCTGCTGCGAGATTTG	
Human	
 GAPDH	GGAGCGAGATCCCTCCAAAAT	GGCTGTTGTCATACTTCTCATGG	
18S	CCTGAGAAACGGCTACCACATC	AGAGTCCTGTATTGTTATTTTTCGTCACT	
 α-SMA	CCAGAGCCA TTGTCACACAC	CAGCCAAGCACTGTCAGG	
 Col1α1	CACACGTCTCGGTCATGGTA	AAGAGGAAGGCCAAGTCGAG	
 TGF-β	GCCCTGGACACCAACTATTGCT	AGGCTCCAAATGTAGGGGCAGG	
 PDGF	CTCGATCCGCTCCTTTGATGA	CGTTGGTGCGGTCTATGAG	
 TIMP-1	CTTCTGCAATTCCGACCTCGT	ACGCTGGTATAAGGTGGTCTG	
 GLS1	GGAAGCCTGCAAAGTAAACCC	CCAAAGTGCAGTGCTTCATCC	
α-SMA, α-smooth muscle actin; Col1α1, collagen type 1α1; GLS1, glutaminase1; IL-1β, interleukin-1β; PDGF, platelet-derived growth factor; TGF-β, transforming growth factor-β1; TNF-α, tumor necrosis factor-α; TIMP-1, tissue inhibitor of metalloproteinase-1.

Protein Extraction and Western Blot Analysis

Total protein was extracted from the cells using RIPA lysis buffer (Beyotime) containing protease and phosphatase inhibitors (Beyotime). Extracted protein was quantified using commercial BCA assay kit (Pierce BCA Protein Assay Kit, Thermo Fisher Scientific). Equal amounts of total protein were separated by SDS-PAGE and transferred onto PVDF membranes (Merck Millipore). The membranes were blocked with 5% non-fat milk for 1 hour at room temperature and incubated with specific primary antibodies, including α-SMA (1:1000; BOSTER), Col1α1 (1:1000, Cell Signaling Technology), GLS1 (1:1000, Proteintech), GAPDH (1:1000; Proteintech), overnight at 4 °C, following incubation for 1 hour with secondary antibodies. Finally, the membranes were detected using a Western enhanced chemiluminescence (ECL) substrate (Clarity Western ECL substrate, Bio-Rad).

Hepatic Macrophages Isolation

Hepatic macrophages were isolated as described previously.15 Mice livers were homogenized, fully digested with a mixture of 0.02 mg/mL DNase I (Sigma-Aldrich), 0.2 mg/mL collagenase type IV (Sigma-Aldrich), and 0.25 mg/mL Pronase E (Solarbio) in 50 mL Hank’s balanced salt solution via rotation at 37 °C for 25 minutes. Thereafter, the lysate was centrifuged at 800 × g (4 °C) for 10 minutes, and the supernatant was discarded. The cell pellet was resuspended in 40% Percoll (Sigma-Aldrich) and slowly dripped onto 80% Percoll (40% Percoll : 80% Percoll (v : v) = 3 : 1). Next, the nonparenchymal cell population was isolated by gradient centrifugation at 800 × g at 4 °C for 30 minutes. The middle cell suspension was cultured in RPMI-1640 supplemented with 1% penicillin-streptomycin and 20% FBS. After 2 hours, adhered hepatic macrophages were collected for subsequent experiments.

Primary Hepatocytes Isolation

Mice livers were perfused and digested through the portal vein by using collagenase type IV and collagenase P (Roche). Then the liver cells were filtered through a 70-μm nylon cell strainer, and the cell suspension was centrifuged at 500 rpm for 1 minute to re-suspend in M199 culture medium, 1% penicillin-streptomycin, and 10% FBS. The solution was added to wells coated with collagen. Cells were cultured with or without 10 μM UroA and collected to determine the ATP level.

Alanine Aminotransferase and ATP Level Assay

Mice serum levels of alanine aminotransferase (ALT) was measured using commercial kits according to the manufacturer’s instructions (Nanjing Jiancheng Bioengineering Institute). Enhanced ATP Assay Kit (Beyotime) was used in the ATP level assay. The cells were collected and lysed, centrifuged at 12,000 × g at 4 ºC for 5 minutes. The supernatant was collected, and ATP level was detected according to the instructions.

GDH, GOT2, and GPT2 Activity Assay

After treating the cells according to the experimental requirements, the cells were harvested, and the absorbance was measured to estimate activities of GDH (Solarbio), GOT2, and GPT2 (Nanjing Jiancheng Bioengineering Institute) according to the manufacturer’s instructions.

GLS1 Plasmid Transfection

Human overexpression wild-type and mutant GLS1 plasmids (rhGLS1D327A and rhGLS1Y394A) were purchased from GeneChem. Plasmids were extracted using an endotoxin-free plasmid extraction kit (TIANGEN). LX-2 cells were transfected with plasmids overexpressing GLS1 or the negative control using Lipofectamine 3000 (Invitrogen) as per the manufacturer’s instructions. The cells were then treated with UroA for 24 hours and collected for subsequent experiments.

SPRi Experiment

The PlexArray HT surface plasmon resonance imaging platform (SPRi) was used to determine the binding affinity of UroA with GLS1 protein (Abnova, Jhongli), BPTES were used as positive control. GLS1, rhGLS1D327A, and rhGLS1Y394A proteins (10 μg) were fixed on the sensors and activated with 10 mM NiSO4. Different concentrations of UroA (2.5, 5, 10, 20 μmol/L) and BPTES (10 μmol/L), diluted with PBS were flown at a rate of 30 μL/min. The reaction signal was collected and analyzed with PlexArray HT specialized software (Plexera Bioscience). Data were analyzed using Origin 9.0 (Origin Lab Corporation) and BIA Evaluation 4.1.1.

Molecular Docking of Compounds With GLS1

The 3D structure of UroA was obtained from PubChem, and the crystal structure of the GLS1 was obtained from the Protein Data Bank (PDB ID: 3VOZ). Schrödinger-Maestro (version 11.1) software was used to predict the binding poses of UroA and GLS1. The initial files of the ligand and protein were prepared and optimized by adding charges and hydrogen atoms and removing water molecules using Maestro. Receptor grid generation was used to choose a grid box that enclosed the whole binding site and dock ligand. Then UroA was docked into the presumptive binding sites (BPTES-binding site). The exported docking images and docking scores were used for analysis.

Bacterial Culture

L. murinus was purchased from BeNa Culture Collection and revived in Man-Rogosa-Sharpe (MRS) medium (HKM) at 37 °C with anaerobic environment.36 The bacteria were cultured for 12 to 16 hours of growth and collected at the late logarithmic phase. The precipitated bacteria were resuspended in PBS and administered to CPH mice by oral gavage (5×10ˆ7 CFU) once a day for 14 weeks.

Bacterial Composition Analysis

Bacterial DNA was extracted from mice feces using a Bacterial Genomic DNA Extraction Kit (Tiangen). The DNA was diluted into 5 ng/μL for quantitative real-time RT-PCR. For the microbial diversity analysis, the 16S rRNA gene V4 region was amplified and sequenced by using the Illumina sequencing platform (Bioacme Biological Technologies Corporation). The raw sequences were quality-controlled using QIIME2. Data were demultiplexed and clustered into species-level operational taxonomic units. Principal component, alpha diversity, and beta diversity analyses were performed using QIIME2.

Transcriptome Analysis

Total RNA was extracted by TRIzol reagent, according to the manufacturer’s instructions. RNA quantification was assessed by the NanoDrop2000 Spectrophotometer (Thermo Fisher). High-quality RNA was used for cDNA libraries construction and sequenced by Bioacme Biological Technologies Corporation. RNA sequencing libraries were generated using the KAPA Stranded RNA-Seq Kit for Illumina with multiplexing primers. Then sequencing was performed on Illumina Nova sequencer. The GO enrichment analysis of differentially expressed genes (DEGs) was implemented using the GOseq R package and GO terms. The genes with a fold change (FC) >1.5 and a P value < .05 when compared between groups were considered as DEGs.

Targeted Metabolomic Analysis

Targeted metabolomic analysis was performed by Metabo-Profile. Each sample was mixed with 120 μL methanol containing internal standard and extracted by ultrasonic method, and centrifuged at 18,000 × g for 20 minutes. Then, analysis and quantitation of samples were continued by using an ultra-performance liquid chromatography coupled to tandem mass spectrometry (UPLC–MS/MS) system (ACQUITY UPLC-Xevo TQ-S, Waters Corp). ACQUITY UPLC BEH C18 1.7 μM VanGuard pre-column (2.1 × 5 mm) and ACQUITY UPLC BEH C18 1.7 μM analytical column (2.1 × 100 mm) were used. Mobile Phases: A = water with 0.1% formic acid; and B = acetonitrile : IPA (v : v=70 : 30). Gradient conditions: 0-1 minutes (5% B), 1-11 minutes (5-78% B), 11-13.5 minutes (78-95% B), 13.5-14 minutes (95-100% B), 14-16 minutes (100% B), 16-16.1 minutes (100-5% B), 16.1-18 minutes (5% B) with 0.4 mL/minutes. The raw data by UPLC-MS/MS were processed using TMBQ software (v1.0, Metabo-Profile) to perform peak integration, calibration, and quantification for each metabolite. Statistical analyses were performed by the selfdeveloped platform iMAP (v1.0, Metabo-Profile).

UroA Analysis

The UroA levels in samples were detected by LC-MS/MS system. Nine-fold w/v of water was added to feces, 4-fold v/v of methhanol was added to plasma samples followed by ultrasonic extraction for 10 minutes, and then homogenate was centrifuged at 13,000 × g, 4 °C for 15 minutes. The supernatant collected and dried under nitrogen, then which was re-dissolved into the mobile phase for analysis. Thermo TSQ Vantage triple quadrupole mass spectrometer was used for detection. The chromatographic column was C18, 2 × 150 mm, 5 μm; with a column temperature of 45 °C. The flow rate was 0.3 mL/min; mobile phase A = water (0.01% TFA) and mobile phase B = acetonitrile (0.01% TFA). Data acquisition and analysis were performed using TraceFinder software (version 3.3 sp1; Thermo Fisher Scientific Corp).

Data Availability of Omics Analysis

Raw sequencing data associated with microbial diversity analysis is accessible at https://ngdc.cncb.ac.cn/gsa/s/0Nx3Xd9b, and the accession number is CRA010472; Raw sequencing data associated with transcriptome analysis is accessible at https://ngdc.cncb.ac.cn/gsa/s/Wmf38of3, and the accession number is CRA010222.

Statistical Analysis

Statistical analyses and mapping were performed using the GraphPad Prism 8 software. All results are expressed as mean ± standard error of mean (SEM). Statistical differences between 2 groups were analyzed using a 2-tailed, unpaired Student t-test, whereas differences between more than 2 groups were analyzed using 1-way analysis of variance (ANOVA). Spearman’s rank correlation test was used to analyze the correlations. The statistical sample sizes (n) are shown in the figure legends. P value < .05 is considered statistically significant.

CRediT Authorship Contributions

Rui Li (Conceptualization: Lead; Data curation: Lead; Funding acquisition: Supporting; Investigation: Lead; Software: Lead; Supervision: Equal; Writing – original draft: Lead)

Zhile Liu (Conceptualization: Supporting; Data curation: Equal; Software: Equal; Validation: Lead; Visualization: Lead; Writing – original draft: Equal)

Wensou Huang (Conceptualization: Supporting; Data curation: Supporting; Investigation: Equal; Resources: Equal; Validation: Equal; Visualization: Equal)

Yongjina Guo (Conceptualization: Equal; Data curation: Equal; Investigation: Equal; Software: Equal; Visualization: Supporting; Writing – original draft: Supporting)

Chan Xie (Data curation: Equal; Formal analysis: Equal; Resources: Equal; Supervision: Equal; Validation: Supporting)

Hongmei Wu (Resources: Equal; Supervision: Equal; Validation: Supporting; Visualization: Supporting)

Jianxin Liu (Data curation: Equal; Formal analysis: Supporting; Validation: Equal; Visualization: Supporting)

Xiaoyang Hong (Software: Supporting; Supervision: Supporting; Validation: Equal; Visualization: Supporting)

Xiaobin Wang (Supervision: Supporting; Validation: Supporting; Visualization: Supporting)

Jingjun Huang (Funding acquisition: Equal; Validation: Supporting)

Mingyue Cai (Funding acquisition: Supporting; Validation: Supporting; Visualization: Supporting)

Zhaoxiong Guo (Software: Equal; Visualization: Equal)

Licong Liang (Software: Supporting; Validation: Supporting)

Liteng Lin (Conceptualization: Equal; Funding acquisition: Equal; Investigation: Supporting; Resources: Equal; Writing – original draft: Supporting; Writing – review & editing: Lead)

Kangshun Zhu (Conceptualization: Lead; Funding acquisition: Lead; Supervision: Lead; Writing – review & editing: Lead)

Conflicts of interest The authors disclose no conflicts.

Funding This work was funded by 10.13039/501100001809 National Natural Science Foundation of China (82001929 , 82172043 , 82001930 , 82302326 ), 10.13039/501100021171 Basic and Applied Basic Research Foundation of Guangdong Province (2020A1515110654 ), Youth Innovative Talents Project of Guangdong Province University (2020KQNCX057 ), 10.13039/501100010031 Postdoctoral Research Foundation of China (2022M720909 ), Plan on Enhancing Scientific Research in GMU (2023.198).
==== Refs
References

1 Ginès P. Krag A. Abraldes J.G. Solà E. Fabrellas N. Kamath P.S. Liver cirrhosis Lancet 398 2021 1359 1376 34543610
2 Gracia-Sancho J. Marrone G. Fernández-Iglesias A. Hepatic microcirculation and mechanisms of portal hypertension Nat Rev Gastroenterol Hepatol 16 2019 221 234 30568278
3 De Gottardi A. Rautou P.E. Schouten J. VALDIG group Porto-sinusoidal vascular disease: proposal and description of a novel entity Lancet Gastroenterol Hepatol 4 2019 399 411 30957754
4 Gunarathne L.S. Rajapaksha H. Shackel N. Angus P.W. Herath C.B. Cirrhotic portal hypertension: from pathophysiology to novel therapeutics World J Gastroenterol 26 2020 6111 6140 33177789
5 Villanueva C. Albillos A. Genescà J. β blockers to prevent decompensation of cirrhosis in patients with clinically significant portal hypertension (PREDESCI): a randomised, double-blind, placebo-controlled, multicentre trial Lancet 393 2019 1597 1608 30910320
6 Yoo J.J. Kim S.G. Kim Y.S. Propranolol plus endoscopic ligation for variceal bleeding in patients with significant ascites: propensity score matching analysis Medicine (Baltimore) 99 2020 e18913
7 Vilaseca M. García-Calderó H. Lafoz E. The anticoagulant rivaroxaban lowers portal hypertension in cirrhotic rats mainly by deactivating hepatic stellate cells Hepatology 65 2017 2031 2044 28142199
8 Brusilovskaya K. Königshofer P. Lampach D. Soluble guanylyl cyclase stimulation and phosphodiesterase-5 inhibition improve portal hypertension and reduce liver fibrosis in bile duct-ligated rats United European Gastroenterol J 8 2020 1174 1185
9 de Franchis R. Bosch J. Garcia-Tsao G. Reiberger T. Ripoll C. Baveno VII Faculty Baveno VII - Renewing consensus in portal hypertension J Hepatol 76 2022 959 974 35120736
10 Albillos A. de Gottardi A. Rescigno M. The gut-liver axis in liver disease: pathophysiological basis for therapy J Hepatol 72 2020 558 577 31622696
11 Tranah T.H. Edwards L.A. Schnabl B. Shawcross D.L. Targeting the gut-liver-immune axis to treat cirrhosis Gut 70 2021 982 994 33060124
12 Zhang X. Coker O.O. Chu E.S. Dietary cholesterol drives fatty liver-associated liver cancer by modulating gut microbiota and metabolites Gut 70 2021 761 774 32694178
13 de Vos W.M. Tilg H. Van Hul M. Cani P.D. Gut microbiome and health: mechanistic insights Gut 71 2022 1020 1032 35105664
14 Gong S. Lan T. Zeng L. Gut microbiota mediates diurnal variation of acetaminophen induced acute liver injury in mice J Hepatol 69 2018 51 59 29524531
15 Li R. Xie L. Li L. The gut microbial metabolite, 3,4-dihydroxyphenylpropionic acid, alleviates hepatic ischemia/reperfusion injury via mitigation of macrophage pro-inflammatory activity in mice Acta Pharm Sin B 12 2022 182 196 35127379
16 García-Villalba R. Giménez-Bastida J.A. Cortés-Martín A. Urolithins: a comprehensive update on their metabolism, bioactivity, and associated gut microbiota Mol Nutr Food Res 66 2022 e2101019
17 D’Amico D. Andreux P.A. Valdés P. Singh A. Rinsch C. Auwerx J. Impact of the natural compound urolithin A on health, disease, and aging Trends Mol Med 27 2021 687 699 34030963
18 Denk D. Petrocelli V. Conche C. Expansion of T memory stem cells with superior anti-tumor immunity by Urolithin A-induced mitophagy Immunity 55 2022 2059 2073.e8 36351375
19 Lee H.J. Jung Y.H. Choi G.E. Urolithin A suppresses high glucose-induced neuronal amyloidogenesis by modulating TGM2-dependent ER-mitochondria contacts and calcium homeostasis Cell Death Differ 28 2021 184 202 32704090
20 Luan P. D’Amico D. Andreux P.A. Urolithin A improves muscle function by inducing mitophagy in muscular dystrophy Sci Transl Med 13 2021 eabb0319
21 Iwakiri Y. Trebicka J. Portal hypertension in cirrhosis: pathophysiological mechanisms and therapy JHEP Rep 3 2021 100316
22 Du K. Hyun J. Premont R.T. Hedgehog-YAP signaling pathway regulates glutaminolysis to control activation of hepatic stellate cells Gastroenterology 154 2018 1465 1479.e13 29305935
23 Choi W.M. Kim H.H. Kim M.H. Glutamate signaling in hepatic stellate cells drives alcoholic steatosis Cell Metab 30 2019 877 889.e7 31474565
24 Singh A. D’Amico D. Andreux P.A. Urolithin A improves muscle strength, exercise performance, and biomarkers of mitochondrial health in a randomized trial in middle-aged adults Cell Rep Med 3 2022 100633
25 Liu S. D’Amico D. Shankland E. Effect of Urolithin A supplementation on muscle endurance and mitochondrial health in older adults: a randomized clinical trial JAMA Netw Open 5 2022 e2144279
26 Singh A. D’Amico D. Andreux P.A. Direct supplementation with Urolithin A overcomes limitations of dietary exposure and gut microbiome variability in healthy adults to achieve consistent levels across the population Eur J Clin Nutr 76 2022 297 308 34117375
27 Tomás-Barberán F.A. González-Sarrías A. García-Villalba R. Urolithins, the rescue of “old” metabolites to understand a “new” concept: metabotypes as a nexus among phenolic metabolism, microbiota dysbiosis, and host health status Mol Nutr Food Res 61 2017
28 Zhang F. Wang F. He J. Regulation of hepatic stellate cell contraction and cirrhotic portal hypertension by Wnt/β-catenin signalling via interaction with Gli1 Br J Pharmacol 178 2021 2246 2265 33085791
29 Jalan R. De Chiara F. Balasubramaniyan V. Ammonia produces pathological changes in human hepatic stellate cells and is a target for therapy of portal hypertension J Hepatol 64 2016 823 833 26654994
30 Tao H. Li W. Zhang W. Urolithin A suppresses RANKL-induced osteoclastogenesis and postmenopausal osteoporosis by, suppresses inflammation and downstream NF-κB activated pyroptosis pathways Pharmacol Res 174 2021 105967
31 Yoo H.C. Yu Y.C. Sung Y. Han J.M. Glutamine reliance in cell metabolism Exp Mol Med 52 2020 1496 1516 32943735
32 Yin X. Peng J. Gu L. Targeting glutamine metabolism in hepatic stellate cells alleviates liver fibrosis Cell Death Dis 13 2022 955 36376267
33 Trivedi P. Wang S. Friedman S.L. The power of plasticity-metabolic regulation of hepatic stellate cells Cell Metab 33 2021 242 257 33232666
34 Simon J. Nuñez-García M. Fernández-Tussy P. Targeting hepatic glutaminase 1 ameliorates non-alcoholic steatohepatitis by restoring very-low-density lipoprotein triglyceride assembly Cell Metab 31 2020 605 622.e10 32084378
35 Li B. Cao Y. Meng G. Targeting glutaminase 1 attenuates stemness properties in hepatocellular carcinoma by increasing reactive oxygen species and suppressing Wnt/beta-catenin pathway EBioMedicine 39 2019 239 254 30555042
36 Hu J. Deng F. Zhao B. Lactobacillus murinus alleviate intestinal ischemia/reperfusion injury through promoting the release of interleukin-10 from M2 macrophages via Toll-like receptor 2 signaling Microbiome 10 2022 38 35241180
37 Pan F. Zhang L. Li M. Predominant gut Lactobacillus murinus strain mediates anti-inflammaging effects in calorie-restricted mice Microbiome 6 2018 54 29562943
38 Mandorfer M. Bota S. Schwabl P. Nonselective β blockers increase risk for hepatorenal syndrome and death in patients with cirrhosis and spontaneous bacterial peritonitis Gastroenterology 146 2014 1680 1690.e1 24631577
39 Tow W.K. Chee P.Y. Sundralingam U. Palanisamy U.D. The therapeutic relevance of urolithins, intestinal metabolites of ellagitannin-rich food: a systematic review of in vivo studies Nutrients 14 2022 3494 36079752
40 Soth M.J. Le K. Di Francesco M.E. Discovery of IPN60090, a clinical stage selective glutaminase-1 (GLS-1) inhibitor with excellent pharmacokinetic and physicochemical properties J Med Chem 63 2020 12957 12977 33118821
41 Boyer-Diaz Z. Aristu-Zabalza P. Andrés-Rozas M. Pan-PPAR agonist lanifibranor improves portal hypertension and hepatic fibrosis in experimental advanced chronic liver disease J Hepatol 74 2021 1188 1199 33278455
42 Cho S.S. Yang J.H. Lee J.H. Ferroptosis contribute to hepatic stellate cell activation and liver fibrogenesis Free Radic Biol Med 193 2022 620 637 36370962
43 Getachew Y. Cusimano F.A. Gopal P. Reisman S.A. Shay J.W. The synthetic triterpenoid RTA 405 (CDDO-EA) halts progression of liver fibrosis and reduces hepatocellular carcinoma size resulting in increased survival in an experimental model of chronic liver injury Toxicol Sci 149 2016 111 120 26443840
44 Lin L. Gong H. Li R. Nanodrug with ROS and pH dual-sensitivity ameliorates liver fibrosis via multicellular regulation Adv Sci (Weinh) 7 2020 1903138
45 Pradere J.P. Kluwe J. De Minicis S. Hepatic macrophages but not dendritic cells contribute to liver fibrosis by promoting the survival of activated hepatic stellate cells in mice Hepatology 58 2013 1461 1473 23553591
46 Gibb A.A. Huynh A.T. Gaspar R.B. Glutamine uptake and catabolism is required for myofibroblast formation and persistence J Mol Cell Cardiol 172 2022 78 89 35988357
