Regulation of Autophagy and Metabolism in Hepatocellular Carcinoma: Involvement of Wnt‐β‐Catenin Pathway
Sanjit K. Roy, Rashmi Srivastava, Nancy Landry, Shivam Srivastava, Anju Shrivastava, Rakesh K. Srivastava
TL;DR
This study shows that riluzole can inhibit liver cancer growth by targeting the Wnt/β-catenin pathway and inducing cell death.
Contribution
The study reveals riluzole's novel mechanism of inhibiting HCC via autophagy and metabolic pathway modulation.
Findings
Riluzole inhibits HCC cell viability and induces autophagy without harming normal hepatocytes.
Riluzole disrupts the Wnt/β-catenin pathway and reduces glucose and glutamine metabolism in HCC cells.
Riluzole increases ROS levels and disrupts mitochondrial function, leading to cell death.
Abstract
Most cancer cells rely on aerobic glycolysis to support uncontrolled proliferation and evade apoptosis and switch to glutamine metabolism to survive under hypoxic conditions. In hepatocellular carcinoma (HCC), the Wnt/β‐catenin pathway acts as a critical driver of metabolic reprogramming and stemness, primarily by enhancing aerobic glycolysis and altering the tumour microenvironment. The Wnt/β‐catenin pathway induces activation of enzymes required for glucose metabolism and regulates the expression of glutamate transporter and glutamine synthetase. The objective of this study is to examine the mechanism by which riluzole inhibits HCC growth and induces autophagy. The results indicate that riluzole inhibits cell viability and colony formation of HCC cells and cancer stem cells (CSCs) and induces apoptosis, while sparing human normal hepatocytes. Riluzole induces autophagic cell death by…
Genes, proteins, chemicals, diseases, species, mutations and cell lines named across the full text — each resolved to its canonical identifier and authoritative record.
Click any figure to enlarge with its caption.
FIGURE 1
FIGURE 2
FIGURE 3
FIGURE 4
FIGURE 5
FIGURE 6
FIGURE 7Peer Reviews
No public reviews on file for this paper yet. If you reviewed it on a platform where reviews are public (OpenReview, ICLR, NeurIPS, ICML), you can paste yours below so the community can read it here.
Videos
No videos yet. Explain this paper in a talk, walkthrough, or lecture? Add one.
Taxonomy
TopicsCancer, Hypoxia, and Metabolism · Autophagy in Disease and Therapy · Cancer, Stress, Anesthesia, and Immune Response
Introduction
1
Hepatocellular carcinoma (HCC) is the 6th most common and 3rd leading cause of cancer‐related death worldwide, often manifesting at the advanced stage when cure is no longer possible. Liver cancer incidence has more than tripled since 1980. Liver cancer death rates have increased by almost 3% per year since 2000. According to the American Cancer Society's 2025 report, approximately 42,220 new cases will be diagnosed, and approximately 30,090 people will die of liver cancer. The 5‐year survival rate is around 10%–20% due to late‐stage diagnosis. HCC is a highly complex and heterogeneous disease. Surgical resection is the primary treatment option for patients with HCC. However, it is often not feasible due to advanced metastatic disease. Despite increased efforts to understand the biology of HCC, we have not made significant progress in treating it. There are no effective drugs for the treatment of HCC. Furthermore, the existence of cancer stem cells (CSCs) in the liver hinders the development of new drugs because CSCs can contribute to therapy failure and drug resistance. Therefore, there is an unmet need to develop effective, non‐toxic anti‐cancer drugs that target both cancer cells and CSCs. Riluzole is an approved drug for amyotrophic lateral sclerosis (ALS) and is under testing in clinical trials as a substance for melanoma therapy. Multi‐kinase inhibitors sorafenib and regorafenib are currently the only approved systemic drugs for HCC, and unfortunately, they are marginally effective in extending survival. Therefore, new targeted therapies for HCC are urgently needed.
Oxidative stress and dysregulated redox signalling are key mechanisms linking metabolic liver disease to HCC [1, 2]. In addition to causing DNA damage, long‐term accumulation of reactive oxygen species (ROS) activates proliferative and survival pathways that drive carcinogenesis [3]. Cancer cells with dysregulated metabolism demonstrate upregulation of glucose uptake (e.g., Glut1 and Glut3) due to increased expression of glucose transporters, increased enzymatic activity (e.g., hexokinase 2, phosphofructokinase‐1, pyruvate kinase M2) of the glycolytic pathway, and mitochondrial inhibition (effectively shunting pyruvate toward lactate production) [4, 5]. In addition to the dependency on glycolysis, cancer cells are addicted to increased rates of hypoxia‐inducible factor (HIF)‐dependent glutamine metabolism [3, 6, 7]. In cancer cells, glutaminolysis acts as a compensatory mechanism to maintain a functional Krebs (Tricarboxylic acid) cycle when glucose is diverted toward lactate production [8, 9, 10]. Furthermore, targeting mitochondrial glutaminase activity inhibits oncogenic transformation [11].
In hypoxic tumour environments, HIF‐1 serves as a master regulator that drives metabolic reprogramming toward glycolysis to ensure cell survival. HIF regulates Lactate Dehydrogenase A (LDHA) and Monocarboxylate Transporter 4 (MCT4), which leads to the secretion of lactate from cells. Increased LDHA activity in tumour cells causes an upregulation of NADH relative to NAD^+^. Because cancer cells exhibit a high glycolytic rate, there is enhanced shuttling of glycolytic intermediates to the pentose phosphate pathway, resulting in augmented NADPH production that mitigates oxidative stress. The ratio of NAD(P)H to NAD(P)^+^ determines ATP production. While normal differentiated cells primarily rely on mitochondrial oxidative phosphorylation to generate energy, most cancer cells rely on aerobic glycolysis. Pyruvate kinase M2 (PKM2) is an essential regulator of the Warburg effect. Specific cancer‐associated mutations enable cancer cells to acquire and metabolise nutrients in a manner that facilitates proliferation rather than efficient ATP production.
In HCC, the Wnt/β‐catenin pathway acts as a critical driver of metabolic reprogramming and stemness, primarily by enhancing glycolysis and altering the tumour microenvironment [12]. It promotes a glycolytic phenotype through several key mechanisms: direct enzyme regulation [increased activation of Pyruvate Dehydrogenase Kinase 1 (PDK1) inhibits the conversion of pyruvate to acetyl‐CoA, thereby shifting glucose metabolism toward lactate production rather than oxidative phosphorylation], target gene activation (c‐Myc‐dependent upregulation of Glut1, Glut3, LDHA and PKM2) and lactate export (increased expression of MCT1 facilitates lactate transport and promotes glycolysis‐mediated tumour growth) [13, 14]. Aberrant activation of this pathway occurs in approximately 30%–40% of HCC cases, leading to enhanced tumour growth, metastasis and drug resistance [15]. Dysregulation generally results from mutations in pathway components or altered expression of Wnt ligands and receptors [16, 17, 18]. In the absence of Wnt ligands, the destruction complex (GSK3, CK1α, axin and APC) promotes the phosphorylation of β‐catenin, resulting in its ubiquitylation and subsequent degradation [19, 20, 21]. The binding of Wnt ligands with Frizzled (Fzd) receptors and the Wnt co‐receptor LRP5 or LRP6 activates the Dishevelled (Dvl) cytoplasmic phosphoprotein, which inhibits β‐catenin phosphorylation and degradation. The translocation of β‐catenin into the nucleus causes binding to TCF/LEF proteins. It acts as a transcriptional co‐activator, modulating the expression of target genes such as Cyclin D1, COX‐2, PDK, MCT‐1 and cMyc [22, 23, 24, 25]. There are no studies examining the effects of riluzole on metabolism through the Wnt signalling pathway in HCC.
The objective of this study is to examine the molecular mechanisms by which riluzole regulates metabolism and autophagy in HCC. Our data demonstrate that riluzole inhibits cell proliferation and induces autophagic cell death through Beclin1 and Atg5. Riluzole inhibits Wnt‐β‐catenin/TCF‐LEF pathway and suppresses the expression of PDK, MCT1, cMyc, AXIN, Bcl‐2 and CyclinD1. Riluzole suppresses glucose transporter (Glut1 and Glut3) expression, lactate dehydrogenase A (LDHA) expression, glucose uptake and NAD+ levels. Riluzole inhibits glutamate release, which reduces the antioxidant glutathione (GSH), leading to increased reactive oxygen species (ROS) and cell death, a mechanism exploited in anti‐cancer therapies. Riluzole disrupts mitochondrial homeostasis by inhibiting Bcl‐2 and upregulating Bax expression. In conclusion, riluzole inhibits HCC growth by regulating glucose and glutamine metabolism and inducing autophagic cell death, thereby highlighting its therapeutic potential for HCC treatment.
Materials and Methods
2
Reagents
2.1
JC‐1 mitochondrial membrane potential assay kit, and Pierce BCA Protein Assay Kit were purchased from Thermo Fisher (Grand Island, NY). BD Matrigel was purchased from BD Bioscience (San Jose, CA). TRIzol and polybrene were purchased from Invitrogen (Grand Island, NY). CellTiter‐Glo Luminescent Cell Viability Assay and Luciferase assay kits were purchased from Promega Corporation (Madison, WI). All other chemicals were purchased from Sigma‐Aldrich (St. Louis, MO).
Cell Culture
2.2
Human HCC cell lines (HepG2, Hep3B, SNU‐382 and SNU‐475) were purchased from American Type Culture Collection (ATCC), Manassas, VA, and were authenticated by the vendor using short tandem repeat (STR) profiling. Cell lines were grown in Dulbecco's Modified Eagle's Medium (DMEM) with 10% Fetal Bovine Serum (HyClone) and antibiotics. Human liver cancer stem cells (CSCs) were isolated from primary tumours and cultured in stem cell culture medium as per the supplier's instructions (Celprogen, Torrance, CA). Human normal hepatocytes were purchased from ATCC. All cells were Mycoplasma free (as per detection kit from Lonza) and used within 3 months of continuous passage.
Lentiviral Particle Production and Transduction
2.3
The protocol for lentivirus production and transduction was described elsewhere [26, 27]. Briefly, 293 T cells were transfected with 4 μg of plasmid and 4 μg of the lentiviral vectors using Lipofectamine‐3000 as per protocol from Invitrogen. After collecting supernatant, PEG‐it virus precipitation solution (SBI System Biosciences) was added, and viral particles were collected through ultracentrifugation. CSCs and cancer cell lines were transduced with lentiviral particles with 6 μg/mL polybrene [28, 29].
Apoptosis
2.4
HCC cells were treated with or without riluzole for various time points. Apoptosis was measured by TUNEL (terminal deoxynucleotidyl transferase (TdT)‐mediated dUTP nick end labelling) assay as per the manufacturer's protocol (Life Technologies, Grand Island, NY).
Cell Cycle Analysis
2.5
Cell cycle analysis was performed as described elsewhere [30]. In brief, HCC cells (5 × 10^5^) were seeded in cell culture dishes. After 24 h, the medium was removed and replaced with fresh medium containing riluzole 0–20 μM for 48 h. Cell cycle analysis was performed by measuring propidium iodide (PI) fluorescence in ethanol‐fixed cells. In brief, cells were harvested by trypsinization and fixed with cold 70% ethanol for 24 h. Cells were rinsed 3 times with ice‐cold PBS and resuspended in 1 mL of permeabilizing solution (Triton X 100 (0.25%), sodium azide (0.01%) and RNAs A (100 μg/μL Sigma‐Aldrich) in PBS for 10 min). Cells were rinsed once with PBS, resuspended with 1 mL of PBS with PI (2.5 mg/mL), and incubated for 15 min at 4°C. Cell cycle analysis was performed using a flow cytometer (Becton Dickinson).
Cell Viability
2.6
The CellTiter‐Glo Luminescent Cell Viability Assay is a homogeneous method for determining the number of viable cells in culture by quantifying ATP, an indicator of metabolically active cells. Cells were seeded in a 96‐well plate and treated with or without riluzole for various time points. Cell viability was measured by CellTiter‐Glo assay as per the manufacturer's protocol (Promega Corporation, Madison, WI).
Colony Formation Assay
2.7
Colony formation assays were performed as described elsewhere [27, 31]. In brief, HCC cells were seeded at a low density into 6‐well plates for about 3 weeks. Following incubation, colonies were fixed with methanol, stained with 0.5% crystal violet, and counted under a microscope.
Measurement of Mitochondrial Membrane Potential
2.8
Mitochondrial membrane potential was measured as we described elsewhere [10]. In brief, HCC cells were seeded in a 96‐well plate and treated with or without riluzole for various time points. MitoProbe JC‐1 Assay Kit measured mitochondrial membrane potential as per manufacturer's protocol (Thermo Fisher Scientific).
Western Blot Analysis
2.9
HCC cells were harvested, washed in phosphate buffer saline (PBS), and then lysed in ice‐cold lysis buffer solution [50 mM Tris–HCl pH 7.4, 15 mM MgCl_2_, 150 mM NaCl, 1 mM EDTA, 0.5% (v/v) sodium deoxycholate, 0.5% (v/v) NP‐40 and 0.1% (v/v) SDS], containing 1 mM sodium orthovanadate, 1 mM phenylmethylsulfonyl fluoride (PMSF), 1 mM sodium fluoride (NaF), 1 mM sodium pyrophosphate and a cocktail of protease inhibitors (Sigma‐Aldrich, St. Louis, MO). Lysis was performed on ice for 30 min, and the samples were centrifuged at 17,000×g for 30 min. The protein content was quantified with the bicinchoninic acid (BCA) Protein Assay kit (Thermo Fisher Scientific). Equal amounts of proteins (40 μg) were fractionated on an SDS polyacrylamide gel (SDS‐PAGE) by electrophoresis and then transferred to PVDF membranes with the Trans‐Blot Turbo Transfer System (Bio‐Rad). Membranes were then blocked for 1 h with 5% bovine serum albumin (BSA) in PBS and incubated with primary antibodies overnight at 4°C. After 1 h incubation time with species‐specific antibodies conjugated with horseradish peroxidase (Bio‐Rad), chemiluminescence was measured by using a chemiluminescent substrate detection system ECL and the iBright Imaging System (Invitrogen).
TCF/LEF Reporter Assay
2.10
TCF/LEF transcriptional activity was measured as we described elsewhere [10]. In brief, lentiviral particles expressing cop‐GFP and luciferase genes (TCF/LEF‐mCMV‐EF1‐Neo) were prepared as described elsewhere [32]. HCC cells were transduced with lentiviral particles containing the gene of interest. Transduced cells (5–10,000 cells per well) were seeded in 96‐well plates and treated with or without riluzole for various time points. At the end of the incubation period, luciferase reporter activity was measured as per the manufacturer's instructions (Promega Corp., Madison, WI).
Glucose Uptake
2.11
Glucose uptake by cancer cells was measured as we described elsewhere [10]. In brief, HCC cells were labelled with 2.5 μg/mL of 2‐deoxy‐2‐[(7‐nitro‐2,1,3‐benzoxadiazol‐4‐yl)amino]‐D‐glucose (2‐NBD Glucose) for 30 min, and treated with riluzole for 24 h. Cells were washed and resuspended in PBS. Fluorescence was measured (excitation 544 nm and emission at 590 nm).
Measurement of NAD+
2.12
NAD+ levels were measured as we described elsewhere [10]. In brief, HCC cells were seeded in 24‐well plate and treated with riluzole (0–10 μM) for 1 h and total cellular NAD+ concentration was measured at 450 nm by a NAD/NADH assay kit (Cayman).
Glutamate Release Assay
2.13
HepG2, Hep3B, SNU‐382 and SNU‐475 cells and CSCs were treated with riluzole (0–10 μM) for 24 h. Glutamate release was measured using the Amplex Red Glutamic Acid/Glutamate Oxidase Assay Kit with a fluorometer, with excitation at 540 nm and emission at 590 nm (Invitrogen).
Measurement of Intracellular GSH
2.14
HepG2, Hep3B, SNU‐382 and SNU‐475 cells and CSCs were treated with riluzole (0–10 μM) for 24 h. Intracellular total GSH was detected by measuring the product of glutathionylated DTNB at 405 nm (GSH assay kit, Cayman Chemical).
Measurement of Reactive Oxygen Species
2.15
HepG2 and Hep3B cells were pre‐treated with NAC (3 mM) for 2 h, followed by treatment with riluzole (0–10 μM) for 24 h. After that, cells were labelled with 2′,7′‐dichlorofluorescein diacetate (DCFDA/H2DCFDA) and ROS production was measured with a fluorometer using excitation at 495 nm and emission at 529 nm (Cellular Reactive Oxygen Species Detection Assay Kit, Abcam).
Quantitative Real‐Time PCR
2.16
RNA extraction, cDNA synthesis and quantitative polymerase chain reaction (PCR) were carried out as previously described [27]. In brief, total RNA was first extracted using TRIzol reagent (Invitrogen), and cDNA was generated using the Reverse Transcription System (Promega) in a 20 μL reaction containing 1 μg of total RNA. A 0.5 μL aliquot of cDNA was amplified by Fast SYBR Green PCR Master Mix (Applied Biosystems) in each 20 μL reaction. PCR reactions were run on the Quant Studio 5 real‐Time PCR system (Applied Biosystems). The gene‐specific primers were described elsewhere [10].
Quantification and Statistical Analysis
2.17
Graph Pad Prism software version 10 (La Jolla, CA, USA) was used for statistical analysis. Statistical significance was estimated using a two‐tailed Student's *t‐*test for comparisons between two groups or one‐way Analysis of Variance (ANOVA) followed by the indicated post hoc analysis for multiple comparisons in each case. The threshold for statistical significance is p < 0.05, unless otherwise stated. The mean ± SD or SEM was calculated for each experimental group, unless otherwise specified.
Results
3
Riluzole Inhibited Cell Viability and Colony Formation, and Induced Apoptosis in HCC Cell Lines and CSCs While Sparing Human Normal Hepatocytes
3.1
Riluzole is being used for the treatment of ALS. However, its preclinical potential for HCC is not well understood. We first examined the effects of riluzole on HCC cells and CSCs. HCC CSCs (CD133^+^ and CD44^+^) were isolated from primary tumours and characterised as we described earlier [27, 33]. HCC CSCs expressed CD44 and CD133 cell surface markers (Figure 1). Riluzole inhibited cell viability in HepG2, Hep3B and CSCs (Figure 1B), but had no effect on the viability of human normal hepatocytes (Figure 1C).
*Inhibition of cell viability and colony formation, and induction of apoptosis by riluzole. (A) Expression of stem cell markers in HCC CSCs. CSCs were stained with CD44 and CD133 antibody and the expression of CD44 and CD133 was measured by immunocytochemistry. (B, C) HCC cell lines (HepG2 and Hep3B), cancer stem cells (CSCs), and human normal hepatocytes were seeded in 96‐well plates, treated with or without riluzole (0–40 μM) for 72 h, and cell viability was measured by CellTiter‐Glo Luminescent Cell Viability Assay (Promega). (D) Colony formation. HCC cell lines (HepG2, Hep3B, SNU‐382 and SNU‐475) and CSCs were treated with riluzole (0–20 μM). Number of colonies formed at 21 days was counted under a microscope. (E) Normal human hepatocytes, HCC cell lines (HepG2 and Hep3B) and CSCs were treated with riluzole (0–20 μM) for 72 h. Apoptosis was measured by TUNEL assay. Data represent mean ± SD (n = 4). , # and % = significantly different from control and each other; p < 0.05.
Since riluzole inhibited HCC cell viability, we next examined its effects on colony formation. Riluzole inhibited colony formation by HepG2, Hep3B, SNU‐382, SNU‐475 and CSCs in a dose‐dependent manner (Figure 1D). Riluzole‐induced apoptosis in HepG2, Hep3B and CSCs (Figure 1E). Interestingly, maximum induction of apoptosis was seen in Hep3B cells, whereas the response of HCC CSCs to riluzole was the lowest. By comparison, riluzole did not affect normal hepatocytes.
Riluzole‐Induced Autophagy in HCC Through Beclin and ATG
3.2
Autophagy is a self‐preservation mechanism in which cells remove/digest damaged intracellular organelles to generate energy for survival [34, 35]. Cancer cells adapt this mechanism to survive and grow in a competitive tumour microenvironment. Autophagosomes serve as essential carriers in the autophagy pathway, enabling cells to adapt to metabolic stress and maintain homeostasis. After formation, autophagosomes deliver cytoplasmic components to the lysosomes. The outer membrane of an autophagosome fuses with a lysosome to form an autolysosome. Lysosomal hydrolases degrade the autophagosome‐delivered contents and their inner membrane [36]. We next examined whether riluzole‐induced cell death occurs through autophagy. HepG2 were transfected with a plasmid encoding mRFP‐GFP‐LC3 and treated with various doses of riluzole. The effects of riluzole on autophagy were assessed by measuring the number of autophagosomal LC3 (GFP+/RFP+) and autolysosomal LC3 (GFP−/RFP+) puncta. Riluzole‐induced the formation of autophagosomal LC3 (GFP+/RFP+) and autolysosomal LC3 (GFP−/RFP+) puncta in HepG2 cells in a dose‐dependent manner (Figure 2).
*Riluzole induces autophagy in HCC through Beclin and ATG. (A) Riluzole induces autophagy. HepG2 were transfected with a plasmid encoding mRFP‐GFP‐LC3 and treated with various doses of riluzole (0–20 μM) for 24 h. Cells were visualised with a fluorescence microscope, and autophagosomal LC3 puncta (GFP+/RFP+) and autolysosomal LC3 puncta (GFP−/RFP+) were quantified. Data represent mean ± SD (n = 4). , @, # and $ = significantly different from control and each other; p < 0.05. (B) Autophagy inhibitors 3‐methyladenine (3‐MA), chloroquine (CQ) and bafilomycin A1 (BA) enhance riluzole‐induced apoptosis. HepG2 were pre‐treated with 3‐MA, CQ, or BA for 2 h followed by treatment with riluzole (0–20 μM) for 48 h. Apoptosis was measured by TUNEL assay. (C) Expression of Beclin and ATG5. HepG2 cells were transfected with either non‐targeting control, Beclin1/Crispr KO, or ATG5/Crispr KO plasmid. After transfection, cells were harvested to isolate mRNA. qRT‐PCR was performed to measure mRNA expression of Beclin1 and ATG. Data represent mean ± SD (n = 4). * = significantly different from control; p < 0.05. (D) Inhibition of Beclin1 or Atg5 enhances riluzole‐induced apoptosis. HepG2/NTC, HepG2/Beclin1/Crispr KO and HepG2/Atg5/Crispr KO cells were treated with riluzole (0–10 μM) for 48 h. Apoptosis was measured by TUNEL assay. Data represent mean ± SD (n = 4). * and # = significantly different from NTC; p < 0.05.
We next confirmed the autophagic cell death by using autophagy inhibitor 3‐methyladenine (3‐MA), chloroquine (CQ) and bafilomycin A1 (BA). HepG2 cells were pre‐treated with 3‐MA, CQ, or BA for 2 h followed by treatment with riluzole (0–20 μM) for 48 h. Autophagy inhibitors 3‐MA, CQ, and BA enhanced riluzole‐induced apoptosis. These data suggest that riluzole‐induced cell death occurs through autophagy.
Beclin1 and Atg5 genes play significant roles in mediating autophagic cell death. We therefore examined the involvement of Beclin1 and ATG in riluzole‐induced autophagy.
HepG2 cells were transfected with either non‐targeting control, Beclin1/Crispr knockout, or ATG5/Crispr knockout plasmid. Transfection of HepG2 cells with Beclin1/Crispr/cas9 or Atg5/Crispr/Cas9 inhibited the expression of Beclin1 or ATG5 gene respectively (Figure 2C).
We next treated HepG2/NTC, HepG2/Beclin1/Crispr KO and HepG2/Atg5/Crispr KO cells with riluzole (0–10 μM) for 48 h. Riluzole alone induced apoptosis in HepG2/NTC cells. Inhibition of Beclin1 or Atg5 by Crispr/Cas9 technique enhanced riluzole‐induced apoptosis. These data suggest that Beclin1 and Atg5 are involved in riluzole‐induced autophagic cell death.
Riluzole Inhibited Components of β‐Catenin/TCF‐LEF Pathway
3.3
The Wnt/β‐catenin signalling pathway plays a crucial role in cell proliferation, differentiation and stem cell maintenance [37]. Since riluzole inhibited cell proliferation and induced apoptosis in HCC, we next sought to examine the effects of riluzole on the Wnt/β‐catenin signalling pathway. We measured the expression of genes involved in this pathway by q‐RT‐PCR and the transcriptional activity of TCF‐LEF1 by luciferase assay. Riluzole inhibited the expression of β‐catenin, Wnt3a, Wnt5a, Axin1, TCF, LEF and GSK3β in HepG2 cells (Figure 3A–G). Since riluzole inhibited gene expression in the β‐catenin pathway, we next sought to measure TCF‐LEF1 transcriptional activity, which is regulated by β‐catenin. Riluzole inhibited TCF‐LEF1 transcriptional activity in HepG2 cells (Figure 3H). Similarly, riluzole inhibited the expression of β‐catenin, Wnt3a, Wnt5a, Axin1, TCF, LEF and GSK3β in HepG2 cells (Figure 3I–O). Furthermore, riluzole inhibited TCF‐LEF1 transcriptional activity in Hep3B cells (Figure 3P). These data suggest that riluzole‐induces autophagic cell death by suppressing the Wnt/β‐catenin signalling pathway.
*Inhibition of β‐catenin/TCF‐LEF1 pathway by riluzole. (A–G) HepG2 cells were treated with riluzole (0–20 μM) for 36 h. RNA was extracted, and the expression of β‐catenin, Wnt3a, Wnt5a, Axin1, TCF, LEF and GSK3β was measured by q‐RT‐PCR. Data represent mean ± SD (n = 4). * and # = significantly different from control and each other; p < 0.05. (H) HepG2 cells were transduced with TCF/LEF1‐responsive GFP/firefly luciferase viral particles (pGreen Fire1‐TCF/LEF1 with EF1, System Biosciences). After transduction, cells were treated with riluzole (0–20 μM) for 36 h. TCF/LEF1 reporter activity was measured by luciferase assay, as we described [38]. Data represent mean ± SD (n = 4). *, # and % = significantly different from control and each other; p < 0.05. (I–O) Hep3B cells were treated with riluzole (0–40 μM) for 36 h. RNA was extracted, and the expression of β‐catenin, Wnt3a, Wnt5a, Axin1, TCF, LEF and GSK3β was measured by q‐RT‐PCR. Data represent mean ± SD (n = 4). * and # = significantly different from control and each other; p < 0.05. (P) Hep3B cells were transduced with TCF/LEF1‐responsive GFP/firefly luciferase viral particles (pGreen Fire1‐TCF/LEF1 with EF1, System Biosciences). After transduction, cells were treated with riluzole (0–20 μM) for 36 h. TCF/LEF1 reporter activity was measured by luciferase assay, as we described [38]. Data represent mean ± SD (n = 4). , # and % = significantly different from control and each other; p < 0.05.
Riluzole Inhibited the Expression of TCF‐LEF1 Target Genes in HCC
3.4
The Wnt/β‐catenin signalling pathway plays a crucial role in cell proliferation, differentiation, and stem cell maintenance [37]. Aberrant activation of this pathway has been found in approximately 30%–40% of HCC cases [15]. The activity of β‐catenin/TCF is often deregulated in HCC, leading to the activation of genes whose dysregulation has significant consequences for HCC development. Therefore, identifying the target genes of Wnt signalling is essential for understanding β‐catenin‐mediated carcinogenesis. Since riluzole inhibited the Wnt/β‐catenin/TCF‐LEF1 pathway, we next sought to examine the effects of riluzole on downstream targets of the Wnt pathway. These downstream targets regulate the cell cycle, cell growth, components of the Wnt pathway, and metabolism. Riluzole inhibited the expression of cyclin D1 in both HepG2 and Hep3B cells (Figure 4A,F), and caused growth arrest at G2/M stage of cell cycle (data not shown). Transcription factor TCF/LEF induces the expression of cMyc, Axin2, cMyc and monocarboxylate lactate transporter 1 (MCT‐1). Riluzole inhibited the expression of Axin2, cMyc, MCT1 and DNMT1 in both HepG2 and Hep3B cells (Figure 4B–FJ). These data suggest that riluzole can regulate the cell cycle, cell proliferation, and glucose metabolism in HCC by modulating the expression of components of the Wnt pathway and its downstream targets.
*Inhibition of TCF‐LEF1 target genes and induction of growth arrest at G2/M stage by riluzole. (A–E) HepG2 cells were treated with riluzole (0–20 μM) for 36 h. RNA was isolated and q‐RT‐PCR was used to measure the expression of cyclin D1, Axin2, cMyc, MCT1 and DNMT1. Data represent mean ± SD (n = 4). *, # and % = significantly different from control and each other; p < 0.05. (F–J) Hep3B cells were treated with riluzole (0–20 μM) for 36 h. RNA was isolated, and the expression of cyclin D1, Axin2, cMyc, MCT1 and DNMT1 was measured by qRT‐PCR. Data represent mean ± SD (n = 4). , # and % = significantly different from control and each other; p < 0.05.
Riluzole Inhibited Bcl‐2, Induced Bax Expression and Disrupted Mitochondrial Membrane Potential in HCC Cells
3.5
Mitochondrial dysfunction contributed to HCC cell death, characterised by an increased BAX/Bcl‐2 ratio and positive TUNEL signals [39, 40]. Bcl‐2 family members play a significant role in regulating mitochondrial functions, including mitochondrial membrane potential [41, 42]. We, therefore, measured the expression of anti‐apoptotic Bcl‐2 and proapoptotic Bax [43]. Riluzole inhibited Bcl‐2 and induced Bax expression in HepG2 cells (Figure 5A,B). Similarly, riluzole inhibited Bcl‐2 and induced Bax expression in Hep3B cells (Figure 5C,D). Since Bcl‐2 and Bax act at the level of mitochondria and regulate permeability transition, we measured the effects of riluzole on mitochondrial membrane potential in HCC cells. Treatment of HepG2 and Hep3B cells with riluzole significantly inhibited mitochondrial membrane potential in a time‐dependent manner (Figure 5E,F). These data suggest that mitochondria are involved in riluzole‐induced apoptosis by modulating Bcl‐2 and Bax expression.
*Inhibition of Bcl‐2, induction of Bax and drop in mitochondrial membrane potential by riluzole. (A–C) HepG2 cells were treated with riluzole (0–20 μM) for 36 h. RNA was isolated, and the expression of Bcl‐2 and Bax was measured by qRT‐PCR. Data represent mean ± SD (n = 4). *, # and % = significantly different from control and each other; p < 0.05. (D–F) Hep3B cells were treated with riluzole (0–20 μM) for 36 h. RNA was isolated, and the expression of Bcl‐2 and Bax was measured by qRT‐PCR. Data represent mean ± SD (n = 4). *, # and % = significantly different from control and each other; p < 0.05. (G) Measurement of mitochondrial membrane potential. HepG2 cells were treated with or without riluzole (20 μM), and mitochondrial membrane potential was measured over time. Data represent mean ± SD (n = 4). , @, #, , %, & and ** = significantly different from control and each other; p < 0.05.
Riluzole Inhibited Glucose Transporter (Glut1 and Glut3), PDK1 and Lactate Dehydrogenase A (LDHA‐A) Expression, Glucose Uptake and Nicotinamide Adenine Dinucleotide (NAD+) Level
3.6
The dysregulated canonical Wnt/β‐catenin pathway can modify metabolic enzymes [44, 45, 46]. Wnt signalling increases the expression of glucose transporters and key metabolic enzymes, thereby enhancing glucose uptake and glycolysis. In metabolic reprogramming of ‘glucose addiction’, liver cancer cells become heavily dependent on glucose to fuel their rapid growth and survival. This shift allows cancer cells to survive even in nutrient‐poor environments. Since cancer cells and CSCs are addicted to aerobic glycolysis and lactate (Warburg effect), we measured the expression of glucose transporter 1 and 3 (Glut1 and Glut3), PDK1, lactate dehydrogenase‐A (LDH‐A) expression, glucose uptake, and nicotinamide adenine dinucleotide (NAD+) level in HepG2 and Hep3B cells. Riluzole inhibited the expression of Glut1, Glut3, PDK1, LDH‐A, and PKM2 in both HepG2 and Hep3B cells (Figure 6A–J).
Riluzole inhibits Glut1, Glut3, PDK1, LDH‐A and PKM2 expression, glucose uptake, and NAD+ level. (A–E) HepG2 cells were treated with riluzole (0–20 μM) for 36 h. RNA was extracted, and the expression of Glut‐1, Glut‐3, PDK1, LDH‐A and PKM2 was measured by q‐RT‐PCR. Data represent mean ± SD (n = 4). * and # = significantly different from control; p < 0.05. (F–J) Hep3B cells were treated with riluzole (0–20 μM) for 36 h. RNA was extracted, and the expression of Glut‐1, Glut‐2, PDK1, LDH‐A and PKM2 was measured by q‐RT‐PCR. Data represent mean ± SD (n = 4). * and # = significantly different from control; p < 0.05. (K) Glucose uptake. HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs were labelled with 2.5 μg/mL of 2‐deoxy‐2‐[(7‐nitro‐2,1,3‐benzoxadiazol‐4‐yl)amino]‐d‐glucose (2‐NBD Glucose) for 30 min and treated with riluzole (0–10 μM) for 24 h. Cells were washed and resuspended in PBS. Fluorescence was measured (excitation 544 nm and emission at 590 nm). Data represent mean ± SD (n = 4). * = significantly different from respective control; p < 0.05. (L) HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs were treated with riluzole (0–10 μM) for 1 h and total cellular NAD+ concentration was measured at 450 nm by a NAD/NADH assay kit (Cayman). Data represent mean ± SD (n = 4). * = significantly different from respective control; p < 0.05.
Liver cancer cells often stop using low‐affinity enzymes (such as GCK) and instead activate high‐affinity enzymes [such as Hexokinase 2 (HK2)], to pull glucose from the bloodstream aggressively. Since riluloze inhibited the expression of Glut1 and Glut3, we sought to examine the effects of riluzole on glucose uptake by HCC. Riluzole inhibited glucose uptake by HepG2, Hep3B, SNU‐382, NU‐475 and CSCs (Figure 6K). Liver cancer cells possess significantly higher NAD+ levels than normal hepatocytes, a crucial adaptation for their rapid growth, proliferation and survival. We therefore examined the effects of riluzole on NAD+ levels in HCC cells. Riluzole inhibited NAD+ levels in HCC cells (Figure 6L). Overall, these data suggest that riluzole can inhibit glucose uptake and lactate production by inhibiting Glut1, Glu3 and LDH‐A expression, respectively.
Glutamine Is Essential for HCC
3.7
Metabolism alterations are frequently found in HCC, including glutamine metabolic reprogramming [47]. HCC are addicted to amino acid glutamine for energy, building blocks (nucleotides, proteins), and protection from oxidative stress, fuelling their proliferation and survival [24, 48, 49, 50]. To test the requirement of glutamine for HCC, we have used two glutaminase (GLS) inhibitors bis‐2‐(5‐phenylacetamido‐1,2,4‐thiadiazol‐2‐yl) ethyl sulphide (BPTES) and 6‐diazo‐5‐oxo‐L‐norleucin (DON). BPTES is a cell‐permeable small molecule that selectively inhibits GLS1, an enzyme crucial for cancer cell survival, by converting glutamine to glutamate, thus blocking cancer growth, and promoting cell death. DON, initially developed as an antibiotic, is a glutamine antagonist that acts as a mechanism‐based inhibitor of enzymes that use glutamine (e.g., GLS) to synthesise building blocks (nucleotides, lipids, proteins). HepG2 cells were grown in glutamine‐free medium and also in the presence of BPTES or DON. As shown in Figure 7A,B, inhibition of GLS by BPTES or DON induced apoptosis in HepG2 cells. As expected, glutamine deprivation also induced apoptosis in HepG2 cells. Interestingly, GLS inhibitors induced apoptosis to the same extent as glutamine deprivation. These data suggest that glutamine is required for survival of HCC.
*Requirement of glutamine, inhibition of glutamate efflux and GSH level, and upregulation of reactive oxygen species (ROS) in HCC. (A) HepG2 cells were grown in the absence or presence of glutaminase (GLS) inhibitor (BPTES or DON, 5 mM) with glutamine (2 mM) in the medium. After 48 h, apoptosis was measured by TUNEL assay. Data represent mean ± SD (n = 4). * = significantly different from control, p < 0.05. (B) HepG2 cells were grown in the presence or absence of glutamine (2 mM) for 48 h. Apoptosis was measured by TUNEL assay. Data represent mean ± SD (n = 4). * = significantly different from control, p < 0.05. (C) Glutamate release. HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs were treated with riluzole (0–10 μM) for 24 h. Glutamate release was measured using the Amplex Red Glutamic Acid/Glutamate Oxidase Assay Kit with a fluorometer, with excitation at 540 nm and emission at 590 nm (Invitrogen). Data represent mean ± SD (n = 4). * = significantly different from control, p < 0.05. (D) HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs were treated with riluzole (0–10 μM) for 24 h. Intracellular total GSH was detected by measuring the product of glutathionylated DTNB at 405 nm (GSH assay kit, Cayman Chemical). Data represent mean ± SD (n = 4). * = significantly different from control, p < 0.05. (E, F) HepG2 and Hep3B cells were pre‐treated with NAC (3 mM) for 2 h, followed by treatment with riluzole (0–10 μM) for 24 h. Cells were labelled with 2′,7′‐dichlorofluorescein diacetate (DCFDA/H2DCFDA) and ROS production was measured for various time points (0–360 min) with a fluorometer using excitation at 495 nm and emission at 529 nm (Cellular Reactive Oxygen Species Detection Assay Kit, Abcam). Data represent mean ± SD (n = 4). , @, #, , %, & and ** = significantly different from control; p < 0.05.
Riluzole Inhibited Glutamate Release and Glutathione (GSH) Level and Elevated Reactive Oxygen Species (ROS) in HCC Cells and CSCs
3.8
The FDA has approved riluzole for the treatment of ALS. It works by inhibiting the release of glutamate [51, 52]. We therefore examine the effects of riluzole on glutamate release, intracellular GSH level, and ROS production in HCC. As shown in Figure 7C, riluzole inhibited glutamate release in HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs.
Glutathione (GSH) is a tripeptide consisting of glutamate, cysteine, and glycine that functions as a key antioxidant in mammalian cells [53]. It modulates redox homeostasis and protects cells against ROS‐induced damage. Since GSH participates in oxidative stress response and blocks cellular toxicity, we sought to examine the effects of riluzole on intracellular GSH and ROS production. Riluzole inhibited intracellular GSH levels in HepG2, Hep3B, SNU‐382, SNU‐475 and HCC CSCs (Figure 7D). Glutathione protects the cellular membrane by preventing lipid peroxidation (LPO) and subsequent ROS generation [54]. Treatment of HepG2 and Hep3B cells with riluzole resulted in increased ROS production, reaching a plateau between 120 and 150 min (Figure 7E,F).
N‐acetyl‐L‐cysteine (NAC) is a precursor to the antioxidant glutathione, which protects cells from free radical damage and oxidative stress. Pre‐treatment of cells with antioxidant N‐acetyl‐L‐cysteine (NAC) inhibited riluzole‐induced ROS production, confirming the involvement of oxidative stress (Figure 7G). These data suggest that riluzole can increase intracellular glutamate, inhibit GSH and elevate ROS, which may be one of the mechanisms of apoptosis induction through mitochondrial dysfunction.
Discussion
4
We have shown for the first time that riluzole inhibits HCC growth and development by regulating glucose, glutamine metabolism and autophagy. Riluzole not only inhibits the growth of cancer cells but also cancer stem cells (CSCs), which are generally responsible for cancer initiation, progression, drug resistance, and chemotherapy failure. Riluzole‐induced autophagic cell death by inducing Beclin1 and Atg5. Furthermore, riluzole affects cancer cell metabolism and mitochondrial homeostasis in HCC. Finally, riluzole inhibited the Wnt/β‐catenin/TCF‐LEF pathway, which regulated cell proliferation, glucose, and glutamine metabolism.
In the present study, riluzole inhibited cell proliferation and induced apoptosis in several HCC cell lines and CSCs. The inhibitory effects of riluzole on HCC CSCs were promising because CSCs often do not respond to anticancer drugs and are responsible for drug resistance and cancer relapse. In addition to apoptosis, riluzole treatment also caused growth arrest at the G2/M stage and reduced the G1 stage of the cell cycle, which was associated with inhibition of cyclin D1 expression. Similarly, in another study, riluzole‐induced caspase‐dependent apoptosis and G2/M cell cycle arrest in HCC SNU449 and Huh7 cell lines [55]. In another study, we demonstrated that riluzole inhibited cyclin D1 expression and caused growth arrest at the G2/M stage of the cell cycle in pancreatic cancer [10]. In the present study, riluzole had no effect in human normal hepatocytes, suggesting its activity was limited to malignant cells and thus offers great hope for the treatment of HCC.
Mitochondrial glutamine is crucial for cell energy, redox balance, and signalling, serving as a key fuel for the TCA cycle, especially in cancer cells and under stress, protecting mitochondria from damage and supporting DNA repair [56, 57]. Glutamine metabolism is vital for cellular adaptation, immune response, and neurological function, making it a therapeutic target in cancer. Glutamine can enter the TCA cycle after conversion to glutamate and then aKG through a process termed glutaminolysis, catalysed by GLS and GDH. Alternatively, alanine or aspartate aminotransferase (ALT and AST, respectively) can convert glutamine to aKG, with concomitant production of alanine or aspartate, respectively. Glutamine oxidises in mitochondria and produces ATP. HCC cells are addicted to glutamate for survival [58, 59, 60]. Riluzole inhibits glutamate release by inactivating voltage‐dependent ion channels [55, 61, 62]. In prostate cancer, serum glutamate levels directly correlate with Gleason score and glutamate blockade decreases proliferation, migration, and invasion and induces cell death [63]. Blocking glutamate release by riluzole inhibits cell proliferation in glioblastoma, melanoma, breast and prostate cancer [61, 64, 65, 66, 67]. Since riluzole inhibited glutamate release and GSH level, and increased ROS production in HCC cells and CSCs, this could be considered as one of the mechanisms of apoptosis induction.
Upregulation of the WNT/β‐catenin pathway induces aerobic glycolysis by activating Glut, PDK1, pyruvate kinase M2 (PKM2), MCT‐1 and LDH‐A, and inactivating the pyruvate dehydrogenase complex. Oncogenic Myc regulates the expression of glycolysis genes, such as PDK1, Glut1, HK2 and LDHA [68, 69]. The WNT/β‐catenin pathway directly regulates Myc expression [70]. Aerobic glycolysis supplies a large portion of glucose as lactate, regardless of oxygen availability. Aerobic glycolysis is less efficient in producing ATP compared to oxidative phosphorylation. Phosphorylation of PDK‐1 inhibits the PDH, and a large part of pyruvate cannot be converted into acetyl‐CoA in mitochondria, and only a part of acetyl‐CoA can enter the TCA cycle. Cytosolic pyruvate is converted into lactate through the enzymatic activity of LDH‐A. In the present study, riluzole inhibited the expression of c‐Myc, Glut1, Glut3, LDHA and PKM2, glucose uptake and NAD^+^ levels. In HCC, PKM2 was significantly upregulated, promoted metastasis by recruiting myeloid‐derived suppressor cells, and was associated with a poor prognosis of HCC patients [71, 72]. Knockdown of PKM2 inhibited HCC cell proliferation, migration, and invasion and tumour growth, suggesting PKM2 as a therapeutic target [71]. Furthermore, knockdown of β‐catenin results in reduced glutamate transporter (Glt‐1) and glutamine synthetase (GS) expression in astrocytes [25]. Similar to the action of riluzole, inhibition of mitochondrial ATP production downregulated Wnt/β‐catenin signalling pathway [73, 74]. These data suggest that targeting PKM2 may represent a practical treatment approach for HCC.
PDKs can regulate the conversion of pyruvate to acetyl‐CoA through the mitochondrial pyruvate dehydrogenase complex. PDKs link glycolysis to the tricarboxylic acid (TCA) cycle. PDK1 is a crucial enzyme in cancer cells, promoting aggressive growth, survival, metabolism, migration, stemness, and resistance to chemotherapy by activating numerous downstream targets (such as AKT, PLK1 and MYC) and influencing metabolic shifts [75, 76, 77, 78, 79]. In the present study, riluzole inhibited PDK1 expression in HCC cells. Furthermore, high PDK1 expression in HCC patients correlates with shorter overall survival and higher rates of recurrence after surgery. Therefore, PDK1 is considered a valuable therapeutic target and biomarker for HCC.
Bcl‐2 family members regulate cell growth, survival and apoptosis [80, 81, 82, 83]. Mainly, anti‐apoptotic members such as Bcl‐2 and Bcl‐X_L_ enhance cell growth and proliferation, whereas pro‐apoptotic members such as Bax and Bad induce apoptosis [84]. Following cellular stress, Bak and/or Bax are activated and compromise the integrity of the outer mitochondrial membrane (OMM), resulting in permeabilization of the mitochondrial outer membrane. As a result of MOMP, pro‐apoptotic proteins (e.g., cytochrome c) move to the cytoplasm where they activate caspases to induce apoptosis. Anti‐apoptotic Bcl‐2 family proteins regulate cellular survival by tightly controlling the interactions between Bak/Bax and the BH3‐only proteins capable of directly inducing Bak/Bax activation. In the present study, mitochondrial dysfunction contributed to HCC cell death, as shown by increased BAX/Bcl‐2 ratio and positive TUNEL signals. Riluzole inhibited Bcl‐2 expression, induced Bax expression, and reduced mitochondrial membrane potential, leading to apoptosis. These data suggest that riluzole can act at the mitochondrial level to regulate autophagic cell death.
In conclusion, the proapoptotic and antiproliferative effects of riluzole are exerted by rewiring mitochondrial signalling specific to HCC cells. Glutamine could be required for the maintenance of mitochondrial membrane potential and integrity, and for supporting the NADPH production needed for redox control and macromolecular synthesis. Riluzole targeting the Wnt/β‐catenin signalling pathway offers a promising therapeutic avenue for HCC. The pathway's central role in HCC progression, along with its frequent dysregulation, makes it an attractive target for intervention.
Author Contributions
Sanjit K. Roy: conceptualization, investigation, writing – original draft, writing – review and editing, visualization, validation, methodology, formal analysis, project administration, data curation. Rashmi Srivastava: conceptualization, formal analysis, visualization. Nancy Landry: conceptualization, methodology, visualization, investigation, writing – original draft, writing – review and editing, validation, formal analysis, supervision, data curation, project administration. Shivam Srivastava: conceptualization, visualization, investigation, writing – review and editing. Anju Shrivastava: writing – review and editing, resources, supervision, project administration, conceptualization. Rakesh K. Srivastava: project administration, resources, supervision, writing – review and editing, conceptualization, writing – original draft, methodology.
Conflicts of Interest
The authors declare no conflicts of interest.
The reference list from the paper itself. Each links out to its DOI / PubMed record.
- 1P. P. Hsu and D. M. Sabatini , “Cancer Cell Metabolism: Warburg and Beyond,” Cell 134 (2008): 703–707.18775299 10.1016/j.cell.2008.08.021 · doi ↗ · pubmed ↗
- 2M. Mossenta , D. Busato , M. Dal Bo , and G. Toffoli , “Glucose Metabolism and Oxidative Stress in Hepatocellular Carcinoma: Role and Possible Implications in Novel Therapeutic Strategies,” Cancers 12 (2020): 12.10.3390/cancers 12061668 PMC 735247932585931 · doi ↗ · pubmed ↗
- 3D. R. Wise and C. B. Thompson , “Glutamine Addiction: A New Therapeutic Target in Cancer,” Trends in Biochemical Sciences 35 (2010): 427–433.20570523 10.1016/j.tibs.2010.05.003PMC 2917518 · doi ↗ · pubmed ↗
- 4J. W. Kim and C. V. Dang , “Cancer's Molecular Sweet Tooth and the Warburg Effect,” Cancer Research 66 (2006): 8927–8930.16982728 10.1158/0008-5472.CAN-06-1501 · doi ↗ · pubmed ↗
- 5O. Warburg , “On the Origin of Cancer Cells,” Science 123 (1956): 309–314.13298683 10.1126/science.123.3191.309 · doi ↗ · pubmed ↗
- 6Z. F. Jiang , M. Wang , J. L. Xu , and Y. J. Ning , “Hypoxia Promotes Mitochondrial Glutamine Metabolism Through HIF 1alpha‐GDH Pathway in Human Lung Cancer Cells,” Biochemical and Biophysical Research Communications 483 (2017): 32–38.28065856 10.1016/j.bbrc.2017.01.015 · doi ↗ · pubmed ↗
- 7M. A. Medina , “Glutamine and Cancer,” Journal of Nutrition 131 (2001): 2539 S–2542 S.11533309 10.1093/jn/131.9.2539 S · doi ↗ · pubmed ↗
- 8J. Zou , K. Du , S. Li , et al., “Glutamine Metabolism Regulators Associated With Cancer Development and the Tumor Microenvironment: A Pan‐Cancer Multi‐Omics Analysis,” Genes (Basel) 12 (2021): 12.10.3390/genes 12091305 PMC 846641834573287 · doi ↗ · pubmed ↗