Background
Metabolic plasticity enables cancer cells to switch between glycolysis and oxidative phosphorylation (OXPHOS) during tumorigenesis and metastasis in response to intracellular signaling pathways and environmental cues. However, it is still largely unknown how cancer cells orchestrate gene regulation to balance their glycolysis and OXPHOS activities. The switch from oxidative to glycolytic metabolism in the presence of oxygen -known as the “Warburg effect”- is a cancer cell strategy to survive by exploiting a rapid glucose-derived source not only for energy but also for anabolism [
1]. Beyond glucose metabolism, the metabolic rearrangement of cancer also involves lipids: the balance between endogenous (synthesis) and exogenous (uptake) fatty acids (FAs) is required in cancer for membranes formation, signaling, post-translational modification of proteins as well as for energy supply through OXPHOS [
2‐
4]. Cells store FAs inside lipid droplets (LDs), complex organelles characterized by a phospholipid layer surrounding a core of triacylglycerols [
5]. Despite being long considered inert energy reservoirs, LD dynamism has been recently disclosed responsible for activating several signaling pathways by regulating intracellular FAs [
6,
7]. Dysregulations of many enzymes that modulate FAs availability (i.e., lipogenesis, lipolysis) have been associated with tumorigenesis and tumor progression [
8]. In particular, all lipases involved in LDs degradation have been associated with cancer and/or related comorbidities (e.g., cachexia) [
9,
10]. Adipose triglyceride lipase (ATGL), the first and rate-limiting enzyme of the lipolytic cascade [
5], has been found altered in several human cancers [
2,
3]. ATGL activity, mediated by the Ser47-Asp166 dyad [
5,
11], is responsible for the hydrolysis of LDs-contained triacylglycerols in diacylglycerols and free FAs, in turn promoting the activity of the downstream lipases, hormone-sensitive lipase (HSL) and monoacylglycerol lipase (MAGL) [
2,
5]. FAs derived from lipase activity feed mitochondrial metabolism, boosting the OXPHOS and consequently the production of reactive oxygen species (ROS). Cancer cells exploit the connection between cellular metabolism and redox homeostasis for their proliferative advantage. In tumor, the levels of ROS are increased, and an enhancement of antioxidant systems occurs to prevent oxidative damage. Nevertheless, several lines of evidence demonstrate that ROS signaling also contributes to a switch from oxidative to glycolytic metabolism for disabling mitochondria as a harmful source of excess ROS [
12,
13]. Many redox-responsive transcription factors orchestrate ROS-dependent regulation of cell cycle, metabolism, differentiation and death in cancer [
14‐
16]. Among these, hypoxia-inducible factor-1α (HIF1α) represents the key regulator of glycolysis: it is transcribed upon intracellular ROS increase, containing functional antioxidant response elements in its enhancer [
17,
18]. HIF1α activity is pointed to restore oxygen homeostasis; among the mechanisms that it employs to promote adaptations to hypoxia, HIF1α forces the metabolic switch from OXPHOS toward glycolysis [
19‐
21]. Considering lipids as the unique energetic substrates that necessarily require mitochondria for their utilization and lipolysis as the way for mobilizing LDs-contained FAs, we over-expressed ATGL to monitor the adaptation of cervical cancer cell lines under forced lipid catabolism and identified a pro-tumor role of ATGL activity. ATGL-mediated lipid catabolism increased ROS production, which in turn promoted HIF1α activation. HIF1α increased the proliferation rate and “Warburg effect”, as well as it triggered BCL2 and adenovirus E1B 19-kDa-interacting protein 3 (BNIP3)-mediated mitophagy as a survival mechanism.
Materials and methods
Materials
2-deoxyglucose (2-DG) (D6134), Albumin (A3782), Chloroquine (C6628), CoCl2 (60818), Cycloheximide (CHX) (C7698), Dimethyl sulfoxide (DMSO, 154938), EDTA (E6758), EGTA (E4378), KH2PO4 (A0261677), MG-132 (474790), MgCl2 (A748033 012), MOPS (M3183), N-acetylcysteine (NAC, A7250), Nicotinamide adenine dinucleotide hydrate (NAD+) (N7004), TRI Reagent (T9424), Oil Red-O (ORO) (O0625), Formalin solution neutral buffered 10% (HT5012), Sodium caproate (C4026), Sodium deoxycholate (D6750), Sodium orthovanadate (S6508), Sodium pyrophosphate tetrabasic decahydrate (30411), Sucrose (S0389), Trichloroacetic acid (TCA) (T6399) and Triton X-100 (T9284) were from Sigma-Aldrich. Trypan Blue 0.4% solution (17-942E) was from Lonza. Goat anti-mouse (172–1011) and anti-rabbit (172–1019) IgG (H + L)-horseradish peroxidase conjugated were from Bio-Rad Laboratories. Alexa Fluor™ 568 donkey anti-mouse IgG (H + L) (A10037), Alexa Fluor® 594 goat anti-rabbit IgG (H + L) (A11012), Hoechst 33342 (H1399), MitoTracker™ Green FM (MTG) (M7514) and MitoTracker™ Red FM (MTG) (M7512) were from Thermo Fisher Scientific. Polyethylenimine (PEI) (23966) were from Polysciences. HIF1A inhibitor, YC-1 (ALX-420-025) was from Enzo Life Sciences. L-lactate dehydrogenase (LDH) was from Roche Applied Science. DTT (281) and protease inhibitors cocktail from AMRESCO. TRIS-base (1610716) and Sodium Dodecyl Sulfate (SDS) (161–0300) from BioRad. Dihydroethidium (DHE) (D1168) is from Molecular probes.
Cell lines, transfections and treatments
HeLa and Me-180 cell lines were purchased from American Type Culture Collection (ATCC). Cells were grown in Dulbecco’s modified Eagle’s medium (DMEM) 1 g/L glucose (EuroClone) supplemented with 10% fetal bovine serum (EuroClone), 10 U/ml penicillin/streptomycin (EuroClone) and 2 mM L-glutamine (EuroClone). Cells were authenticated and characterized by the supplier. Mycoplasma test was routinely carried out according to protocols from our laboratory. Cells were cultured at 37 °C in an atmosphere of 5% CO2 in air and plated at a 1 × 105 cells/mL density for all the experiments. After 24 h plating, cells were transiently transfected with pcDNA™4/HisMaxC and pcDNA™4/HisMaxC-ATGL and pcDNA™4/HisMaxC-ATGL (Ser47Ala), for the 48 h, through the Polyethylenimine (PEI) reagent, according to manufacturer’s instructions. pcDNA™4/HisMaxC and pcDNA™4/HisMaxC-ATGL plasmids were kindly provided by Prof. Rudolf Zechner, Institute of Molecular Biosciences, Karl-Franzens-Universität Graz, Graz (Austria); pcDNA™4/HisMaxC-ATGL (Ser47Ala) has been obtained using the Phusion Site-Directed Mutagenesis Kit (Thermo Scientific). Ser47Ala mutation was introduced in ATGL cDNA using the following primers: forward 5′-CACATCTACGGCGCCGCGGCCGGGGCGCTCACGG-3′, reverse 5′-CCGTGAGCGCCCCGGCCGCGGCGCCGTAGATGTG-3′. The final construct was verified by DNA sequencing. The transfection efficiency was determined by transfecting cells with pATGL-EGFP. The percentage of transfected cells was assessed > 85%.
Cells were treated with 10 mM 2-deoxyglucose (2DG) (Sigma-Aldrich, cat. Number D6134) for 24 h, 2.5 mM and 5 mM caproate (Sigma-Aldrich cat. Number C4026) for 24 h, 30 μM chloroquine (CQ) (Sigma-Aldrich, cat number C6628) for 2 h, 10 μg/mL cycloheximide (CHX) (Sigma-Aldrich, cat. Number C7698) for 8 h, 150 μM CoCl2 (Sigma-Aldrich, cat. Number 60818) for 6 h, 50 μM dihydroethidium (DHE) (Molecular probes, cat. Number D1168) for 30 min, 2 μM MG132 (Sigma-Aldrich, cat. Number 474790) for 8 h, 200 μM MitoTracker™ Green FM (MTG) (M7514) for 30 min, 200 μM MitoTracker™ Red FM (MTG) (M7512) for 30 min, 5 mM N-acetylcysteine (NAC) (Sigma-Aldrich, cat. Number A7250) for 24 h, 100 μM YC-1 (Enzo Life Sciences, cat. Number ALX-420-025) for 24 h.
Western blot analysis
At the end of experiment, cells were resuspended in lysis buffer (50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 10 mM NaF, 5 mM Sodium Pyrophosphate, 2 mM Sodium Orthovanadate) supplemented with protease inhibitor cocktail (AMRESCO) to obtain protein lysates. The following sonication of samples was performed through the instrument of BRANSON ULTRASONICS CORPORATION. Protein concentration was determined by the method of Lowry et al. [
22] before performing electrophoresis by SDS-PAGE and blotting onto a nitrocellulose membrane (Bio-Rad).
The following primary antibodies were used: Active β-catenin (Cell signaling, cat. Number 8814, diluted 1:1000), ATGL (Cell Signaling Technology®, cat. Number 2138S, diluted 1:1000), BNIP3 (Sigma-Aldrich, cat. Number B7931, diluted 1:1000), cleaved caspase-3 (Cell signaling, cat. Number 9664S, diluted 1:1000), caspase-3 (Sigma-Aldrich, cat. Number C8487, diluted 1:1000), cyclin D1 (Santa Cruz Biotechnology, cat. Number sc-6281, diluted 1:1000), DRP1 (Cell signaling, cat. Number 14647S, diluted 1:1000), DPR1-p S616 (Cell signaling, cat. Number 3455S, diluted 1:1000), GAPDH (Santa Cruz Biotechnology, cat. Number sc-47724, diluted 1:1000); GLUT-1 (Santa Cruz Biotechnology, cat. Number sc-7903, diluted 1:1000), HIF1α (Santa Cruz Biotechnology, cat number sc-10790, diluted 1:1000), HK-2 (Abnova, cat. Number H00003099-M01, diluted 1:1000), Lamin-B1 (Santa Cruz Biotechnology, cat. Number sc-377000, diluted 1:1000), LC3 (Sigma-Aldrich, cat. Number L7543, diluted 1:1000), LDH-A (Santa Cruz Biotechnology, cat. Number sc-33781, diluted 1:1000), N-cadherin (Sigma Aldrich, cat. Number C-3678, diluted 1:1000), PDHE1α-p S300 (Sigma-Aldrich, cat. Number AP1064, diluted 1:1000), TFAM (Santa Cruz Biotechnology cat. Number 166965, diluted 1:1000), TOM20 (Santa Cruz Biotechnology, cat. Number sc-11021, diluted 1:1000), Ubiquitin (Sigma-Aldrich, cat. Number MAB1510, diluted 1:1000), β-Actin (Cell Signaling Technology®, cat. Number #4970S, diluted 1:1000). Through the incubation with specific secondary antibodies (Bio-Rad), the signaling was identified using a Fluorchem Imaging System (Alpha Innotech), after incubation with LiteAblot® TURBO (EuroClone). Fluorchem Imaging System (Alpha Innotech) was also used to perform densitometry analyses.
Nuclear fraction isolation
Nuclear extraction was performed by incubation of cells for 20 min on ice with nuclei isolation buffer (10 mM Tris-HCl at pH 7.8, 4 mM MgCl2, 1 mM EDTA, 0.5 mM DTT, 1% Triton X-100, 0.25 M Sucrose, 10 mM NaF, 5 mM Sodium Pyrophosphate, 2 mM Sodium Orthovanadate); after centrifugation to isolate nuclei, they were washed with isolation buffer without Triton X-100 and centrifuged two times. Each buffer was supplemented with protease inhibitor cocktail (AMRESCO).
Total cell extracts, cytosolic and nuclear fractions were analyzed by western blot for purity determination. Antibodies against LDH-A and Lamin B1 were used as purity control of cytosolic and nuclear fraction, respectively.
Mitochondrial fraction isolation
Following detachment and washing of the cells, mitochondrial purification was obtained by resuspending in mitochondria isolation buffer (MIB) formed by 1 mM EGTA, pH 7.4, 5 mM MOPS, 5 mM KH2PO4, 0.1% BSA and 0.3 M sucrose, 10 mM NaF, 5 mM Sodium Pyrophosphate, 2 mM Sodium Orthovanadate. Cells were disrupted mechanically with a tight pestle with 30 strokes on ice. The homogenate was centrifuged at 2500×g for 5 min at 4 °C, and the supernatant containing mitochondria was collected while the pellet was processed as before three times to enrich mitochondrial fraction. Supernatants were then centrifuged at 15,000×g for 10 min at 4 °C. The supernatant was collected as cytoplasm faction, the pellet was washed three times with MIB and then centrifuged at 15,000×g for 10 min at 4 °C.
Quantitative real-time PCR (RT-qPCR)
RNA extraction was performed by using TRItidy G (PanReact AppliChem, cat. Number T9424) according to the manufacturer’s instructions. Synthesis of cDNA was obtained from 1 μg of total RNA by using PrimeScript™ RT Reagent Kit (Perfect Real Time) (Takara), and RT-qPCR reaction was performed by using the SYBR®Premix Ex Taq (Tli RNase H Plus) (Takara) on QuantStudio™ 3 real-time PCR System (Thermo Fisher Scientific).
RT-qPCR was performed in triplicates by using designed and tested primer with primer-BLAST (NCBI). Primers used were obtained from Sigma-Aldrich. Human primer sequences are listed used are as follow: BNIP3 forward: 5′-TCTGGACGGAGTAGCTCCAA-3′, reverse: 5′-CTTCCTCAGACTGTGAGCTGT-3′; Cpt1a forward: 5′-GACTCTGGAAACGGCCAACT-3′, reverse: 5′-ATCTTGCCGTGCTCAGTGAA-3′; E-CAD forward: 5′-CGTCCTGGGCAGAGTGAAT-3′, reverse: 5′-TCATTATGTGTTCTCGTGCAG-3′; Fibronectin forward: 5′-CGACACATTCCACAAGCGTC-3′, reverse: 5′-CATTGGTCGACGGGATCACA-3′; HIF1α, forward: 5′-ATTTTGGCAGCAACGACACAG-3′, reverse: 5′-TTTTTCGTTGGGTGAGGGGAG-3′; N-CAD forward: 5′-AGGCTTCTGGTGAAATCGCA-3′, reverse: 5′-GCAGTTGCTAAACTTCACATTG-3′; SLC2A1/GLUT-1, forward: 5′-TTCACTGTCGTGTCGCTGTT-3′, reverse: 5′-TGAGTATGGCACAACCCGC-3′; SLC16A1/MCT-1, forward: 5′-TGCGTGGGTACTGGAACAAG-3′, reverse: 5′-TGCAGGTCAAATCCAAATATCGT-3′; SLC16A3/MCT-4, forward: 5′-CTTCGTTTTTGTGCAGGTCCC-3′, reverse: 5′-ACGAAAGCCCCAAGAATGGA-3′; PNPLA2, forward: 5′-TCGTGTTTCAGACGGAGAGAA-3′, reverse: 5′-CAGACATTGGCCTGGATGAG-3′; VEGF forward: 5′-AGGCCAGCACATAGGAGAGA-3′, reverse: 5′-ACGCGAGTCTGTGTTTTTGC-3′; ACTB, forward: 5′-GGCCGAGGACTTTGATTGCA-3′, reverse: 5′-GGGACTTCCTGTAACAACGCA-3′. The relative mRNA levels were determined by using the 2−ΔΔCt method. The fold changes were relative to a control after normalization to the internal standard ACTB. PNPLA2 was used as an over-expression control.
Cell proliferation assays
Cell proliferation was evaluated by Trypan blue exclusion test procedure, by bromodeoxyuridine (BrdU) incorporation assay and MTT colorimetric assay using the Cell counting Kit-8 (CCK-8, Sigma-Aldrich cat. Number 96992) according to the manufacturer’s protocol.
For BrdU incorporation assay, cells were incubated with 10 μM BrdU for 24 h and then fixed for 30 min in ethanol:acetic acid:di-deionized water solution (18:1:1). DNA was denaturized by incubation on ice for 10 min with 1 N HCl and then 10 min with 2 N HCl. Thanks to the treatment with PBS/0.4% Triton X-100 solution for 10 min the cells were permeabilized. The blocking was done with PBS/3%BSA for 1 h. The incubation with an anti-BrdU antibody (Santa Cruz Biotechnology cat. Number sc-32323) was performed overnight, followed by 1 h of incubation with an AlexaFluor™568 donkey anti-mouse IgG (H + L) secondary antibody; nuclei were stained with 1 μg/mL of Hoechst 33342 for 10 min. Images of cells were obtained with a Delta Vision Restoration Microscopy System (Applied Precision, Issaquah, WA) equipped with an Olympus IX70 fluorescence microscope (Olympus Italia, Segrate, Milano, Italy).
Wound healing assay
HeLa cells were seeded into 24-well plates, at strict confluence, and incubated for 24 h period in complete medium to form a confluent monolayer. The culture well surface was scratched with 200 μL pipette tip to create a “wound”. The serum-starvation was applied to block cell proliferation and wound healing was followed for 16 h. Images were captured with a Delta Vision Restoration Microscopy System (Applied Precision, Issaquah, WA).
Fluorescence microscopy analyses
Cell medium was removed and cells rinsed with PBS and fixed with formalin solution neutral buffered 10% (4% paraformaldehyde containing) for 1 h, blocked with PBS/10% fetal bovine serum solution for 1 h and incubated overnight with an anti-HIF1α (Sigma Aldrich, cat. Number sc-10790) antibody. The secondary antibody was Alexa Fluor® 594 goat anti-rabbit IgG (H + L) (A11012) that is incubated for 1 h at room temperature; nuclei were stained with 1 μg/ml of Hoechst 33342 for 10 min. Delta Vision Restoration Microscopy System (Applied Precision, Issaquah, WA) equipped with an Olympus IX70 fluorescence microscope (Olympus Italia, Segrate, Milano, Italy) was used to acquire fluorescent images of cells.
Oil Red-O staining
Oil Red-O staining was performed to identify the content of lipid droplets inside the cells as previously described [
23]. Briefly, culture medium was removed, and cells were washed in PBS. Cells were fixed with formalin solution neutral buffered 10% (4% paraformaldehyde containing) for 10 min and then immediately for 1 h at room temperature, without intermediate washes. Formalin solution was removed, and cells were washed three times with PBS. Cells were incubated for 5 min at room temperature with 60% isopropanol and after complete drying, cells were incubated for 10 min with the Oil Red-O working solution (filtered 6:4 dilution of a pre-filtered 0.35% Oil Red-O stock solution in 100% isopropanol with di-deionized water). Cells were washed with di-deionized water and completely dried. In order to evaluate lipid droplets content spectrophotometrically, Oil Red-O incorporated in lipid droplets was eluted with a unique wash in 100% isopropanol for 10 min at room temperature by gentle shaking and relative absorbance measured at 510 nm with an Eppendorf BioSpectrometer®.
Determination of mitochondrial mass and potential
In order to identify mitochondrial mass, cells were incubated with 200 nM MTG (Mito Tracker Green, Invitrogen) 30 min before the end of the experimental time, washed with PBS and re-suspended in PBS, and the fluorescence intensity immediately analyzed cytofluorometrically by recording FL-1 fluorescence by means of a FACScalibur instrument. 10,000 events were counted, and data expressed as arbitrary units. To verify the potential of mitochondria, cells were treated with 200 nM MTR (Mito Tracker Red, Invitrogen) for 30 min and it was followed the same protocol as that of MTG, analyzing samples through cytofluorimeter by recording FL-2.
Extracellular lactate assay was performed considering 500 μl of cell medium, that was precipitated with 250 μl of 30% trichloroacetic acid (TCA). Media were collected at − 20° for at least 1 h and then centrifuged at 14,000×g for 20 min at 4 °C. 10 μl of supernatant were incubated for 30 min at 37 °C in 290 μl of reaction buffer (0.2 M glycine, 0.2 M hydrazine buffer pH 9.2 with freshly added 0.6 mg/ml NAD+ and 17 U/ml LDH enzyme). After 30 min at 37 °C, LDH-mediated conversion of lactate into pyruvate with the reduction of NAD+ into NADH is completed. The reaction forms nmoles of NADH, stoichiometrically equivalent to extracellular lactate, allowing to assess the amount of lactate by measuring the amount of NADH at 340 nm using an Eppendorf BioSpectrometer®. Thus, relative absorbance values were converted to lactate concentration using an extinction coefficient of 6220 M− 1 cm− 1 at 340 nm for NADH. Concentrations were normalized on total proteins in the samples.
ROS evaluation
Thirty min before the end of the experimental time, cells were incubated with 50 μM dihydroethidium (DHE) (Molecular probes, cat. Number D1168) at 37 °C. Cells were then washed and re-suspended in PBS. The fluorescence intensity of ethidium, formed by the reaction of DHE with ROS, was analyzed cytofluorometrically by recording FL-2 fluorescence. 10,000 events were counted for each sample, in which the amount of superoxide was quantified based on the shift of the fluorescence intensity of the cell population marked with dihydroethidium.
Apoptosis assay
Cells transfected with ATGL and BNIP3 ΔTM plasmids were seeded in a 12-well plate. After 48 h of over-expression, the cells were collected and stained with an Annexin V-FITC/propidium iodide (PI) following manufacturer’s protocol of Annexin V FITC Assay kit (Cayman Chemical, cat. Number 600300). After an incubation for 10 min at room temperature in the dark, the apoptotic rate of the cells was detected by flow cytometry (CytoFLEX S, Beckman Coulter, USA). CytExpert software was used to analyze the results.
Measurement of hexokinase (HK) activity
HK activity was spectrophotometrically determined. Cells were wash in PBS and lysed for 30 min on ice in lysis buffer (50 mM Tris-HCl, pH 7.5, 1 mM EDTA, 150 mM NaCl, 1% NP-40, 1 mM DTT, protease inhibitor cocktail). The optical absorbance was measured at 340 nm every 15 s for 10 min with an Eppendorf BioSpectrometer® after incubation of 50 μg of protein lysate in reaction buffer (50 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 0.6 mM ATP, 100 mM glucose, 0.2 mM NADP+, 0.1 U/mL of glucose-6-phosphate dehydrogenase) for 30 min at 37 °C. Enzyme activity was represented as change in absorbance per minute (U), normalized on protein.
PNPLA2 expression in the different grades of cervical cancer was assessed by Gene Expression Omnibus (GEO;
http://www.ncbi.nlm.nih.gov/geo, accession number GSE63514) through an Affymetrix Human Genome Array (100 samples of cervical cancer subdivided for grade vs normal cervical epithelium; normal = 24, low grade = 14, moderate grade = 22, high grade = 40).
Data analysis
Data were from at least 3 independent experiments. The results are presented as means ± SD. Statistical evaluation was conducted by the Student t-test for comparison of only two variables and one-way ANOVA with post hoc Tukey for multiple comparisons, by using the GraphPad Prism 7 software. Comparisons were statistically considered significant at p ≤ 0.05 (*), very statistically significant at p ≤ 0.01 (**) and extremely statistically significant at p ≤ 0.001 (***). Statistics are reported in every figure and were determined according to published literature.
Discussion
Lipid metabolism is part of the metabolic rewiring occurring in tumorigenesis and tumor metastasis [
30]. Here, we dissected the effects of boosting lipid catabolism through ATGL over-expression in cervical cancer cells. Using HeLa and Me-180 cell lines we showed that ATGL over-expression promoted cervical cancer cells proliferation demonstrating a pro-tumor feature of ATGL that was also observed in prostate cancer, pancreatic ductal adenocarcinoma, and breast cancer [
31‐
33]. The switch from a proliferative to a migratory state needs the selective release of specific FAs that can function as signalling molecules (mainly deriving from membrane lipids) and as constituents for membranes that are remodeled to facilitate changes in fluidity and thus cell migration (mainly intracellular lipids) [
34]. Indeed, supplementation with FAs increases cell migration [
35]. Moreover, the lipolytic lipase MAGL was extensively associated with the EMT phenotype and aggressiveness in several tumor types [
10,
36]. Our data on EMT highlighted that also ATGL upregulation exacerbated the aggressiveness of HeLa cells promoting cell morphology changes, the expression of mesenchymal and invasion markers and migratory capacity. In line with this, the data from a publicly available microarray database show that the expression of ATGL positively correlates with the aggressiveness of cervical cancer. In our system, the increased proliferation of ATGL over-expressing cells was dependent on a higher glycolytic rate, as the proliferative advantage was lost when they were treated with the glycolysis inhibitor 2DG. The enhancement of glycolysis was determined through the increase of GLUT1 expression and of the level/activity of HK-2, the first check-point enzyme of the glycolytic pathway. The high activity of HK-2 is a typical feature of malignant cells and is responsible for the accelerated glycolytic metabolism contributing to tumorigenesis and metastasis [
37]. The sustained glycolytic flux in ATGL-overexpressing cells culminated in the exacerbation of the “Warburg effect” as demonstrated by the upregulation of the lactate transporters MCT1/MCT4 and the higher concentration of extracellular lactate.
The phenotypic effects we observed were dependent on the metabolic adaptation driven by FAs availability that could feed the Krebs cycle. Indeed, FAs oxidation is accompanied by inhibitory phosphorylation of PDH [
38], the enzyme that catalyzes the aerobic conversion of pyruvate to acetyl-CoA, and that increased after ATGL over-expression thus contributing to diverge glucose toward the “Warburg effect”. The importance of FAs was validated by the use of ATGL catalytic mutant that did not alter the proliferation rate of HeLa cells, nor it causes the glycolysis-related changes. The increase in FAs oxidation was supported by the upregulation of the mitochondrial long-chain fatty acids transporter Cpt1a, which couples acyl chains to carnitine for transport into the mitochondrial matrix and subsequent degradation. FAs oxidation is usually associated with increased ROS production that could be detrimental for cells if not adequately buffered. This is particularly true for cancer cells, normally characterized by higher levels of intracellular ROS [
12,
39]. ROS increase occurring downstream of ATGL-mediated FAs release was harmful to mitochondria and affected mitochondrial potential. Cells coped with oxidative damage by upregulating the removal of dysfunctional mitochondria by mitophagy. Mitophagy has frequently been demonstrated to mitigate oxidative stress deriving from enhanced mitochondrial oxidative metabolism during cancer metabolic adaptations [
40,
41]. We have identified BNIP3 as the mediator of mitophagy in our conditions. We excluded this is only the consequence of the absence of Parkin in HeLa cells as BNIP3 was also upregulated in Me-180 cells after ATGL over-expression [
42]. Caproate, a short-chain fatty acid that is easily oxidized into mitochondria, was sufficient to induce BNIP3, whereas the ATGL catalytic mutant failed, confirming that BNIP3 was transcriptionally induced in response to increased lipid catabolism. In support of the connection between lipid catabolism and BNIP3-mediated mitophagy, there is also in vivo evidence of decreased β-oxidation and accumulation of LDs in BNIP3 null mice [
43]. Inhibition of BNIP3 through its dominant-negative mutant reduced proliferation and increased apoptotic cell death in ATGL over-expressing cells, highlighting the key role of the BNIP3-mediated mitophagy as a pro-survival event. The findings so far discussed unveil two main effects of boosting lipid catabolism in HeLa cells: the induction of glycolysis and the promotion of mitophagy as mediators of increased proliferation. The activation of HIF1α was responsible for this phenotype. Indeed, HIF1α is known to promote the metabolic switch towards glycolysis in the hypoxic condition, when OXPHOS is inefficient, as well as the upregulation of BNIP3 by binding the hypoxia response elements (HRE) on BNIP3 promoter [
44,
45]. Both the promotion of anaerobic production of ATP and the degradation of mitochondria, which are the main consumers of cellular oxygen and the primary source of ROS, allow the metabolic adaptation to oxygen deprivation and exacerbate the malignancy of cancer cells [
46]. However, the activation of HIF1α occurs frequently in normoxia conditions, in an oxygen-independent manner, a phenomenon defined as pseudo-hypoxia [
47]. ATGL mediated pseudo-hypoxic response is mimicked by caproate treatment, while ATGL catalytic mutant failed to activate HIF1α, supporting the involvement of the enhanced lipolysis in the induction of HIF1α. HIF1α activation and the association with the “Warburg effect” were also demonstrated in prostate cancer cells that receive lipids from marrow adipocytes [
48].
The enhanced ROS production, which is known to be involved in HIF1α activation [
49] is the event upstream of the pseudo-hypoxic response. Indeed, several studies demonstrated the association between abundant ROS generation by mitochondria and HIF1α induction. Moreover, ROS can trigger cancer metabolic changes (e.g.
, “Warburg effect”) aimed to reduce mitochondrial oxidative stress and to prolong cancer cell viability. Despite several pieces of evidence showed that ROS can stabilize HIF1α protein even in normoxia, ROS-mediated transcriptional up-regulation of HIF1α can also occur [
47]. We showed that upon ATGL over-expression HIF1α increase was reverted by inhibition of protein synthesis, suggesting that the translation of HIF1α mRNA, higher in ATGL transfected cells, was necessary to observe the protein increase. Future studies will be dedicated to investigating the molecular events behind this result focusing both on redox-dependent transcription factors and the mTOR/p70S6K/RPS6 signaling pathway known to promote HIF1α mRNA translation [
50]. Moreover, the use of additional cell lines is necessary to broaden the significance of ATGL expression in cervical cancer aggressivness and to clarify whether the different effects of ATGL function are cancer-specific as a consequence of the antioxidant equipment or the intracellular LDs content of the cell. Indeed, along with evidence of ATGL tumor-promoting functions in this and other tumors, the antineoplastic role of ATGL was even demonstrated [
2,
51]. We described that increased levels of ATGL in hepatocellular carcinoma cell lines diminished cell proliferation. In that cellular context, we did not observe any increase in ROS levels following ATGL over-expression even though the oxidative metabolism was efficiently activated. It is possible that the prominent antioxidant capacity of liver cells compared to others, like cervical cancer cells, allows them to efficiently face oxidative stress deriving from enhanced lipid catabolism, providing a rationale for the tumor-specific effect of ATGL.
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