International Journal of Molecular Sciences Article Cerium Oxide Nanoparticles Protect against Oxidant Injury and Interfere with Oxidative Mediated Kinase Signaling in Human-Derived Hepatocytes Silvia Carvajal 1 , Meritxell Perramón 1 , Gregori Casals 1, * , Denise Oró 1 , Jordi Ribera 1 , Manuel Morales-Ruiz 1,2 , Eudald Casals 3 , Pedro Casado 4 , Pedro Melgar-Lesmes 1,2 , Guillermo Fernández-Varo 1,2 , Pedro Cutillas 4 , Victor Puntes 5,6,7 and Wladimiro Jiménez 1,2 1 Biochemistry and Molecular Genetics Service, Hospital Clínic Universitari, IDIBAPS, CIBERehd, 08036 Barcelona, Spain; silviacr_87@hotmail.com (S.C.); mperramon@clinic.cat (M.P.); dnise_2@hotmail.com (D.O.); jordi.ribera@ciberehd.org (J.R.); morales@clinic.cat (M.M.-R.); pmelgar@clinic.cat (P.M.-L.); guillermo.fernandez@ciberehd.org (G.F.-V.); wjimenez@clinic.cat (W.J.) 2 Department of Biomedicine, University of Barcelona, 08036 Barcelona, Spain 3 School of Biotechnology and Health Sciences, Wuyi University, Jiangmen 529020, China; eudaldcm@gmail.com 4 Cell Signalling and Proteomics Group, Centre for Haemato-Oncology, Barts Cancer Institute, Queen Mary University of London, London EC1M 6BQ, UK; p.m.casado-izquierdo@qmul.ac.uk (P.C.); p.cutillas@qmul.ac.uk (P.C.) 5 Institut Català de Recerca i Estudis Avançats, (ICREA), 08010 Barcelona, Spain; victor.puntes@icn.cat 6 Vall d’Hebron Insitute of Research (VHIR), 08035 Barcelona, Spain 7 Institut Català de Nanociència i Nanotecnologia (ICN2), 08193 Bellaterra, Spain * Correspondence: casals@clinic.cat; Tel.: +34-932275400-2667  Received: 2 October 2019; Accepted: 20 November 2019; Published: 27 November 2019  Abstract: Cerium oxide nanoparticles (CeO2 NPs) possess powerful antioxidant properties, thus emerging as a potential therapeutic tool in non-alcoholic fatty liver disease (NAFLD) progression, which is characterized by a high presence of reactive oxygen species (ROS). The aim of this study was to elucidate whether CeO2 NPs can prevent or attenuate oxidant injury in the hepatic human cell line HepG2 and to investigate the mechanisms involved in this phenomenon. The effect of CeO2 NPs on cell viability and ROS scavenging was determined, the differential expression of pro-inflammatory and oxidative stress-related genes was analyzed, and a proteomic analysis was performed to assess the impact of CeO2 NPs on cell phosphorylation in human hepatic cells under oxidative stress conditions. CeO2 NPs did not modify HepG2 cell viability in basal conditions but reduced H2 O2 - and lipopolysaccharide (LPS)-induced cell death and prevented H2 O2 -induced overexpression of MPO, PTGS1 and iNOS. Phosphoproteomic analysis showed that CeO2 NPs reverted the H2 O2 -mediated increase in the phosphorylation of peptides related to cellular proliferation, stress response, and gene transcription regulation, and interfered with H2 O2 effects on mTOR, MAPK/ERK, CK2A1 and PKACA signaling pathways. In conclusion, CeO2 NPs protect HepG2 cells from cell-induced oxidative damage, reducing ROS generation and inflammatory gene expression as well as regulation of kinase-driven cell survival pathways. Keywords: cerium oxide nanoparticles; oxidative stress; human hepatic cells; phosphoproteomics; NAFLD 1. Introduction During the last few years, it has been suggested that antioxidants such as superoxide dismutase (Mn-SOD), resveratrol, colchicine, eugenol or vitamin E and C exert beneficial effects in chronic Int. J. Mol. Sci. 2019, 20, 5959; doi:10.3390/ijms20235959 www.mdpi.com/journal/ijms Int. J. Mol. Sci. 2019, 20, 5959 2 of 22 liver disease [1–5]. This hypothesis was raised based on the concept that the root of many hepatic disturbances involves an imbalance in reactive oxygen species (ROS) metabolism. Accordingly, any maneuver towards reversing this imbalance should improve liver functionality and diminish injury. In this regard, the therapeutic effects induced by antioxidants (vitamin C and SOD) on portal hypertension have already been shown in patients with liver disease [6] and CCl4 -treated rats with portal hypertension [1]. Currently, the European and American guidelines for the management of patients with non-alcoholic fatty liver disease (NAFLD) support the use of vitamin E in nondiabetic adults with biopsy-proven NASH [7,8]. More recently a new player, namely cerium oxide nanoparticles (CeO2 NPs), was incorporated to the group of antioxidant substances with therapeutic properties in experimental liver disease. CeO2 NPs are a rare element belonging to the lanthanide series, with both Ce3+ and Ce4+ oxidation states that result in an autoregenerative redox cycle between the two states, followed by the capture or release of oxygen [9]. As a consequence, the oxygen storage capacity of CeO2 NPs becomes very useful to eliminate free radicals while generated in situations of ROS imbalance [10,11]. Recently, we have demonstrated that CeO2 NPs reduce steatosis, portal hypertension and display anti-inflammatory properties in rats with experimental liver fibrosis [12]. Also, we observed an antilipogenic and anti-inflammatory effect in the liver of rats subjected to a methionine and choline deficient diet for six weeks [13]. A major difference between antioxidants such as SOD or vitamin C and CeO2 NPs is that the former two are rapidly oxidized or degraded (metabolized) whereas CeO2 NPs act as self-renewal catalysts. In addition, CeO2 NPs only display biological effects in the case of ROS overproduction; otherwise, they behave as inert inorganic material [14]. For this reason, CeO2 NPs are expected to behave as permanent vitamin C or SOD-like effectors and perform better than natural antioxidants, being specifically active in the case of inflammation. This makes CeO2 NPs especially helpful to arrest NAFLD progression, which is critically dependent on chronic effects of ROS in the liver. It is therefore important to understand the mechanisms of action of CeO2 NPs to ensure safe use in human liver disease. In this investigation, we address this issue by assessing whether CeO2 NPs are able to prevent or attenuate the oxidant-mediated injury induced by H2 O2 or lipopolysaccharide (LPS) in HepG2 cells, which is a cell line derived from human hepatocytes that preserve most of the morphological and metabolic characteristics of the original hepatic cells [15,16]. The aim of the study was to further elucidate whether the therapeutic effect of CeO2 NPs already observed in experimental liver disease can be translated to human cells and to investigate the molecular mechanisms involved in this beneficial effect. 2. Results 2.1. Cerium Oxide Nanoparticles Characterization The CeO2 NPs used in this work and their evolution in the physiological media have been already described extensively in our previous works [12,17], and the results presented here are similar to the ones already presented, i.e., the nanoparticles (NPs) are stable during their time in the physiological media and a protein corona is made on their surface during that time. This can be observed in Figure 1 and Table 1 as size distribution from Transmission Electron Microscopy (TEM) images is only slightly modified, Z-potential decreases to the value of the fetal calf serum, while the hydrodynamic diameter measured by Dynamic Light Scattering (DLS) increases and Ultraviolet-Visible (UV-VIS) spectra show some modifications (both from the slight modification of the size distribution and the absorption of proteins). Int. J. Mol. Sci. 2019, 20, 5959 3 of 22 Int. J. Mol. Sci. 2019, 20, x 3 of 20 Figure Figure1. 1.Characterization Characterization of of thetheCeOCeO2NPs used 2 NPs usedin inthis work. this work. (A)(A)NPs NPsafter afterpurification purification and and resuspension resuspension in in Tetramethylammonium Tetramethylammonium hydroxide hydroxide (TMAOH) (TMAOH) 10 10 mM.mM. Scale barbar Scale is 50 nm. is 50 nm. (B)(B) NPs NPs after 2 days after 2 daysininCell CellCulture Culture Medium Medium (CCM), Dulbecco’s Dulbecco’sModified ModifiedEagleEagleMedium Medium (DMEM) (DMEM) + 10% + 10%Fetal Fetal Calf Serum (FCS). Scale bar is 50 nm. Although TEM images of the NPs Calf Serum (FCS). Scale bar is 50 nm. Although TEM images of the NPs in the CCM are not as clear in the CCM are not as clear as TMAOH, as in in TMAOH, duedue to to thethe presence presence of of other other components components of of thethe media, media, NPsNPscancan still still bebe observed observed and and measured. measured. (C)(C) Evolution Evolution of theofUV-VIS the UV-VIS spectraspectra of CeO2of NPsCeO 2NPs in the in the CCM. NPsCCM. are stableNPsthroughout are stablethe throughout time of thethe time of theand experiments experiments presence ofand presence large of largecan agglomerates agglomerates be ruled outcan sincebethose ruledagglomerates out since those wouldagglomerates increase the would increase absorbance the wavelengths. at larger absorbance at (D)larger and (E)wavelengths. Size distribution (D) using and ImageJ (E) Size free distribution using ImageJ software (Available free https://imagej.nih.gov/ij/) online: software (Available online: https://imagej.nih.gov/ij/) counting more than 500 NPscounting of differentmoreTEM than 500 NPs images of theof NPs different TEM images in TMAOH and inofCCM.the NPs in TMAOH A slight increaseand caninbeCCM. A slight observed in theincrease can be size distribution observed (standardin deviation the size distribution increases from (standard 1.2 to 2.0)deviation probably dueincreases from 1.2ofto to dissolution NPs 2.0) in probably due to the more aggressive media of of dissolution theNPs CCM. in However, after 2 days,media the more aggressive mean ofsize theis CCM. unaltered. (F) X-ray However, Diffraction after 2 days, mean (XRD) spectra size is of the CeO NPs in TMAOH 10 mM showing the characteristic peaks unaltered. (F)2 X-ray Diffraction (XRD) spectra of the CeO2NPs in TMAOH 210 mM showing the of CeO crystals. characteristic peaks of CeO2 crystals. Table 1. Dynamic light scattering (DLS) and Zeta potential (Z-Pot) values of CeO2 NPs purified and resuspended in TMAOH 10 mM and after exposure to DMEM + 10% FCS (CCM). A decrease of the Table 1. Dynamic light scattering (DLS) and Zeta potential (Z-Pot) values of CeO2NPs purified and Z-Potential towards the value of the FCS (Z-Pot = −10 mV) and an increase of the hydrodynamic resuspended in TMAOH 10 mM and after exposure to DMEM + 10% FCS (CCM). A decrease of the diameter due to the absorption of proteins can be observed. The increase of DLS and maintenance of Z-Potential towards the value of the FCS (Z-Pot = −10 mV) and an increase of the hydrodynamic size distribution observed by TEM images after 2 days in CCM is an indication of the stability of the diameter due to the absorption of proteins can be observed. The increase of DLS and maintenance of NPs and the protein corona formation in the CCM. size distribution observed by TEM images after 2 days in CCM is an indication of the stability of the NPs and the protein corona formation in CeO NPs (TEM = 4–5 nm) the2CCM. Table Header DLS (Z-average, nm) CeO2NPs (TEM = 4–5 nm) Z-Pot (mV) NPs after purification Table Header 33.0DLS Z-Pot−47.0 (in TMAOH 10 mM) (Z-average, nm) (mV) 0 d in cCCM 70 −35.4 NPs after purification 33.0 −47.0 1 d (in in TMAOH cCCM 10 mM) 70 −36.4 2 d in cCCM 0 d in cCCM 72.9 70 −35.4−35.0 15 d in cCCM 1 d in cCCM 79 70 −36.4−16.9 30 d in cCCM 2 d in cCCM 100.272.9 −35.0 −9.6 15 d in cCCM 79 −16.9 2.2. CeO2 NPs Protect HepG2 Cells from H2 O2 - and LPS-induced Cytotoxicity 30 d in cCCM 100.2 −9.6 To demonstrate that the CeO2 NPs employed are non-toxic to HepG2, cells were exposed to CeO 2 NPs, 2.2. CeO 2NPs and cellHepG2 Protect survival wasfrom Cells assessed H2O2-using the MTS assay. and LPS-induced As shown in Figure 2A, we did not find Cytotoxicity significant differences in cell death between control and CeO2 NPs-exposed cells up to concentrations To demonstrate that the CeO2NPs employed are non-toxic to HepG2, cells were exposed to CeO2NPs, and cell survival was assessed using the MTS assay. As shown in Figure 2A, we did not Int. J. Mol. Sci. 2019, 20, 5959 4 of 22 of 10 µg/mL of CeO2 NPs. Next, we further certified the ability of CeO2 NPs to shield HepG2 cells from oxidative damage. Cells were incubated with H2 O2 or LPS to promote oxidative stress and were then treated with different doses of CeO2 NPs. Figure 2B,D,E show the effect of CeO2 NPs on cell viability and ROS production, respectively. As anticipated, the presence of H2 O2 significantly increased the oxidation of dichlorofluorescin diacetate (DCF-DA) in HepG2 cells, in association with a reduction in cell viability. Remarkably, both effects were reverted when HepG2 cells were coincubated with 10 µg/mL of CeO2 NPs. Similar results were obtained on measuring glutathione (GSH) concentration. H2 O2 treatment reduced GSH concentrations to 37% ± 5% from the basal value, whereas subsequent administration of CeO2 NPs recovered these figures to 64% ± 5% (p < 0.01). Furthermore, cellular morphological visualization using light microscopy (Figure 2C) showed that most of the HepG2 cells lost their normal morphology when stimulated with H2 O2 , whereas this change was absent when cells were treated with CeO2 NPs. A similar pattern of response was found when cells were stimulated with LPS. Indeed, LPS increased ROS production and decreased cell viability, and CeO2 NPs prevented these effects in HepG2 cells (Figure 3). These results indicate that CeO2 NPs treatment reduces ROS accumulation and the associated cell death induced by H2 O2 and LPS in HepG2 cells. Int. J. Mol. Sci. 2019, 20, 5959 5 of 22 Int. J. Mol. Sci. 2019, 20, x 5 of 20 Figure 2. CeO Figure 2 NPs 2. CeO inhibited 2NPs inhibitedHH 2O 2 -induced 2O cytotoxicityininHepG2 2-induced cytotoxicity HepG2 cells. cells. (A)(A) Viability Viability of HepG2 of HepG2 cells cells after after treatment treatment withwith CeOCeO2 2NPs NPs (10 (10 µg/mL) µg/mL) determined determined using using the MTS the MTS assay (MTS) assay (MTS)at indicated at time indicated time points. Quadruplicates of each group were used in each independent experiment. points. Quadruplicates of each group were used in each independent experiment. The results are The results are expressed expressed as percentage as percentage ofofcontrol control cells cells for forthe thetimes timesindicated. (B) HepG2 indicated. cells were (B) HepG2 cellsexposed to 1.5 were exposed to mM H2O2 and treated with 10 µg/mL of CeO2NPs for 1.5 h. Cell viability was detected using MTS 1.5 mM H2 O2 and treated with 10 µg/mL of CeO2 NPs for 1.5 h. Cell viability was detected using and expressed as percentage of control cells. Data are the mean ± S.E. of triplicate experiments. ** p < MTS and expressed as percentage of control cells. Data are the mean ± S.E. of triplicate experiments. 0.01 vs. control. + presence; − absence. (C) Representative phase contrast light microscopy images of ** p < 0.01 vs. control. + presence; − absence. (C) Representative phase contrast light microscopy HepG2 cells at 1.5 h after H2O2 treatment. (D) Reactive oxygen species (ROS) production was images of HepG2 determined bycells at 1.5 hspectrophotometry fluorescence after H2 O2 treatment. using (D) Reactive oxygen the oxidant-sensitive species dye (ROS) production 2’,7’-DCF-HDA. The was determined results were expressed as percentage of control cells for the treatments indicated. *** p2’,7’-DCF-HDA. by fluorescence spectrophotometry using the oxidant-sensitive dye < 0.001 vs. The results control;were †† p 2 corresponds to a linear p value of < 0.01). (B) Heatmap showing the enrichment of substrate groups for the different kinases calculated by the KSEA algorithm with the PhosphoSite database and the z-score method of calculating enrichment. The extent of enrichment was calculated as the abundance of substrate phosphorylation peptide in HepG2 cells under one condition divided by its abundance in another condition. Specifically, the first column shows the result of KSEA for cells exposed to H2 O2 vs. control cells treated with vehicle, the second column shows KSEA of cells exposed to H2 O2 and CeO2 NPs vs. control treated with vehicle, and the third column shows KSEA of cells exposed to H2 O2 and CeO2 NPs vs. HepG2 exposed to H2 O2 . Column “q” indicates the number of phosphopeptides used to estimate the enrichment in kinase activity for the indicate kinase. An arbitrary cut-off value of 6 phosphopeptides has been used. (C) Heatmap showing phosphopeptides significantly affected by exposure of HepG2 cells to H2 O2 and H2 O2 + CeO2 NPs in proteins linked to mTOR, ERK, CK2A1 and PKA signaling pathways. Statistical significant was assessed using t-test in (A,C) or z-test (B). * p < 0.05; ** p < 0.01; *** p < 0.001. Int. J. Mol. Sci. 2019, 20, 5959 11 of 22 Table 3. Phosphopeptides with significantly decreased phosphorylation with a Log2 fold ≤ −2 after treatment with CeO2 NPs in HepG2 cells exposed to H2 O2 . Protein Phosphopeptide H2 O2 vs Control. Fold H2 O2 + NPs vs H2 O2 Fold AT-rich interactive domain-containing protein 1A ARID1A seq: 1182–1202 + Phospho (ST) 3.0099 * −2.727 * Charged multivesicular body protein 2b CHMP2B pS199 9.963 * −9.964 * C-Jun-amino-terminal kinase-interacting protein 4 SPAG9 seq: 223–241 + Phospho (ST) 10.117 * −10.11 * Deoxynucleotidyltransferase terminal-interacting protein 2 DNTTIP2 pS141 10.135 * −10.13 * MCM3 seq: 696–724 + Gln- > pyro-Glu DNA replication licensing factor MCM3 8.390 * −8.391 * (N-term Q); Phospho (ST) DNA-dependent protein kinase catalytic subunit PRKDC seq: 3197–3232 + Phospho (ST) 8.6913 * −8.692 * Double-stranded RNA-binding protein Staufen homolog 1 STAU1 pS390 8.9250 * −8.926 * E3 ubiquitin-protein ligase UBR5 UBR5 seq: 636–654 + Phospho (ST) 10.128 * −10.12 * Eukaryotic translation initiation factor 4E-binding protein 1 EIF4EBP1 pS65 pT70 10.841 * −10.84 * Kanadaptin SLC4A1AP pS466 9.2633 * −9.264 * SLC4A1AP seq: 324–362 + Oxidation Kanadaptin 9.164 * −9.165 * (M); Phospho (ST) MKL/myocardin-like protein 2 MKL2 seq: 535–562 + Phospho (ST) 9.971 * −9.973 * PCM1 seq: 1923–1972 + Phospho (ST); Pericentriolar material 1 protein 10.677 * −10.67 * Phospho (Y) Prolyl 3-hydroxylase OGFOD1 OGFOD1 seq: 381–427 + Phospho (ST) 9.012 * −9.013 * PRRC2A seq: 1103–1128 + Gln- > Protein PRRC2A 9.2358 * −9.236 * pyro-Glu (N-term Q); Phospho (ST) R3H domain-containing protein 1 R3HDM1 seq: 295–314 + Phospho (ST) 8.592 * −8.593 * Ras-responsive element-binding protein 1 RREB1 seq: 1636–1665 + Phospho (ST) 9.296 * −9.297 * Stress-70 protein. mitochondrial HSPA9 pM370 pM389 11.353 * −11.35 * Telomeric repeat-binding factor 2 TERF2 seq: 404–447 + Phospho (ST) 8.8833 * −8.884 * YAP1 seq: 162–181 + Gln- > pyro-Glu Transcriptional coactivator YAP1 3.3580 * −5.372 * (N-term Q); Phospho (ST) Apoptotic chromatin condensation inducer in the nucleus ACIN1 seq: 463–506 + 3 Phospho (ST) 5.9139 * −11.89 ** Paxillin PXN seq: 298–317 + Phospho (ST) 1.8510 * −2.535 ** Poly(rC)-binding protein 1 PCBP1 pM186 pS190 1.9716 * −2.646 ** Int. J. Mol. Sci. 2019, 20, 5959 12 of 22 Table 3. Cont. Protein Phosphopeptide H2 O2 vs Control. Fold H2 O2 + NPs vs H2 O2 Fold Nascent polypeptide-associated complex subunit alpha NACA pS2029 1.284 * −3.119 ** RNA-binding protein 25 RBM25 pS703 11.804 * −4.246 ** PRPSAP1 seq: 193–220 + Oxidation (M); Phosphoribosyl pyrophosphate synthase-associated protein 1 12.440 ** −4.758 * Phospho (ST) Serine/arginine repetitive matrix protein 2 SRRM2 seq: 2275–2301 + Phospho (ST) 9.8054 ** −9.80 * Cell division cycle protein 23 homolog CDC23 pT562 8.7452 ** −8.746 ** Centrosomal protein of 131 kDa CEP131 seq: 45–56 + Phospho (ST) 9.3197 ** −9.320 ** Deoxynucleotidyltransferase terminal-interacting protein 2 DNTTIP2 pS381 9.412 ** −9.41 ** Nuclear receptor coactivator 2 NCOA2 pS771 2.5932 ** −2.630 ** Oxysterol-binding protein 1 OSBP seq: 377–395 + 2 Phospho (ST) 2.8700 ** −2.558 ** RNA-binding motif protein. X chromosome RBMX pS208 9.8237 ** −9.824 ** Transcription factor Sp5 SP5 seq: 43–68 + Phospho (ST) 4.5004 ** −10.69 ** Uncharacterized protein C6orf106 C6orf106 seq: 264–287 + Phospho (ST) 8.2914 ** −2.875 ** Epsin-1 EPN1 seq: 412–445 + Phospho (ST) 13.834 ** −4.391 *** Pinin PNN pS66 8.894 *** −4.875 * Protein LYRIC MTDH pS298 8.390 *** −8.391 ** POU domain. class 2. transcription factor 1 POU2F1 seq: 273–293 + Phospho (ST) 8.691 *** −8.69 *** Statistical significance was assessed using the t-test. *. p < 0.05; **. p < 0.01; ***. p < 0.001. Int. J. Mol. Sci. 2019, 20, 5959 13 of 22 CK2A1 phosphorylates a wide array of substrates and regulates several cellular processes, including cell cycle progression and transcription. HepG2 exposure to CeO2 NPs also resulted in dephosphorylation of several peptides known to be substrates of CK2A1, such as MCM2 at S139, MYH10 at S1956 and CLIP at S1364 (Figure 7C). In addition to the PhosphoSite annotated substrates, other sites linked to CK2A1 like kanadaptin at S466, MCM3 at S711 and T713, CHMP2B at S199, and RBMX at S208 also followed the same trend (Table 3). Following CeO2 NPs treatment, we also observed dephosphorylation of POU2F1, a substrate of PKACA, a kinase that phosphorylates a wide variety of substrates in the cytoplasm and the nucleus and regulates trafficking of compartmentalized pools of its regulatory subunits (Figure 7C). In summary, an assessment of the phosphoproteome in cells exposed to oxidative stress indicates that CeO2 NPs negatively interfere with the signaling of mTOR, ERK 1/2, CK2A1 and PKACA. Moreover, CeO2 NPs also dephosphorylated TERF2 and ARID1A, which are major therapeutic targets in HCC. 3. Discussion The recent description that nanoceria could be therapeutically useful in pathological conditions characterized by enhanced oxidative stress and inflammation [20], including liver disease, raised the possibility of using this material in patients. However, assessment of the beneficial effects of CeO2 NPs in humans should be made with caution. As a first step to address this issue, the current study sought to investigate the effect of CeO2 NPs in a human hepatic cell line when challenged with a well characterized pro-oxidant or proinflammatory agent such as H2 O2 or LPS. Oxidative stress and inflammation are considered key mechanisms of progression to NASH, fibrosis and/or hepatocellular carcinoma in patients with NAFLD. In the current investigation, exposure of human HepG2 cells to 10 µg/mL of CeO2 NPs did not alter cell viability under normal conditions. This is coincident with previous investigations demonstrating that at the concentration used these nanoparticles are non-toxic to numerous mammalian cells, including endothelial, breast and fibrosarcoma cells [21,22]. This feature seems to be characteristic of CeO2 NPs since other metal oxide nanoparticles, such as zinc oxide and titanium (exposed to UV-light), have displayed remarkable toxicity [23]. CeO2 NPs have been proposed as a potential treatment for clinical conditions in which increased oxidative stress plays a significant pathogenic role. Actually, CeO2 NPs display superoxide dismutase (SOD) and catalase mimetic activities [10,24,25] and present hydroxyl radical scavenging properties, thus resulting in a reduction of ROS [24,25]. This has been further supported by experimental evidence demonstrating the ROS-scavenging potential of CeO2 NPs in medicine [26]. Oxidative stress plays a critical role in the development of chronic liver damage and stimulates its progression. It is well established that oxidative stress constitutes the background of viral, alcoholic liver diseases, non-alcoholic steatohepatitis and participates in the fibrogenic response of the liver [27]. Therefore, this investigation was addressed to assess whether CeO2 NPs could protect hepatocyte injury induced by a direct prooxidant stimulus such as H2 O2 . Our results showed that CeO2 NPs reduced the cellular cytotoxicity induced by H2 O2 , which was associated with a decrease in cellular oxidative stress. Next, we further investigated the ability of CeO2 NPs to decrease the endogenous production of ROS by stimulating HepG2 cells with LPS. This bacterial wall-derived product is a well-known inducer of ROS production in several cell lines [28,29]. In line with these investigations, we showed that LPS treatment increased ROS production in HepG2 cells (Figure 3A). Furthermore, the presence of CeO2 NPs blocked LPS-induced ROS production in these cells. This phenomenon occurred in the absence of significant modifications in cell viability. Altogether, these results show that CeO2 NPs also protect HepG2 cells from LPS-induced oxidative stress. Although the physicochemical properties of CeO2 NPs as a ROS scavenger are well described, the mechanisms of their effects on biological systems remain largely unknown. In order to investigate the specific molecular mechanisms by which CeO2 NPs exert the protective effects observed in HepG2 cells, we studied the pattern of expression of a wide array of genes involved in oxidative stress Int. J. Mol. Sci. 2019, 20, 5959 14 of 22 and antioxidant defense. As expected, treatment of HepG2 cells with H2 O2 significantly altered the biological response to ROS with significant changes in the expression of 30% of the genes studied (25 of 84 genes). Interestingly, CeO2 NPs treatment of H2 O2 -stimulated cells allowed the identification of five specific genes related to the cellular response of CeO2 NPs to ROS. These include important oxidative genes such as myeloperoxidase (MPO), prostaglandin-endoperoxide synthase 1 (PTGS1), also known as cyclooxygenase 1 (COX1), neutrophil cytosolic factor 2 (NCF2, also known as P67PHOX), and inducible nitric oxide synthase (iNOS). The MPO enzyme catalyzes the conversion of hydrogen peroxide to hypochlorite and hypochlorous acid, and its activation has been related to the proapoptotic and profibrotic pathway of progression in non-alcoholic fatty liver disease [30,31]. The PTGS1 enzyme catalyzes the conversion of arachidonic acid to prostaglandins and thromboxane, and it is widely known that PTGS1 disruption translates into reduced inflammatory response [32,33]. NCF2 encodes a 67 kDa cytosolic subunit that is required for activation of NADPH oxidase to produce superoxide anions. iNOS is a major enzyme in the synthesis of nitric oxide. Overproduction of nitric oxide by iNOS is a critical mediator of inflammation contributing to tissue injury. It is interesting to note that iNOS transfers electrons from NADPH in the NO synthesis reaction and that iNOS expression requires NADPH oxidase-dependent redox signaling [34,35]. In addition, iNOS can also catalyze the production of superoxide ion. In our experiments, the 5.3- 3.7- and 8.7-fold induction of MPO, PTGS1, and iNOS genes, respectively, caused by H2 O2 was almost normalized when HepG2 cells were treated with CeO2 NPs. Furthermore, a significant reduction in NCF2 expression was also detected in H2 O2 -exposed cells when treated with CeO2 NPs. Our results therefore suggest that, besides the inherent antioxidant properties of the CeO2 NPs chemistry, the cytoprotective effects induced by CeO2 NPs are mediated by the reduction in the expression of these oxidative enzymes. It is noteworthy that increased expression of these genes was also found in the liver of rats with CCl4 -induced fibrosis [12]. Moreover, treatment with CeO2 NPs significantly reduced the hepatic expression of iNOS and NCF2 in this model of liver disease [12]. The similarity of the biological effects observed after CeO2 NPs administration in the injured liver of experimental animals and that observed in the current study in HepG2 cells cultured under pro-oxidant conditions further support the potential therapeutic usefulness of CeO2 NPs in human liver disease. The impact of CeO2 NPs on cell phosphorylation in human hepatic cells under oxidative stress conditions has not been systematically investigated using untargeted MS-based proteomics. Therefore, we studied the effect of CeO2 NPs in an oxidative stress model that involves exposing cells to H2 O2 (1.5 mM). In other words, we investigated how CeO2 NPs affect the H2 O2 -induced phosphoproteome changes in human-derived hepatocytes. We found that 10% of all the peptides assessed were phosphorylated as a result of exposure to H2 O2 . However, the effect of CeO2 NPs was considerably more selective. Actually, a reduction in phosphorylation was significantly observed in 39 out of the 1037 peptides affected by H2 O2 exposure. A significant number of proteins dephosphorylated as a result of exposure to CeO2 NPs were linked to cell proliferation and gene transcription, including ACIN1, YAP1 and 4E-BP1. The activities of kinases are linked to the wiring of signaling networks [36], thus several tools have been developed to link phosphorylation data to upstream kinases based on phosphorylation motifs [37,38]. In the current investigation we have inferred kinase pathway activation based on values of substrate group enrichment obtained from previous knowledge of the kinase-substrate relationship. As a result, we found that H2 O2 -exposed cells treated with CeO2 NPs had decreased amounts of phosphorylated substrates of mTOR, ERK 1/2 and PKACA, all of which have known roles in promoting cell growth, angiogenesis and carcinogenesis. Moreover, CeO2 NPs also dephosphorylated TERF2 and ARID1A, which play crucial roles in the initiation and development of HCC [39–42]. As a limitation of the study, the toxicity of CeO2 NPS was not evaluated in a normal hepatic cell line. However, Gaiser et al. reported no toxicity of CeO2 NPs in the C3A human hepatocyte cell line at doses up to 100 times higher than those used in our study [43]. More recently, Singh et al. also did not find toxicity of CeO2 NPs in the human hepatic cell line WRL-68 [44]. In addition, although no in vivo or functional experiments were performed to verify the efficacy of CeO2 NPs in reducing oxidative Int. J. Mol. Sci. 2019, 20, 5959 15 of 22 damage and inflammatory response, recent results in animal models of liver disease have consistently shown antioxidant and anti-inflammatory effects of CeO2 NPs in the liver [12,13,45]. Finally, further studies are necessary to confirm that the specific mechanism of oxidative damage reduction identified in this study can be translated to primary human hepatic cells and in in vivo conditions. 4. Material and Methods 4.1. Synthesis and Characterization of CeO2 NPs CeO2 NPs were synthesized by the chemical precipitation of cerium (III) nitrate hexahydrated (Sigma-Aldrich, St. Louis, MO, USA) in a basic aqueous solution [17]. By modifying the pH conditions, different sizes can be obtained. Here, we used 4 nm NPs at a concentration of 1 mg/mL. In a first step, 10 mM of cerium (III) nitrate hexahydrate was dissolved in 100 mL of absolute ethanol at room temperature. The solution was left under stirring for about 30 min. One mL of TMAOH (1.0 ± 0.02 M in H2 O) was added to the 100 mL solution at a final concentration of 10 mM, and the mixture was left under stirring. NPs were purified using centrifugation and resuspended in aqueous solution of 10 mM TMAOH, which acts as a stabilizer. The surface charge of the NPs was characterized in a Z-sizer (Malvern, Worcestershire, UK), while the crystal size was characterized using high-resolution TEM (HR-TEM) in a Tecnai G2 F20 at 200 kV (FEI, Hillsboro, Oregon, USA) and XRD (Xpert Pannalytical, Westborough, MA, USA), and the light interaction was characterized using UV-VIS spectroscopy (Shimadzu, Kyoto, Japan). Size distribution was computer analyzed by ImageJ (National Institutes of Health, Bethesda, MD, USA). CeO2 NPs were kept at 4 ◦ C until used. CeO2 NPs were then diluted with DMEM (Life technologies, Carlsbad, CA, USA) to the final concentration 0.1, 1 or 10 µg/mL. 4.2. Cell Culture and Treatment All studies were conducted with HepG2 human hepatocytes derived from a liver hepatocellular carcinoma obtained from the American Type Culture Collection (ATCC Cat# HB-8065, RRID: CVCL_0027; Manassas, VA, USA). This immortalized, stable cell line can be repeatedly frozen, thawed and propagated. Cells were grown to confluence for 24 h in DMEM, supplemented with 50 U/mL penicillin, 50 µg/mL streptomycin and 10% FCS, in a humidified atmosphere in 5% CO2 at 37 ◦ C. Thereafter, cells were switched to serum-free medium for 24 h. For cell stimulation and treatment, the old medium was removed and replaced with medium containing 1.5 mM H2 O2 (Sigma-Aldrich, St. Louis, MO, USA) and CeO2 NPs (10 µg/mL) or vehicle (TMAOH, 0.17 mM), respectively. In preliminary experiments we observed that concentrations of CeO2 NPs higher than 20 µg/mL may result in a significant diminution of cell viability. On the other hand, the concentration of H2 O2 was selected based on preliminary experiments showing that 60 min after H2 O2 loading the 1.5 mM dose resulted in a maximal effect on DCFH concentration without affecting cell survival. Higher concentrations of H2 O2 , however, resulted in a dramatic decrease in survival. Cells were incubated for the indicated time points and then harvested for biochemical or molecular assays. All experiments were repeated at least three times. Cell viability analysis: Cell viability was determined using MTS methodology (CellTiter 96; Promega, Madison, WI, USA) according to the manufacturer’s instructions. In brief, cells were seeded (1 × 105 cells/well) in 96-well plates and treated with H2 O2 or LPS (10 µg/mL) and CeO2 NPs as described above. Cells were washed with Hank’s Balanced Salt Solution (HBSS), and CellTiter reagent was added to each well. After incubation for 3 h at 37 ◦ C to allow cells to bioreduce MTS into formazan, the absorbance of the formazan was measured with a spectrophotometer (FLUOstar OPTIMA; BMG LABTECH, Ortenberg, Germany) at 492 nm. The quantity of formazan is directly proportional to the number of living cells in culture. ROS measurement: Fluorescence spectrophotometry was used to measure ROS, with 2’,7’-DCF-DA as the probe (Master Probes, Invitrogen Labs, Thermo Fisher Scientific, Waltham, MA, USA). DCF-DA readily diffuses through the membrane and is enzymatically hydrolyzed by intracellular esterases Int. J. Mol. Sci. 2019, 20, 5959 16 of 22 to the nonfluorescent DCFH, which can then be rapidly oxidized to fluorescent DCF in the presence of ROS. Cells incubated alone or treated with H2 O2 or LPS in the presence of CeO2 NPs or vehicle were washed with HBSS and incubated with 10 µM DCF-DA in DMEM for 40 min at 37 ◦ C in the dark. The cells were trypsinized and diluted followed by staining with 0.02% trypan blue. The number of cells stained with trypan blue was counted under a light microscope. The supernatant was collected to measure ROS production, and the intensity of fluorescence was immediately read in a fluorescence spectrophotometer (FLUOstar OPTIMA; BMG LABTECH, Ortenberg, Germany) at 485 nm for excitation and at 520 nm for emission. Additionally, in these experiments we also measured GSH concentration. GSH was determined using the Abcam Inc. detection assay kit (ab138881) (Abcam, Cambridge, UK) according to the manufacturer’s guidelines. The samples were prepared by lysis of total cell proteins in PBS/0.5% NP-40 lysis buffer followed by a 1:50 dilution. Serial dilutions of GSH standards were prepared along with assay mixtures for the detection of GSH using 100× Thiol green solution, incubated for 30 min and read at 490/520 nm. mRNA expression of inflammatory genes in cultured cells: HepG2 cells were seeded (8 × 105 cells per well) in 12-well plates and incubated alone or treated with H2 O2 in the presence of CeO2 NPs (10 µg/mL) or vehicle for 6 and 24 h. Total RNA from cultured cells was extracted using the commercially available kit: TRIZOL (TRI Reagent; Sigma-Aldrich, St. Louis, MO, USA). The RNA concentration was determined using spectrophotometric analysis (ND-100 spectrophotometer; Thermo Fisher Scientific, Waltham, MA, USA). One microgram of total RNA was reverse-transcribed using a cDNA synthesis kit (High-Capacity cDNA Reverse Transcription Kit; Applied Biosystems, Foster City, CA, USA). Primers and probes for human TNFα (left: 5’-CGCTCCCCAAGAAGACAG-3’, right: 5’-CTGCCACGATCAGGAAGG-3’; probe number 73), inducible nitric oxide synthase (iNOS) (left: 5’-TGCATGGATAAGTACAGGCTGA-3, right: 5’-CCATTGCCAAACGTACTGGT-3’; probe number 66), myeloperoxidase (MPO) (left: 5’-CGTCAACTGCGAGACCAG-3’, right: 5’-GTCATTGGGCGGGATCTT-3’; probe number 66), prostaglandin-endoperoxide synthase 1 (PTGS1) (left: 5’-TTCTCTCGCCAGATTGCTG-3’, right: 5’-CCGAGACTCCCTGATGACA-3’; probe number 76) and hypoxanthine-guanine phosphoribosyltransferase (HPRT) used as an endogenous standard (left: 5’-TGACCTTGATTTATTTTGCATACC-3’, right: 5’-CGAGCAAGACGTTCAGTCCT-3’; probe number 73) were designed according to human TNFα, iNOS, MPO, PTGS1 and HPRT sequences (GenBank NM_000594.2, NM_000625.4, NM_000250.1, NM_16931.3, NM_000962.2 and NM_000194.2, respectively) to include intron spanning using the Universal ProbeLibrary Assay Design Center through ProbeFinder version 2.5 software (Roche Diagnostics, Indianapolis, IN, USA; Available online: http://lifescience.roche.com/shop/en/mx/overviews/brand/universal-probe-library). Real-time quantitative polymerase chain reaction was analyzed in duplicate and performed with the LightCycler 480 (Roche Diagnostics, Basel, Switzerland), as previously described [46,47]. A 10-µL total volume reaction of diluted 1:8 cDNA, 200 nM primer dilution, 100 nM prevalidated 9-mer probe (Universal ProbeLibrary) and FastStart TaqMan Probe Master (Roche Diagnostics, Basel, Switzerland) were used in each PCR. A fluorescence signal was captured during each of the 45 cycles (denaturizing for 10 s at 95 ◦ C, annealing for 20 s at 60 ◦ C, and extension for 1 s at 72 ◦ C). Water was used as a negative control. Relative quantification was calculated using the comparative threshold cycle (CT), which is inversely related to the abundance of mRNA transcripts in the initial sample. The mean CT of duplicate measurements was used to calculate ∆CT as the difference in CT for target and reference. The relative quantity of product was expressed as fold induction of the target gene compared with the reference gene according to the formula 2−∆∆CT , where ∆∆CT represents ∆CT values normalized with the mean ∆CT of control samples. 4.3. Oxidative Stress Gene Expression PCR Array in Cultured Cells HepG2 cells were seeded (8 × 105 cells per well) in 12-well plates and incubated alone or treated with H2 O2 in the presence of CeO2NPs (10 µg/mL) or vehicle for 24 h to assess changes in oxidative stress pathways. Total RNA was extracted using TRIZOL as described above. To remove residual DNA, Int. J. Mol. Sci. 2019, 20, 5959 17 of 22 RNA preparations were treated with RNase-Free DNAse set (Qiagen, Hilden, Germany). First-strand cDNA was synthesized from 1 µg total RNA using an RT2 first-strand kit (Qiagen, Hilden, Germany), and PCR arrays were performed according to the manufacturer’s protocols (SABiosciences, Frederick, MD, USA). Real-time PCR array was performed using the Human Oxidative Stress RT2 Profiler™ PCR array, (SABiosciences, Frederick, MD, USA) according to the manufacturer’s protocol. This PCR array combines the quantitative performance of SYBR Green-based real-time PCR with the multiple gene profiling capabilities of microarray to profile the expression of 84 key genes involved in oxidative stress. PCR array plates were processed in a Light Cycler 480 (Roche Diagnostics, Basel, Switzerland) using automated baseline and threshold cycle detection. Gene expression was normalized to internal controls to determine the fold change in gene expression between test and control samples. The relative quantity of product was expressed as fold-induction of the target gene compared with the reference gene according to the formula 2−∆∆CT . Data were interpreted using the SABiosciences’ web-based PCR array data analysis tool. Statistical significance was obtained after performing a Student’s t-test analysis compared to control samples (Available online: http://pcrdataanalysis.sabiosciences.com/pcr/ arrayanalysis.php). 4.4. Statistical Analysis Quantitative data were analyzed using GraphPad Prism 5 (GraphPad Software Inc., San Diego, CA, USA), and statistical analysis of the results was performed using one-way analysis of variance (ANOVA) with the Newman-Keuls post hoc test and the Kruskal-Wallis test with the Dunn post hoc test when appropriate. Results are expressed as mean ± SE and considered significant at a p level less than 0.05. The study was performed according to the criteria of the Investigation and Ethics Committee of the Hospital Clínic Universitari of Barcelona. 4.5. Phosphoproteomic Analysis Large-scale phosphoproteomics was used to gain further insight on the kinase signaling pathways mainly affected by the CeO2 NPs treatment. Thousands of phosphorylation sites were simultaneously quantified to estimate changes in kinase activity induced by CeO2 NPs in HepG2 cells under oxidation-induced conditions. HepG2 cells were cultured as described above. Thereafter, the cells were switched to serum free medium for 24 h. Then medium was replaced and the cells were exposed to 1.5 mM H2 O2 (Sigma-Aldrich, St. Louis, MO, USA), 1.5 mM H2 O2 and 10 µg/mL CeO2 NPs or vehicle (TMAOH, 0.17 mM) for 1 h. Confluent cells were washed three times with cold PBS supplemented with 1 mM Na3 VO4 and 1 mM NaF and lysed in urea buffer (8 M urea in 20 mM HEPES, pH: 8.0, supplemented with 1 mM Na3 VO4 , 1 mM NaF, 1 mM Na4P2O7 and 1 mM β-glycerophophate). Five independent biological replicates were collected for each condition and kept at −80 ◦ C. Cell lysates were further homogenized by sonication (30 cycles of 30 s on 30 s off; Diagenode Bioruptor® Plus, Liege, Belgium) and insoluble material was removed by centrifugation. Protein was quantified using BCA (Thermo Fisher Scientific, Waltham, MA, USA). For tryptic digestion, protein extracts (500 µg) were subjected to cysteine alkylation using sequential incubation with 10 mM dithiothreitol (DDT) and 16.6 mM iodoacetamide (IAM) for 1 h and 30 min respectively at 25 ◦ C and agitation. The urea concentration was then reduced to 2 M by the addition of 20 mM HEPES (pH: 8.0). Then, 100µL of equilibrated trypsin beads ((50% slurry of TLCK-trypsin (Thermo-Fisher Scientific; Cat. #20230, Waltham, MA, USA) were added, and samples were incubated overnight at 37 ◦ C. Trypsin beads were equilibrated by 3 washes with 20 mM HEPES (pH: 8.0). Trypsin beads were removed by centrifugation and the resulting peptide solutions were desalted with C-18-Oasis cartridges as indicated by the manufacturer. Briefly, oasis cartridges were conditioned with 1 mL acetonitrile (ACN) and equilibrated with 2.5 mL of wash solution (0.1% trifluoroacetic acid (TFA) and 2% ACN). Peptides were loaded in the cartridges and washed with 1 mL of wash solution. Finally, peptides were eluted with 0.5 mL of glycolic acid buffer 1 (1 M glycolic acid, 5% TFA, 50 % ACN). Enrichment of phosphorylated peptides was performed with TiO2 . The eluents were normalized to 1 mL with glycolic acid buffer 2 (1 M glycolic acid, 5% TFA, 80 % ACN) and incubated with 50 µl of TiO2 buffer (a 50% slurry in 1% TFA) for 5 min at room temperature. TiO2 beads were packed by centrifugation in Int. J. Mol. Sci. 2019, 20, 5959 18 of 22 empty spin columns (Gygen Corporation; Cat. TT2EMT) previously equilibrated using glycolic acid, 50% ACN and ammonium bicarbonate buffer (20 mM NH4 HCO3 (pH: 6.8) in 50% ACN). For phosphopeptide elution, beads were incubated for 1 min at room temperature with 50 µL of 5% NH4 OH in 50% ACN and centrifuged. This step was repeated three times. Finally, samples were snap frozen, dried in a SpeedVac, and the pellets were stored at −80 ◦ C. Phosphopetide pellets were resuspended in 9 µL of reconstitution buffer (20 fmol/µL enolase in 3% ACN, 0.1% TFA) and 5.0 µL were loaded onto an LC-MS/MS system consisting of a Dionex UltiMate 3000 RSLC directly coupled to an Orbitrap Q-Exactive Plus mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). Phosphopeptides were loaded in a µ-pre-column (Acclaim™ PepMap™ 100 C18 LC; Cat 160454, Thermo Fisher Scientific, Waltham, MA, USA) and separated in an analytical column (Acclaim™ PepMap™ 100 C18 LC; Cat. 164569, Thermo Fisher Scientific, Waltham, MA, USA) using a gradient that runs from 3% to 23% B over 120 min. The UPLC system delivered a flow of 2 µL/min (loading) and 300 nL/min (gradient elution). Solvent A consists of 3% ACN: 0.1% FA and solvent B consists of 100% ACN; 0.1% FA. The Q-Exactive Plus acquired full scan survey spectra (m/z 375–1500) with a 70,000 FWHM resolution followed by data-dependent acquisition in which the 20 most intense ions were selected for HCD (higher energy collisional dissociation) and MS/MS scanning (200–2000 m/z) with a resolution of 17,500 FWHM. Peptide identification was performed by matching of the MS/MS data to the SwissProt database, restricted to human entries with the Mascot search engine [48]. Phosphopeptides with a mascot expectancy of < 0.005 (~2% false discovery rate) were included in a database of sites quantifiable by MS. Pascal software [49–51] was then used to obtain peak areas of extracted ions chromatograms of phosphorylated peptides in this database across all the samples compared. The significance of the differences in the log2-transformed data across samples was assessed using the Student’s t-test. Inference of kinase activities from the phosphoproteomic data was performed using kinase substrate enrichment analysis (KSEA), as described previously [18]. 5. Conclusions Our study shows that CeO2 NPs directly protect human-derived hepatocytes from oxidative damage, reducing ROS generation and inflammatory gene expression, thus opening new avenues to use CeO2 NPs in human liver diseases. Additionally, specific cell kinase driven signaling pathways downregulated by CeO2 NPs treatment in cells under oxidative stress have been identified for the first time. Further experiments with appropriate inhibitors are needed to demonstrate a direct cause-effect relationship between CeO2 NPs and the modulation of mTOR, ERK and PKACA pathways. Moreover, additional in vivo validations are necessary to determine the potential clinical applications of these findings. Author Contributions: conceptualization, S.C., D.O., G.C., G.F.-V., M.M.-R., P.C. (Pedro Cutillas)and W.J.; methodology, S.C., M.P., D.O., E.C., J.R., P.M.-L. and P.C. (Pedro Casado); formal analysis, S.C., M.P., D.O. and P.C. (Pedro Casado); data curation, S.C., M.P., D.O. and P.C. (Pedro Casado); writing—original draft preparation, S.C., D.O. and W.J.; writing—review and editing, S.C., G.C., D.O., E.C., P.C. (Pedro Cutillas), G.F.-V., V.P. and W.J.; supervision, G.C. and W.J.; funding acquisition, G.C., V.P and W.J. Funding: This research was funded by grants to W. Jiménez from Ministerio de Economia y Competitividad [grants, SAF2015-64126-R, RTI2018-094734-B-C21], to M. Morales-Ruiz [grant SAF2016-75358-R] and to G. Casals [grant PI-15/00777]; cofinanced by FEDER, European Union, a way of making Europe, Agència de Gestió d’Ajuts Universitaris i de Recerca [grant SGR 2014/219]; and Fundació La Marató de TV3 [grant Marató 120930]; Wuyi University Funding for Hight Talents Introduction, grant number 2018TP010 and Foundation from Department of Education of Guangdong Province, grant number 2016KCXTD005 and 2017KSYS010, to Eudald Casals The Centro de Investigación Biomédica en Red de Enfermedades Hepáticas y Digestivas (CIBERehd) is funded by the Instituto de Salud Carlos III. Conflicts of Interest: The authors declare no conflicts of interest. Int. J. Mol. Sci. 2019, 20, 5959 19 of 22 Abbreviations 2’,7’-DCF-DA 20 ,70 -dichlorofluorescin diacetate CeO2 NPs Cerium Oxide Nanoparticles DDT Dithiothreitol DMEM Dulbecco’s Modified Eagle Medium FCS Fetal Calf Serum HBSS Hank’s Balanced Salt Solution HCC Hepatocellular Carcinoma HCD Higher Energy Collisional Dissociation HR-TEM High Resolution Transmission Electron Microscopy KSEA Kinase-Substrate Enrichment Analysis LC-MS/MS Liquid Chromatography Tandem Mass Spectrometry LPS Lipopolysaccharide MS Mass Spectrometry MS/MS Tandem mass spectrometry NPs Nanoparticles PCR Polymerase Chain Reaction ROS Radical Oxygen Species RT-PCR Real Time PCR SOD Superoxide Dismutase UPLC Ultrahigh Pressure Liquid Chromatography UV-VIS Ultraviolet-Visible XRD X-ray Diffraction References 1. Laviña, B.; Gracia-Sancho, J.; Rodríguez-Vilarrupla, A.; Chu, Y.; Heistad, D.D.; Bosch, J.; García-Pagán, J.C. Superoxide dismutase gene transfer reduces portal pressure in CCl4 cirrhotic rats with portal hypertension. Gut 2009, 58, 118–125. [CrossRef] [PubMed] 2. Ma, Z.; Zhang, Y.; Li, Q.; Xu, M.; Bai, J.; Wu, S. Resveratrol improves alcoholic fatty liver disease by downregulating HIF-1α expression and mitochondrial ROS production. PLoS ONE 2017, 12, e0183426. [CrossRef] [PubMed] 3. Rojkind, M.; Kershenobich, D. Effect of colchicine on collagen, albumin and transferrin synthesis by cirrhotic rat liver slices. Biochim. Biophys. Acta 1975, 378, 415–423. [CrossRef] 4. Yogalakshmi, B.; Viswanathan, P.; Anuradha, C.V. Investigation of antioxidant, anti-inflammatory and DNA-protective properties of eugenol in thioacetamide-induced liver injury in rats. Toxicology 2010, 268, 204–212. [CrossRef] [PubMed] 5. Houglum, K.; Venkataramani, A.; Lyche, K.; Chojkier, M. A pilot study of the effects of d-alphatocopherol on hepatic stellate cell activation in chronic hepatitis C. Gastroenterology 1997, 113, 1069–1073. [CrossRef] 6. Singal, A.K.; Jampana, S.C.; Weinman, S.A. Antioxidants as therapeutic agents for liver disease. Liver Int. 2011, 31, 1432–1448. [CrossRef] 7. Chalasani, N.; Younossi, Z.; Lavine, J.E.; Charlton, M.; Cusi, K.; Rinella, M.; Harrison, S.A.; Brunt, E.M.; Sanyal, A.J. The diagnosis and management of nonalcoholic fatty liver disease: Practice guidance from the American Association for the Study of Liver Diseases. Hepatology 2017, 67, 328–357. [CrossRef] 8. European Association for the Study of the Liver (EASL); European Association for the Study of Diabetes (EASD); European Association for the Study of Obesity (EASO). EASL–EASD–EASO Clinical Practice Guidelines for the management of non-alcoholic fatty liver disease. J. Hepatol. 2016, 64, 1388–1402. [CrossRef] 9. Wason, M.S.; Zhao, J. Cerium oxide nanoparticles: Potential applications for cancer and other diseases. Am. J. Transl Res. 2013, 5, 126–131. 10. Heckert, E.G.; Karakoti, A.S.; Seal, S.; Self, W.T. The role of cerium redox state in the SOD mimetic activity of nanoceria. Biomaterials 2008, 29, 2705–2709. [CrossRef] Int. J. Mol. Sci. 2019, 20, 5959 20 of 22 11. Barkam, S.; Ortiz, J.; Saraf, S.; Eliason, N.; Mccormack, R.; Das, S.; Gupta, A.; Neal, C.; Petrovici, A.O.; Hanson, C.; et al. Modulating the Catalytic Activity of Cerium Oxide Nanoparticles with the Anion of the Precursor Salt. J. Phys. Chem. C 2017, 121, 20039–20050. [CrossRef] 12. Oró, D.; Yudina, T.; Fernández-Varo, G.; Casals, E.; Reichenbach, V.; Casals, G.; de la Presa, B.G.; Sandalinas, S.; Carvajal, S.; Puntes, V.; et al. Cerium oxide nanoparticles reduce steatosis, portal hypertension and display anti-inflammatory properties in rats with liver fibrosis. J. Hepatol. 2016, 64, 691–698. [CrossRef] 13. Carvajal, S.; Perramón, M.; Oró, D.; Casals, E.; Fernández-Varo, G.; Casals, G.; Parra, M.; de la Presa, B.G.; Ribera, J.; Pastor, Ó.; et al. Cerium oxide nanoparticles display antilipogenic effect in rats with non-alcoholic fatty liver disease. Sci. Rep. 2019, 9, 12848. [CrossRef] [PubMed] 14. Xu, C.; Qu, X. Cerium oxide nanoparticle: A remarkably versatile rare earth nanomaterial for biological applications. NPG Asia Mater. 2014, 6, e90. [CrossRef] 15. Raza, H.; John, A.; Benedict, S. Acetylsalicylic acid-induced oxidative stress, cell cycle arrest, apoptosis and mitochondrial dysfunction in human hepatoma HepG2 cells. Eur. J. Pharm. 2011, 668, 15–24. [CrossRef] 16. Melgar-Lesmes, P.; Pauta, M.; Reichenbach, V.; Casals, G.; Ros, J.; Bataller, R.; Morales-Ruiz, M.; Jiménez, W. Hypoxia and proinflammatory factors upregulate apelin receptor expression in human stellate cells and hepatocytes. Gut 2011, 60, 1404–1411. [CrossRef] 17. Casals, E.; Pfaller, T.; Duschl, A.; Oostingh, G.J.; Puntes, V.F. Hardening of the nanoparticle-protein corona in metal (Au, Ag) and oxide (Fe3 O4 , CoO, and CeO2 ) nanoparticles. Small 2011, 7, 3479–3486. [CrossRef] 18. Casado, P.; Rodriguez-Prados, J.C.; Cosulich, S.C.; Guichard, S.; Vanhaesebroeck, B.; Joel, S.; Cutillas, P.R. Kinase-Substrate Enrichment Analysis provides insights into the heterogeneity of signaling pathway activation in leukemia cells. Sci. Signal. 2012, 6, 268. [CrossRef] 19. Angulo-Urarte, A.; Casado, P.; Castillo, S.D.; Kobialka, P.; Kotini, M.P.; Figueiredo, A.M.; Castel, P.; Rajeeve, V.; Milà-Guasch, M.; Millan, J.; et al. Endothelial cell rearrangements during vascular patterning require PI3-kinase-mediated inhibition of actomyosin contractility. Nat. Commun. 2018, 9, 4826–4842. [CrossRef] 20. Niu, J.; Wang, K.; Kolattukudy, P.E. Cerium oxide nanoparticles inhibit oxidative stress and nuclear factor-κB activation in H9c2 cardiomyocytes exposed to cigarette smoke extract. J. Pharm. Exp. Ther. 2011, 338, 53–61. [CrossRef] 21. Akhtar, M.J.; Ahamed, M.; Alhadlaq, H.A.; Khan, M.A.; Alrokayan, S.A. Glutathione replenishing potential of CeO2 nanoparticles in human breast and fibrosarcoma cells. J. Colloid Interface Sci. 2015, 453, 21–27. [CrossRef] [PubMed] 22. Chen, S.; Hou, Y.; Cheng, G.; Zhang, C.; Wang, S.; Zhang, J. Cerium oxide nanoparticles protect endothelial cells from apoptosis induced by oxidative stress. Biol. Trace Elem. Res. 2013, 154, 156–166. [CrossRef] [PubMed] 23. Xia, T.; Kovochich, M.; Liong, M.; Mädler, L.; Gilbert, B.; Shi, H.; Yeh, J.I.; Zink, J.I.; Nel, A.E. Comparison of the mechanism of toxicity of zinc oxide and cerium oxide nanoparticles based on dissolution and oxidative stress properties. ACS Nano 2008, 2, 2121–2134. [CrossRef] [PubMed] 24. Korsvik, C.; Patil, S.; Seal, S.; Self, W.T. Superoxide dismutase mimetic properties exhibited by vacancy engineered ceria nanoparticles. Chem. Commun. 2007, 1056–1058. [CrossRef] [PubMed] 25. Pirmohamed, T.; Dowding, J.M.; Singh, S.; Wasserman, B.; Heckert, E.; Karakoti, A.S.; King, J.E.; Seal, S.; Self, W.T. Nanoceria exhibit redox state-dependent catalase mimetic activity. Chem. Commun. 2010, 46, 2736–2738. [CrossRef] [PubMed] 26. Nelson, B.C.; Walker, M.L.; Riley, K.R.; Sims, C.M. Antioxidant Cerium Oxide Nanoparticles in Biology and Medicine. Antioxidants 2016, 5, 15. [CrossRef] 27. Cichoz-Lach, H.; Michalak, A. Oxidative stress as a crucial factor in liver diseases. World J. Gastroenterol. 2014, 20, 8082–8091. [CrossRef] 28. Zhao, G.; Yu, R.; Deng, J.; Zhao, Q.; Li, Y.; Joo, M.; van Breemen, R.B.; Christman, J.W.; Xiol, L. Pivotal role of reactive oxygen species in differential regulation of lipopolysaccharide-induced prostaglandins production in macrophages. Mol. Pharm. 2013, 83, 167–178. [CrossRef] 29. Pflüger, P.; Viau, C.M.; Coelho, V.R.; Berwig, N.A.; Staub, R.B.; Pereira, P.; Saffi, J. Gamma-decanolactone inhibits iNOS and TNF-alpha production by lipopolysaccharide-activated microglia in N9 cells. Eur. J. Pharm. 2016, 780, 38–45. [CrossRef] Int. J. Mol. Sci. 2019, 20, 5959 21 of 22 30. Pulli, B.; Ali, M.; Iwamoto, Y.; Zeller, M.W.; Schob, S.; Linnoila, J.J.; Chen, J.W. Mieloperoxidase-hepatocyte-stellate cell cross talk promotes hepatocyte injury and fibrosis in experimental nonalcoholic steatohepatitis. Antioxid. Redox Signal. 2015, 23, 1255–1269. [CrossRef] 31. Rensen, S.S.; Slaats, Y.; Nijhuis, J.; Jans, A.; Bieghs, V.; Driessen, A.; Malle, E.; Greve, J.W.; Buurman, W.A. Increased hepatic myeloperoxidase activity in obese subjects with nonalcoholic steatohepatitis. Am. J. Pathol. 2009, 175, 1473–1482. [CrossRef] [PubMed] 32. Langenbach, R.; Morham, S.G.; Tiano, H.F.; Loftin, C.D.; Ghanayem, B.I.; Chulada, P.C.; Mahler, J.F.; Lee, C.A.; Goulding, E.H.; Kluckman, K.D.; et al. Prostaglandin synthase 1 gene disruption in mice reduces arachidonic acid-induced inflammation and indomethacin-induced gastric ulceration. Cell 1995, 83, 483–492. [CrossRef] 33. Choi, H.S.; Bosetti, F. Cyclooxygenase-1 null mice show reduced neuroinflammation in response to β-amyloid. Aging 2009, 1, 234–244. [CrossRef] [PubMed] 34. Förstermann, U.; Sessa, W.C. Nitric oxide synthases: Regulation and function. Eur. Heart J. 2012, 33, 829–837. [CrossRef] [PubMed] 35. Wu, F.; Tyml, K.; Wilson, J.X. iNOS expression requires NADPH oxidase-dependent redox signaling in microvascular endothelial cells. J. Cell Physiol. 2008, 217, 207–214. [CrossRef] [PubMed] 36. Pawson, T.; Linding, R. Network medicine. FEBS Lett. 2008, 582, 1266–1270. [CrossRef] 37. Linding, R.; Jensen, L.J.; Ostheimer, G.J.; Van Vugt, M.A.; Jørgensen, C.; Miron, M.I.; Diella, F.; Colwill, K.; Taylor, L.; Elder, K.; et al. Systematic discovery of in vivo phosphorylation networks. Cell 2007, 129, 1415–1426. [CrossRef] 38. Schwartz, D.; Gygi, S.P. An iterative statistical approach to the identification of protein phosphorylation motifs from large-scale data sets. Nat. Biotechnol. 2005, 23, 1391–1398. [CrossRef] 39. Kim, H.; Yoo, J.E.; Cho, J.Y.; Oh, B.K.; Yoon, Y.S.; Han, H.S.; Lee, H.S.; Jang, J.J.; Jeong, S.H.; Kim, J.W.; et al. Telomere length; TERT and shelterin complex proteins in hepatocellular carcinomas expressing “stemness”-related markers. J. Hepatol. 2013, 59, 746–752. [CrossRef] 40. Oh, B.K.; Kim, Y.J.; Park, C.; Park, Y.N. Up-regulation of telomere-binding proteins; TRF1; TRF2; and TIN2 is related to telomere shortening during human multistep hepatocarcinogenesis. Am. J. Pathol. 2005, 166, 73–80. [CrossRef] 41. Gu, D.L.; Chen, Y.H.; Shih, J.H.; Lin, C.H.; Jou, Y.S.; Chen, C.F. Target genes discovery through copy number alteration analysis in human hepatocellular carcinoma. World J. Gastroenterol. 2013, 19, 8873–8879. [CrossRef] [PubMed] 42. Sun, X.; Wang, S.C.; Wei, Y.; Luo, X.; Jia, Y.; Li, L.; Gopal, P.; Zhu, M.; Nassour, I.; Chuang, J.C.; et al. Arid1a has context-dependent oncogenic and tumor suppressor functions in liver cancer. Cancer Cell 2017, 32, 574–589. [CrossRef] [PubMed] 43. Gaiser, B.K.; Fernandes, T.F.; Jepson, M.A.; Lead, J.R.; Tyler, C.R.; Baalousha, M.; Biswas, A.; Britton, G.J.; Cole, P.A.; Johnston, B.D.; et al. Interspecies comparisons on the uptake and toxicity of silver and cerium dioxide nanoparticles. Environ. Toxicol. Chem. 2012, 31, 144–154. [CrossRef] [PubMed] 44. Singh, R.; Singh, S. Redox-dependent catalase mimetic cerium oxide-based nanozyme protect human hepatic cells from 3-AT induced acatalasemia. Colloids Surf. B Biointerfaces 2019, 175, 625–635. [CrossRef] [PubMed] 45. Ni, D.; Wei, H.; Chen, W.; Bao, Q.; Rosenkrans, Z.T.; Barnhart, T.E.; Ferreira, C.A.; Wang, Y.; Yao, H.; Sun, T.; et al. Ceria Nanoparticles Meet Hepatic Ischemia-Reperfusion Injury: The Perfect Imperfection. Adv. Mater. 2019, 31, e1902956. [CrossRef] 46. Reichenbach, V.; Ros, J.; Fernández-Varo, G.; Casals, G.; Melgar-Lesmes, P.; Campos, T.; Makriyannis, A.; Morales-Ruiz, M.; Jiménez, W. Prevention of fibrosis progression in CCl4 -treated rats: Role of the hepatic endocannabinoid and apelin systems. J. Pharm. Exp. Ther. 2012, 340, 629–637. [CrossRef] 47. Melgar-Lesmes, P.; Casals, G.; Pauta, M.; Ros, J.; Reichenbach, V.; Bataller, R.; Morales-Ruiz, M.; Jimenez, W. Apelin mediates the induction of profibrogenic genes in human hepatic stellate cells. Endocrinology 2010, 151, 5306–5314. [CrossRef] 48. Perkins, D.N.; Pappin, D.J.; Creasy, D.M.; Cottrell, J.S. Probability-based protein identification by searching sequence database using mass spectrometry data. Electrophoresis 1999, 20, 3551–3567. [CrossRef] 49. Casado, P.; Cutillas, P.R. A self-validating quantitative mass spectrometry method for assessing the accuracy of high-content phosphoproteomic experiments. Mol. Cell. Proteom. 2011, 10, M110.003079. [CrossRef] Int. J. Mol. Sci. 2019, 20, 5959 22 of 22 50. Cutillas, P.R.; Venhaesebroeck, B. Quantitative profile of five murine core proteomes using label-free functional proteomics. Mol. Cell. Proteom. 2007, 6, 1560–1573. [CrossRef] 51. Casado, P.; Bilanges, B.; Rajeeve, V.; Vanhaesebroeck, B.; Cutillas, P.R. Environmental stress affects the activity of metabolic and growth factor signaling networks and induces autophagy markers in MCF7 breast cancer cells. Mol. Cell. Proteom. 2014, 13, 836–848. [CrossRef] [PubMed] © 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).