Folic Acid Homeostasis and Its Pathways Related to Hepatic Oxidation in Adolescent Rats Exposed to Binge Drinking

  Citation: Gallego-Lopez, M.d.C.; Ojeda, M.L.; Romero-Herrera, I.; Nogales, F.; Carreras, O. Folic Acid Homeostasis and Its Pathways Related to Hepatic Oxidation in Adolescent Rats Exposed to Binge Drinking. Antioxidants 2022,11, 362. https://doi.org/10.3390/ antiox11020362 Academic Editors: Marco Fiore and Mauro Ceccanti Received: 15 December 2021 Accepted: 8 February 2022 Published: 11 February 2022 Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations. Copyright: © 2022 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 (https:// creativecommons.org/licenses/by/ 4.0/). antioxidants Article Folic Acid Homeostasis and Its Pathways Related to Hepatic Oxidation in Adolescent Rats Exposed to Binge Drinking María del Carmen Gallego-Lopez , María Luisa Ojeda * , Inés Romero-Herrera, Fátima Nogales and Olimpia Carreras Department of Physiology, Faculty of Pharmacy, Seville University, 41012 Seville, Spain; [email protected] (M.d.C.G.-L.); iner[email protected] (I.R.-H.); [email protected] (F.N.); [email protected] (O.C.) *Correspondence: [email protected]; Tel.: +34-954556518 Abstract: Chronic ethanol consumption and liver disease are intimately related to folic acid (FA) homeostasis. Despite the fact that FA decreases lipid oxidation, its mechanisms are not yet well elucidated. Lately, adolescents have been practising binge drinking (BD), consisting of the intake of a high amount of alcohol in a short time; this is a particularly pro-oxidant form of consumption. The aim of this study is to examine, for the first time, FA homeostasis in BD adolescent rats and its antioxidant properties in the liver. We used adolescent rats, including control rats and rats exposed to an intermittent intraperitoneal BD model, supplemented with or without FA. Renal FA reabsorption and renal FA deposits were increased in BD rats; hepatic deposits were decreased, and heart and serum levels remained unaffected. This depletion in the liver was accompanied by higher transaminase levels; an imbalance in the antioxidant endogenous enzymatic system; lipid and protein oxidation; a decrease in glutathione (GSH) levels; hyper-homocysteinemia (HHcy); an increase in NADPH oxidase (NOX) 1 and NOX4 enzymes; an increase in caspase 9 and 3; and a decrease in the anti-apoptotic metallopeptidase inhibitor 1. Furthermore, BD exposure increased the expression of uncoupled endothelial nitric oxide synthase (eNOS) by increasing reactive nitrogen species generation and the nitration of tyrosine proteins. When FA was administered, hepatic FA levels returned to normal levels; transaminase and lipid and protein oxidation also decreased. Its antioxidant activity was due, in part, to the modulation of superoxide dismutase activity, GSH synthesis and NOX1, NOX4 and caspase expression. FA reduced HHcy and increased the expression of coupled eNOS by increasing tetrahydrobiopterin expression, avoiding nitrosative stress. In conclusion, FA homeostasis and its antioxidant properties are affected in BD adolescent rats, making it clear that this vitamin plays an important role in the oxidative, nitrosative and apoptotic hepatic damage generated by acute ethanol exposure. For this, FA supplementation becomes a potential BD therapy for adolescents, preventing future acute alcohol-related harms. Keywords: binge drinking; folic acid; oxidative stress; apoptosis; nitrosative stress 1. Introduction The relationship between folic acid (FA), chronic ethanol consumption (Chr-EtOH) and alcoholic liver disease (ALD) is well recognized in clinical and experimental research. ChrEtOH suppresses key biological processes directly relevant to FA and folate metabolism, such as folate intestinal absorption, liver uptake and storage, urinary excretion, and folate enzymes synthesis and methylation; it also leads to lower folate intake [ 1 ]. Therefore, currently, FA is supplied to Chr-EtOH patients in order to prevent ALD, as this is the main vitamin deficiency from which they suffer [2]. The mechanisms of Chr-EtOH injury are multifactorial, involving several pathways, most of them related to its oxidative metabolism, which takes place mainly in the liver. The principal product of this metabolism is acetaldehyde, which leads to changes in the Antioxidants 2022,11, 362. https://doi.org/10.3390/antiox11020362 https://www.mdpi.com/journal/antioxidants Antioxidants 2022,11, 362 2 of 22 reduced and oxidized nicotinamide adenine dinucleotide (NADH-NAD + ) ratio, increasing mitochondrial oxidation. One of these EtOH metabolizer enzymes is CYP2E1, which produces acetaldehyde but also reactive oxygen species (ROS) directly, increasing oxidative stress (OS) [ 3 ]. Biomolecular oxidative damage is considered one of the main mechanisms related to ALD. Synthetic FA, together with natural folates, is a member of the B9 vitamin family and is essential for amino acid metabolism. FA is taken up from the diet and bio-transformed mainly in the liver by the enzyme dihidrofolate reductase (DHFR). It functions as a coenzyme in reactions of one-carbon transfer, needed for the biosynthesis of several essential molecules [ 4 ]. Adequate intake of FA is vital for metabolism, cellular homeostasis and DNA synthesis [ 5 ]. In addition, in the last meta-analysis displayed, it is suggested that FA supplementation improves the antioxidant defense system by increasing serum concentrations of glutathione (GSH) and decreasing lipid oxidation [ 6 ]. Despite the fact that different authors have described this property, all studies approach the issue from a onesided point of view. For instance, “ in vitro ” studies describe FA as a novel treatment to combat hepatocellular carcinoma by modulating ROS sinker proteins and maintaining mitochondrial redox homeostasis [ 7 ]. Other “ in vitro ” studies suggest a direct action of FA on OS generation as a ROS scavenger, acting as a reducing agent [ 8 ]; however, this action is not clear “ in vivo ” [ 9 , 10 ]. Most authors relate the antioxidant properties of FA to its ability to lower the pro-oxidant homocysteine (Hcy). Hyper-homocysteinemia (HHcy) increases hepatic and renal nicotinamide adenine dinucleotide phosphate (NADPH) oxidase (NOX) activity, which in turn leads to high amounts of ROS production, increasing lipid peroxidation, and it also affects the mRNA levels of the mitochondrial NOX4, the mitochondrial Complex II and calcium levels, contributing to apoptosis [ 11 – 13 ]. In the kidneys, Hcy also increases oxidation by affecting the activities of the pro-oxidant xanthine oxidase and the antioxidant superoxide dismutase (SOD) [ 12 ]. FA metabolism is deeply related to the Hcy metabolism through the methionine cycle. Therefore, in the liver, FA supplementation inhibits NOX-ROS production, decreasing OS, by decreasing Hcy [ 11 ]. Recently, Ju et al. suggested that FA, by inducing the mitochondrial enzyme methylene tetrahydrofolate dehydrogenase 2 (MTHFD 2 ), could modulate NOX4 and redox homeostasis together to modulate apoptosis in colorectal cancer [ 14 ]. However, all these authors point to other FA antioxidant effects not related to Hcy remethylation. FA stimulates the activity of the enzyme γ -glutamylcysteine synthetase ( γ -GCS), increasing the transsulfuration pathway, and the synthesis of GSH. GSH is an endogenous antioxidant that is also necessary for the correct function of the endogenous antioxidant enzyme glutathione peroxidase (GPx) [ 15 ]. Finally, different authors studying endothelial cells point to FA as an important protector from hypoxia by decreasing both ROS and apoptosis, linked to NOX4 pathways by the induction of nitric oxide (NO) production. This action is attributed to the fact that FA increases endothelial NO synthase (eNOS) in its coupled form, which is the antioxidant form [ 16 ]. This mechanism is linked to the upregulation of the enzyme DHFR in response to FA administration, which enhances tetrahydrobiopterin (BH4), promoting eNOS recoupling [ 17 – 19 ]. In this regard, there is not a well-elucidated overview of all these mechanisms in the same study. Binge drinking (BD) is an acute way of consuming EtOH, defined by the National Institute on Alcohol Abuse and Alcoholism (NIAAA) as a pattern of drinking alcohol that brings blood alcohol concentration (BAC) to 0.08% or higher ( ≥ 0.08 g/dL) during a short period of time [ 20 ]. Before and during the COVID-19 pandemic, this strong method of EtOH consumption has been the most widespread among teenagers [ 21 , 22 ], being a public health concern since adolescence is a stage that is especially vulnerable to the toxic effects of EtOH. Acute EtOH consumption leads to different biological effects than Chr-EtOH, since in its metabolism, it produces higher amounts of ROS by increasing CYP2E1 activity [ 23 ]. BD exposure is an especially potent pro-oxidant that could produce different changes in FA homeostasis, compromising the oxidative balance even more. Nevertheless, there are few studies related to acute EtOH consumption and FA homeostasis. In human volunteers, Antioxidants 2022,11, 362 3 of 22 serum FA levels fell and returned rapidly to normal values after acute alcohol exposure [ 24 ]. The mechanisms for these quick phenomena remain unexplained. In this context, our research group found that in adolescent rats, 12 h after BD exposure, serum FA levels were unaltered and peripheral oxidation took place. When these animals were supplemented with FA, serum FA levels increased and lipid and DNA oxidation significantly decreased [ 25 ]. However, plasma levels of metabolites are not a simple reflection of changes in liver levels of the same metabolites [ 26 ]; therefore, FA tissue deposits should be analyzed. In this study, for the first time, FA tissue levels are measured together with FA intake and FA in serum and urine in order to increase knowledge about FA homeostasis in adolescent rats exposed to BD. Moreover, to understand the possible repercussion of this on the oxidative disruption provoked by BD exposure, different oxidative pathways will be analyzed at the same time “ in vivo ”, also in FA-supplemented adolescent rats. We will obtain a general overview of FA involvement in ROS and reactive nitrogen species (RNS) over-production, OS/nitrosative stress (NS) balance and biomolecular damage which could induce apoptosis during BD exposure. 2. Materials and Methods 2.1. Experimental Design A total of 24 Wistar male adolescent rats (Centre of Production and Animal Experimentation, Vice-Rector’s Office for Scientific Research, University of Seville) were used to execute this experiment. These animals were received at 21 days old and housed in groups of 2 rats per cage for 1 week with free access to food and drink to acclimatize them to housing and handling conditions. The animals were kept in the animal house of the Faculty of Pharmacy at an automatically controlled temperature (22–23 ◦ C) and a 12 h light–dark cycle (07:00–19:00). At 28 days of age, when Wistar rats initiate the adolescence age, the animals were randomly divided into 4 experimental groups (n= 6): control group (C): rats were fed with commercial feed (2 ppm FA) and water ad libitum and, 3 days a week, they received an intraperitoneal (i.p.) injection of physiological saline solution (PSS); binge drinking group (BD): rats were fed with commercial feed (2 ppm FA) and water ad libitum and, 3 days a week, they received an i.p. injection of ethanol (3 g/kg); control folic acid group (CF): rats were fed with commercial feed supplemented with FA (8 ppm FA) and water ad libitum and, 3 days a week, they received an i.p. injection of PSS; and binge drinking folic acid group (BDF): rats were fed with commercial feed supplemented with FA (8 ppm FA) and water ad libitum and, 3 days a week, they received an i.p. injection of ethanol (3 g/kg). The animals remained under this treatment for 3 weeks (47 days old), at which point they were humanely sacrificed. Throughout the complete experimental period (28 to 47 days old), the rats suffered the typical-age alterations characteristic of adolescence, including those of the puberty phase; according to the review of Spear [ 27 ], there is an abundance of evidence that confirms that the animal model used in this experiment can be extrapolated to human adolescents, such as behaving like an adolescent or initiating alcohol and the consumption of other drugs. Animal care procedures and experimental protocols were in accordance with EU regulations (Council Directive 86/609/EEC, 24 November 1986 ) and were approved by the Ethics Committee of the University of Seville (CEEA-US2019-4). 2.2. Diet Used The diet used during the experimental protocol was a standard pellet diet (2014 Teklad Global 14% Protein Rodent Maintenance Diet, Harlan Laboratories, Barcelona, Spain) that covered all their nutritional and energy requirements, with an FA content of 2 ppm. Supplemented groups (CF and BDF) received an FA-supplemented diet prepared by adding a fine FA powder (ACOFARMA, Barcelona, Spain) at 8 ppm to the crushed standard pellet diet. Later on, supplemented biscuits were made by reconstituting the mixture with water and were dried at room temperature. Fresh batches of diet were made each week. Antioxidants 2022,11, 362 4 of 22 2.3. Binge Drinking Treatment EtOH exposed groups (BD and BDF) received the alcohol treatment called “binge drinking” by the administration of EtOH (3 g/kg) in PSS at 20% (v/v) via i.p. during 3 consecutive days per week for 3 weeks [ 28 ]. No i.p. injections were given during the remaining four days of each week. This i.p. forced BD method has been chosen in order to avoid digestive interferences with FA oral supplementation and to analyze the direct effect of this drug on the body. On the other hand, control groups (C and CF) were administered an equivalent volume of PSS i.p. over the same period. The injections were given starting at 7:00 pm when the dark cycle began. 2.4. Nutritional Control Each day during the three weeks of treatment, the food intake (g/day) was determined by measuring the difference between the feed placed and the feed remaining the next day using an analytical balance. FA intake ( µ g/day) was calculated from the data of the food intake, taking into account that the feed for the supplemented animals was prepared with a concentration of 8 ppm of FA and the base diet contained 2 ppm. At the same time, to monitor the development of the animals during the experiment, they were weighed every day using an analytical balance. The measurements were always taken at the same time, between 9:00–10:00 a.m., to avoid differences due to the circadian rhythm. 2.5. Samples Once the experimental period was over, the rats fasted for 12 h in individual metabolic cages, where urine samples were collected. Following that, 24 h after their last treatment with EtOH or PSS, the animals were weighed and anesthetized with an i.p. injection of 28% w/vurethane (Sigma-Aldrich, Madrid, Spain) (0.5 mL/100 g of body weight). Then, blood was obtained by intracardiac puncture through the thorax and placed in tubes. The clot was allowed to retract for 30 min at room temperature, and then it was centrifuged at 1300 × gfor 15 min to prepare the serum. Finally, when the blood was removed, the abdomen of each rat was opened by a midline incision, and the whole livers were extracted, debrided of adipose and connective tissue in ice-cold saline and weighed in order to calculate the liver somatic index (LSI% = liver weight/total rat weight). The kidneys and heart were also removed. All samples were frozen in liquid nitrogen and stored at − 80 ◦ C for future biochemical determinations. 2.6. Serum Measurements The serum levels of Hcy and the transaminases alanine aminotransferase (ALT) and aspartate aminotransferase (AST) were measured with an automated analyzer (Technicon RA-1000, Bayer Diagnostics, Leverkusen, Germany). Metallopeptidase inhibitor 1 (TIMP-1) was analyzed by the Luminex xMAP (Millipore, Darmstadt, Germany), a system that combines three basic forms of xMAP technology. 2.7. Folic Acid Homeostasis Serum, urine, liver, kidneys and heart FA levels were analyzed by ELISA using the “Folic Acid ELISA Kit” from CELL BIOLABS, INC. (MET-5068). This analysis was carried out with the reagents supplied by the manufacturer and according to their specifications in a 96-well plate. The Folic Acid ELISA Kit is a competitive enzyme immunoassay developed for rapid detection and quantification of FA. The FA standards and the samples are added to a FA conjugate preabsorbed microplate. After a brief incubation, an anti-folic acid polyclonal antibody is added, followed by an HRP conjugated secondary antibody. The amount of FA in the samples is quantified by comparing its absorbance at 450 nm with the FA standard curve. Regarding the preparation of samples, the tissues (liver, kidneys, heart) were homogenized (1:10 w/v) in ice-cold PBS using a Potter homogenizer (Pobel 245432, Madrid, Spain). The homogenates were incubated at 4 ◦ C for 20 min, and then Antioxidants 2022,11, 362 5 of 22 they were centrifugated at 12,000 × gfor 20 min. The urine samples were diluted (1:5) in Assay Diluent. 2.8. Activity of Antioxidant Enzymes and Oxidative Stress Markers in the Liver Liver tissue samples were homogenized (1:4 w/v) using a Potter homogenizer (Pobel 245432, Madrid, Spain) in a sucrose buffer (15 mM Tris/HCl, pH 7.4, 250 mM sucrose, 1 mM EDTA and 1 mM dithiothreitol) in an ice bath. Then, the homogenates were centrifuged at 3000 r.p.m for 20 min at 4 ◦ C and kept at − 80 ◦ C until the time of analysis. The activity of SOD (U/mg protein) was determined by the Fridovich method, which is based on the ability of SOD to inhibit the reduction of cytochrome c induced by the xanthine-xanthine oxidase system; it is measured by the absorbance increase at 550 nm for 3 min, due to the reduction of cytochrome c by adding xanthine oxidase [29]. Catalase (CAT) activity (U/mg protein) was determined using H 2 O 2 as substrate by the assay of Beers and Sizer, where the disappearance of H 2 O 2 was followed spectrophotometrically at 240 nm for 3 min [ 30 ]. The GPx activity (mU/mg) was determined using the method described by Lawrence and Burk [ 31 ]. In this assay, the oxidized glutathione (GSSG) formed by the action of GPx is coupled to the reaction that catalyzes the glutathione reductase (GR) enzyme, measuring the absorbance decrease at 340 nm for 3 min due to the oxidation of NADPH. The GR activity (mU/mg protein) was analyzed according to the spectrophotometric method described by Worthington and Rosemeye using GSH as substrate, where the absorbance decrease at 340 nm during 3 min due to the oxidation of NADPH was measured [ 32 ]. In order to measure the OS markers in the liver, the amount of lipid peroxidation was evaluated by the quantification of malondialdehyde (MDA) (mol/mg protein), the end-product of oxidative degradation of lipids, using a colorimetric reaction with thiobarbituric acid (TBA) at 535 nm as described by Draper and Hadley [ 33 ]. The protein oxidation was measured by the detection of carbonyl groups (CG) (nmol/mg protein) at 366 nm by the method described by Reznick and Packer, where the reaction of 2,4-dinitrophenylhydrazine (DNPH) with CG takes place [34]. 2.9. GSH Hepatic Levels To measure GSH hepatic deposits, liver homogenates were prepared in 50 mmol/L potassium phosphate buffer (pH 7.5) containing 1 mmol/L EDTA and 1 mg/mL bovine serum albumin with the Potter homogenizer (Pobel 245432, Madrid, Spain). Then, they were centrifuged at 3000 r.p.m for 20 min using the resulting supernatant fraction. Total GSH levels in the samples were determined by a colorimetric method using a commercial kit (Bioxytech GSH/GSSG-412, Oxis Research, Beverly Hills, CA, USA). 2.10. Nitric Oxide Hepatic Levels Livers were homogenized in ice-cold PBS (1:10) using a Potter homogenizer (Pobel 245432, Madrid, Spain). Homogenates were incubated at 4 ◦ C for 20 min and centrifuged for 20 min at 12,000 × g. Before the assay, the homogenates were filtered with 10,000 MWCO polysulfone filters (Vivaspin 500 SARTORIUS, Stonehouse, UK). The NO levels in the liver were analyzed by the Nitric Oxide Detection Kit (ADI-917-010) from Enzo Life Sciences Inc. (Barcelona, Spain). This kit quantitatively determines nitrate and nitrite concentrations using the enzyme nitrate reductase, which converts nitrate into nitrite, followed by the colorimetric detection at 540 nm of nitrite as a colored azo dye product from the Griess reaction. The NO levels in a system are measured by the determination of both nitrate and nitrite concentrations in the sample. 2.11. Immunoblotting Analysis in the Liver The livers of adolescent rats were homogenized (1:10 w/v) in 50 mM phosphate buffer (K 2 HPO 4 50 mM, KH 2 PO 4 50 mM, EDTA 0.01 mM, protease inhibitor 1:10 (Complete Protease Inhibitor Cocktail Tablets, ROCHE, Madrid, Spain) using the Potter homogenizer Antioxidants 2022,11, 362 6 of 22 (Pobel 245432, Madrid, Spain). Then, homogenates were centrifuged at 2000 r.p.m at 4 ◦ C for 10 min, and the final supernatant was aliquoted and frozen at −80 ◦C until analysis. The expression of the hepatic proteins NOX1, NOX4, eNOS, DHFR, nitrotyrosine, caspase-9, caspase-3 and β -actin (as load control) in the previously described homogenates was carried out using the protein immunodetection technique or Western blot. After determining the protein content of the homogenates, the appropriate volumes (100 µ g/lane) were mixed with an equal volume of sample buffer (Sample Buffer, Laemmli 2X Concentrate SIGMA) plus 5% of 2-mercaptoethanol (BIORAD, Madrid, Spain). Proteins were separated by SDS-polyacrylamide gel electrophoresis (PAGE) using acrylamide gels (6% for eNOS and 12% for the rest of proteins). Samples were heated at 95 ◦ C for 5 min before electrophoresis to denature proteins, except for dimer and monomer eNOS. In this case, in order to separate the dimer and the monomer of eNOS, samples were not heated. They did not contain 2-mercaptoethanol, and the temperature of the gel was maintained below 15 ◦ C during electrophoresis (low-temperature SDS-PAGE). The proteins were then transferred onto a nitrocellulose membrane using a blot system for 120 min. Membranes were blocked next for 1 h at room temperature with blocking buffer composed of TBS (10 mM Tris-HCl, 100 mM NaCl, pH 7.5) and 5% milk powder (BIORAD, Madrid, Spain ). For immunodetection of the proteins, membranes were incubated overnight at 4 ◦ C with specific primary antibodies (mouse monoclonal IgG, Santa Cruz Biotechnology, Heidelberg, Germany ) in a dilution of 1:1000. Monoclonal mouse anti β -actin (IgG1A5441, Sigma-Aldrich, Madrid, Spain ) was used to detect β -actin as a loading control with a dilution of 1:10000. The next day, after 54 min washes with TBST (TBS + 0.1% Tween 20, PanReac Applichem, Barcelona, Spain), probed membranes were incubated with the secondary antibody (Goat Anti-Mouse IgG (H + L)-HRP Conjugate, BIORAD, Madrid, Spain) in a dilution of 1:2500 for all proteins and 1:8000 for β -actin. The membranes were incubated for 1 min with the commercial developer solution Luminol ECL reagent (GE Healthcare and Lumigen Inc., Buckinghamshire, UK) and analyzed with the Amersham Imager 600 (GE Healthcare, Buckinghamshire, UK). Then, the quantification of the blots was performed by densitometry with the ImageJ program. The results were expressed as percent arbitrary relative units, referring to values in control animals which were defined as 100%. 2.12. Ratios of Measurements The hepatic ratios of GR/GPx, GR/GSH and cleaved caspase-3/caspase-3 were calculated. The results were expressed as percentages with regard to the values in control animals, defined as GR and cleaved caspase-3 100%. 2.13. Statistical Analysis The results are expressed as the mean ± standard error of the mean (SEM), where “n” is the number of samples used. The means shown were obtained from 12 data ( n= 12 ) for biochemical determinations or from 6 data (n= 6) for proteins expression. To compare the different variable objects of our study in the four experimental groups, the data were analyzed using the analysis of variance (ANOVA) in the program GraphPad InStat 3 (San Diego, CA, USA) . Subsequently, the Tukey–Kramer test was used to determine the significant differences between the means, considering statistically significant differences with values of p< 0.05. 3. Results 3.1. Nutritional and Hepatic Parameters Intermittent i.p. BD exposure to adolescent rats did not alter food intake, body weight gain, relative liver weight or hepatic protein content. However, transaminase levels, specifically of AST and ALT, were significantly increased (F(3, 20) = 7.7, p< 0.001; (F(3, 20) = 4.8 ,p< 0.05 for AST and ALT, respectively) (Table 1). FA supplementation did not affect any of those parameters, but in BD-exposed rats it decreased AST (F(3,20) = 3.5, p< 0.05) and especially ALT (F(3,20) = 5.1, p< 0.01) levels. Antioxidants 2022,11, 362 7 of 22 Table 1. Nutritional and hepatic parameters. Parameters C BD CF BDF Body weight increase (g/day) 5.72 ±0.17 5.29 ±0.14 5.80 ±0.21 5.20 ±0.53 Food intake (g/day) 14.29 ±1.31 13.84 ±1.09 14.35 ±1.85 13.95 ±1.04 LSI (%) 3.48 ±0.17 3.43 ±0.067 3.65 ±0.054 3.65 ±0.12 Liver protein content (mg/g liver wet tissue) 8.95 ±0.33 9.53 ±0.50 9.37 ±0.50 9.19 ±0.89 Serum AST (U/L) 118 ±13 210 ±8 *** a 112 ±13 168 ±13 + Serum ALT (U/L) 29.4 ±1.4 38.8 ±2.2 * aa 27.6 ±1.7 29.12 ±2.2 The results are expressed as mean ± SEM and analysed by a multifactorial one-way ANOVA followed by Tukey’s test. The number of animals in each group is 6 and n= 12. LSI: liver somatic index. AST: aspartate aminotransferase. ALT: alanine aminotransferase. SEM: standard error of the mean. Groups: C: control group, BD: binge drinking group, CF: control folic group, BDF: binge drinking folic group. Signification: C vs. BD: *p< 0.05 , *** p< 0.001; BD vs. BDF: a: p< 0.05, aa p< 0.01; BDF vs. CF: + p< 0.05. 3.2. Folic Acid Homeostasis With respect to FA homeostasis (Figure 1), BD exposition did not affect FA intake or serum FA levels, but it did lead to significantly low renal FA excretion (F(3.20) = 13.6, p< 0.001 ). This rise in FA renal reabsorption increased renal FA deposits (F(3.20) = 3.2, p< 0.05 ); however, hepatic FA levels were decreased (F(3.20) = 3.5, p< 0.05) and heart levels were unaffected. After FA supplementation, FA intake significantly increased in control and BD animals, together with higher levels in serum, the liver and, mainly, in the kidneys. 3.3. Oxidative Hepatic Balance In the liver, BD exposure significantly increased SOD and CAT activities ( F(3.20) = 3.2 and 3.9, p< 0.05 to SOD and CAT, respectively) and decreased GPx activity ( F(3.20) = 3.8 , p< 0.05) leading to protein ( F(3.20) = 4.3 ,p< 0.05) and lipid oxidation (F(3.20) = 10.29, p< 0.001 ). FA supplementation to BD animals reduced SOD activity (F(3.20) = 6.1, p< 0.01 ) and decreased lipid (F(3.20) = 5.2, p< 0.01) and protein oxidation (F(3.20) = 4.1, p< 0.05 ) (Figure 2). BD exposition also significantly decreased GSH hepatic levels (F(3.20) = 9.9, p< 0.001); however the ratio of GR/GPx was unaffected, and the GR/GSH ratio was significantly increased (F(3.20) = 12.5, p< 0.001). FA supplementation to BD rats significantly increased hepatic GSH levels (F(3.20) = 9.6, p< 0.001), leading to control levels of the ratios GR/GPx and GR/GSH (Figure 3). 3.4. Serum Homocysteine and NOX Hepatic Expression After BD consumption, serum Hcy levels significantly rise (F(3.20) = 6.4, p< 0.01), together with NOX1 and NOX4 liver expression (F(3.20) = 4.7 and 4.3, p< 0.05 to NOX1 and NOX4, respectively). FA supplementation to control animals did not affect any of these parameters; however, FA supplementation to BD rats decreased Hcy levels (F(3.20) = 10.52, p< 0.001) and NOX1 and NOX4 expression (F(3.20) 4.3, p< 0.05 to NOX1, F(3.20) = 9.65, p< 0.001 to NOX4) (Figure 4). 3.5. Hepatic eNOS Expression Figure 5analyses total eNOS (eNOSt), eNOS dimer or couple eNOS, which generates NO, and eNOS monomer or uncoupled eNOS with a pro-oxidative function. The expression of the enzyme DHFR, which increased BH4, promoting eNOS recoupling, was also measured in the liver. BD exposure increased eNOS in its total (F(3.20) = 6.0, p< 0.01), dimer (F(3.20) = 8.8, p< 0.001) and monomer (F(3.20) = 7.7, p< 0.001) forms, but decreased DHFR expression (F(3.20) = 7.8, p< 0.001). FA administration to control rats significantly increased eNOS dimer form (F(3.20) = 7.3, p< 0.001). FA supplemented to BD rats decreased eNOSt (F(3.20) = 4.2, p< 0.05), mainly by reducing eNOS monomer form (F(3.20) = 9.8, p< 0.001), since it increased DHFR expression (F(3.20) = 5.8, p< 0.01). Antioxidants 2022,11, 362 8 of 22 Antioxidants 2022, 11, x FOR PEER REVIEW 8 of 22 Figure 1. Folic acid homeostasis in the rats’ bodies. Intake levels (A), serum levels (B), urine levels (C) and tissue deposit levels in the liver (D), kidneys (E) and heart (F). The results are expressed as mean ± SEM and analysed by a multifactorial one-way ANOVA followed by Tukey’s test. The number of animals in each group is 6 and n = 12. SEM: standard error of the mean. Groups: C: control group, BD: binge drinking group, CF: control folic group, BDF: binge drinking folic group. Signification: C vs. BD: * p < 0.05, *** p < 0.001; C vs. CF: c p < 0.05, ccc p < 0.001; BD vs. BDF: aa p < 0.01, aaa p < 0.001. B D E A C F Figure 1. Folic acid homeostasis in the rats’ bodies. Intake levels ( A ), serum levels ( B ), urine levels ( C ) and tissue deposit levels in the liver ( D ), kidneys ( E ) and heart ( F ). The results are expressed as mean ± SEM and analysed by a multifactorial one-way ANOVA followed by Tukey’s test. The number of animals in each group is 6 and n= 12. SEM: standard error of the mean. Groups: C: control group, BD: binge drinking group, CF: control folic group, BDF: binge drinking folic group. Signification: C vs. BD: * p< 0.05, *** p< 0.001; C vs. CF: c p< 0.05, ccc p< 0.001; BD vs. BDF: aa p< 0.01 , aaa p< 0.001. Antioxidants 2022,11, 362 9 of 22 Antioxidants 2022, 11, x FOR PEER REVIEW 9 of 22 Figure 2. Hepatic oxidative balance: activity of antioxidant enzymes and oxidative stress markers. Superoxide dismutase (SOD) (A), catalase (CAT) (B), glutathione peroxidase (GPx) (C), glutathione reductase (GR) (D), lipid oxidation expressed by the levels of malondialdehyde (MDA) (E) and protein oxidation expressed by the levels of carbonyl groups (CG) (F). The results are expressed as mean ± SEM and analysed by a multifactorial one-way ANOVA followed by Tukey’s test. The number of animals in each group is 6 and n = 12. SEM: standard error of the mean. Groups: C: control group, BD: binge drinking group, CF: control folic group, BDF: binge drinking folic group. Signification: C vs. BD: * p < 0.05, *** p < 0.001; BD vs. BDF: a p < 0.05; aa p < 0.01. A B C D E F Figure 2. Hepatic oxidative balance: activity of antioxidant enzymes and oxidative stress markers. Superoxide dismutase (SOD) ( A ), catalase (CAT) ( B ), glutathione peroxidase (GPx) ( C ), glutathione reductase (GR) ( D ), lipid oxidation expressed by the levels of malondialdehyde (MDA) ( E ) and protein oxidation expressed by the levels of carbonyl groups (CG) ( F ). The results are expressed as mean ±SEM and analysed by a multifactorial one-way ANOVA followed by Tukey’s test. The number of animals in each group is 6 and n= 12. SEM: standard error of the mean. Groups: C: control group, BD: binge drinking group, CF: control folic group, BDF: binge drinking folic group. Signification: C vs. BD: * p< 0.05, *** p< 0.001; BD vs. BDF: a p< 0.05; aa p< 0.01. Antioxidants 2022,11, 362 16 of 22 proved that Chr-EtOH exposure induced NOX4 expression in the mitochondrial fraction from hepatic cells [ 55 ]. Therefore, inhibition of NOX4 activity protects against alcoholinduced fat accumulation and activation of intrinsic apoptosis via improving mitochondrial function. It is known that dysfunctional mitochondria release cytochrome C to the cytosol after alcohol exposure, leading to apoptosis activation. These authors confirm that this release was due in part to the ROS generated by NOX4. Once more, when FA is administered to BD rats, HHcy is avoided, and NOX1 and NOX4 hepatic expressions are decreased, contributing to reduced lipid and protein oxidation and apoptosis. This is probably due to FA contributing to restoring Hcy remethylation to methionine or to catabolizing it through the transsulfuration pathway, increasing GSH hepatic levels. It is also interesting to bear in mind that FA decreases NOX4 expression in particular. It could perhaps be acting directly on hepatic NOX4 expression, which is also backed by the results obtained by Ju et al. in colon cells, by inducing the mitochondrial MTHFD 2 enzyme [ 14 ]. It is also recognized that FA inhibits NOX activity in hepatic cells by reducing the mRNA levels of several essential NOX subunits [ 56 ]. Moreover, some authors concluded that FA could decrease NOX4 activity in endothelial cells by the induction of eNOS [ 16 ]. In fact, a new NOX4/DHFR/eNOS uncoupling pathway related to FA in the diet has recently been described in aortic cells [57]. 4.5. eNOS, NO and NS The eNOS enzyme is a dimer that depends on the cofactor BH4 to produce NO. When either cofactor bioavailability is limited or when OS appears, the eNOS dimer is destabilized and uncoupled, resulting in the production of O 2•− rather than NO. In this context, BD increases total hepatic eNOS expression, but either forms coupled or dimer (NO generator) and uncoupled or monomer (O 2•− generator). BH4 generation depends on the enzyme DHFR, which transforms dihydrobiopterin (BH2) to BH4 [ 19 ]. DHFR is decreased in the liver of BD adolescent rats. The enzyme DHFR depends on FA levels, connecting with the lower FA hepatic deposits found in these animals [ 58 ]. BH4 depletion is the main cause of the eNOS uncoupling process [ 59 ], even in adolescents exposed to high amounts of EtOH [ 60 ]. It is transformed to its inactive form BH2 in pro-oxidant ambients, such as that which is generated in the liver during BD exposure. Therefore, the hepatic OS generated by BD exposure could also be contributing to lower eNOS recoupling. Moreover, the eNOS form depends on the pair GSH/GSSG; when this ratio decreases, S-glutathionylation appears, and the enzyme changes into its uncoupled form [ 61 ]. Since BD decreases GSH deposits, the ratio is downregulated, and eNOS S-glutathionylation could be affecting eNOS function, leading to O2•− generation. The presence of NO and ROS in the liver of BD adolescent rats involves the formation of nitrosylating intermediates, mainly peroxinitrite (ONOO − ), which contributes to the formation of S-nitrosylated proteins (protein-SNOs), indicating that NS is also established [ 62 , 63 ]. In the liver, the balance between coupled and uncoupled eNOS is very important for NO homeostasis and ROS production in hepatic sinusoidal endothelial cells. Its imbalance has been observed in cirrhotic livers, being implicated in inflammation and fibrosis [ 64 ]. Therefore, this is another piece of biomolecular data that implies that BD exposure during adolescence has an important impact on liver function. FA therapy by restoring DHFR protein hepatic expression leads to recoupled eNOS expression in BD rats. It increases NO production and decreases protein-SNOs in the liver, avoiding NS. Different mechanisms are related to FA and eNOS activity in endothelial cells, including greater bioavailability of BH4 through stabilization of BH4 and/or recycling from BH2, mainly by upregulating the activity of DHFR in the biopterin recycling pathway, direct interaction with eNOS, and directly decreasing ROS generation [ 65 ]. Moreover, since FA increases GSH synthesis, eNOS S-glutathionylation could be avoided, contributing to stabilizing its coupled form. Antioxidants 2022,11, 362 17 of 22 4.6. Apoptosis Caspases are a family of protease enzymes essential in programmed cell death via apoptosis. Caspase-9 is an initiator caspase related to the intrinsic apoptotic pathway, which appears when mitochondrial cytochrome c is released into the cytosol after cell stress. This cytochrome c recruits caspase-9, then caspase-9 activates other executioner caspases, such as caspase-3. Finally, caspase-3 leads to the degradation of cellular components [ 66 ]. BDexposed rats present with significantly high caspase-9 expression in the liver, which implies that mitochondria are under a stressor condition. This is in agreement with the NOX4 upregulation found in the liver, which generates ROS specifically in this organelle [ 55 ]. The ratio of cleaved caspase-3/caspase-3 is also significantly increased in these animals, confirming that apoptosis is established. Moreover, TIMP-1 serum levels are reduced in BD rats. Apart from modulating extracellular matrix remodeling by controlling the activity of matrix metalloproteinases (MMPs), this protein also acts by signaling molecules with cytokine-like activity, having anti-apoptotic functions [ 67 ]. It is clear that i.p. BD exposure during adolescence leads to important biomolecular changes in the liver by generating OS and NS. It mainly affects NOX4 expression and specifically damages mitochondrial function, which contributes to increased intrinsic apoptotic pathways. After FA supplementation, caspase-9 expression is decreased according to the lower levels of NOX4 found in the liver of these animals. This indicates that FA protects liver mitochondria from EtOH damage. Moreover, the ratio of cleaved caspase-3/caspase-3 is also reduced, indicating that apoptosis is decreased when FA hepatic levels are balanced, and that this vitamin plays an important role during BD liver damage. FA is known for its anti-apoptotic properties in different types of cells [13,68–71]. These authors attribute this action to different mechanisms: through decreasing NOX activity-ROS generationmitochondrial complex II; decreasing OS-, preventing telomeric DNA oxidation and attrition; regulating the expression of apoptosis-related genes (decreased caspase-3 and upregulated BCL2/BAX ratio); modulating microRNA-34a, associated with Bcl-2 signaling; or decreasing caspase-9 activity. In this context, FA supplementation also increases TIMP-1 serum levels. Among the mechanisms attributed to TIMP-1 in apoptosis is its relationship to the anti-apoptotic protein of the Bcl-family, Bcl-2. This protein increases TIMP-1 expression, and TIMP-1 also stabilizes Bcl-2/BAX, resulting in an upregulation of Bcl-2 and a decrease of the pro-apoptotic Bcl-2-associated X (Bax) protein, avoiding cytochrome-c release by mitochondria [ 67 ]. This correlation between FA supplementation and TIMP-1 increase in BD-exposed rats reinforces the idea that FA plays an important role in avoiding mitochondrial damage and apoptosis. 5. Conclusions The present study described for the first time that BD exposure during adolescence in male rats leads to a depletion of FA hepatic deposits together with a low GSH synthesis and higher Hcy serum levels. This implies that, apart from its oxidative damage by generating ROS and affecting the antioxidant enzyme system, the methionine cycle and transsulfuration pathway are affected after BD exposure, in part by the depletion of FA, increasing hepatic damage. It has also been found that BD leads to NS by increasing eNOs in its uncoupled form, either by its direct pro-oxidant action or by decreasing the GSH/GSSG ratio. Finally, this OS leads to apoptosis mainly by increasing NOX4 activity and mitochondrial damage (Figure 8A). When FA depletion is avoided using a supplemented FA diet, most of these situations are avoided, showing that this vitamin plays a pivotal role in BD hepatic damage. Collectively, we add to the literature new data to strengthen the notion that FA is a vitamin that confers a novel role to combat ROS/RNS insults after BD exposure in adolescents since FA decreases lipid and protein hepatic oxidation, NS and apoptosis by different simultaneous mechanisms. Some of these mechanisms depend on its own antioxidant properties, such as the decrease of NOX4 expression, and others are related to its action on the methionine cycle and transsulfuration pathway, which contributes to increased hepatic GSH (antioxidant) levels and decreased Antioxidants 2022,11, 362 18 of 22 serum Hcy (pro-oxidant) levels. Furthermore, FA contributes to recoupling the eNOS enzyme by restoring DHFR protein abundance and reducing ROS production, avoiding NS. It is important to remark that in vivo treatment with FA completely attenuates NOX4 expression after BD exposure, decreasing mitochondrial damage and apoptosis (Figure 8B). Therefore, FA seems to be a suitable supplement to avoid hepatic damage after BD exposure in adolescents. Antioxidants 2022, 11, x FOR PEER REVIEW 18 of 22 5. Conclusions The present study described for the first time that BD exposure during adolescence in male rats leads to a depletion of FA hepatic deposits together with a low GSH synthesis and higher Hcy serum levels. This implies that, apart from its oxidative damage by generating ROS and affecting the antioxidant enzyme system, the methionine cycle and transsulfuration pathway are affected after BD exposure, in part by the depletion of FA, increasing hepatic damage. It has also been found that BD leads to NS by increasing eNOs in its uncoupled form, either by its direct pro-oxidant action or by decreasing the GSH/GSSG ratio. Finally, this OS leads to apoptosis mainly by increasing NOX4 activity and mitochondrial damage (Figure 8A). When FA depletion is avoided using a supplemented FA diet, most of these situations are avoided, showing that this vitamin plays a pivotal role in BD hepatic damage. Collectively, we add to the literature new data to strengthen the notion that FA is a vitamin that confers a novel role to combat ROS/RNS insults after BD exposure in adolescents since FA decreases lipid and protein hepatic oxidation, NS and apoptosis by different simultaneous mechanisms. Some of these mechanisms depend on its own antioxidant properties, such as the decrease of NOX4 expression, and others are related to its action on the methionine cycle and transsulfuration pathway, which contributes to increased hepatic GSH (antioxidant) levels and decreased serum Hcy (pro-oxidant) levels. Furthermore, FA contributes to recoupling the eNOS enzyme by restoring DHFR protein abundance and reducing ROS production, avoiding NS. It is important to remark that in vivo treatment with FA completely attenuates NOX4 expression after BD exposure, decreasing mitochondrial damage and apoptosis (Figure 8B). Therefore, FA seems to be a suitable supplement to avoid hepatic damage after BD exposure in adolescents. (A) EtOH (BD) HEPATOCYTE ↓γ-GCS ADH ↑↑ CYP2E1 Acetaldehyde Acetate ALDH Mitochondria KC Ox-Ph Acetyl CoA ATP H2O ↑↑ ROS ↑↑ ROS ↑NOX 4 ↑NOX 1 NADH/NAD+ ↑ Cytochrome C ↑Caspase-9 ↑Caspase-3 Apoptosis ↑Homocysteine SAH SAM Methionine Methionine metabolism 5-MTHF 5,10-MTHF THF Folate metabolism ↓DHFR Cystathionine Cysteine ↓GSH GSSG GR ↓GPx ↑SOD H2O + O2 H2O2 ↑CAT O2•- BH2 BH4 ↑eNOS uncoupled ↑eNOS coupled NO O2•- ROS ONOO- ↑↑ RNS CAT Alcohol metabolism ↑Lipid and protein oxidation EtOH (BD) ↓Folic acid ↑Protein nitration ↑Lipid and protein oxidation Transsulfuration pathway Antioxidants 2022, 11, x FOR PEER REVIEW 19 of 22 (B) Figure 8. (A) BD in adolescence increases the hepatic production of ROS and RNS, causing oxidative and nitrosative stress leading to apoptosis. BD stimulates the pro-oxidant enzymes CYP2E1 and NOX1, increasing cytosolic ROS generation, and NOX4 activity, increasing ROS mitochondrial generation and cytochrome C expulsion, contributing to apoptosis. BD affects the antioxidant enzymatic defense system, increasing SOD and CAT activities and decreasing GPx activity, but lipid and protein oxidation appears. BD affects the methionine cycle and the transsulfuration pathway, leading to depletion in GSH levels and HHcy. BD reduces the expression of DHFR, decreasing the cofactor BH4 and increasing the uncoupled form of eNOS, producing RNS and protein nitration. BD also leads to a hepatic depletion of FA, which could be implicated in all these actions. (B) FA antioxidant pathways against hepatic oxidation induced by BD in adolescence. FA supplementation to BD rats manages to reduce the oxidation of lipids and proteins, as well as protein nitration and apoptosis. It decreases the expression of the pro-oxidant enzymes NOX1 and NOX4, avoiding ROS generation and apoptosis. FA restores the balance of the antioxidant enzymatic defense system. Due to its participation in the methionine cycle and the transsulfuration pathway, it increases the levels of the antioxidant GSH and decreases the pro-oxidant Hcy. FA also increases the expression of DHFR and BH4 viability, increasing the stability of coupled eNOS, avoiding RNS generation and nitrosative damage. Author Contributions: Conceptualization, M.L.O. and M.d.C.G.-L.; methodology, F.N. and M.d.C.G.-L.; validation, M.L.O., F.N. and O.C.; formal analysis, F.N., M.L.O. and M.d.C.G.-L.; investigation, I.R.-H., F.N. and M.d.C.G.-L.; resources, O.C.; data curation, M.L.O.; writing—original draft preparation, M.d.C.G.-L. and M.L.O.; writing—review and editing, M.d.C.G.-L., I.R.-H. and M.L.O.; supervision, M.L.O., F.N. and O.C.; management of references, M.d.C.G.-L.; figure design, M.d.C.G.-L. and M.L.O.; funding acquisition, O.C. All authors have read and agreed to the published version of the manuscript. Funding: This research was funded by the Andalusian Regional Government for its support to the CTS-193 research group. Institutional Review Board Statement: Animal care procedures and experimental protocols were conducted according to the EU regulations (Council Directive 86/609/EEC, 24 November 1986), and they were approved by the Ethics Committee of the University of Seville. Informed Consent Statement: Not applicable. Data Availability Statement: Data is contained within this article. EtOH (BD) HEPATOCYTE ADH ↑↑ CYP2E1 Acetaldehyde Acetate ALDH Mitochondria KC Ox-Ph Acetyl CoA ATP H2O ↓ ROS ROS ↓NOX 4 ↓NOX 1 NADH/NAD+NADH/NAD+ Cytochrome C ↓Caspase-9 ↓Caspase-3 Apoptosis ↓Homocysteine SAH SAM Methionine Methionine metabolism 5-MTHF 5,10-MTHF THF Folate metabolism ↑DHFR ↑GSH GSSG GR GPx ↓SOD H2O + O2 H2O2 CAT O2•- BH2 BH4 ↓eNOS uncoupled eNOS coupled NO O2•- ↓ROS ↓ RNS CAT Alcohol metabolism ↓Lipid and protein oxidation EtOH (BD) ↑Folic acid ↓Protein nitration Transsulfuration pathway ↓Lipid and protein oxidation FA FA FA FA FA FA FA γ-GCS Cystathionine Cysteine ONOOFigure 8. ( A ) BD in adolescence increases the hepatic production of ROS and RNS, causing oxidative and nitrosative stress leading to apoptosis. BD stimulates the pro-oxidant enzymes CYP2E1 and NOX1 , Antioxidants 2022,11, 362 19 of 22 increasing cytosolic ROS generation, and NOX4 activity, increasing ROS mitochondrial generation and cytochrome C expulsion, contributing to apoptosis. BD affects the antioxidant enzymatic defense system, increasing SOD and CAT activities and decreasing GPx activity, but lipid and protein oxidation appears. BD affects the methionine cycle and the transsulfuration pathway, leading to depletion in GSH levels and HHcy. BD reduces the expression of DHFR, decreasing the cofactor BH4 and increasing the uncoupled form of eNOS, producing RNS and protein nitration. BD also leads to a hepatic depletion of FA, which could be implicated in all these actions. ( B ) FA antioxidant pathways against hepatic oxidation induced by BD in adolescence. FA supplementation to BD rats manages to reduce the oxidation of lipids and proteins, as well as protein nitration and apoptosis. It decreases the expression of the pro-oxidant enzymes NOX1 and NOX4, avoiding ROS generation and apoptosis. FA restores the balance of the antioxidant enzymatic defense system. Due to its participation in the methionine cycle and the transsulfuration pathway, it increases the levels of the antioxidant GSH and decreases the pro-oxidant Hcy. FA also increases the expression of DHFR and BH4 viability, increasing the stability of coupled eNOS, avoiding RNS generation and nitrosative damage. Author Contributions: Conceptualization, M.L.O. and M.d.C.G.-L.; methodology, F.N. and M.d.C.G.-L. ; validation, M.L.O., F.N. and O.C.; formal analysis, F.N., M.L.O. and M.d.C.G.-L.; investigation, I.R.-H., F.N. and M.d.C.G.-L.; resources, O.C.; data curation, M.L.O.; writing—original draft preparation, M.d.C.G.-L. and M.L.O.; writing—review and editing, M.d.C.G.-L., I.R.-H. and M.L.O.; supervision, M.L.O., F.N. and O.C.; management of references, M.d.C.G.-L.; figure design, M.d.C.G.-L. and M.L.O.; funding acquisition, O.C. All authors have read and agreed to the published version of the manuscript. Funding: This research was funded by the Andalusian Regional Government for its support to the CTS-193 research group. Institutional Review Board Statement: Animal care procedures and experimental protocols were conducted according to the EU regulations (Council Directive 86/609/EEC, 24 November 1986), and they were approved by the Ethics Committee of the University of Seville. Informed Consent Statement: Not applicable. Data Availability Statement: Data is contained within this article. Acknowledgments: To The Center for Research, Technology and Innovation (CITIUS) of the University of Seville for their technical support. Conflicts of Interest: The authors declare no conflict of interest. References 1. Medici, V.; Halsted, C.H. Folate, Alcohol, and Liver Disease. Mol. Nutr. Food Res. 2013,57, 596–606. [CrossRef] [PubMed] 2. Halsted, C.H. Alcohol and Folate Interactions: Clinical Implications. In Folate in Health and Disease; Marcel Dekker: New York, NY, USA, 1995; pp. 313–328. 3. Hernández-Rodríguez, S.; Gutiérrez-Salinas, J.; García-Ortíz, L.; Mondragón-Terán, P.; Ramírez-García, S.; Núñez-Ramos, N.R. Estrésoxidativo y Nitrosativo Como Mecanismo de Daño al Hepatocito Producido Por El Metabolismo Del Etanol. Med. Interna Mex. 2014,30, 295–308. 4. Warzyszynska, J.E.; Kim, Y.-I.J. Folate in Human Health and Disease. In eLS; John Wiley & Sons: Hoboken, NJ, USA, 2014; ISBN 9780470015902. 5. Sharma, J.; Krupenko, S.A. Folate Pathways Mediating the Effects of Ethanol in Tumorigenesis. Chem.-Biol. Interact. 2020 ,324, 109091. [CrossRef] [PubMed] 6. Asbaghi, O.; Ghanavati, M.; Ashtary-larky, D.; Bagheri, R.; Rezaei Kelishadi, M.; Nazarian, B.; Nordvall, M.; Wong, A.; Dutheil, F.; Suzuki, K.; et al. Effects of Folic Acid Supplementation on Oxidative Stress Markers: A Systematic Review and Meta-analysis of Randomized Controlled Trials. Antioxidants 2021,10, 871. [CrossRef] 7. Lai, K.G.; Chen, C.F.; Ho, C.T.; Liu, J.J.; Liu, T.Z.; Chern, C.L. Novel Roles of Folic Acid as Redox Regulator: Modulation of Reactive Oxygen Species Sinker Protein Expression and Maintenance of Mitochondrial Redox Homeostasis on Hepatocellular Carcinoma. Tumor Biol. 2017,39, 1010428317702649. [CrossRef] 8. Joshi, R.; Adhikari, S.; Patro, B.S.; Chattopadhyay, S.; Mukherjee, T. Free Radical Scavenging Behavior of Folic Acid: Evidence for Possible Antioxidant Activity. Free. Radic. Biol. Med. 2001,30, 1390–1399. [CrossRef] 9. Stanger, O.; Wonisch, W. Enzymatic and Non-Enzymatic Antioxidative Effects of Folic Acid and Its Reduced Derivates. In Water Soluble Vitamins, Subcellular Biochemistry; O. Stanger: London, UK, 2012; Volume 56, pp. 131–161. ISBN 9789400721999. Antioxidants 2022,11, 362 20 of 22 10. Cano, M.J.; Ayala, A.; Murillo, M.L.; Carreras, O. Protective Effect of Folic Acid against Oxidative Stress Produced in 21-Day Postpartum Rats by Maternal-Ethanol Chronic Consumption during Pregnancy and Lactation Period. Free. Radic. Res. 2001 ,34, 1–8. [CrossRef] 11. Woo, C.W.H.; Prathapasinghe, G.A.; Siow, Y.L.; Karmin, O. Hyperhomocysteinemia Induces Liver Injury in Rat: Protective Effect of Folic Acid Supplementation. Biochim. Biophys. Acta 2006,1762, 656–665. [CrossRef] 12. Hwang, S.Y.; Siow, Y.L.; Au-Yeung, K.K.W.; House, J.; Karmin, O. Folic Acid Supplementation Inhibits NADPH Oxidase-Mediated Superoxide Anion Production in the Kidney. Am. J. Physiol. Ren. Physiol. 2011,300, 189–198. [CrossRef] 13. Hsu, H.C.; Chang, W.M.; Wu, J.Y.; Huang, C.C.; Lu, F.J.; Chuang, Y.W.; Chang, P.J.; Chen, K.H.; Hong, C.Z.; Yeh, R.H.; et al. Folate Deficiency Triggered Apoptosis of Synoviocytes: Role of Overproduction of Reactive Oxygen Species Generated via NADPH Oxidase/Mitochondrial Complex II and Calcium Perturbation. PLoS ONE 2016,11, e0146440. [CrossRef] 14. Ju, H.Q.; Lu, Y.X.; Chen, D.L.; Zuo, Z.X.; Liu, Z.X.; Wu, Q.N.; Mo, H.Y.; Wang, Z.X.; Wang, D.S.; Pu, H.Y.; et al. Modulation of Redox Homeostasis by Inhibition of MTHFD2 in Colorectal Cancer: Mechanisms and Therapeutic Implications. J. Natl. Cancer Inst. 2019,111, 584–596. [CrossRef] [PubMed] 15. Gaucher, C.; Boudier, A.; Bonetti, J.; Clarot, I.; Leroy, P.; Parent, M. Glutathione: Antioxidant Properties Dedicated to Nanotechnologies. Antioxidants 2018,7, 62. [CrossRef] [PubMed] 16. Cheng, F.; Lan, J.; Xia, W.; Tu, C.; Chen, B.; Li, S.; Pan, W. Folic Acid Attenuates Vascular Endothelial Cell Injury Caused by Hypoxia via the Inhibition of ERK1/2/NOX4/ROS Pathway. Cell Biochem. Biophys. 2016,74, 205–211. [CrossRef] [PubMed] 17. Gao, L.; Chalupsky, K.; Stefani, E.; Cai, H. Mechanistic Insights into Folic Acid-Dependent Vascular Protection: Dihydrofolate Reductase-Mediated Reduction in Oxidant Stress in Endothelial Cells and Angiotensin II-Infused Mice: A Novel HPLC-Based Fluorescent Assay for DHFR Activity. J. Mol. Cell. Cardiol. 2009,47, 752–760. [CrossRef] 18. Chalupsky, K.; Kraˇcun, D.; Kanchev, I.; Bertram, K.; Görlach, A. Folic Acid Promotes Recycling of Tetrahydrobiopterin and Protects Against Hypoxia-Induced Pulmonary Hypertension by Recoupling Endothelial Nitric Oxide Synthase. Antioxid. Redox Signal. 2015,23, 1076–1091. [CrossRef] 19. Schneider, M.P.; Schlaich, M.P.; Harazny, J.M.; Raff, U.; Ritt, M.; Ott, C.; Schmieder, R.E. Folic Acid Treatment Normalizes Nos-Dependence of Vascular Tone in the Metabolic Syndrome. Obesity 2011,19, 960–967. [CrossRef] 20. National Institute on Alcohol Abuse and Alcoholism What Is Binge Drinking? Available online: https://www.niaaa.nih.gov/ publications/brochures-and-fact-sheets/binge-drinking (accessed on 23 March 2021). 21. Bonar, E.E.; Parks, M.J.; Gunlicks-Stoessel, M.; Lyden, G.R.; Mehus, C.J.; Morrell, N.; Patrick, M.E. Binge Drinking before and after a COVID-19 Campus Closure among First-Year College Students. Addict. Behav. 2021,118, 106879. [CrossRef] 22. Muzi, S.; Sansò, A.; Serena Pace, C. What’s Happened to Italian Adolescents During the COVID-19 Pandemic? A Preliminary Study on Symptoms, Problematic Social Media Usage, and Attachment: Relationships and Differences with Pre-Pandemic Peers. Front. Psychiatry 2021,12, 590543. [CrossRef] 23. Lu, Y.; Cederbaum, A.I. CYP2E1 and Oxidative Liver Injury by Alcohol. Free. Radic. Biol. Med. 2008,44, 723–738. [CrossRef] 24. Eiciner, E.R.; Haliman, R.S. Effect of Alcohol on Serum Folate Level. J. Clin. Investig. 1973,52, 584–591. [CrossRef] 25. Ojeda, M.L.; Rua, R.M.; Nogales, F.; Díaz-Castro, J.; Murillo, M.L.; Carreras, O. The Benefits of Administering Folic Acid in Order to Combat the Oxidative Damage Caused by Binge Drinking in Adolescent Rats. Alcohol Alcohol. 2016 ,51, 235–241. [CrossRef] [PubMed] 26. Duncan, T.M.; Reed, M.C.; Frederik Nijhou, H. A Population Model of Folate-Mediated One-Carbon Metabolism. Nutrients 2013 , 5, 2457–2474. [CrossRef] [PubMed] 27. Spear, L. Modeling Adolescent Development and Alcohol Use in Animals. Alcohol. Res. Health 2000,24, 115. 28. Callaci, J.J.; Himes, R.; Lauing, K.; Roper, P. Long-Term Modulations in the Vertebral Transcriptome of Adolescent-Stage Rats Exposed to Binge Alcohol. Alcohol Alcohol. 2010,45, 332–346. [CrossRef] 29. Fridovich, I. Cytochrome c. In CRC Handbook of Methods for Oxygen Radical Research; CRC Press: Boca Raton, FL, USA, 1985; pp. 213–215. 30. Beers, R.F.; Sizer, I.W. A Spectrophotometric Method for Measuring the Breakdown of Hydrogen Peroxide by Catalase. J. Biol. Chem. 1952,195, 133–140. [CrossRef] 31. Lawrence, R.A.; Burk, R.F. Glutathione Peroxidase Activity in Selenium-Deficient Rat Liver. Biochem. Biophys. Res. Commun. 1976 , 71, 952–958. [CrossRef] 32. Worthington, D.J.; Rosemeyer, M.A. Human Glutathione Reductase: Purification of the Crystalline Enzyme from Erythrocytes. Eur. J. Biochem. 1974,48, 167–177. [CrossRef] 33. Draper, H.H.; Hadley, M. Malondialdehyde Determination as Index of Lipid Peroxidation. Methods Enzymol. 1990 ,186, 421–431. [CrossRef] 34. Reznick, A.Z.; Packer, L. Oxidative Damage to Proteins: Spectrophotometric Method for Carbonyl Assay. Methods Enzymol. 1994 , 233, 357–363. [CrossRef] 35. Mathews, S.; Xu, M.; Wang, H.; Bertola, A.; Gao, B. Animals Models of Gastrointestinal and Liver Diseases. Animal Models of Alcohol-Induced Liver Disease: Pathophysiology, Translational Relevance, and Challenges. Am. J. Physiol.-Gastrointest. Liver Physiol. 2014,306, 819–823. [CrossRef] 36. Binder, C.; Knibbe, K.; Kreissl, A.; Repa, A.; Thanhaeuser, M.; Greber-Platzer, S.; Berger, A.; Jilma, B.; Haiden, N. Does Acute Alcohol Intoxication Cause Transaminase Elevations in Children and Adolescents? Alcohol 2016 ,51, 57–62. [CrossRef] [PubMed] Antioxidants 2022,11, 362 21 of 22 37. Pereira-Fernandes, L.M.; Lopes, K.S.; Santana, L.N.S.; Fontes-Júnior, E.A.; Ribeiro, C.H.M.A.; Silva, M.C.F.; de Oliveira Paraense, R.S.; Crespo-López, M.E.; Gomes, A.R.Q.; Lima, R.R.; et al. Repeated Cycles of Binge-like Ethanol Intake in Adolescent Female Rats Induce Motor Function Impairment and Oxidative Damage in Motor Cortex and Liver, but Not in Blood. Oxid. Med. Cell. Longev. 2018,2018, 3467531. [CrossRef] 38. Ojeda, M.L.; Rua, R.M.; Murillo, M.L.; Carreras, O.; Nogales, F. Binge Drinking during Adolescence Disrupts Se Homeostasis and Its Main Hepatic Selenoprotein Expression. Alcohol. Clin. Exp. Res. 2015,39, 818–826. [CrossRef] [PubMed] 39. Nyblom, H.; Berggren, U.; Balldin, J.; Olsson, R. High AST/ALT Ratio May Indicate Advanced Alcoholic Liver Disease Rather than Heavy Drinking. Alcohol Alcohol. 2004,39, 336–339. [CrossRef] [PubMed] 40. Ojeda, M.L.; Carreras, O.; Sobrino, P.; Murillo, M.L.; Nogales, F. Biological Implications of Selenium in Adolescent Rats Exposed to Binge Drinking: Oxidative, Immunologic and Apoptotic Balance. Toxicol. Appl. Pharmacol. 2017,329, 165–172. [CrossRef] 41. Shaw, S.; Jayatilleke, E.; Herbert, V.; Colman, N. Cleavage of Folates during Ethanol Metabolism. Role of Acetaldehyde/Xanthine Oxidase-Generated Superoxide. Biochem. J. 1989,257, 277–280. [CrossRef] 42. Stallons, L.J.; Whitaker, R.M.; Schnellmann, R.G. Suppressed Mitochondrial Biogenesis in Folic Acid-Induced Acute Kidney Injury and Early Fibrosis. Toxicol. Lett. 2014,224, 326–332. [CrossRef] 43. Nogales, F.; Rua, R.M.; Ojeda, M.L.; Murillo, M.L.; Carreras, O. Oral or Intraperitoneal Binge Drinking and Oxidative Balance in Adolescent Rats. Chem. Res. Toxicol. 2014,27, 1926–1933. [CrossRef] 44. Vatsalya, V.; Gala, K.S.; Hassan, A.Z.; Frimodig, J.; Kong, M.; Sinha, N.; Schwandt, M.L. Characterization of Early-Stage Alcoholic Liver Disease with Hyperhomocysteinemia and Gut Dysfunction and Associated Immune Response in Alcohol Use Disorder Patients. Biomedicines 2021,9, 7. [CrossRef] 45. Murray, T.V.A.; Dong, X.; Sawyer, G.J.; Caldwell, A.; Halket, J.; Sherwood, R.; Quaglia, A.; Dew, T.; Anilkumar, N.; Burr, S.; et al. NADPH Oxidase 4 Regulates Homocysteine Metabolism and Protects against Acetaminophen-Induced Liver Damage in Mice. Free. Radic. Biol. Med. 2015,89, 918–930. [CrossRef] 46. Škovierová, H.; Vidomanová, E.; Mahmood, S.; Sopková, J.; Drgová, A.; ˇ Cerveˇnová, T.; Halašová, E.; Lehotský, J. The Molecular and Cellular Effect of Homocysteine Metabolism Imbalance on Human Health. Int. J. Mol. Sci. 2016 ,17, 1733. [CrossRef] [PubMed] 47. Avila, M.A.; Berasain, C.; Prieto, J.; Mato, J.M.; García-Trevijano, E.R.; Corrales, F.J. Influence of Impaired Liver Methionine Metabolism on the Development of Vascular Disease and Inflammation. Current medicinal chemistry. Cardiovasc. Hematol. Agents 2005,3, 267–281. [CrossRef] [PubMed] 48. Kharbanda, K.K. Alcoholic Liver Disease and Methionine Metabolism. Semin. Liver Dis. 2009 ,29, 155–165. [CrossRef] [PubMed] 49. Perna, A.F.; Ingrosso, D.; Lombardi, C.; Acanfora, F.; Satta, E.; Cesare, C.M.; Violetti, E.; Romano, M.M.; de Santo, N.G. Possible Mechanisms of Homocysteine Toxicity. Kidney Int. 2003,63, S137–S140. [CrossRef] 50. Karolczak, K.; Watala, C. Melatonin as a Reducer of Neuroand Vasculotoxic Oxidative Stress Induced by Homocysteine. Antioxidants 2021,10, 1178. [CrossRef] 51. Dumitrescu, R.G. Alcohol-Induced Epigenetic Changes in Cancer. Methods Mol. Biol. 2018,1856, 157–172. [CrossRef] 52. Lan, T.; Kisseleva, T.; Brenner, D.A. Deficiency of NOX1 or NOX4 Prevents Liver Inflammation and Fibrosis in Mice through Inhibition of Hepatic Stellate Cell Activation. PLoS ONE 2015,10, e0129743. [CrossRef] 53. Ishida, K.; Kaji, K.; Sato, S.; Ogawa, H.; Takagi, H.; Takaya, H.; Kawaratani, H.; Moriya, K.; Namisaki, T.; Akahane, T.; et al. Sulforaphane Ameliorates Ethanol plus Carbon Tetrachloride-Induced Liver Fibrosis in Mice through the Nrf2-Mediated Antioxidant Response and Acetaldehyde Metabolization with Inhibition of the LPS/TLR4 Signaling Pathway. J. Nutr. Biochem. 2021,89, 108573. [CrossRef] 54. Andueza, A.; Garde, N.; García-Garzón, A.; Ansorena, E.; López-Zabalza, M.J.; Iraburu, M.J.; Zalba, G.; Martínez-Irujo, J.J. NADPH Oxidase 5 Promotes Proliferation and Fibrosis in Human Hepatic Stellate Cells. Free. Radic. Biol. Med. 2018 ,126, 15–26. [CrossRef] 55. Sun, Q.; Zhang, W.; Zhong, W.; Sun, X.; Zhou, Z. Pharmacological Inhibition of NOX4 Ameliorates Alcohol-Induced Liver Injury in Mice through Improving Oxidative Stress and Mitochondrial Function. Biochim. Biophys. Acta-Gen. Subj. 2017 ,1861, 2912–2921. [CrossRef] 56. Sarna, L.K.; Wu, N.; Wang, P.; Hwang, S.Y.; Siow, Y.L.; Karmin, O. Folic Acid Supplementation Attenuates High Fat Diet Induced Hepatic Oxidative Stress via Regulation of NADPH Oxidase. Can. J. Physiol. Pharmacol. 2012 ,90, 155–165. [CrossRef] [PubMed] 57. Huang, K.; Wang, Y.; Siu, K.L.; Zhang, Y.; Cai, H. Targeting Feed-Forward Signaling of TGF β /NOX4/DHFR/ENOS Uncoupling/TGF β Axis with Anti-TGF β and Folic Acid Attenuates Formation of Aortic Aneurysms: Novel Mechanisms and Therapeutics. Redox Biol. 2021,38, 101757. [CrossRef] [PubMed] 58. Gao, L.; Siu, K.L.; Chalupsky, K.; Nguyen, A.; Chen, P.; Weintraub, N.L.; Galis, Z.; Cai, H. Role of Uncoupled Endothelial Nitric Oxide Synthase in Abdominal Aortic Aneurysm Formation: Treatment with Folic Acid. Hypertension 2012 ,59, 158–166. [CrossRef] [PubMed] 59. Aircart-Ramos, C. Papel Del Módulo Carboxi-Terminal De Las Óxido Nítrico Sintasas En La Regulación De La Síntesis De No: Caracterización Del Receptor Nuclear E75; Un Sensor De Óxido Nítrico: Madrid, Spain, 2013. 60. Hwang, C.-L.; Bian, J.-T.; Thur, L.A.; Peters, T.A.; Piano, M.R.; Phillips, S.A. Tetrahydrobiopterin Restores Microvascular Dysfunction in Young Adult Binge Drinkers. Alcohol. Clin. Exp. Res. 2020,44, 407–414. [CrossRef] Antioxidants 2022,11, 362 22 of 22 61. Fleming, I. NO Signaling Defects in Hypertension. In Nitric Oxide: Biology and Pathobiology; Elsevier Inc.: Amsterdam, The Netherlands , 2017; pp. 301–311. ISBN 9780128042731. 62. Htet Hlaing, K.; Clément, M.v. Formation of Protein S-Nitrosylation by Reactive Oxygen Species. Free Radic. Res. 2014 ,48, 996–1010. [CrossRef] 63. Pignatelli, P.; Menichelli, D.; Pastori, D.; Violi, F. Oxidative Stress and Cardiovascular Disease: New Insights. Kardiol. Pol. 2018 , 76, 713–722. [CrossRef] 64. Tai, Y.; Zhao, C.; Zhang, L.; Tang, S.; Jia, X.; Tong, H.; Liu, R.; Tang, C.; Gao, J. Celecoxib Reduces Hepatic Vascular Resistance in Portal Hypertension by Amelioration of Endothelial Oxidative Stress. J. Cell. Mol. Med. 2021,25, 10389–10402. [CrossRef] 65. Stanhewicz, A.E.; Kenney, W.L. Role of Folic Acid in Nitric Oxide Bioavailability and Vascular Endothelial Function. Nutr. Rev. 2017,75, 61–70. [CrossRef] 66. Ting-Jun, F.; Li-Hui, H.; Ri-Shan, C.; Jin, L. Caspase Family Proteases and Apoptosis. Acta Biochim. Biophys. Sin. 2005 ,37, 719–727. [CrossRef] 67. Ries, C. Cytokine Functions of TIMP-1. Cell. Mol. Life Sci. 2014,71, 659–672. [CrossRef] 68. Li, W.; Ma, Y.; Li, Z.; Lv, X.; Wang, X.; Zhou, D.; Luo, S.; Wilson, J.X.; Huang, G. Folic Acid Decreases Astrocyte Apoptosis by Preventing Oxidative Stress-Induced Telomere Attrition. Int. J. Mol. Sci. 2020,21, 62. [CrossRef] [PubMed] 69. Li, W.; Li, Z.; Zhou, D.; Zhang, X.; Yan, J.; Huang, G. Maternal Folic Acid Deficiency Stimulates Neural Cell Apoptosis via MiR-34a Associated with Bcl-2 in the Rat Foetal Brain. Int. J. Dev. Neurosci. 2019,72, 6–12. [CrossRef] [PubMed] 70. Cui, S.; Li, W.; Wang, P.; Lv, X.; Gao, Y.; Huang, G. Folic Acid Inhibits Homocysteine-Induced Cell Apoptosis in Human Umbilical Vein Endothelial Cells. Mol. Cell. Biochem. 2018,444, 77–86. [CrossRef] [PubMed] 71. Yao, Z.; Chen, D.; Wang, A.; Ding, X.; Liu, Z.; Ling, L.; He, Q.; Zhao, T. Folic Acid Rescue of ATRA-Induced Cleft Palate by Restoring the TGF-βSignal and Inhibiting Apoptosis. J. Oral Pathol. Med. 2011,40, 433–439. [CrossRef]