1521-0111/89/5/560–574$25.00 http://dx.doi.org/10.1124/mol.116.103697MOLECULAR PHARMACOLOGY Mol Pharmacol 89:560–574, May 2016Copyright ª 2016 by The American Society for Pharmacology and Experimental Therapeutics

All-Trans-Retinoic Acid Enhances Mitochondrial Function inModels of Human Liver s

Sasmita Tripathy, John D Chapman, Chang Y Han, Cathryn A Hogarth, Samuel L.M. Arnold,Jennifer Onken, Travis Kent, David R Goodlett, and Nina IsoherranenDepartments of Pharmaceutics (S.T., S.L.M.A., N.I.), Medicinal Chemistry (J.D.C., D.R.G.), and Diabetes Obesity Center forExcellence and the Department of Medicine, Division of Metabolism, Endocrinology and Nutrition (C.Y.H.), University ofWashington, Seattle, Washington; School of Molecular Biosciences and The Center for Reproductive Biology, Washington StateUniversity, Pullman, Washington (C.A.H., J.O., T.K.); and School of Pharmacy, University of Maryland, Baltimore, Maryland (D.R.G.)

Received February 6, 2016; accepted February 25, 2016

ABSTRACTAll-trans-retinoic acid (atRA) is the active metabolite of vitamin A.The liver is the main storage organ of vitamin A, but activation ofthe retinoic acid receptors (RARs) in mouse liver and in humanliver cell lines has also been shown. Although atRA treatmentimprovesmitochondrial function in skeletal muscle in rodents, itsrole in modulating mitochondrial function in the liver is contro-versial, and little data are available regarding the human liver. Theaim of this study was to determine whether atRA regulates he-patic mitochondrial activity. atRA treatment increased the mRNAand protein expression of multiple components of mitochon-drial b-oxidation, tricarboxylic acid (TCA) cycle, and respiratorychain. Additionally, atRA increased mitochondrial biogenesis inhuman hepatocytes and in HepG2 cells with and without lipidloading based on peroxisomeproliferator activated receptor gammacoactivator 1a and 1b and nuclear respiratory factor 1 mRNA and

mitochondrial DNA quantification. atRA also increased b-oxidationand ATP production in HepG2 cells and in human hepatocytes.Knockdown studies of RARa, RARb, and PPARd revealed thatthe enhancement of mitochondrial biogenesis and b-oxidationby atRA requires peroxisome proliferator activated receptordelta. In vivo in mice, atRA treatment increased mitochondrialbiogenesis markers after an overnight fast. Inhibition of atRAmetabolism by talarozole, a cytochrome P450 (CYP) 26 specificinhibitor, increased the effects of atRA on mitochondrial bio-genesis markers in HepG2 cells and in vivo in mice. Thesestudies show that atRA regulates mitochondrial function andlipid metabolism and that increasing atRA concentrations inhuman liver via CYP26 inhibition may increase mitochondrialbiogenesis and fatty acid b-oxidation and provide therapeuticbenefit in diseases associated with mitochondrial dysfunction.

IntroductionVitamin A (retinol) is stored predominantly in the liver as

retinyl esters. However, the active metabolite of retinol, all-trans-retinoic acid (atRA), appears to also regulate genetranscription in the liver (O’Byrne and Blaner, 2013). Mostactions of atRA are mediated via interactions with retinoicacid receptors (RARs) (Wolf, 2010; Yu et al., 2012), but atRAalso activates the peroxisome proliferator-activated receptor(PPAR)-b/d (Shaw et al., 2003; Berry and Noy, 2007; Wolf,2010). In addition, atRA induces the expression of the tran-scriptional repressor small heterodimer partner in the liver,resulting in broad effects on gene transcription, especially of

genes responsible for bile acid metabolism and transport (Kohet al., 2014; Mamoon et al., 2014; Yang et al., 2014). Theactivation of multiple pathways by atRA likely leads to diverseeffects on liver physiology, including altered lipid homeostasis.In humans both excessive intake of vitamin A or retinoids andvitamin A deficiency lead to hypertriglyceridemia, elevatedtotal cholesterol, and decreased high-density lipoprotein cho-lesterol (Bershad et al., 1985; Lawrence et al., 2001; Staels,2001; Brelsford and Beute, 2008; Lilley et al., 2013). However,likely because of a dichotomy of the effects of vitamin A on theliver, the endogenous role of atRA in regulating lipid and fattyacid homeostasis and role of atRA in liver disease is not wellunderstood.In rodents atRA appears to regulate various processes in

lipid homeostasis in a species-specific manner. Data fromseveral mouse studies suggest that atRA signaling via RARis needed to maintain liver mitochondrial fatty acid oxida-tion and lipid homeostasis. Hepatocyte-specific RAR knockout

This research was supported by grants from National Institutes of HealthNational Institute of General Medical Sciences [Grants R01 GM111772, R01GM081569, and R01 GM081569-S1] and National Institute of Diabetes andDigestive and Kidney Diseases [Grant P30 DK035816].

dx.doi.org/10.1124/mol.116.103697.s This article has supplemental material available at molpharm.

aspetjournals.org.

ABBREVIATIONS: ACSL, acyl CoA synthase ligase; ATGL, adipose triglyceride lipase; atRA, all-trans-retinoic acid; CPT1a, carnitinepalmitoyltransferase 1a; CYP, cytochrome P450; DMEM, Dulbecco’s modified essential medium; DMSO, dimethylsulfoxide; ECH1, enoylCoA hydratase 1; FASN, fatty acid synthase; FGF21, fibroblast growth factor 21; IDH2, isocitrate dehydrogenase; NAFLD, nonalcoholic fattyliver disease; NRF1, nuclear respiratory factor 1; PGC1a, peroxisome proliferator activated receptor gamma coactivator 1a; PGC1b,peroxisome proliferator activated receptor gamma coactivator 1b; PPAR, peroxisome proliferator activated receptor; RA, retinoic acid; RAR,retinoic acid receptor; SDHA, succinate dehydrogenase subunit A; SREBP1, sterol regulatory element binding protein 1.

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mice develop microvesicular steatosis, hepatocellular car-cinoma, decreased mitochondrial fatty acid oxidation, andincreased peroxisomal and microsomal fatty acid oxidationpresumably due to deficient atRA signaling (Yanagitani et al.,2004). Similarly, in normal mice atRA enhances fatty acidoxidation and ketogenesis via RAR activation and FGF21induction (Amengual et al., 2012; Li et al., 2013). In agreementwith a role of atRA in maintaining lipid homeostasis in theliver, decrease of hepatic atRA concentrations via inhibition ofatRA synthesis in mice led to microvesicular vacuolation butwithout a change in liver triglycerides (Paik et al., 2014).However, in diet-induced obese mice atRA treatment de-creased hepatic triglyceride content (Berry and Noy, 2009)and hepatic lipid accumulation (Kim et al., 2014). In contrast,in rats vitamin A deficiency led to increased expression ofgenes involved in fatty acidmetabolism in the liver (McClinticket al., 2006) and decreased liver total phospholipid content andphosphatidylcholine synthesis (Oliveros et al., 2007), demon-strating opposite effects to those observed in mice. Low retinoldiet also significantly decreased stellate cell free fatty acids andtotal lipids, whereas high retinol diet increased triglycerides,cholesteryl esters, free fatty acids, and total lipids in rat stellatecells (Moriwaki et al., 1988). How well these observations inmice or rats reflect atRA signaling in healthy or diseasedhuman liver is not known.Hepatic vitamin A stores are depleted in alcoholic liver

disease, and vitamin A deficiency is believed to play a role inthe development and progression of the disease (Bell et al.,1989; Ward and Peters, 1992; Lee and Jeong, 2012; Clugstonet al., 2013). Current data also strongly suggest that alteredvitamin A homeostasis and retinoid metabolism is present innonalcoholic fatty liver disease (NAFLD) and in subsequentsteatosis. In liver biopsies from patients with NAFLD, themRNA of retinaldehyde dehydrogenases involved in atRAformation and the mRNA of cytochrome P450 (CYP) 26A1responsible for atRA metabolism was increased (Ashla et al.,2010). In addition, in a panel of human livers with increasinglevel of steatosis, retinol and retinyl ester concentrations andRARb mRNA decreased significantly with increasing steatosis(Trasino et al., 2015). Finally, in a study of patients withNAFLD and steatosis, retinoic acid (RA) concentrations weresignificantly lower in plasma from patients than in healthycontrols (Liu et al., 2015). Yet detailed studies of retinoidsignaling in models of NAFLD have not been conducted.Because NAFLD is at least in part a mitochondrial diseaseassociated with decreased mitochondrial gene expression andfunction (Pessayre, 2007; Wei et al., 2008; Aharoni-Simonet al., 2011; Betancourt et al., 2014), regulation of mitochon-drial fatty acid oxidation and lipid homeostasis by atRA couldplay a role inNAFLDprogression. The aim of this studywas todetermine whether atRA regulates mitochondrial functionand lipid homeostasis in the healthy and fatty human liverand to establish how well mouse and rat models of atRAsignaling in the liver correlate with models of human liver.

Materials and MethodsChemicals and Reagents. Dulbecco’smodified essentialmedium

(DMEM)with high glucose (25mM) and fetal calf serumwere obtainedfrom Invitrogen (Carlsbad, CA). All-trans-retinoic acid (atRA) wasobtained from Sigma-Aldrich, St. Louis, MO. GW0742 (PPARdagonist) and TTNPB (RAR-pan agonist) were obtained from Tocris,

Minneapolis, MN. Talarozole (R115866) was obtained from Med-Chem Express, Princeton, NJ. siRNAs for RARa, RARb, andPPARd were obtained from Ambion, Life Science Technologies,Grand Island, NY. ATP assay kit was purchased from BiovisionInc., Milpitas, CA. Primer and probe pairs for human peroxisomeproliferator activated receptor gamma coactivator 1a (PGC1a)(Hs01016719_m1, FAM), PGC1b (Hs00991677_m1, FAM), nuclearrespiratory factor 1(NRF1) (Hs00192316_m1, FAM), carnitinepalmitoyltransferase 1a (CPT1a; (Hs00912671_m1, FAM), PPARa(Hs00947536_m1, FAM), PPARd (Hs04187066_g1, FAM), PPARg(Hs01115513_m1, FAM), GAPDH (Hs99999905_m1, VIC), RARa(Hs00940446_m1, FAM), RARb (Hs00233407_m1, FAM), adiposetriglyceride lipase (ATGL; Hs00386101_m1, FAM), diacylglycerolO-acyl transferase 2 (Hs01045913_m1, FAM), microsomal triglyc-eride transfer protein (Hs00165177_m1, FAM), and sterol regula-tory element binding protein 1(SREBP1; Hs01088691_m1, FAM)were obtained from Applied Biosystems, Carlsbad, CA. Primer andprobe pairs for mouse Rarb (Mm01319677_m1, FAM), Cyp26a1(Mm00514486_m1, FAM), Pgc1a (Mm01208835_m1, FAM),Cpt1a (Mm01231183_m1, FAM), Atgl (Mm00503040_m1, FAM),Pgc1b (Mm00504720_m1), Nrf1 (Mm01135606_m1), and b-Actin(Mm00607939_s1, FAM) were obtained from Applied Biosystems.Mitochondrial genes and nuclear gene primer set #7246 (for human)and #RR290 (for mouse) were purchased from Takara Clontech,Mountain View, CA.

HepG2 Cell Culture. HepG2 cells were cultured in 6-well plates(Corning Life Sciences, Corning, NY) with a cell density 1�106 cellsper well in DMEM with 25 mM glucose media with 10% fetal calfserum in a 5% carbon dioxide environment in a humidified incubatorat 37°C. All media contained penicillin and streptomycin, and all thestudies were conducted in triplicate. Lipid-loaded HepG2 cells wereused as a model for fatty liver. For lipid loading, HepG2 cells weretreatedwith 200mMoleic acid for 3 days. Every 24 hours, 200mMoleicacid was added to the media. The lipid loading was confirmed by OilRed O staining (BioVision Inc.) according to manufacturer’srecommendations.

Cell Viability Assay. HepG2 cell growth was measured usingWST-1 reagent (Roche, Indianapolis, IN). Briefly, cells were grown ina 96-well plate cultured in a 5% carbon dioxide environment in ahumidified incubator at 37°C. By using six replicates per treatmentand a 0.1% dimethylsulfoxide (DMSO) control, the effect of atRAtreatment (0.001–50 mM) on cell proliferation was measured after72 hours of treatment. Every 24 hours, the media were removed, cellswere washed with PBS, and the media containing atRAwere replacedto maintain constant atRA concentrations. At 72 hours, 10 ml of WST-1reagent were added to each well and incubated at 37°C for 1 hour tomeasure cell viability and growth according to manufacturer’sinstructions. The absorbance was measured at 450 nm with back-ground subtraction at 680 nm. Each treatment was normalized to thecontrol cells, and cell viability was expressed as a percentage of thecontrol.

Human Hepatocyte Experiments. Cryopreserved human he-patocytes from five donors (3women and 2men aged 13-52 years) wereobtained from Xenotech (Lenexa, KS). Hepatocytes were thawed at37°C, placed into thawing media (K2000; Xenotech), and cell viabilitywas confirmed using trypan blue. Hepatocytes were diluted in platingmedia (Williams Emedia plus hepatocytes supplement pack CM3000,LifeTechnologies, Grand Island, NY) at 1.2 � 106 live cells. Platingmedia (330ml) containing hepatocytes were added to each well in 12 or24-well collagen type I-coated plates (LifeTechnologies) and allowed toattach for ∼8 hours. The plating media were then replaced withWilliamsEmedia containing cellmaintenance supplements (CM4000;LifeTechnologies). Hepatocytes from three donors were plated andtreated with 1 mM atRA or ethanol as a vehicle control for 24 hours,and at the end of the 24-hour treatment, cells were harvested forPGC1a, PGC1b, and NRF1 mRNA measurements or for quantifica-tion of ATP production or mitochondrial DNA. Hepatocytes weretreated with 1 mM atRA or ethanol as a vehicle control for 48 hours for

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SDHA protein quantification. For lipid loading, human hepatocyteswere loaded with 600 mM oleic acid once for 24 hours and then treatedwith 1 mM atRA or ethanol as a vehicle control for 24 hours. At the endof the 24-hour treatment, the cells were harvested for PGC1a, PGC1b,and NRF1 mRNA measurements or analyzed for ATP production. Alltreatments were conducted in triplicate.

Proteomic Analysis. Proteomic changes driven by atRA inHepG2 cells were evaluated using the PAcIFIC shotgun proteomicmethod (Chapman et al., 2016). HepG2 cells were treated with eitheratRA (10 mM) or 0.1% DMSO (vehicle control) for 72 hours andharvested. For proteomic analysis, HepG2 cells were cultured inDMEM media supplemented with 10% fetal bovine serum and 1%penicillin/streptomycin in 10 cm2 dishes and cells were grown to70% confluence. The cells were treated with either atRA (10 mM) or0.1% DMSO (control) for 72 hours. The media were replaced every24 hours with fresh treatmentmedia containing either atRA orDMSOuntil harvest. Cells were harvested by washing them twice with icecold PBS and scraping the cells into 0.5 ml of 100 mM ammoniumbicarbonate for shotgun proteomics analysis. Harvested cell suspen-sion was sonicated to lyse the cells and subsequently centrifuged at5,000 g to pellet cellular debris. Supernatant from each sample wascollected and placed in a clean Eppendorf tube on ice. Proteinquantification was performed with a BCA protein quantification kitfollowing the manufacturer’s protocol (Thermo Scientific, Waltham,MA). Protein quantity was normalized for each sample before furtherproteomics sample preparation. The samples with 200 mg protein per100 ml were denatured with the addition of 6 M urea. Subsequently,samples were buffered with the addition of 7 ml 1.5 M Tris pH 8.8,reduced with 2.5 ml of 200 mM tris-2-carboxyethyl phosphine for1 hour at 37°C, alkylated with 20 ml of 200 mM iodoacetamide for1 hour at room temperature in the dark, and then quenchedwith 20 mlof 200 mM dithiothreitol. Before addition of sequencing-grade porcinetrypsin (Promega, Madison, WI) at a protein to enzyme ratio of 50:1,samples were diluted with 900 ml of 50 mM ammonium bicarbonateand 200 ml of MeOH. After an overnight incubation, peptides weredesalted on a Vydac C18 macrospin column (The Nest Group, South-borough, MA) according to the manufacturer’s protocol. Resultingeluent was concentrated on a SPD 111V SpeedVac (Thermo Savant,San Jose, CA) and stored until further use. Peptide digests wereanalyzed on an LTQ Oribtrap (Thermo Fisher, San Jose, CA) coupledwith anano-electrospray ionization in positive ionmode. For thePAcIFICacquisition, Captive spray ionization mass spectrometry was performedusing a Waters NanoAquity (Milford, MA). The mass spectrometerparameters were set as defined previously (Panchaud et al., 2009,2011). A fused silica 200 mm i.d. trapping column purchased from NewObjective (Woburn, MA) and packed with 2 cm of 200 Å, 5 mm MagicC18AQ particles (Michrom, Auburn, CA) was used. Successive analyticalseparation was performed on a fused silica 200 mm i.d. trapping columnpurchased from New Objective and packed with 15 cm of 100 Å, 5 mmMagic C18AQ particles (Michrom). For each sample injection, 1 mg of thepeptide sample was loaded on the trapping column at 6 ml/min with 95%water/5% acetonitrile/0.1% formic acid for 2.5 minutes. Trapped peptideswere then eluted from the trapping column onto the analytical columnusing a variable gradientwith a flow rate of 2.5ml/min. The gradient usedtwomobile phase solutions: A, water/0.1% formic acid andB, acetonitrile/0.1% formic acid. For mass spectrometry, ion source conditions wereoptimized using the tuning and calibration solution suggested by theinstrument manufacturer and under conditions recommended by theCaptive spray ionizationmanufacturer (Bruker/Michrom).Data acquiredwere converted from Thermo’s RAW format to the universal mzXMLformat and searched against the IPI human database v3.49 (http://www.ebi.ac.uk/IPI/IPIhuman.html) using SEQUEST (Havilio et al., 2003). Theprecursor ion tolerancewas set to 3.75, and additional search parametersfor SEQUEST included trypsin enzyme specificity, cysteines modifiedwith iodoacetamide and the variable option formethionines in reduced oroxidized form. Results were analyzed with Peptide Prophet (Keller et al.,2002; Nesvizhskii et al., 2003) ensuring that peptide hits with a proba-bility of .0.99 were accepted and linked to protein entries. Protein

identifications were assigned relative quantification values using in-housespectral counting software (Ryu et al., 2008). The relative quantificationvalue for each protein is determined by summing all peptide tandemmassspectrometry spectra correlated to a respective protein (Liu et al., 2004).After spectral counting, the data were filtered using a Student’s t test anda P value cutoff of 0.05 to ensure that protein changes were consistentthroughout the biologic triplicate and technical duplicate analyses.

mRNA Analysis and Quantitative RT-PCR. RT-PCR was usedto quantify mRNA expression of various genes (StepOnePlus, AppliedBiosystems, Carlsbad, CA) as previously described (Tay et al., 2010).For mRNA extraction, 300 ml of TRI reagent (Invitrogen) was added toeach well containing cultured HepG2 cells or hepatocytes after aspira-tion of all media, and mRNA extracted according to the manufacturer’srecommendations. Total RNA was quantified using the Nanodrop 2000cSpectrophotometer (Thermo Fisher Scientific). cDNA was synthesizedfrom 1 mg mRNA using TaqMan Gene expression reagents (AppliedBiosystems). TaqMan real-time gene expression master mix and PCRprimers and fluorescent probes were obtained from Applied Biosystems.Probes were labeled with the 59-reporter dye 5-carboxyfluorescein and anonfluorescent black hole quencher on the 39-end. GAPDH was used asthe housekeeping gene, and all assays were done as multiplexes. Nochanges in GAPDH mRNA levels were observed in any of the experi-ments. All biological triplicate samples were analyzed in duplicate.Changes in target mRNA were measured using relative quantification(fold-difference) and the DDCT method (Tay et al., 2010).

Effects of atRA on Mitochondrial Number, Function, andProtein Expression. The effect of atRA on mitochondrial number,function, and protein expression was determined by measuring ATPproduction, fatty acid oxidation, quantity of mitochondrial DNA, andexpression of succinate dehydrogenase subunit A (SDHA). All exper-iments were conducted as biological triplicates.

For Western blotting, proteins were extracted as whole cell extractfrom HepG2 cells treated with vehicle (ethanol) or 1 mM atRA for48 hours using whole cell lysis buffer in the presence of protease (RocheApplied Science) and phosphatase inhibitors (1 mM b-glycerol phos-phate, 2.5 mM Na-pyrophosphate, 1 mM Na3VO4). Protein concen-trations were measured by BCA protein assay. Whole cell proteinextracts were separated by SDS-polyacrylamide gel electrophoresis(NuPAGE-Novex 4–12% polyacrylamide Bis-Tris, Life Science Tech-nologies) and transferred to nitrocellulose membranes. Blots wereincubated with primary antibodies against SDHA and b-actin over-night at 4°C. Next day blots were washed and incubated with secondaryantibody for 1 hour at room temperature. Antigen-antibody reactionswere detected and quantified using LiCor Odyssey scanner andsoftware (Licor Inc., Lincoln, NE). SDHA (product #5839) was pur-chased from Cell Signaling, Danvers, MA. b-Actin antibody (product#AC-15) was obtained from Abcam, Cambridge, MA. The secondaryantibodies, IRDye 680 (antimouse) and IRDye 800, (anti-rabbit) wereobtained from LiCor Inc. All Western blots were conducted as technicalduplicates.

To measure b-oxidation, [1-14C] palmitate (American RadiolabeledChemicals Inc.) was used. HepG2 cells were gently detached withnonenzymatic dissociation solution (Sigma) and suspended in Krebs-Ringer buffer containing 1 nM insulin and placed in vials. Wellscontaining cylinders of Whatman filter paper were suspended abovethe HepG2 cells containing solution in vials. The vials were stopperedwith rubber caps, gassed with 95% O2-5% CO2, and 0.5 mCi of [1-14C]palmitate for b-oxidation was injected into the vials. After incubationfor 3 hours at 37°C, 200 ml hyamine hydroxide was injected into thewells containing the filter paper, 200 ml of 1N HCl was added to theHepG2 cells, and the vials were incubated overnight at 37°C. The filterpaper in the wells was then transferred to 10 ml of scintillation fluidand counted.

To measure ATP production, 105 HepG2 cells per well were platedin a 12-well plate. HepG2 cells were treated with 0.01–10 mM atRA for24 hours. Cells were then harvested for the ATP assay, and ATP wasquantified from the cell lysate using a commercially availablecolorimetric kit purchased from Biovision Inc. Total ATP production

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was normalized to milligram protein. For ATP assay in normal andlipid loaded human hepatocytes, hepatocytes were treated with 1 mMatRA for 24 hours and then cells were harvested for the ATP assay asdescribed above.

To quantify mitochondria number, the mitochondrial DNA wasquantified using qPCR. Genomic DNA from HepG2 cells and humanhepatocytes was extracted by using DNAzol. Total DNA was quanti-fied using the Nanodrop 2000c Spectrophotometer (ThermofisherSci.). Genomic DNA (100 ng) per reaction was used to perform qPCR.The mitochondrial and nuclear genes primer set were obtained fromTakara Clontech. No changes in nuclear DNA levels were observed inany of the experiments. All biologic triplicate samples were analyzedin duplicate. Changes in mitochondrial DNA copy number weremeasured using relative quantification (copy number) versus nuclearDNA and the DDCT method.

Role of RAR and PPARd on RA Signaling in the Liver. Todetermine the nuclear receptors potentially responsible for atRAeffects on mitochondrial function, HepG2 cells were first treated with1 mM atRA, 50 nM TTNPB (RAR pan agonist), 200 nM GW0742(PPARd agonist) or vehicle (ethanol) for 24 hours, and the mRNAexpression of PGC1a, PGC1b, and NRF1 was measured using RT-PCR as described above. Second RARa, RARb, and PPARd siRNAwasused in normal and lipid-loaded HepG2 cells together with 1 mM atRAor vehicle treatment of 48–72 hours to determine the effects of eachnuclear receptor knock down on PGC1a, PGC1b, and NRF1 mRNAand on ATP production, SDHA protein content, mitochondrial number,and b-oxidation. siRNAs for RARa (s11801), RARb (s11803), andPPARd (s10883) were purchased from Ambion, Thermo Fisher Scien-tific. Lipofectamine RNAiMAX transfection reagent (13778-075) waspurchased from Invitrogen, ThermoFisher Scientific. For the siRNAexperiments, 106 HepG2 cells were grown in 6-well plates as describedabove. Cells were transfected with either siRNA for RARa, RARb, andPPARd or control siRNA (scrambled RNA) at 25 pmol/well togetherwith 7.5 ml/well of the lipofectamine RNAiMAX transfection reagentaccording to the manufacturer’s recommendations. Forty-eight hours

after transfection, 1 mM atRA or vehicle was added to the media. After24 hours of atRA treatment, cells were harvested, and either RNAextracted and the expression of PGC1a, PGC1b, and NRF1 measured,or ATP production, SDHA protein, b-oxidation, or mitochondrialnumber were measured as described above. The knock down of PPARdand RARb was confirmed by Western blot of PPARd and RARb proteinexpression. Antibodies for PPARd (ab137724) and RARb (EPR2017)were purchased from Epitomics-Abcam. Western blotting was conductedas described above using whole cell lysate.

Effect of CYP26 Inhibition on atRA Potency. Normal andlipid loaded HepG2 cells were treated with 1 mM atRA or ethanol(vehicle) in the presence and absence of 1 mM talarozole (a CYP26specific inhibitor) for 24 hours. Media were collected for atRA andatRA metabolite (4-OH-atRA and 4-oxo-atRA) analysis by liquidchromatography tandem mass spectrometry as previously described(Topletz et al., 2015) using an AB Sciex QTRAP 4500 mass spectrom-eter operated on a negative ion electrospray detection and coupledwith an LC-20ADultra-fast liquid chromatography system (ShimadzuCo.,Kyoto, Japan) andAgilent ZorbaxC18 column (3.5mm,2.1� 100mm).Cells were harvested for RNA extraction and mRNA expression ofPGC1a, PGC1b, and NRF1 was measured as described above. In aseparate study, HepG2 cells were treated with 1 mM atRA or ethanol(vehicle) in the presence and absence of 1 mM talarozole for 24 hours,and ATP production was measured as described above.

Effect of RA and Talarozole Treatment on MitochondrialBiogenesis in Mice. All animal experiments were approved byWashington State University Animal Care and Use Committee andwere conducted in accordance with the guiding principles for the careand use of research animals of the National Institutes of Health(Bethesda, MD). Mixed background (129/C57BL/6) mice housed in atemperature- and humidity-controlled environment with food andwater provided ad libitum were used for the studies. Before tissuecollection, the animals were euthanized by CO2 asphyxiation, followedby cervical dislocation, and livers were removed and snap frozen inlight-protected tubes until analysis.

Fig. 1. atRA increases the protein and mRNA expression of enzymes involved in fatty acid oxidation in HepG2 cells. (A) Cell viability after treatment ofHepG2 cells with increasing concentrations of atRA. Cell proliferation is shown as mean 6 S.D., n = 6. (B) Heat map for the effects of 10 mM atRAtreatment in comparisonwith vehicle (DMSO) on expression of enzymes involved in differentmetabolic pathways as determine by the proteomic analysis.(C) The effect of atRA treatment (1 and 10 mM) on the mRNA expression of enzymes involved in lipid metabolism. mRNA results are expressed asmean 6 S.D., n = 3 and are representative of at least 3 separate studies. *P , 0.05 compared with vehicle-treated controls Student’s t test.

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TABLE 1Relative proteomic analysis of the effects of atRA on protein expression in HepG2 cellsThe changes were quantified via spectral counting. Significant differences are underlined. Only proteins for which the same peptide was consistently detected in all replicatecontrol and treated samples are shown.

UniProt ID RA/Con P value Protein Name Gene Name

Transcription Q92804 1.00 1.00 TATA-binding protein-associated factor 2N TAF15P12004 0.93 0.23 Proliferating cell nuclear antigen PCNAP20290 1.21 0.02 Transcription factor BTF3 BTF3Q13185 0.88 0.02 Chromobox protein homolog 3 CBX3Q08211 1.51 0.04 ATP-dependent RNA helicase A DHX9

RNA Processing O43776 1.15 0.33 Asparagine–tRNA ligase, cytoplasmic NARSQ99714 0.63 0.15 3-hydroxyacyl-CoA dehydrogenase type-2 HSD17B10O75643 1.70 0.004 U5 small nuclear ribonucleoprotein 200 kDa helicase SNRNP200P31942 1.91 0.008 Heterogeneous nuclear ribonucleoprotein H3 HNRNPH3Q1KMD3 1.36 0.01 Heterogeneous nuclear ribonucleoprotein U-like protein 2 HNRNPUL2P55795 0.70 0.01 Heterogeneous nuclear ribonucleoprotein H2 HNRNPH2P23246 1.20 0.03 Splicing factor, proline- and glutamine-rich SFPQQ16630 1.30 0.03 Cleavage and polyadenylation specificity factor subunit 6 CPSF6P49588 1.32 0.03 Alanine-tRNA ligase, cytoplasmic AARSQ15637 1.40 0.04 Splicing factor 1 SF1

Translation P46779 0.52 0.01 60S ribosomal protein L28 RPL28P56537 0.96 0.77 Eukaryotic translation initiation factor 6 EIF6P62241 0.80 0.02 40S ribosomal protein S8 RPS8Q9NR30 1.32 0.01 Nucleolar RNA helicase 2 DDX21P24534 1.28 0.02 Elongation factor 1-beta EEF1B2P26373 0.73 0.03 60S ribosomal protein L13 RPL13

Protein integrity P61604 0.78 0.27 10 kDa heat shock protein, mitochondrial HSPE1Q9Y3C6 0.95 0.42 Peptidyl-prolyl cis-trans isomerase-like 1 PPIL1P13667 1.18 0.02 Protein disulfide-isomerase A4 PDIA4P27797 1.24 0.03 Calreticulin CALRP08107 2.70 0.03 Heat shock 70 kDa protein 1A/1B HSPA1AQ9Y4L1 1.50 0.03 Hypoxia upregulated protein 1 HYOU1P11021 0.93 0.04 78 kDa glucose-regulated protein HSPA5P08238 1.13 0.04 Heat shock protein HSP 90-beta HSP90AB1

Protease activity P49720 0.90 0.69 Proteasome subunit beta type-3 PSMB3Q99436 0.75 0.29 Proteasome subunit beta type-7 PSMB7P35998 1.20 0.02 26S protease regulatory subunit 7 PSMC2P55036 1.21 0.04 26S proteasome non-ATPase regulatory subunit 4 PSMD4

Cellular trafficking Q9Y678 0.85 0.11 Coatomer subunit gamma-1 COPG1Q8TEX9 1.28 0.03 Importin-4 IPO4Q14974 1.15 0.004 Importin subunit beta-1 KPNB1P51149 1.37 0.03 Ras-related protein Rab-7a RAB7AO00151 1.30 0.04 PDZ and LIM domain protein 1 PDLIM1

Metabolism P23526 1.05 0.59 Adenosylhomocysteinase AHCYO60701 0.93 0.16 UDP-glucose 6-dehydrogenase UGDHP51659 1.30 0.49 Peroxisomal multifunctional enzyme type 2 HSD17B4P40925 1.01 0.72 Malate dehydrogenase, cytoplasmic MDH1Q15181 1.22 0.26 Inorganic pyrophosphatase PPA1Q9NR45 0.98 0.89 Sialic acid synthase NANSO43175 1.03 0.53 D-3-phosphoglycerate dehydrogenase PHGDHQ8NF37 1.00 1.00 Lysophosphatidylcholine acyltransferase 1 LPCAT1P54819 1.03 0.64 Adenylate kinase 2, mitochondrial AK2Q06830 1.02 0.685 Peroxiredoxin-1 PRDX1P14324 1.52 0.01 Farnesyl pyrophosphate synthase FDPSQ99541 2.57 0.001 Perilipin-1 PLIN2P49327 1.27 0.002 Fatty acid synthase FASNP06737 2.46 0.00 Glycogen Phosphorylase, liver form PYGLQ13011 1.27 0.006 Delta(3,5)-Delta(2,4)-dienoyl-CoA isomerase, mitochondrial ECH1Q95573 1.56 0.01 Long-chain-fatty-acid-CoA ligase 3 ACSL3P17174 2.11 0.01 Aspartate aminotransferase, cytoplasmic GOT1P49419 1.74 0.01 Alpha-aminoadipic semialdehyde dehydrogenase ALDH7A1Q86SX6 2.26 0.01 Glutaredoexin-related protein 5, mitochondrial GLRX5P12268 1.50 0.01 Inosine-59-monophosphate dehydrogenase 2 IMPDH2Q53HR2 1.68 0.01 Acyl-Coenzyme A dehydrogenase ACADP25705 1.56 0.02 ATP synthase subunit alpha, mitochondrial ATP5A1Q8NBX0 1.35 0.02 Saccharopine dehydrogenase-like oxidoreductase SCCPDHO60448 1.38 0.02 Long-chain-fatty-acid-CoA ligase 4 ACSL4P05091 1.24 0.03 Aldehyde dehydrogenase, mitochondrial ALDH2P48735 1.67 0.03 Isocitrate dehydrogenase [NADP], mitochondrial IDH2P51648 1.88 0.03 Fatty aldehyde dehydrogenase ALDH3A2P52209 1.44 0.03 6-phosphogluconate dehydrogenase, decarboxylating PGDP00387 1.29 0.03 NADH-cytochrome b5 reductase 3 CYB5R3P12277 1.38 0.04 Creatine kinase B- type CKBP80404 2.00 0.04 4-aminobutyrate aminotransferase, mitochondrial ABATP78417 1.34 0.04 Glutathione S-transferase omega-1 GSTO1P50897 1.62 0.04 Palmitoyl-protein thioesterase 1 PPT1

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To determine the effect of atRA treatment on mitochondrialbiogenesis in mice, young adult (∼60 dpp) female mice were treatedonce daily intraperitoneally with 5 mg/kg atRA (Sigma Aldrich)dissolved in 15% DMSO/85% sesame oil or vehicle for 5 consecutivedays. Twelve hours after the final dose, all food was removed and allmice were fasted for 12 hours. After the fast, the mice were split intofasted and refed treatment groups. Animals in the fasted group wereeuthanized and livers collected 24 hours after the final RA or vehiclecontrol dose, immediately after the 12-hour fast. For the refed group,food was added to cages after the 12-hour fasting period and the animalswere euthanized and livers collected 4 hours later after 4-hour exposureto food (28 hours after the final RA dose). To determine whetherinhibition of CYP26 enzymes has similar effects as atRA administra-tion onmouse liver, adult (∼90 dpp)malemicewere treated twice daily(every 12 hours) for 3 consecutive days with 2.5 mg/kg talarozole(Active Biochem, Maplewood NJ) dissolved in polyethylene glycol 300or vehicle control. Twelve hours after the final dose, the animals wereeuthanized and livers collected. mRNAwas extracted from 50–100mgliver tissue and qRT-PCR was used to determine changes in targetmRNA as described above. b-Actin was used as the housekeeping gene.To quantify mitochondrial number, SDHA expression was determinedby Western blotting as described above, and mitochondria number wasquantified from genomic DNA using qPCR as described above.

Statistical Analysis. Statistical analyses were conducted usingGraphPad prism v.5 statistical software (San Diego, CA). All differ-ences involving multiple comparisons were tested using one-wayanalysis of variance with Tukey test as the post hoc analysis, whereassingle comparisons were done using Student’s t test. Data areexpressed as mean 6 S.D.; P , 0.05 was considered significant.

ResultsProteomic Analysis of atRASignaling in the Liver. atRA

treatment had no effect on HepG2 cell proliferation orviability at atRA concentrations from 0.001 to 50 mM overthe 72-hour treatment (Fig. 1A), confirming that alterationsin protein or mRNA expression were not due to alterednumber of cells. Based on the proteomic analysis of HepG2

cells treated with 10 mM atRA for 72 hours, the expression of59 proteins in several functional categories was significantlyaltered by atRA treatment (Table 1). This was a smallfraction (,0.1%) of all proteins detected in the experiment.Table 1 lists those proteins for which the same peptides weredetected reliably in all analyzed samples. Almost half of theproteins (24 of 59, 41%) whose expression was significantlyaltered were proteins involved in lipid metabolism (Fig. 1B),suggesting that one of the major effects of atRA in the liveris modulating fatty acid oxidation and lipid metabolism.Based on the proteomic analysis, atRA treatment signifi-cantly increased expression of enzymes involved in fatty acidb-oxidation such as acyl CoA synthase ligase (ACSL) 3,ACSL4, fatty acyl CoA dehydrogenase and ECH1, isocitratedehydrogenase (IDH2), a key enzyme involved in tricarbox-ylic acid cycle (TCA), and ATP synthase (ATP synthasesubunit a), the mitochondrial respiratory chain complex V(Fig. 1B). In agreement with proteomic findings, the mRNAlevels of CPT1a, the rate limiting enzyme for fatty acidoxidation andATGL, a triglyceride specific lipase involved intriglyceride hydrolysis were significantly increased (Fig. 1C)in HepG2 cells at 1 and 10 mM atRA treatment. Interest-ingly, atRA treatment at 1 mM had no effect on mRNA levelsof SREBP1c and diacylglycerol O-acyl transferase 2, whichare involved in fatty acid and triglyceride synthesis, nordid atRA treatment alter the mRNA levels of microsomaltriglyceride transfer protein, an enzyme involved in very lowdensity lipoprotein synthesis and secretion (Fig. 1C). How-ever, at 10 mM, atRA increased SREBP1c mRNA (a keytranscription factor for de novo fatty acid synthesis; Fig. 1C)and FASN protein, which is regulated by SREBP1 (Fig. 1B).Taken together, the proteomics and mRNA analysis suggestthat atRA increases mitochondrial fatty acid oxidation inthe HepG2 cells at physiologically relevant concentrationsand may increase fatty acid synthesis at supraphysiologicconcentrations.

TABLE 1—Continued

UniProt ID RA/Con P value Protein Name Gene Name

P11586 1.28 0.04 C-1-tetrahydrofolate synthase, cytoplasmic MTHFD1Cell cycle regulation P42166 0.85 0.67 Thymopoietin isoform alpha TMPO

P33991 1.45 0.13 DNA replication licensing factor MCM4 MCM4P33993 0.94 0.67 DNA replication licensing factor MCM7 MCM7P49736 1.47 0.004 DNA replication licensing factor MCM2 MCM2P62826 1.14 0.04 GTP-binding nuclear protein Ran RANP18754 1.35 0.01 Regulator of chromosome condensation RCC1

Cell survival O75874 1.20 0.08 Isocitrate dehydrogenase [NADP] cytoplasmic IDH1P38646 1.21 0.04 Stress-70 protein, mitochondrial HSPA9Q13442 1.78 0.01 28 kDa heat- and acid-stable phosphoprotein PDAP1

Apoptosis P10599 1.00 1.00 Thioredoxin TXNP21796 1.28 0.01 Voltage-dependent anion-selective channel protein 1 VDAC1P09382 0.95 0.64 Galectin-1 LGALS1Q9Y265 1.00 1.00 RuvB-like 1 RUVBL1

Structural integrity P15924 0.93 0.28 Desmoplakin DSPQ09666 1.64 0.19 Neuroblast differentiation-associated protein AHNAK AHNAKQ15149 0.85 0.32 Plectin PLECQ01082 1.02 0.89 Spectrin beta chain SPTBN1P63313 0.72 0.10 Thymosin beta-10 TMSB10Q9Y2B0 1.09 0.39 Protein canopy homolog 2 CNPY2P35579 2.61 0.02 Myosin-9 MYH9P67936 2.58 0.01 Tropomyosin alpha-4 chain TPM4

Misc. P08758 1.12 0.31 Annexin A5 ANXA5Q13283 1.19 0.41 Ras GTPase-activating protein-binding protein 1 G3BP1Q96CN7 0.72 0.23 Isochorismatase domain-containing protein 1 ISOC1Q9H3K6 1.38 0.30 BolA-like protein 2 BOLA2

ACAD, fatty acyl CoA dehydrogenase; ATP5A1, ATP synthase subunit a.

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Effects of atRA on Mitochondrial Biogenesis Markersin Models of Healthy Human Liver. On the basis of theproteomic data that suggested that mitochondrial fatty acidoxidation was increased, the mRNA expression of key mito-chondrial biogenesis markers PGC1a, PGC1b, and NRF1was measured in HepG2 cells treated with 1 mM atRA for24 hours. The mRNA of PGC1a, PGC1b, and NRF1 wasincreased two- to threefold (P , 0.05) in the HepG2 cellssuggesting increased mitochondrial biogenesis in responseto atRA treatment (Fig. 2A). In agreement with the mRNAdata, ATP production was significantly increased aftertreatment with 1 mM atRA (Fig. 2B), suggesting mitochondrialfunction was also increased.

Considerable depletion of atRA was observed in the treatedcells at 24 hours of treatment with 1 mM atRA. atRAconcentrations decreased by 87% to 129 6 19.15 nM after24 hours (Table 2). To test whether inhibition of CYP26 (themain enzyme metabolizing atRA) increases atRA concentra-tions and potency, HepG2 cells were treatedwith atRA togetherwith talarozole, a CYP26 inhibitor, and atRA concentrationsand mitochondrial biogenesis markers and ATP productionwere measured. In the presence of talarozole, atRA con-centrations were 2.6-fold higher at the end of the 24-hourtreatment than when cells were treated with atRA alone,whereas the concentrations of the metabolites of atRA weredecreased upon talarozole treatment (Table 2). The increased

Fig. 2. atRA increases mitochondrial biogenesis, ATP production, and fatty acid b-oxidation in normal HepG2 cells. (A) The effect of atRA (1 mM) andtalarozole cotreatment (TLZ) on mRNA expression of mitochondrial biogenesis genes. (B) The effects of atRA and TLZ cotreatment on ATP production.(C and D) Induction of SDHA protein after treatment with atRA as measured by Western blotting. (C) Quantification of the representative Western blotshown in (D). The blot shows analysis of three independent biologic replicates. (E) The effects of atRA treatment on palmitate oxidation as a measure ofincreased b-oxidation. (F) The effect of atRA treatment on mitochondrial DNA copy number. All experiments were conducted as described in MaterialsandMethods. Results are expressed as mean6 S.D., n = 3. Comparisons between vehicle (ethanol)- and atRA-treated cells were made by Student’s t test:*P , 0.05 compared with vehicle-treated controls. #P , 0.05 compared with 1 mM atRA-treated controls.

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exposure of the cells to atRA was reflected in two- to threefoldhigher PGC1a, PGC1b, and NRF1 mRNA induction in thepresence of talarozole together with atRA comparedwith atRAalone (Fig. 2A). Talarozole treatment with atRA also resultedin approximately twofold greater ATP production in HepG2cells compared with atRA treatment alone (Fig. 2B). Surpris-ingly, talarozole alone also increased ATP production, PGC1b,and NRF1 mRNA expression but not PGC1a mRNA.To confirm that the observed changes in the mRNA of

mitochondrial biogenesis genes after atRA treatment result inimprovedmitochondrial function and increasedmitochondrialnumber and fatty acid oxidation, the protein expression ofSDHA, a marker for increase in mitochondrial content wasmeasured together with mitochondrial DNA and fatty acidb-oxidation. In HepG2 cells 1 mM atRA treatment increasedSDHA protein expression twofold (Fig. 2, C and D) and

1-10 mM atRA treatment increased fatty acid b-oxidation two-to threefold compared with vehicle-treated cells (Fig. 2E).Mitochondrial DNA was increased four- to fivefold after atRAtreatment (Fig. 2F).To further translate the findings fromHepG2 cells to human

liver, the effect of atRA on mitochondrial function and biogen-esis was evaluated in human hepatocytes from three donors.Similar to the HepG2 cells, in human hepatocytes the mRNAexpression of PGC1a, PGC1b, and NRF1 was increased two-to fivefold (P , 0.05) compared with vehicle treated group(Fig. 3A). In addition, atRA increased ATP production by two-to threefold when compared to vehicle-treated human hepa-tocytes. atRA treatment also increased SDHA protein expressiontwo- to threefold (Fig. 3C) and mitochondrial DNA two- tosevenfold (Fig. 3D), demonstrating that atRA increases mito-chondria number and function in human hepatocytes.

Fig. 3. atRA increases mitochondrial biogenesis and ATP production in normal human hepatocytes. (A) Induction of the mRNA expression of keymitochondrial biogenesis genes (PGC1a, PGC1b, and NRF1) by 1 mM atRA. All data are shown as fold increase in comparison with vehicle control. (B)Induction of ATP production by 1 mM atRA in three independent human hepatocyte donors. (C) Induction of SDHA protein in three individual humanhepatocyte donors after treatment with 1 mM atRA as measured by Western blotting. The quantification of the replicate Western blots and arepresentative blot of duplicate treatments in three donors is shown. (D) The effect of atRA on mitochondrial DNA copy number in three humanhepatocyte donors. All experiments were conducted as described in Materials and Methods. Results are expressed as mean 6 S.D., n = 3. Comparisonsbetween vehicle (ethanol)- and atRA-treated cells were made by Student’s t test: *P , 0.05 compared with vehicle-treated controls.

TABLE 2Cell media atRA, 4-OH-atRA, and 4-oxo-atRA concentrations at the end of the 24-hour treatment withvehicle, atRA, talarozole, or the combination of atRA and talarozole

Treatment atRA 4-OH-atRA 4-oxo-atRA

nM

Vehicle (ethanol) ND ND NDatRA (1 mM) 129 6 19 7.5 6 1.2 7.4 6 1.2Talarozole (1 mM) ND ND NDatRA (1 mM) + talarozole (1 mM) 334.3 6 26.8 ND 1.5 6 0.1

ND, not detected.

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To explore the potential nuclear receptors responsible forthe effects of atRA on mitochondrial biogenesis, the effect ofatRA on the mRNA expression of RARs and PPARs wastested. atRA significantly increased the mRNA expression of

RARb and PPARd but had no effect on RARa, PPARa, orPPARg (Supplementary Fig. 1A). This suggests that the effectof atRA on mitochondrial biogenesis may be a combination ofRARb and PPARd activation. The role of RARb and PPARd in

Fig. 4. The increase of mitochondrial biogenesis, fatty acid b-oxidation, and ATP production by atRA treatment in HepG2 cells requires PPARd. (A) Theinfluence of RARa, RARb, and PPARd siRNA on the induction ofmitochondrial biogenesis genes by atRA. (B) The effect of RARb andPPARd siRNA on theinduction of ATP production by atRA (C and D). The effect of RARb and PPARd siRNA on the induction of SDHA protein expression by atRA asmeasuredby Western blotting. (C) The quantification of replicate Western blots and (D) representative Western blot. The representative blot shows analysis ofbiologic duplicates for each treatment. (E and F) The effect of RARb and PPARd siRNA on fatty acid b-oxidation (E) and induction of mitochondrialDNA copy number (F) by atRA in HepG2 cells. Results for the mRNA, mitochondrial DNA, fatty acid oxidation, and ATP production studies arepresented as fold change compared with vehicle (ethanol). Results for protein studies are presented as ratio of target protein over loading controlb-actin. Results are expressed as mean6 S.D., n = 3. *P, 0.05 compared with the vehicle. #P, 0.05 compared with the scrambled siRNA plus atRA-treated group.

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the induction of the genes and processes of interest wasfurther studied using selective RAR and PPAR agonists incomparison to atRA. Although the PPARd agonist GW0742induced PGC1b and NRF1, PGC1a was not changed. Incontrast the RAR pan agonist, TTNPB had no effect onPGC1a, PGC1b, and NRF1 mRNA expression alone or incombination with atRA (Supplemental Fig. 1B). Interestingly,when GW0742 was used together with atRA, the mRNA

induction of PGC1b and NRF1 was greater than with atRAalone, whereas the induction of PGC1a was similar. Togetherthese results suggest that RARs are not involved in theinduction of mitochondrial biogenesis or function by atRA.To further explore the role of specific nuclear receptors in

the increase in mitochondrial biogenesis markers by atRA,RARa, RARb, and PPARd siRNAs were used. The RARa,RARb, and PPARd siRNA decreased the corresponding

Fig. 5. atRA increases mitochondrial biogenesis and ATP production in lipid loaded HepG2 cells and human hepatocytes. (A) Oil Red O staining ofcontrol-treated and lipid-loaded HepG2 cells. Oil Red O staining was measured by quantifying the absorbance at 492 nMwavelength. The measured ODvalue for vehicle treated cells was 1.203 and for oleic acid loaded cells 2.642. (B) Induction of the mRNA expression of key mitochondrial biogenesis genes(PGC1a, PGC1b, and NRF1) by atRA, talarozole, or their combination in the lipid-loaded HepG2 cells. (C) Induction of ATP production by atRA,talarozole (TLZ), or their combination in lipid-loaded HepG2 cells. (D) Induction of SDHA protein expression after treatment of lipid-loaded HepG2 cellswith atRA asmeasured byWestern blotting. The quantification of the replicateWestern blots and a representative blot of triplicate treatments is shown.(E) Induction of the mRNA expression of key mitochondrial biogenesis genes by 1 mM atRA in lipid loaded human hepatocytes from three individualdonors. (F) Induction of ATP production by 1 mM atRA in lipid-loaded human hepatocytes from three individual donors. Results for mRNA studies arepresented as fold change in mRNA abundance relative to vehicle (ethanol). mRNA results, ATP assay, and protein expression results are expressed asmean 6 S.D., n = 3. Comparisons between vehicle (ethanol)- and atRA-treated cells were made by Student’s t test: *P , 0.05 versus vehicle. #P , 0.05versus 1 mM atRA. †P , 0.05 versus 100 nM atRA.

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nuclear receptor mRNA levels by 75, 80, and 60%, respec-tively (Supplemental Fig. 2A). The RARb and PPARd siRNAalso decreased RARb and PPARd protein expression respec-tively by 50–60% in atRA treated cells compared with atRAplus scrambled siRNA treated controls (Supplemental Fig. 2,B–D). In contrast, RARb siRNA had no effect on the induc-tion of PPARd by atRA and PPARd siRNA had no effect on theinduction of RARb by atRA (Supplemental Fig. 2, B–D).RARa and RARb siRNA had no effect on the induction ofPGC1a, PGC1b, and NRF1 mRNA by atRA (Fig. 4A). Incontrast, PPARd siRNA significantly decreased the inductionof PGC1a, PGC1b, and NRF1 mRNA by atRA compared withscrambled siRNA control (Fig. 4A), suggesting that theseeffects of atRA in the HepG2 cells are mediated by PPARd. Inagreement with the role of PPARd in the regulation ofmitochondrial function and biogenesis, PPARd siRNA elim-inated the increase in ATP production by atRA as well as theinduction of SDHA protein expression and increase inb-oxidation (Fig. 4, B–E). PPARd siRNA also significantlydeceased the atRA-mediated induction of mitochondrial DNA(Fig. 4F).Effects of atRA on Mitochondrial Biogenesis Markers

in Models of Human Fatty Liver. Lipid-loaded HepG2cells and primary human hepatocytes were used as a model ofhuman fatty liver. Lipid accumulation was confirmed via OilRed O staining (Fig. 5A). In lipid-loaded HepG2 cells, PGC1a,PGC1b, and NRF1 mRNA were increased two- to threefold

(P, 0.05) in response to atRA treatment (Fig. 5B), a finding inexcellent agreement with the normal HepG2 cells. In lipid-loaded HepG2 cells, 1 mM atRA treatment resulted in a dose-dependent increase in ATP production (Fig. 5C). Similarlyin lipid-loaded HepG2 cells atRA treatment also increasedSDHA protein expression twofold (Fig. 5D). The effect oftalarozole on the potency of atRA as an inducer of mitochon-drial biogenesis genes and ATP production was also similar inlipid-loaded HepG2 cells (Fig. 5, B and C) as in normal HepG2cells. The induction of PGC1a, PGC1b, and NRF1 mRNA andATP production was significantly greater when lipid-loadedHepG2 cells were cotreated with talarozole and atRA thanwith atRA alone at 100 nM (Fig. 5, B and C). In excellentagreement with the lipid-loaded HepG2 cell data and normalhuman hepatocyte experiments, in lipid-loaded human he-patocytes from three donors, the mRNA expression ofPGC1a, PGC1b and NRF1 was increased two- to fourfold(P, 0.05) and ATP production was increased two- to threefoldcompared with vehicle-treated group in each of the threedonors (Fig. 5, E and F).Similar to normal HepG2 cells, atRA significantly increased

the transcription of both RARb and PPARd in lipid-loadedHepG2 cells (Fig. 6A). Interestingly, in lipid-loaded HepG2cells, RARawas also induced by atRA treatment.When siRNAfor each of the nuclear receptors was used in the lipid-loadedHepG2 cells, only PPARd knock down attenuated the atRA-mediated induction of PGC1a, PGC1b, and NRF1 mRNA

Fig. 6. atRA requires PPARd in the reg-ulation of mitochondrial biogenesis inlipid-loaded HepG2 cells. (A) The induc-tion of nuclear receptor mRNA by 1 mMatRA in lipid-loaded HepG2 cells. (B) Theinfluence of RARa siRNA on the inductionof mitochondrial biogenesis genes byatRA. (C) The influence of RARb siRNAon the induction of mitochondrial biogen-esis genes. (D) The influence of PPARdsiRNA on the induction of mitochondrialbiogenesis genes. Results are presented asfold change in mRNA abundance relativeto vehicle (ethanol). mRNA results areexpressed as mean 6 S.D., n = 3, and arerepresentative of at least 3 separate stud-ies. *P , 0.05 versus vehicle. # P , 0.05versus scrambled plus atRA.

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(Fig. 6, B–D). RARb and RARa knock down had no effect onthe induction of PGC1a, PGC1b, and NRF1mRNA by atRA inthe lipid-loaded cells.Effect of Increased atRA Concentrations on the

Expression of Mitochondrial Biogenesis In Vivo inMouse Liver. To evaluate whether the in vitro findings ofthe effects of atRA on hepatic mitochondrial biogenesis inhuman liver models translate to in vivo, mice were treatedwith vehicle or atRA for 5 days, and hepatic mRNA expressionof Pgc1a, Pgc1b, Nrf1, Cpt1a, and Atgl was measured after anovernight fast or after feeding (Fig. 7). atRA treatmentresulted in up to sixfold (P , 0.05) increase in the mRNA ofPgc1a, Pgc1b, Nrf1, and Cpt1a (Fig. 7A) but had no effect onAtgl expression in fasted condition. The induction of Pgc1a,Pgc1b, Nrf1, and Cpt1a was accompanied with increasedmitochondrial DNA and SDHA expression in the mouseliver after atRA treatment in fasted condition (Fig. 7, B–D).

However, 4 hours later, after refeeding of the mice, atRA nolonger affected any of the above mentioned markers (Fig. 7,A–D). The induction of the marker genes of RAR activation,Cyp26a1 and Rarb, wasmuchmoremodest than the inductionof mitochondrial biogenesis marker genes. Rarb was inducedapproximately 2.5-fold (P , 0.05), but there was no signifi-cant increase in Cyp26a1 mRNA (1.7-fold, P . 0.05) afteratRA treatment (Fig. 7E). Unlike mitochondrial biogenesismarkers, there was no difference in Rarb induction betweenfed and fasted groups (Fig. 7), a finding in agreement withdifferent induction mechanisms and mRNA half-lives forthese target genes.To increase the exposure of endogenous atRA in the liver,

male C57BL/6X129 mice were treated with talarozole, aninhibitor of Cyp26, and mRNA expression of key mitochondrialbiogenesis genes was measured. Similar to atRA treatment,talarozole treatment increased mRNA expression of Pgc1b and

Fig. 7. Effect of atRA and talarozoletreatment on mitochondrial biogenesismarkers and fatty acid oxidation genesin vivo in mice. (A) Hepatic mRNA ex-pression of Pgc1a, Pgc1b, Nrf1, Atgl andCpt1a in mice treated with atRA. (B)Quantification of mitochondrial DNA inatRA and vehicle-treated mice after anovernight fast (fasted group) or 4 hourslater after feeding. (C and D) Induction ofliver SDHA protein expression in atRAand vehicle-treated mice as measured byWestern blotting. (C) Quantification ofthe representative Western blot shownin (D). The blot shows analysis of threeindividual mice for each treatment. (E)Hepatic mRNA expression of Rarb andCyp26a1 in atRA- or vehicle-treated mice.(F) Induction of hepatic mitochondrialbiogenesis genes in mice after talarozole(TLZ) treatment compared with vehicle-treatedmice. AllmRNAdataare presentedas fold change in mRNA abundance rela-tive to control mice. b-Actin was used asthe housekeeping gene. For protein data,results are expressed as ratio of SDHA tob-actin. Results are expressed as mean 6S.D., n = 4. Comparisons between groupswere made by Student’s t test: *P , 0.05versus vehicle or control.

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Nrf1 five- to tenfold (P , 0.05), but the fourfold increase inPgc1a mRNA was not significant (P . 0.05; Fig. 7F).

DiscussionTreatment of most cancer cell lines in vitro or neuroblas-

toma and acute promyelocytic leukemia in vivo with RA leadsto apoptosis and cell cycle arrest manifesting the classic RAR-mediated effects of atRA regulating cell proliferation. Inter-estingly, in models of human liver, treatment with atRA hadno effect of cell viability or proliferation at concentrations thattypically lead to cell cycle arrest and apoptosis. This result ledus to evaluate atRA signaling in HepG2 cells using aproteomic approach. The proteomic studies showed thatinstead of cell cycle regulation, atRA treatment predomi-nantly altered the expression of proteins responsible forregulating lipid and fatty acid metabolism in the human livermodels. These findings led to the identification of a novel effectof atRA signaling in the liver. The indication of the proteomicstudy that atRA alters lipid and fatty acid metabolism wasintriguing, because use of retinoids andRA isomers isotretinoin(13-cisRA) and alitretinoin (9-cisRA) in dermatology and incancer treatment is associated with changes in lipid andlipoprotein metabolism, including hypertriglyceridemia, hy-percholesterolemia, increased total cholesterol, and decreasedhigh-density lipoprotein-cholesterol in about one-third of thetreated patients (Lawrence et al., 2001; Staels, 2001; Brelsfordand Beute, 2008; Lilley et al., 2013). The induction of SREBP1mRNA observed in this study after treatment with 10 mMatRA is in agreement with the clinical findings, as the 10 mMatRA treatment closely mimics the RA exposures observedafter retinoid therapy. The 1 mM treatments used in thesubsequent experiments in this study are expected to closelymimic endogenous atRA concentrations in the liver, becausethe measured media concentrations during treatment wereapproximately 100 nM. The low media concentrations mea-sured are likely due to extensive metabolism of atRA inthe cells as reported previously (Topletz et al., 2015) andpartitioning and nonspecific binding of atRA into cells. Strik-ingly, at the concentrations of atRA (100 nM and 1 mM) thatreflect changes in endogenous atRA concentrations, SREBP1mRNA was not increased despite consistent changes in otherproteins and mRNA. This suggests that at different concentra-tions of atRA, different nuclear receptors and signaling pathwaysare activated, leading to different effects on lipid metabolism.In addition to SREBP1 induction, the proteomic study

identified an increase in a complement of proteins thatcontribute to fatty acid oxidation, mitochondrial function, andmitochondrial energy production, leading to a hypothesis thatatRA increases mitochondrial biogenesis and fatty acidb-oxidation in the liver. The subsequent mechanistic studiesshown here consistently support a beneficial regulatory roleof endogenous atRA on lipid homeostasis. The studies showthat atRA increases mitochondrial number (mitochondrialDNA and SDHA expression), mitochondrial function (ATPproduction), and fatty acid b-oxidation (14C palmitate oxida-tion) in normal and fatty human liver. The collective pathwaysand proteins affected by atRA treatment are summarized inFig. 8. It should, however, be noted that the increaseobserved in palmitate oxidation may also be partially dueto increased peroxisomal fatty acid oxidation, and further

studies are needed to characterize the complete effects ofatRA on liver lipid homeostasis.Previous mouse studies suggested that atRA increases fatty

acid oxidation and decreases hepatic lipid accumulation pre-dominantly via RAR-mediated processes. atRA has beenshown to suppress PPARg2 expression and decrease hepaticlipid accumulation in diet-induced obese mice by activatingRARa (Kim et al., 2014). After adenoviral overexpression ofRARb, atRA treatment increased the expression of fatty acidoxidative genes CPT1a and MCAD in mouse liver and inHepG2 cells (Li et al., 2013). atRA also enhanced fatty acidoxidation and ketogenesis via RAR activation and FGF21induction in HepG2 cells (Amengual et al., 2012; Li et al.,2013). Although the data presented in this study in humanhepatocytes, HepG2 cells, and inmice are in agreement with arole of atRA at endogenous concentrations in increasing fattyacid b-oxidation, ATP production, mitochondrial function, andmitochondrial biogenesis, the data do not support a role ofRARs in how atRA regulates these processes. While atRAclearly activates RARs in the HepG2 cells based on theobserved induction of RARb mRNA, the siRNA studies andchemical agonist experiments show that the regulation ofmitochondrial function by atRA is likely mediated by PPARd.

Fig. 8. Mitochondrial biogenesis and fatty acid oxidation pathwaysinduced by atRA in the liver models and a proposed molecular mechanismfor the observed effects. (A) Collective representation of fatty acid metab-olism and mitochondrial function proteins and mRNA that were inducedby atRA in the models of human liver. Green boxes and circles indicateproteins and mRNA that were shown to increase in expression in responseto atRA treatment. (B) Proposed molecular mechanism of atRA regulationof mitochondrial function.

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This conclusion is supported by the fact that only PPARdsiRNA decreased the atRA-mediated induction of mitochon-drial biogenesis genes, mitochondrial DNA content, ATPproduction, and fatty acid oxidation. Importantly, PPARdsilencing had similar consistent effects on all markers ofmitochondrial biogenesis and fatty acid b-oxidation, suggestingthat atRA mediates all of these processes via similar mecha-nism. The proposed mechanism via PPARd is shown in Fig. 8.The in vivo studies in mice are in agreement with thisinterpretation of predominant role of PPARd, because atRAtreatment resulted in only modest effects on the mRNA levelsof RAR target genes CYP26A1 and RARb but had significanteffects on the expression of mitochondrial biogenesis genesthat are believed to be PPARd targets. Yet, it is possible thatthe modest effects on CYP26A1 and RARb mRNA in mice aredue to the short mRNA half-life of these genes and atRAbecause the induction was analyzed 24–28 hours after lastatRA dose. The findings of the role of PPARd in RA signalingare in agreement with previous studies that documented thatcellular retinoic acid binding protein II and fatty acid bindingprotein-5 direct atRA to the nuclear receptors RAR andPPARb/d, respectively, and that the pleotropic effects of atRAare due to activation of both of these receptors (Napoli, 1996;Schug et al., 2007; Berry and Noy, 2012; Bonet et al., 2012;Obrochta et al., 2015). However, we also show that atRAtreatment increases PPARb/d mRNA, and it is possible thatthis induction in part or solely contributes to the observedeffects on mitochondrial biogenesis without atRA bindingdirectly to PPARb/d.Altered vitamin A homeostasis is well documented in

alcoholic liver disease, but recent reports associate alteredvitamin A homeostasis also with NAFLD and steatosis (Ashlaet al., 2010; Kim et al., 2014; Liu et al., 2015; Trasino et al.,2015). For example, human liver biopsies of NAFLD andnonalcoholic steatohepatitis patients demonstrated increasedexpression of the main enzyme metabolizing atRA, CYP26A1,suggesting that atRA is depleted in livers of these patients(Ashla et al., 2010). This is likely deleterious in NAFLD,because we show that atRA increases mitochondrial biogen-esis and fatty acid oxidation in models of fatty liver andNAFLD. Because NAFLD is associated with decreased mito-chondrial function (Sookoian et al., 2010; Aharoni-Simonet al., 2011; Handa et al., 2014), these data suggest thatdecreasing atRA concentrations in NAFLD may contribute todecreasing mitochondrial function and disease progressionand that increasing atRA concentrations in NAFLD may bebeneficial and result in improvement of mitochondrial func-tion and fatty acid oxidation. Such increases in endogenousatRA concentrations could be achieved for example via inhi-bition of the CYP26 enzymes as shown here with talarozoletreatment in HepG2 cells and in mice. In the HepG2 cells,talarozole decreased atRA metabolism increasing atRA con-centrations and thus potentiating the effects of atRA onmitochondrial biogenesis. Similarly, in vivo inmice, talarozoletreatment increased the expression of mitochondrial biogen-esis genes, suggesting that increasing endogenous atRAconcentrations in the liver is sufficient to obtain the effectsobserved in HepG2 cells and human hepatocytes after atRAtreatment. Consistent with the human liver models, atRAtreatment increased the expression of PGC1a, PGC1b, NRF1,and CPT1a under fasting conditions in mice and increasedmitochondrial number and SDHA protein. As expected from

the energy needs of fed mice, after refeeding the mice had lowexpression of PGC1a, PGC1b, NRF1, ATGL, and CPT1acompared with fasted mice, and atRA did not have equivalenteffects on mitochondrial number and function after feeding asin the fasted mice. Yet, it is also possible that because of thelonger time interval between last atRA dose and collection ofsamples in the fed mice compared with fasted, the effects ofatRA were no longer observed in the fed mice. Additionalstudies are needed to fully explore the differences in atRAsignaling under fasted and fed conditions.In summary, this study shows that atRA increases mito-

chondrial biogenesis, fatty acid b-oxidation, expression ofmitochondrial respiratory chain components, and ATP pro-duction in models of healthy and fatty human liver and in vivoin mouse liver, and these effects are predominantly regulatedthrough PPARd signaling instead of RAR activation. More-over, increasing atRA concentration by CYP26 inhibitionpotentiated atRA effects on mitochondrial biogenesis andATP production in cell models and in vivo in mice. Takentogether the increased expression of CYP26A1 in NAFLD andthe observed efficacy of a CYP26 inhibitor in mice and inhuman liver models suggest that CYP26 inhibition would bebeneficial in NAFLD and result in increased mitochondrialfatty acid oxidation and mitochondrial biogenesis.

Acknowledgments

The authors thank Dr. Alan Chait of NORC, University ofWashington, for advice on the fatty acid oxidation assay and FaithStevison, MS, for skillful assistance with retinoid analysis.

Authorship Contributions

Participated in research design: Tripathy, Chapman, Hogarth, andIsoherranen.

Conducted experiments: Tripathy, Chapman, Han, Hogarth, Arnold,Onken, and Kent.

Contributed new reagents or analytical tools: Chapman andGoodlett.

Wrote or contributed to the writing of the manuscript: Tripathy,Chapman, Han, Hogarth, and Isoherranen.

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Address correspondence to: Nina Isoherranen, Department of Pharmaceu-tics, School of Pharmacy, University of Washington, Health Science Building,Room H-272M, Box 357610, Seattle, WA 98195-7610. E-mail: ni2@uw.edu

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