Oxidative Stress Attenuates Lipid Synthesis and Increases Mitochondrial Fatty Acid Oxidation in Hepatoma Cells Infected with Hepatitis C Virus^*

Materials

Fatty acid-free bovine serum albumin (BSA), oleic acid (OA), glycerol, leupeptin, phenylmethanesulfonyl fluoride (PMSF), protease inhibitor mixture, cholesteryl oleate, l-α-phosphatidylcholine, heptane, isopropyl ether, GW7647, α-tocopherol, N-acetylcysteine (NAC), and digitonin were obtained from Sigma. Thin layer chromatography plates (catalog no. WHT4860-820) were from Fisher. Dulbecco's modified Eagle's medium (DMEM), TRIzol, Platinum Quantitative PCR SuperMix-UDG, Lipofectamine 2000, 2′,7′-dichlorodihydrofluorescein diacetate (H₂DCFDA), Click-iT EdU Alexa Fluor 488 flow cytometry kit, and penicillin/streptomycin mixture 2000 were purchased from Invitrogen. OxiSelect hydrogen peroxide/peroxidase assay kit was from Cell Biolabs, Inc. (San Diego). The antibodies were obtained from the following sources: rabbit anti-human CPT1A and rabbit anti-human ACADL polyclonal antibodies were from GeneTex Inc. (Irvine, CA); mouse anti-core C7-50 monoclonal antibody and mouse monoclonal IgG1 (isotype control, ab170190) were from Affinity BioReagents (Rockford, IL); mouse anti-NS5A NE10 monoclonal antibody (NE10) was a gift (Timothy L. Tellinghuisen, Dept. of Infectious Diseases, The Scripps Research Institute); rabbit anti-human PPARα polyclonal antibody was purchased from Applied Biological Materials Inc. (Richmond, British Columbia, Canada); rabbit anti-histone H2B (ab18977) was from Abcam (San Francisco, CA), rabbit anti-AMPKα and anti-phospho AMPKα (Thr-172) monoclonal antibodies (D5A2 and 40H9, respectively), acetyl-CoA carboxylase rabbit monoclonal antibody (C83B10), and phospho-acetyl-CoA carboxylase (Ser-79) (D7D11) rabbit mAb were from Cell Signaling Technology, Inc. (Danvers, MA); anti-human actin (clone C4) mouse monoclonal antibody was from Millipore (Billerica, MA); HRP-conjugated anti-mouse IgG antibody was from MP Biomedical (Solon, OH); HRP-conjugated donkey anti-rabbit IgG was from Jackson ImmunoResearch (West Grove, PA); Alexa Fluor goat anti-rabbit IR dye 680 and goat anti-mouse IR dye 750 were from Invitrogen. Micro BCA protein assay reagent kit was obtained from Pierce; [9,10-³H]oleic acid, [1-¹⁴C]oleic acid, and [³H]acetic acid were obtained from PerkinElmer Life Sciences. Western blotting detection reagents and Hyperfilm^TM ECL were from GE Healthcare. PPRE Cignal^TM Reporter system was from SABiosciences via Qiagen (Toronto, Ontario, Canada). ENLITEN ATP assay system bioluminescence detection kit and Dual-Luciferase reporter assay system were from Promega (Madison, WI). qScript^TM cDNA SuperMix was from Quanta Biosciences (Gaithersburg, MD). All DNA primer synthesis was performed by IDT (Coralville, IA). Cell proliferation kit I (MTT) was from Roche Applied Science.

Cells and Cell Culture

Huh7.5 cells, a mutant line of Huh7 cells that support HCV replication with high efficiency (56, 57), and pJFH-1, used to transfect Huh7.5 cells to generate infectious virus stocks, were generously provided by Dr. Charles M. Rice (Rockefeller University, New York). Unless otherwise indicated, cells were maintained in DMEM containing 10% FBS, 100 units/ml penicillin, and 100 μg/ml streptomycin at 37 °C in humidified air containing 5% CO₂.

Generation of Viral Stocks and Infected Cell Cultures

Viral stocks were collected from Huh7.5 cells transfected with Japanese fulminant hepatitis (JFH)-1 HCV RNA between 3 and 40 days post-transfection essentially as described (53, 58). Infected cell cultures were generated by inoculating low passage Huh7.5 cells with viral stocks at a multiplicity of infection (m.o.i.) = 0.01 infectious virions/cell essentially as described (53, 58). The percentage of HCV-infected cells and the production of viral RNA and infectivity titers were determined periodically by immunocytochemistry with anti-HCV core immunostaining, quantitative RT-PCR, and limiting dilution assays essentially as described (58, 59). No split was necessary after 10 days due to cytopathic effects (53). Instead, these were enrolled into experiments as “infected” cells. “Naive” cells used in experiments were derived from the same Huh7.5 cell stocks as infected cells (58). Sequencing of the viral genome (Molecular Biology Services Unit, Dept. of Biological Sciences, University of Alberta) confirmed the absence of adaptive mutation G451R previously demonstrated to confer enhanced infectivity (53) from all infected cells enrolled into experiments.

Flow Cytometry Analysis

The number of cells progressing through S-phase was determined by flow cytometry using Click-iT EdU Alexa Fluor 488 flow cytometry kit according to the supplied protocol. Flow cytometry was performed on a BD FACSCalibur machine (BD Biosciences) with analysis done using FlowJo software (version 8.7.1).

Cellular Viability

Cellular viability was estimated using cell proliferation kit I (MTT) according to the manufacturer's instructions.

Detection of ROS/RNS

Production of ROS/RNS was monitored using the cell-permeant H₂DCFDA according to the manufacturer's instructions. Upon cleavage of the acetate groups by intracellular esterases and oxidation by intracellular H₂O₂, low molecular weight peroxides, peroxynitrites, and nitric oxide (60–64), the nonfluorescent H₂DCFDA is converted to the highly fluorescent 2′,7′-dichlorofluorescein. Thus, fluorescence intensity is proportional to the amount of oxidants produced by the cells. Cellular H₂O₂ levels were determined using the OxiSelect hydrogen peroxide/peroxidase assay kit according to the manufacturer's instructions. In the presence of HRP, the 10-acetyl-3,7-dihydroxyphenoxazine reagent reacts with H₂O₂ in a 1:1 stoichiometry to produce highly fluorescent resorufin. Fluorescence values are proportional to the H₂O₂ within the samples. The H₂O₂ content in unknown samples was determined by comparison with its respective standard curve.

Quantitative Immunoblot Analysis

Whole cell lysates were obtained by directly lysing cell pellets with RIPA buffer followed by sonication. Post-nuclear supernatants and nuclear extracts were prepared essentially as described (65). Whole cell lysates, post-nuclear supernatants, or nuclear extracts were separated on SDS-polyacrylamide gel, transferred to nitrocellulose membranes, and blocked with 3% BSA in PBS. Membranes were then incubated for 1 h at room temperature with appropriate dilutions of primary antibodies followed by 1:10,000 appropriate IR dye-conjugated IgG secondary antibodies. All immunoreactive bands were quantified using the Odyssey CLx Infrared Imaging System from LI-COR Biosciences (Lincoln, NE).

Malonyl-CoA Assay

Reversed phase high performance liquid chromatography was used for the determination of malonyl-CoA levels. Details of this method are supplied elsewhere (66).

De Novo Lipogenesis

Cells were seeded in 60- mm culture dishes and allowed to adhere for 4 h prior to incubating for 4 h in 2 ml of DMEM containing 10 μCi of [³H]acetate and 100 μm sodium acetate. Cells were washed twice with PBS and collected. Lipids were extracted and resolved by TLC in neutral lipid solvent, and the bands corresponding to triacylglycerol (TAG), free cholesterol (FC), cholesteryl ester (CE), free fatty acids (FFAs), and phospholipids (PL) were scraped and analyzed as described below under “Lipid Analysis.”

Release of ³H₂O from the β-Oxidation of [9,10-³H]Oleic Acid

These experimental conditions were adapted from studies conducted by Hansson et al. (67). Cells were seeded in 60-mm culture dishes and allowed to adhere for 4 h prior to pulse labeling for 4 h with 2 ml of serum-free DMEM containing 5 μCi of [9,10-³H]OA and 0.4 mm OA complexed to 0.4% BSA (pulse) to stimulate neutral lipid synthesis. After 4 h, the pulse media were removed by aspiration and retained for analysis, and cells were washed three times with serum-free DMEM containing 0.4% fatty acid-free BSA and then incubated with 2 ml of serum-free DMEM for an additional 4 h (chase) prior to collecting the chase media for analysis. Pulse and chase media samples were centrifuged at 2,500 × g for 5 min to remove cell debris. Labeled fatty acids were precipitated out from media samples by the addition of 30 μl of 20% BSA and 16 μl of 70% perchloric to 200 μl of culture media. Media were then centrifuged at 25,000 × g for 5 min before an aliquot of the supernatant was counted for radioactivity. The cells were washed with ice-cold PBS, and cell homogenates were prepared in the same buffer, and protein concentration was measured.

¹⁴CO₂ production from the complete β-oxidation of [1-¹⁴C]oleic acid was performed as described previously (68). Briefly, [1-¹⁴C]oleic acid stock (2.5 μCi) bound to 0.4 mm OA, 0.5% BSA in 10 ml DMEM was placed in T150 flasks containing cell cultures and fitted with a center vial with a filter paper moistened with 1 m KOH to absorb CO₂. The flasks were sealed with rubber septa, and the cells were incubated at 37 °C. After 4 h of incubation, the cells were lysed by the injection of 1 ml of perchloric acid through the flask seal, which also results in the liberation of CO₂. Then 0.4 ml of 1 m KOH was added to a center well suspended in the sealed flask. The incubations were continued for a further 1 h at room temperature to allow the CO₂ to be absorbed onto the filter paper and KOH. The filter papers and KOH were removed, added to 4 ml of scintillation fluid, and counted.

Fatty Acid Uptake

Cumulative uptake of oleic acid was measured as described previously (69). Adherent cells were incubated with [9,10-³H]oleic acid (2.5 μCi/ml) mixed with 0.4 mm OA, 0.5% FFA-free BSA at 37 °C for 1, 5, 10, 20, or 30 min. Cells were washed with ice-cold 0.5% BSA in PBS three times and then in PBS alone three times. Cells were lysed with 1 m NaOH, and aliquots were used for scintillation counting and protein determination.

ATP Assay

ATP was measured in cells using the Promega ENLITEN ATP assay system bioluminescence detection kit according to manufacturer's instructions.

PPRE Reporter Assay

PPRE reporter assay was performed according to the Cignal^TM reporter assay handbook. Briefly, cells were transfected with tandem PPRE-firefly luciferase and control Renilla luciferase using Lipofectamine 2000. GW7647 was added 24 h after the start of transfection for an additional 24 h, at which time the cells were harvested and processed according to the Dual-Luciferase reporter assay manufacturer's instructions. Luminescence was measured with the Enspire 2300 multilabel reader (PerkinElmer Life Sciences), and firefly luciferase signal was normalized to internal control Renilla luciferase signal.

Quantitative RT-PCR

Primer sequences used for quantitative RT-PCR are immediately below. Total RNA was isolated using TRIzol reagent and reverse-transcribed with a mixture of random hexamers and oligo(dT) using qScript^TM cDNA SuperMix according to manufacturer's instructions. The transcripts were detected by real time PCR using a Qiagen Rotorgene 3000 instrument. Reaction mixtures contained Platinum Quantitative-PCR SuperMix-UDG with 400 nm concentrations for each primer in a total volume of 20 μl. Following a 3-min initial denaturation at 94 °C, PCR amplification of all genes proceeded for 40 PCR cycles of 95 °C for 20 s, 56 °C for 20 s, and then 72 °C for 20 s. Data analysis was performed with the Corbett Rotor-Gene software using the standard curves method. All transcripts values were normalized to β-actin.

The primers used for quantitative RT-PCR are as follows: CPT1 F, 5′-CGTCTTTTGGGATCCACGATT-3′, and CPT1 R, 5′-TGTGCTGGATGGTGTCTGTCTC-3′; actin F, 5′-CCAACCGCGAGAAGATGA-3′, and actin R, 5′-TCCATCACGATGCCAGTG-3′; VLCAD F, 5′-AAAATTCACAACTTTGGGCTGA-3′, and VLCAD R, 5′-CCCTGGTCCATGTTAGCACT-3′; LCAD F, 5′-TGCCCATGGTATTAGCCTTT-3′, and LCAD R, 5′-CAAAGAATAGTTCTGCGGTATCC-3′; MCAD F, 5′-GCACAAAGAGCTTTGGATGA-3′, and MCAD R, 5′-CAGCATAAATGATATTGCTTGGTG-3′; ACO F, 5′-GGCACCACTGCTCAGAAAG-3′, and ACO R, 5′-AGCCTCGAAGGTGGGTTC-3′; and PPARα F, 5′-ACTGAGAAGCTGTCACCACAGT-3′, and PPARα R, 5′-GCCGAGCTCCAAGCTACTC-3′.

Animal Care and Procedures

Severe combined immunodeficiency disorder, (SCID)/albumin (Alb)-urokinase plasminogen activator (uPA) (SCID/Alb-uPA) mice were housed in a virus- and antigen-free facility supported by the Health Sciences Laboratory Animal Services at the University of Alberta and were cared for in accordance with the Canadian Council on Animal Care guidelines. All experimental protocols involving mice were reviewed and approved by the University of Alberta Health Sciences Animal Welfare Committee. Serum from HCV-infected patients was obtained following informed consent of all donors and with ethics approval from the University of Alberta Faculty of Medicine and Dentistry Research Ethics Review Board. Littermates were transplanted at 10–14 days old with cryopreserved human hepatocytes from a single donor as described previously (70). Replacement of the mouse liver parenchyma with human hepatocytes was estimated by serum levels of human α1 anti-trypsin (hAAT) (71) beginning at 8 weeks post-transplant. Eight transplanted mice having similar and high levels of liver repopulation with human hepatocytes were selected for infection studies and were segregated into naive (n = 4) and infected (n = 4) treatment groups. Four mice were inoculated with infectious serum obtained from a genotype 1A HCV-infected patient. Naive mice did not receive the inoculation. After 2 weeks, all experimental mice were fasted for 16 h prior to blood draw for serum analysis of ketone bodies (see below), hAAT, and HCV RNA essentially as described (72–74).

Plasma Ketone Bodies

Plasma ketone body levels were measured via the cyclic enzymatic method using Autokit Total Ketone Bodies (Wako Chemicals, code no. 415-73301).

Indirect Immunohistochemistry

Engrafted human hepatocytes in paraffin-embedded liver sections were identified using the Super Sensitive ISH detection system (Biogenex, San Ramon, CA) with fluoresceinated Alu probe (catalog no. HK844-2K, Biogenex, San Ramon, CA) according to the manufacturer's instructions. For simultaneous detection of HCV antigens and human hepatocytes, the primary antibody solution provided with the ISH kit was supplemented with anti-core C7–50 and anti-NS5A NE10 antibodies (1:50 of each). For detection of only HCV antigens, the primary antibody solution was replaced with anti-HCV antibodies (core and NS5A) in a PBS solution (containing 1% BSA as carrier, same solution as that provided for anti-HRP mouse monoclonal from Biogenex). Mouse monoclonal IgG1 was used as an isotype control.

Lipid Analysis

Cells were harvested in phosphate-buffered saline (PBS), sonicated on ice, and centrifuged at 1,500 rpm for 5 min in a microcentrifuge to pellet cell debris. Supernatant (cell homogenate) was collected, and lipids were extracted from cell homogenates with chloroform/methanol (2:1, v/v) in the absence (mass determination) or presence (metabolic studies) of nonradioactive lipid carriers (phosphatidylcholine, oleic acid, dioleoylglycerol, trioleoylglycerol, and cholesteryl oleate) as described previously (75). The chloroform phase containing lipids was dried under nitrogen and re-dissolved in a small volume of chloroform for scintillation counting. Alternatively, the re-dissolved lipids were applied to Silica Gel H thin layer chromatography plates. The plates were developed in heptane/isopropyl ether/acetic acid 60:40:4 (by volume) to separate neutral lipids (FA, TG, free cholesterol, and cholesteryl esters). Lipid classes were visualized by exposure to iodine vapor; bands were scraped, and the associated radioactivity was determined by scintillation counting. To analyze lipid mass, lipid extractions were carried out with cell homogenates in the absence of lipid carriers and in the presence of known amounts of tridecanoylglycerol (internal standard). Lipids were processed and analyzed by gas chromatography as described previously (65).

Fatty Acid Composition

For analysis of total lipid-associated FAs, total cell lipids were extracted with chloroform/methanol (2:1, v/v). The organic (chloroform) phases were dried under nitrogen and dissolved in chloroform containing 5 μg of internal standard C17:0. The FAs were converted to FA methyl esters by incubation with 1.5 ml of 6% H₂SO₄ in methanol for 2 h at 80 °C. Resulting FA methyl esters were analyzed by gas chromatography as described previously (75).

Statistical Analysis

Unless stated otherwise, means ± S.E. were determined for a minimum of n = 4 independent experiments. The results were statistically analyzed by the Student's t test or two-way analysis of variance. p values less than 0.05 were considered to be significant.

Other Methods

Protein concentrations were determined using the Pierce micro BCA assay Kit according to the supplied protocol.

HCV-induced Cell Cycle Arrest Is Associated with ROS Accumulation

A mutant line of Huh7 cells (Huh7.5) that supports JFH-1 (genotype 2a) HCV infection with high replication efficiency (56) and exhibits HCV-mediated growth arrest (7, 9, 53, 54) was used to examine whether oxidative stress was associated with this stage of infection. After HCV infection, the percentage of HCV-infected cells and the production of viral RNA and infectious virions were determined periodically by immunocytochemistry with anti-HCV core immunostaining, quantitative RT-PCR, and limiting dilution assays essentially as described (58, 59). Infected cell cultures were enrolled into experiments as “infected cells” at 10 days post-infection (m.o.i. =0.01), when they appeared to grow more slowly relative to their naive counterparts, with more dead cells appearing in the culture media. These cultures were characterized by high levels of HCV infection, producing >10⁷ IU of HCV RNA per mg of cellular RNA with infectivity titers >10⁵ focus-forming units/ml. Importantly, >95% of the infected cells were HCV core-positive (Fig. 1A), thereby minimizing the contribution of non-infected cells present in infected cultures to experimental outcomes (76). The numbers of cells progressing through S-phase were enumerated by flow cytometry (Fig. 1B). This analysis revealed that >80% of the infected cells were growth-arrested, because <20% exhibited evidence of EdU incorporation, as opposed to >75% with naive cells (Fig. 1B). Despite their reduced proliferative capacity, the surviving infected cells remained sufficiently healthy with viabilities >90% (Fig. 1C). H₂DCFDA is sensitive to both H₂O₂ and NO-based radicals and can be oxidized to 2′,7′-dichlorofluorescein by various ROS and RNS, including H₂O₂, although the OxiSelect reagent is specifically sensitive to H₂O₂ (60–64). Intracellular ROS/RNS levels were examined using H₂DCFDA as the detection reagent (Fig. 1D). This analysis revealed that infected cells have ∼5-fold more intracellular ROS/RNS than naive cells, indicating that HCV-induced cell cycle arrest is associated with more oxidative/nitrosative stress. At least some of this increase is due to elevated H₂O₂ production, because infected cells were also found to have higher H₂O₂ levels than naive cells.

HCV-induced Cell Cycle Arrest Is Associated with ROS/RNS-mediated AMPK Activation

Full activation of AMPK requires specific phosphorylation (at Thr-172) within the catalytic domain of the α-subunit by upstream kinases (38), and this signaling system is activated by oxidative stress in various cell types, including liver cells (37). The possibility that AMPK might be activated in infected cells was examined by immunoblot analysis (Fig. 2A). The levels of AMPK were found to be similar between naive and infected cells, but infected cells contained substantially more Thr-172 phosphorylated AMPK (p-AMPK) than naive cells, indicating that more of the AMPK is activated in these cells. To examine the contribution of ROS/RNS to AMPK activation, two different antioxidants were used (Fig. 2B). α-Tocopherol is membrane-soluble and functions primarily as an inhibitor of lipid peroxidation by virtue of its chain-breaking and free radical scavenger actions. In contrast, NAC is a small water-soluble molecule directly providing –SH groups for adduction or oxidation, and it is a precursor of glutathione (77). Pre-treatment of infected cells with either α-tocopherol or NAC led to a reduction in cellular ROS/RNS production by ∼30% (Fig. 2B). The cellular levels of p-AMPK were also reduced by ∼30% when infected cells were pre-treated with α-tocopherol, whereas total AMPK levels remained unchanged (Fig. 2C). This is an indication that oxidative/nitrosative stress plays some role in the activation of AMPK in Huh7.5 cells during HCV-induced cell cycle arrest. Although the levels of p-AMPK were also found to be reduced by ∼30% in infected cells pre-treated with NAC, these reduced levels did not reach statistical significance (data not shown).

HCV-induced Cell Cycle Arrest Is Associated with Attenuated DNL Activity

Acetyl-CoA carboxylase (ACC) catalyzes the conversion of acetyl-CoA to malonyl-CoA, which is the first committed intermediate formed during fatty acid synthesis. Two separate genes encode different ACC isoforms in hepatocytes as follows: ACC1 is localized in the cytosol and is committed to DNL, and ACC2 is anchored to the mitochondrial membrane and is involved in the control of fatty acid oxidation via allosteric inhibition of CPT1, which is also present on the mitochondrial surface, by locally produced malonyl-CoA (38). Both ACC isoforms are phosphorylated by AMPK (at Ser-79 and Ser-220 for ACC1 and ACC2, respectively) in hepatocytes (including Huh7 cells (78)), thereby inhibiting malonyl-CoA production in both compartments (79, 80). Because HCV-mediated growth arrest was associated with activated AMPK, the expression of ACC1/2 and the presence of their AMPK-phosphorylated derivatives were examined by immunoblot analysis using antibodies that were selected based on their ability to detect both ACC1 (265 kDa) and ACC2 (280 kDa) isoforms and their AMPK-phosphorylated derivatives (Fig. 3A). However, only a single ACC species (∼265 kDa) was detected in naive and infected Huh7.5 cells when these antibodies were used. Although this is consistent with what has previously been reported with Huh7.5 cells when the same antibodies were used (81), both ACC isoforms are present in Huh7 cells and have predictable mobility on SDS-PAGE (78). The reason for this disparity is not understood, but because alternative promoter usage and exon splicing can result in several ACC1 and ACC2 variants ranging from 253 to 280 kDa (82), we also performed immunoblot analysis using ACC1- and ACC2-specific antibodies (data not shown). This analysis revealed that both ACC species are present in naive and infected cells, albeit with similar mobility on SDS-PAGE (i.e. ∼265 kDa). Furthermore, there was no apparent difference in the levels of ACC1 or ACC2 between naive and infected cells. In agreement with having higher p-ACC levels, infected cells had just 50% of the malonyl-CoA present in naive cells (Fig. 3B).

HCV replication has been shown to suppress glucose uptake through the down-regulation of cell surface glucose transporters (83). To rule out the possible contribution of altered substrate supply to the drop in malonyl-CoA levels, infected cells were supplied with [³H]acetate. This offers an alternate pathway of producing acetyl-CoA via acetyl-CoA synthetase instead of from pyruvate via the pyruvate dehydrogenase complex. Thus, any change in the incorporation of [³H]acetate into cellular lipids is due to altered catalytic efficiency, for which ACC activity has major influence. As expected with higher p-ACC levels, infected cells incorporated less [³H]acetate into FFA and their esterified derivatives (TAG, PL, and CE) than naive cells (Fig. 4). In addition, infected cells incorporated less [³H]acetate into free cholesterol (FC) than naive cells, indicating that these cells also have reduced cholesterol biosynthetic activity. These results demonstrate unequivocal association between HCV-induced cell cycle arrest and attenuated DNL activity.

HCV-induced Cell Cycle Arrest Is Associated with Enhanced β-Oxidation Activity

A drop in malonyl-CoA is expected to increase β-oxidation activity through the disinhibition of CPT1 (79, 80, 84, 85). To examine this possibility, mitochondrial β-oxidation was monitored by the release of ³H₂O from [9,10-³H]oleic acid (Fig. 5A), an assay for long- and medium-chain fatty acid oxidation (86). By the end of the labeling period (pulse), infected cells had released significantly more ³H₂O into the media than had naive cells. Naive and infected cells released substantially less ³H₂O into the media after oleic acid was withdrawn from the culture media (chase), indicating that exogenous fatty acids enter a pool that is tightly coupled to mitochondrial β-oxidation. However, infected cells continued to release more ³H₂O into the media than did naive cells even after fatty acids were withdrawn from the culture media. This indicates that the mobilization of TAG in preformed storages pools (e.g. lipid droplets) for β-oxidation occurs more readily with the infected cells. Infected cells also released considerably more ¹⁴CO₂ due to the complete β-oxidation of [1-¹⁴C]oleic acid than naive cells (Fig. 5B), indicating that the complete oxidation of long-chain fatty acids is more efficient in the infected cells (86). It is noteworthy that infected and naive cells had taken up similar levels of labeled oleic acid within the first 10 min of the labeling period (Fig. 5C), ruling out the possibility that enhanced β-oxidation observed with the infected cells was due to altered cellular uptake of labeled oleic acid. Reduced levels of labeled oleic acid in infected cells observed at later time points (10–20 min) is likely a reflection of enhanced β-oxidation rather than reduced uptake. As expected with enhanced mitochondrial β-oxidation, infected cells exhibited higher cellular ATP levels than their naive counterparts (Fig. 5D). These results demonstrate an unequivocal association between HCV-induced cell cycle arrest and enhanced β-oxidation activity.

HCV-induced Cell Cycle Arrest Is Associated with Elevated Nuclear PPARα and PPARα Target Gene Expression

PPARα is a type II nuclear receptor localized to the nucleus as part of an inhibitory complex prior to ligand-induced activation (87). Ligand-induced activation of PPARα leads to the enhanced expression of PPARα target genes, many of which have important roles in β-oxidation (including CPT1) (87). However, several kinase-dependent processes also appear to lead to the activation of PPARα, including AMPK (88). Because AMPK was activated in infected cells, PPARα and CPT1 expression levels were examined by immunoblot analysis using nuclear and whole cell extracts, respectively. PPARα was readily detected in nuclear extracts of infected cells (Fig. 6A). By contrast, very little (if any) PPARα was detected in nuclear extracts from naive cells (Fig. 6A). Infected cells contained ∼2-fold more CPT1 than naive cells (Fig. 6B). To examine the activation status of nuclear PPARα, a PPARα-selective agonist (GW7647) was administered to cells following transient transfection with a PPAR-responsive luciferase reporter gene (Fig. 6C). Without GW7647, naive and infected cells exhibited similar activities of the luciferase reporter gene. It is not known which of the three PPAR (α, β/δ, and γ) family members contribute to this baseline activity because each of these is present in liver (87). Furthermore, there is evidence that HCV infection can lead to the increased expression and activation of PPARγ in hepatoma cells (41). Importantly, a 1.5-fold increase in reporter activity was observed with naive cells in response to GW7647, indicating that at least some of the PPARα was unactivated prior to administration of GW7647. Despite having substantially more nuclear PPARα than naive cells, no appreciable change in reporter activity was observed when GW7647 was administered to infected cells. This is an indication that PPARα is constitutively active in infected cells. In support of this, a significant up-regulation of several PPARα gene targets with important roles in fatty acid oxidation was observed with infected cells compared with naive cells (Fig. 6D). These included CPT1, medium- and long-chain acyl-CoA dehydrogenase (MCAD and LCAD, respectively), acyl-CoA oxidase (ACO), and PPARα genes.

HCV Infection Is Associated with Enhanced β-Oxidation in the Humanized Livers of SCID/uPA Mice

The SCID/Alb-uPA mouse model is the first small animal model capable of supporting the entire HCV life cycle. Their use in HCV research and other areas of biomedical research has been extensively reviewed, and they continue to be the preferred model for in vivo studies of HCV infection (89, 90). A genomic study of the host response to HCV infection with this model has uncovered a potential link between disturbances in lipid metabolism and oxidative stress (8). Specifically, mice that showed evidence of HCV-mediated oxidative stress also exhibited induction of several genes with important roles in β-oxidation, including PPARα (8). Steatosis is frequently observed in the histologically integrated human hepatocytes that make up the humanized livers of these mice, independent from HCV infection (Fig. 7A). This has been attributed to enhanced DNL arising from an inherent deficiency in this mouse model for human growth hormone (91). Although this is expected to limit interpretations regarding the impact of HCV infection on DNL, we considered using this model to examine hepatic β-oxidation during HCV infection because in the starved state, hepatic β-oxidation results in the production of ketone bodies, which in turn are used as an energy source, particularly by the brain (92). Eight mice (111–115 days old) that were transplanted with donor-matched cryopreserved human hepatocytes (10–14 days of age) and having similar high levels of liver repopulation with human hepatocytes (as determined by serum-based ELISA for hAAT at weaning) were selected for infection studies and then segregated into naive (n = 4 mice) and infected (n = 4 mice) treatment groups. For the infected group, mice were inoculated with serum obtained from a genotype 1A HCV-infected patient. Naive mice did not receive the inoculation. At 2 weeks post-inoculation, all experimental mice were fasted for 16 h prior to blood draw for serum analysis of ketone bodies, hAAT, and HCV RNA titers and liver sampling for detection of HCV antigens (core and NS5A) using immunohistochemistry (Fig. 7B). Following 16 h of fasting, the circulating levels of ketone bodies were found to be ∼2-fold higher in infected mice versus naive mice (Fig. 7B). Hepatocytes expressing HCV-specific antigen have been detected in liver biopsies previously using immunohistochemistry or fluorescence microscopy, but HCV antigens are recognized by either technique only at or near the limits of sensitivity (93–96). Using immunohistochemistry, we identified cells that labeled positively for HCV antigens (core and NS5A) in the livers from all infected mice. This diffuse and weak labeling was localized to histologically integrated human hepatocytes (brown nuclei) and was absent when immunohistochemistry was performed using an isotype control antibody. A more sensitive method, with larger numbers of mice, is required to examine whether there is a correlation between the proportion of HCV-infected human hepatocytes in the chimeric livers of SCID/Alb-uPA mice and the production of ketone bodies in the starved state. The results of this experiment provide encouraging in vivo evidence that hepatic mitochondrial β-oxidation is enhanced during HCV infection.

Cellular Lipids Accumulate in Huh7.5 Cells during HCV-induced Cell Cycle Arrest

HCV infection is associated with the hepatic accumulation of lipids, and this feature contributes to the development of hepatic steatosis in the clinical setting and to various aspects of the viral life cycle in vitro (41). Because the combination of attenuated DNL and enhanced β-oxidation is not conducive to lipid accumulation, cellular lipid levels were investigated in naive versus infected cells. Cellular levels of TAG, PL, CE, and FC were determined by gas chromatography (Fig. 8A). Surprisingly, each of these lipid classes was significantly elevated in infected cells when compared with naive cells. A comprehensive fatty acid profile of total lipids was also obtained (Table 1). This analysis revealed significant alterations in the fatty acid composition of cellular lipids in infected versus naive cells (boldface). The ratios of specific fatty acids, typically regarded as indices of desaturation/elongation (97), were also determined (Fig. 8B). Relative to naive cells, infected cells had higher 18:0/16:0 and 20:4/18:2 ratios indicating increases in elongase and Δ6 desaturase activities, respectively. The lower 18:1/18:0 ratio observed with infected cells is an indication of reduced stearoyl-CoA desaturase activity. Monounsaturated oleic acid (18:1) and saturated palmitic and stearic acids (16:0 and 18:0) were the major cellular fatty acids, accounting for ∼70% of the fatty acids present in naive and infected cells. Compared with naive cells, the fatty acid composition of infected cells was characterized by slight increases in saturated and monounsaturated fatty acids, respectively, and a larger increase in PUFAs (∼2-fold) (Fig. 8C).