Hepatitis C Virus Enhances Lysosome‐Associated Membrane Protein 2 A Transcription Through Nuclear Factor Erythroid 2‐Related Factor 2 to Support Viral Replication

ABSTRACT

Hepatitis C virus (HCV) establishes persistent infection by rewiring host stress‐response pathways. Chaperone‐mediated autophagy (CMA) contributes to HCV replication, but it remains unclear whether HCV regulates lysosome‐associated membrane protein 2 A (LAMP‐2A), the rate‐limiting receptor for CMA. Here, we examined LAMP‐2A regulation in HCV‐infected Huh‐7.5 cells. HCV infection increased LAMP‐2A promoter activity, mRNA, and protein abundance, indicating transcriptional upregulation. Among candidate stress‐responsive transcription factors, nuclear factor erythroid 2‐related factor 2 (NRF2), hypoxia‐inducible factor 1α (HIF‐1α), and nuclear factor of activated T cells 1 (NFAT1) were elevated in infected cells. However, promoter mutagenesis identified NRF2 as the principal direct regulator. Mutation of the NRF2‐responsive antioxidant response element markedly reduced basal and HCV‐induced LAMP‐2A promoter activity. Chromatin immunoprecipitation assays revealed NRF2 association with the LAMP‐2A promoter, and HCV infection increased nuclear accumulation and Ser40 phosphorylation of NRF2. Functionally, shRNA‐mediated knockdown of LAMP‐2A reduced intracellular HCV RNA and protein levels. These findings identify an NRF2‐LAMP‐2A regulatory axis engaged during HCV infection and support a model in which HCV upregulates LAMP‐2A to establish a cellular environment favorable for viral replication.

Abbreviations

AREs

Antioxidant response elements

CMA

Chaperone‐mediated autophagy

DGAT1

O‐acyltransferase 1

dpi

Days post‐infection

eMI

Endosomal microautophagy

FFU

Focus‐forming units

HCV

Hepatitis C virus

HIF‐1α

Hypoxia‐inducible factor 1α

HNF‐1α

Hepatocyte nuclear factor 1 alpha

HSC70

Heat shock cognate protein of 70 kDa

KPNA2

Karyopherin subunit alpha 2

LAMP‐2A

Lysosome‐associated membrane protein 2 A

MOI

Multiplicity of infection

NFAT1

Nuclear factor of activated T cells 1

NRF2

Nuclear factor erythroid 2‐related factor 2

TBK1

TANK‐binding kinase 1

1. Introduction

Hepatitis C virus (HCV) is a positive‐sense RNA virus belonging to the genus Hepacivirus within the family Flaviviridae and establishes chronic infection, remaining a major cause of liver disease worldwide [1, 2]. Its genome, approximately 9.6 kb in length, is translated into a single large precursor polyprotein of about 3,000 amino acids. This precursor is subsequently processed by both viral‐encoded proteases and host proteolytic enzymes to generate the mature structural components (core, E1, and E2) as well as the non‐structural proteins (p7, NS2, NS3, NS4A, NS5A, and NS5B) that are required for viral replication [3]. Successful viral replication depends on extensive remodeling of host cellular pathways, including stress responses and intracellular degradation systems [4, 5, 6]. Increasing evidence suggests that HCV modulates autophagy‐related pathways to establish a favorable intracellular environment for viral persistence [7, 8].

Chaperone‐mediated autophagy (CMA) is a selective form of autophagy that mediates the lysosome‐dependent degradation of specific cytosolic proteins and plays a critical role in cellular quality control [9]. Unlike macroautophagy, CMA does not involve vesicle formation but instead directly translocates substrate proteins across the lysosomal membrane. In this pathway, target proteins containing a pentapeptide KFERQ‐like motif are recognized by the cytosolic chaperone, heat shock cognate protein of 70 kDa (HSC70), which delivers the substrate proteins to the lysosomal surface. The substrate proteins then bind to the lysosomal membrane receptor, lysosome‐associated membrane protein 2 A (LAMP‐2A), the rate‐limiting component of CMA [9, 10, 11]. Upon substrate binding, LAMP‐2A undergoes multimerization to form a translocation complex, allowing the unfolded substrate protein to cross the lysosomal membrane. After translocation into the lysosomal lumen, the substrate is rapidly degraded by resident lysosomal proteases. CMA activity is tightly regulated at multiple levels, including LAMP‐2A expression, stability, and assembly dynamics [12]. This pathway is upregulated under stress conditions such as nutrient deprivation, oxidative stress, and hypoxia, thereby contributing to cellular adaptation and survival [13]. Dysregulation of CMA has been associated with aging and various pathological conditions, including cancer, neurodegenerative diseases, and metabolic disorders. Furthermore, emerging evidence suggests that certain viruses exploit or modulate CMA components to facilitate their replication and persistence, highlighting the importance of CMA in virus‐host interactions [11, 14].

In our previous studies, we demonstrated that HCV NS5A stimulates HSC70‐dependent selective lysosomal degradation pathways, including CMA‐mediated degradation of hepatocyte nuclear factor 1 alpha (HNF‐1α) [15, 16, 17, 18] and endosomal microautophagy (eMI)‐mediated degradation of diacylglycerol O‐acyltransferase 1 (DGAT1) [19]. These findings suggest that HCV exploits HSC70‐dependent lysosomal degradation pathways to remodel host metabolic processes and cell homeostasis.

Despite these advances, the upstream regulatory mechanisms by which HCV controls selective lysosomal degradation pathways remain incompletely understood. It remains unclear whether HCV modulates the expression of LAMP‐2A, and how such regulation is induced at the transcriptional level. Given that CMA activity is closely associated with cellular stress responses, redox imbalance, and metabolic adaptation, virus‐induced stress signaling pathways may contribute to the transcriptional regulation of LAMP‐2A during infection. However, the molecular mechanisms underlying the link between viral infection and CMA activation remain to be clarified.

In this study, we investigated whether HCV regulates LAMP‐2A expression and elucidated the transcriptional mechanisms responsible for its induction. By clarifying how HCV regulates LAMP‐2A expression, we sought to define a mechanistic link between virus‐induced stress signaling and CMA‐related lysosomal regulation.

2. Materials and Methods

2.1. Cell Culture and Viruses

The human hepatoma cell line Huh‐7.5 was obtained from Dr. Charles M. Rice (The Rockefeller University, New York, NY, USA) [20]. Cells were maintained in high‐glucose Dulbecco's modified Eagle's medium (DMEM) containing l‐glutamine and phenol red (Fujifilm Wako Pure Chemical Industries, Osaka, Japan) supplemented with 10% heat‐inactivated fetal bovine serum (Biowest, Nuaillé, France), 50 IU/mL penicillin, 50 μg/mL streptomycin (Gibco, Grand Island, NY, USA), and 0.1 mM non‐essential amino acids (Invitrogen, Carlsbad, CA, USA). Cells were cultured at 37°C in a humidified atmosphere containing 5% CO₂. Phosphate‐buffered saline without calcium and magnesium (PBS[‐]) (Nissui, Tokyo, Japan) was used for washing cells. Plasmid transfections were performed using FuGENE 6 transfection reagent (Promega, Madison, WI, USA) according to the manufacturer's instructions. The pFL‐J6/JFH1 plasmid, encoding the full‐length genome of the chimeric hepatitis C virus genotype 2a strain J6/JFH1, was obtained from Dr. Charles M. Rice [21]. Full‐length HCV genomic RNA was synthesized in vitro from linearized pFL‐J6/JFH1 and introduced into Huh‐7.5 cells by electroporation as described previously [22, 23]. Virus‐containing culture supernatants were collected, clarified by centrifugation, and stored at −80°C until use. Infectious titers were determined by an immunofluorescence‐based focus‐forming assay as previously described. Briefly, serial 10‐fold dilutions of viral supernatants were used to infect Huh‐7.5 cells seeded in 24‐well plates (2 × 10⁵ cells per well). At 24 h post‐infection, cells were fixed and subjected to immunofluorescence staining using an anti‐HCV core monoclonal antibody. HCV‐positive foci were counted microscopically, and viral titers were expressed as focus‐forming units per milliliter (FFU/mL). The multiplicity of infection (MOI) used in subsequent experiments was calculated based on these titers. Unless otherwise indicated, HCV infection was performed at an MOI of 1. For the immunofluorescence analysis in Figure 5A, an MOI of 2 was used to increase the proportion of infected cells and facilitate visualization of NRF2 intracellular localization.

Figure 5.

HCV infection promotes nuclear translocation and phosphorylation of NRF2. (A, upper panel) Huh‐7.5 cells were transfected with pCAG‐FLAG‐NRF2 plasmid and mock‐infected or infected with HCV J6/JFH1 at multiplicity of infection (MOI) of 2. HCV infection was performed at an MOI of 2 in this experiment to facilitate visualization of NRF2 localization in infected cells. At 24 h post‐infection, the cells were fixed and subjected to immunofluorescence staining with anti‐FLAG mouse monoclonal antibody followed by Alexa Fluor 594‐conjugated goat anti‐mouse IgG (red) to detect exogenous NRF2 and anti‐NS5A Rabbit polyclonal antibody followed by Alexa Fluor 488‐conjugated goat anti‐rabbit IgG (green) to identify HCV‐infected cells. Nuclei were counterstained with Hoechst 33342 (blue). Scale bar: 20 μm. Representative images from three independent experiments are shown. (A, lower panel) Quantification of NRF2 subcellular localization. The proportion of cells showing cytoplasmic plus nuclear or nuclear‐only NRF2 signal was determined by visual scoring of at least 50 cells per condition across 15 fields of view per coverslip. Grey bars: Mock + FLAG‐NRF2; Black bars: HCV + FLAG‐NRF2. Data represent mean ± SEM from three independent experiments. Statistical significance was determined by Student's t‐test. *p < 0.05; **p < 0.01. (B, C) Huh7.5 cells mock‐infected or infected with HCV J6/JFH1 at an MOI of 1 and transfected with pCAG‐FLAG‐NRF2. At 3 days after infection, the cells were harvested and subjected to subcellular fractionation. Cytoplasmic (B) and nuclear (C) fractions were subjected to immunoblot analysis with antibodies against NRF2, phopho‐NRF2 (Ser40), NS5A, IκBα (cytoplasmic fraction marker), and Histone H3 (nuclear fraction marker). Band intensities were quantified by densitometric analysis using ImageJ. Numbers below NRF2 and p‐NRF2 panels indicate quantification of band intensities normalized to IκBα (cytoplasmic fraction) or Histone H3 (nuclear fraction), relative to the mock‐infected control. Representative blot images from three independent experiments are shown.

2.2. Expression Plasmids

Total RNA was extracted from Huh‐7.5 cells and reverse‐transcribed into cDNA. The cDNA fragment was amplified by PCR using gene‐specific primers (LAMP‐2A forward, 5′‐AAGCGGCCGCACCATGGTGTGCTTCCGCCTCTTC‐3′; LAMP‐2A reverse, 5′‐TTGCGGCCGCTTACTTATCGTCGTCATCCTTGTAATCAAATTGCTCATATCCAGCATG‐3′, HIF‐1α forward, 5′‐TCGAGCTCAGCGGCCGCCATGGAGGGCGCCGGCGGC‐3′; HIF‐1α reverse, 5′‐AGTGAATTCGCGGCCGCTCAGTTAACTTGATCCAA‐3′, NFAT1 forward, 5′‐TCGAGCTCAGCGGCCGCCATGAACGCCCCCGAGCGG‐3′; NFAT1 reverse, 5′‐AGTGAATTCGCGGCCGCTCATAATATGTTTTGTAT‐3′) and cloned into the indicated expression vector. The PCR product was purified, digested with Not I, and inserted into the Not I site of the pCAG or pCAG‐FLAG expression vector using the In‐Fusion HD Cloning Kit (Takara Bio USA, Inc., 639649) according to the manufacturer's instructions. The resulting construct encodes LAMP‐2A fused to a C‐terminal FLAG tag. The pCAG‐FLAG‐NRF2 plasmid was as previously described [24]. The human LAMP‐2A promoter region (nt −1077 to +87 relative to the transcription start site) was amplified from human genomic DNA using the following primers: forward, 5′‐CTAACTGGCCGGTACCTGCATTAGCGTGAACAGGCA‐3′; reverse, 5′‐TCTTGATATCCTCGAGGCACTGATGACCACCGAC‐3′. The amplified fragment was digested with Kpn I and Xho I and cloned into the corresponding sites of the pGL4.10‐basic luciferase reporter vector (Promega). Putative NRF2‐ and HIF‐1α‐binding sites within the LAMP‐2A promoter were predicted using bioinformatics analysis. The NRF2‐binding site mutant construct was generated by fusion PCR using the following primers: forward, 5′‐CCCCTAGTCTTACACCTCATACTGAAGCGCCGA‐3′; reverse, 5′‐TCGGCGCTTCAGTATGAGGTGTAAGACTAGGGG‐3′. The HIF‐1α‐binding site mutant construct was generated similarly using primers: forward, 5′‐ATTTGACGCATGCACCAATATGTTTATTGCAGC‐3′; reverse, 5′‐GCTGCAATAAACATATTGGTGCATGCGTCAAAT‐3′. All constructs were verified by DNA sequencing (Eurofins Genomics, Tokyo, Japan).

2.3. Antibodies and Reagents

The mouse monoclonal antibodies (mAbs) used in this study were anti‐NRF2 (A‐10) mAb (sc‐365949; Santa Cruz Biotechnology, Dallas, TX, USA), anti‐HIF‐1α (28b) mAb (sc‐13515; Santa Cruz Biotechnology), anti‐NFATc2(NFAT1)(4G6‐G5) mAb (sc‐7296; Santa Cruz Biotechnology), anti‐HCV core mAb (clone 2H9) [22], anti‐β‐actin mAb (A‐5441; Sigma‐Aldrich, St. Louis, MO, USA), anti‐Histone H3 (1G1) mAb (sc‐517576; Santa Cruz Biotechnology). The rabbit polyclonal antibodies (pAbs) used in this study were anti‐LAMP‐2A pAb (ab125068; Abcam, Cambridge, UK), anti‐NRF2 mAb (phospho S40) antibody [EP1809Y] (ab76026; Abcam), anti‐NS5A (2914‐1) pAb (a kind gift from T. Wakita, Japan Institute for Health Security [JIHS], Tokyo, Japan) and anti‐IκBα pAb (sc‐371; Santa Cruz). Horseradish peroxidase (HRP)‐conjugated anti‐mouse IgG (7076S; Cell Signaling Technology, Danvers, MA, USA) and HRP‐conjugated anti‐rabbit IgG (7074S; Cell Signaling Technology) were used as secondary antibodies.

2.4. Immunoblot Analysis

Immunoblot analysis was performed as described previously [25, 26]. Cell lysates were resolved on 10% or 15% sodium dodecyl sulfate‐polyacrylamide gel electrophoresis (SDS‐PAGE) and transferred to 0.45 μm polyvinylidene difluoride membrane (PVDF) Immobilon‐P (Millipore, Billerica, MA, USA). Membranes were probed with primary antibodies, followed by horseradish peroxidase (HRP)‐conjugated secondary antibody. Immunoreactive bands were visualized using Amersham enhanced chemiluminescence (ECL) western blotting detection reagents (Cytiva, Marlborough, MA, USA). Band intensities were quantified using ImageJ software (version 1.54r).

2.5. Generation of Stable Knockdown Cells

Short hairpin RNA (shRNA) targeting human LAMP‐2A and a non‐targeting scramble control were designed with the following sequences: shLAMP‐2A, 5′‐GGCAGGAGTACTTATTCTA‐3′; shScramble, 5′‐GGACATCGACGGCTTTATA‐3′. Complementary oligonucleotides encoding each shRNA were synthesized, annealed, and cloned into the pSilencer 2.1‐U6 hygro vector (Ambion, Austin, TX, USA) according to the manufacturer's instructions [27]. Huh‐7.5 cells were transfected with the indicated shRNA plasmids using FuGENE 6 reagent (Promega). Stable knockdown cell lines were generated by selection with hygromycin B (200 μg/mL; Nacalai Tesque, Kyoto, Japan). Drug‐resistant colonies were pooled and maintained under continuous hygromycin selection. Knockdown efficiency was confirmed by immunoblotting.

2.6. Quantitative Real‐Time RT‐PCR

Total RNA was extracted using the ReliaPrep RNA Cell Miniprep System (Promega) according to the manufacturer's instructions. First‐strand cDNA was synthesized using the GoScript Reverse Transcription System (Promega). Quantitative real‐time PCR was performed on a StepOnePlus Real‐Time PCR System (Applied Biosystems, Foster City, CA, USA) using TB Green Premix Ex Taq II (Tli RNaseH Plus) (Takara Bio, Shiga, Japan) with SYBR Green chemistry. Relative mRNA expression levels were normalized to human GAPDH. HCV RNA levels were quantified using J6/JFH1‐specific primers. The primer sequences were as follows: LAMP‐2A (forward: 5′‐CACAAGGAAAGTATTCTACAGCTCA‐3′; reverse: 5′‐CAGCATGATGGTGCTTGAGAC‐3′), J6/JFH1 (forward: 5′‐AGACGTATTGAGGTCCATGC‐3′; reverse: 5′‐CCGCAGCGACGGTGCTGATAG‐3′), and GAPDH (forward: 5′‐GCCATCAATGACCCCTTCATT‐3′; reverse: 5′‐TCTCGCTCCTGGAAGATGG‐3′).

2.7. Dual‐Luciferase Reporter Assay

Huh‐7.5 cells were seeded in 24‐well plates and transiently co‐transfected with wild‐type or mutant LAMP‐2A promoter reporter plasmids (pGL4.10[luc2]) together with the Renilla luciferase control plasmid pRL‐TK (Promega). At 48 h post‐transfection, cells were harvested and lysed, and luciferase activities were measured using the Dual‐Luciferase Reporter Assay System (Promega) according to the manufacturer's instructions. Firefly and Renilla luciferase activities were quantified using a GloMax 96 Microplate Luminometer (Promega). Firefly luciferase activity was normalized to Renilla luciferase activity for each sample. All experiments were performed in triplicate.

2.8. Chromatin Immunoprecipitation (ChIP) Assay

ChIP assays were performed using the SimpleChIP Plus Enzymatic Chromatin IP Kit (Agarose Beads) (Cell Signaling Technology) according to the manufacturer's instructions [24]. Huh7.5 cells were transfected with pCAG‐FLAG‐NRF2 or the empty vector as indicated. At 48 h post‐transfection, cells were crosslinked with 1% formaldehyde for 10 min at room temperature and quenched with glycine. Following cell lysis and chromatin digestion, immunoprecipitation was carried out using a rabbit anti‐NRF2 monoclonal antibody (Cell Signaling Technology). Normal rabbit IgG served as a negative control. After reversal of crosslinks and DNA purification, the enriched DNA fragments were analyzed by quantitative real‐time PCR using primers specific for the LAMP‐2A promoter region (forward: 5′‐GCGGAGATTGGCTGTAAGCA‐3′; reverse: 5′‐AGCCTTGCAAAAGCCAGGAA‐3′).

2.9. Immunofluorescence Staining

Huh‐7.5 cells grown on glass coverslips were fixed with 4% paraformaldehyde for 15 min at room temperature and permeabilized with 0.1% Triton X‐100 in PBS for 15 min. After being washed with PBS, cells were blocked with 1% bovine serum albumin (BSA) (Nacalai Tesque) in PBS for 1 h. Cells were then incubated for 1 h at room temperature with a mouse monoclonal anti‐FLAG antibody and a rabbit polyclonal anti‐NS5A antibody diluted in 1% BSA in PBS. After three washes with PBS, cells were incubated for 1 h with Alexa Fluor 594‐conjugated anti‐mouse IgG (Invitrogen) and Alexa Fluor 488‐conjugated anti‐rabbit IgG (Invitrogen) diluted in blocking buffer. After four additional washes with PBS, coverslips were mounted onto glass slides and images were acquired using a confocal laser scanning microscope (LSM 700) (Carl Zeiss, Oberkochen, Germany).

2.10. Cell Fractionation Assay

Cytoplasmic and nuclear fractions were prepared from HCV‐infected and mock‐infected Huh‐7.5 cells using the NE‐PER Nuclear and Cytoplasmic Extraction Reagents (Thermo Fisher Scientific, Waltham, MA, USA) according to the manufacturer's instructions [25, 28]. Protease inhibitors were included in all extraction buffers. The resulting fractions were subjected to immunoblot analysis. The purity of cytoplasmic and nuclear extracts was verified using IκBα and Histone H3 as cytoplasmic and nuclear marker proteins, respectively.

2.11. Statistical Analysis

All data are presented as mean ± standard error of the mean (SEM) from at least three independent experiments. Statistical analyses were performed using GraphPad Prism 9 (GraphPad Software, San Diego, CA, USA). Comparisons between two groups were conducted using a two‐tailed Student's t‐test. For multiple‐group comparisons, one‐way or two‐way analysis of variance (ANOVA) was applied as appropriate. A p value < 0.05 was considered statistically significant. Statistical significance is indicated as follows: *p < 0.05; **p < 0.01; ***p < 0.001. ns, not significant.

3. Results

3.1. HCV Infection Induces LAMP‐2A Expression at the Transcriptional and Protein Levels

To determine whether HCV infection regulates LAMP‐2A expression, we first examined LAMP‐2A promoter activity using a luciferase reporter assay. HCV infection significantly increased LAMP‐2A promoter activity compared with mock‐infected cells at 2, 4, and 6 days post‐infection (Figure 1A). Consistent with the enhanced promoter activity, quantitative reverse transcription (RT)‐PCR analysis revealed that LAMP‐2A mRNA levels were markedly upregulated following HCV infection, particularly at 4 and 6 days post‐infection (Figure 1B). Furthermore, Western blot analysis demonstrated that LAMP‐2A protein levels were also increased in HCV‐infected cells compared with mock‐infected cells (Figure 1C, first panel, lanes 4 and 6). These results indicate that HCV infection induces LAMP‐2A expression at both the transcriptional and protein levels.

Figure 1.

HCV infection enhances LAMP‐2A expression at the transcriptional and protein levels. (A) Huh‐7.5 cells were transfected with a LAMP‐2A promoter‐driven luciferase reporter and subsequently infected with HCV J6/JFH1 at a multiplicity of infection (MOI) of 1. Cells were cultured and harvested at 2, 4 and 6 days post‐infection (dpi) and subjected to a dual‐luciferase reporter assay using a full‐length LAMP‐2A promoter construct (−1077 to +87 bp). Firefly luciferase activity was normalized to Renilla luciferase activity. Data represent mean ± SEM from three independent experiments. The value for day 2 in the mock‐infected group was arbitrarily expressed as 1.0. Statistical significance was determined by Student's t‐test. **p < 0.01; ***p < 0.001. (B) LAMP‐2A mRNA levels were quantified by quantitative RT‐PCR and normalized to GAPDH mRNA. The value for day 2 in the mock‐infected group was arbitrarily expressed as 1.0. Data represent mean ± SEM from three independent experiments. Statistical significance was determined by Student's t‐test. **p < 0.01; ***p < 0.001. (C) Cell lysates were subjected to immunoblot analysis with antibodies against LAMP‐2A, NS5A (as a marker for HCV infection), and β‐actin (loading control). Representative blot images from three independent experiments are shown. Band intensities were quantified by densitometric analysis using ImageJ. Numbers below each panel indicate quantification of band intensities relative to β‐actin, normalized to the mock‐infected control at the corresponding time point.

3.2. LAMP‐2A Supports HCV RNA Accumulation and Protein Expression

To investigate whether LAMP‐2A contributes to HCV replication, we established Huh‐7.5 cells stably expressing shLAMP‐2A or shScramble control and confirmed efficient knockdown of LAMP‐2A (Figure 2A). After HCV infection, intracellular HCV RNA levels were significantly reduced in shLAMP‐2A cells at all examined time points (Figure 2B). Consistently, HCV core and NS5A protein levels were markedly decreased by LAMP‐2A knockdown (Figure 2C, second and third panel, lane 4, 6, 8). These results indicate that LAMP‐2A supports intracellular HCV RNA accumulation and viral protein expression.

Figure 2.

LAMP‐2A supports HCV RNA accumulation and viral protein expression. (A) LAMP‐2A mRNA levels were quantified by quantitative RT‐PCR and normalized to GAPDH in stable Huh‐7.5 cells expressing shLAMP‐2A or shScramble (control), confirming efficient knockdown of LAMP‐2A at 12, 24, 36 and 48 h post‐infection with HCV (MOI = 1). Data represent mean ± SEM from three independent experiments. Statistical significance was determined by Student's t‐test. *p < 0.05; **p < 0.01. (B) Intracellular HCV RNA levels were quantified by quantitative RT‐PCR and normalized to GAPDH. Data are expressed as fold change relative to the shScramble control at 12 h post‐infection and represent mean ± SEM from three independent experiments. Statistical significance was determined by Student's t‐test. **p < 0.01. (C) Cell lysates were subjected to immunoblot analysis with antibodies against LAMP‐2A, NS5A, core and β‐actin. Lanes 1 and 2: 12 h post‐infection (shScramble and shLAMP‐2A); Lanes 3 and 4: 24 h post‐infection (shScramble and shLAMP‐2A); Lanes 5 and 6: 36 h post‐infection (shScramble and shLAMP‐2A); Lanes 7 and 8: 48 h post‐infection (shScramble and shLAMP‐2A). Numbers below each panel indicate quantification of band intensities relative to β‐actin, normalized to the shScramble control at the corresponding time point. Representative blot images from three independent experiments are shown.

3.3. HCV Infection Upregulates Transcription Factors and Enhances LAMP‐2A Promoter Activity

Previous reports indicate that LAMP‐2A transcription can be regulated by stress‐responsive transcription factors, including NRF2, HIF‐1α, and NFAT1 [29, 30, 31, 32]. We therefore examined whether these transcription factors are altered during HCV infection and contribute to LAMP‐2A induction (Figure 3A). Immunoblot analysis revealed that HCV infection increased the protein levels of NRF2, HIF‐1α, and NFAT1 compared with mock‐infected cells at 4 and 6 dpi (Figure 3B, first, second, and third panel, lanes 2 and 4). To assess their functional relevance, we performed promoter‐reporter assays following transcription factor overexpression. NRF2 markedly enhanced LAMP‐2A promoter activity, HIF‐1α showed a modest stimulatory effect, whereas NFAT1 did not significantly alter promoter activity under our experimental conditions. (Figure 3C). These results suggest that HCV infection may enhance LAMP‐2A transcription through the activation of transcription factors NRF2 or HIF‐1α.

Figure 3.

HCV infection upregulates NRF2 and HIF‐1α and enhances LAMP‐2A promoter activity. (A) Schematic representation of predicted transcription factor‐binding motifs in the human LAMP‐2A promoter. Putative NRF2, HIF‐1α, and NFAT1 motifs are shown based on sequence analysis; functional promoter activation was evaluated experimentally in Figure 3C. (B) Huh‐7.5 cells were infected with HCV J6/JFH1 at a multiplicity of infection (MOI) of 1 or mock‐infected. The cells were cultured and harvested at 4 and 6 days post infection (dpi). Cell lysates were subjected to immunoblot analysis with antibodies against NRF2, HIF‐1α, NFAT1, NS5A (HCV infection marker), and β‐actin (loading control). Representative blot images from three independent experiments are shown. (C) Luciferase reporter assay of the LAMP‐2A promoter after overexpression of the indicated transcription factors. Huh‐7.5 cells were infected with HCV J6/JFH1 at a MOI of 1 and co‐transfected with a LAMP‐2A promoter luciferase reporter together with expression plasmids encoding NRF2, HIF‐1α, or NFAT1, or an empty vector (−) as a control. LAMP‐2A promoter activity was measured by dual‐luciferase assays and normalized to Renilla luciferase activity. At 48 h post‐transfection, luciferase activities were measured. Gray bars: Mock‐infected; Black bars: HCV‐infected. Data represent mean ± SEM from three independent experiments. Statistical analysis was carried out using the Student's t‐test. **p < 0.01. NRF2 strongly increased promoter activity, HIF‐1α showed a weaker effect, and NFAT1 did not significantly alter promoter activity.

3.4. NRF2 Binds to the LAMP‐2A Promoter, and Mutation of Its Binding Site Abolishes HCV‐Induced LAMP‐2A Promoter Activation

To determine whether NRF2 or HIF‐1α directly regulates LAMP‐2A transcription during HCV infection, we analyzed putative NRF2‐ and HIF‐1α‐binding sites in the LAMP‐2A promoter region identified by in silico prediction using the JASPAR database. A schematic representation of the predicted binding sites and mutant constructs is shown in Figure 4A. Mutation of the NRF2‐binding site (−29 to −20, ATGACTCGCA → MT‐1, ACACCTCATA) significantly reduced promoter activity under both mock and HCV‐infected conditions (Figure 4B). In contrast, mutation of the predicted HIF‐1α‐binding site (−696 to −689, ACACGTAT → MT‐2, ACCAATAT) did not significantly alter promoter activity. ChIP‐qPCR further showed association of NRF2 with the LAMP‐2A promoter under mock conditions, and this association was reduced when the NRF2‐binding site was mutated (Figure 4C). NRF2 association with the promoter was also detected in HCV‐infected cells and was diminished by mutation of the NRF2‐binding site (Figure 4D). Together, these results indicate that NRF2 directly engages the LAMP‐2A promoter and that the identified antioxidant response element is required for basal and HCV‐responsive promoter activation. These results suggest that HIF‐1α indirectly engages the LAMP‐2A promoter.

Figure 4.

NRF2 directly binds to the LAMP2A promoter, and mutation of the NRF2‐binding site abolishes HCV‐induced promoter activation. (A) Schematic representation of the LAMP‐2A promoter region (−1077 to +87 bp). The wild‐type (WT) construct contains the full‐length promoter driving firefly luciferase. The putative NRF2‐binding site (antioxidant responsive element, ARE; −29 to −20; wild‐type: ATGACTCGCA) was mutated in MT‐1 (mutated sequence: ACACCTCATA). The putative HIF‐1α‐binding site (hypoxia response element, HRE; −696 to −689; wild‐type: ACACGTAT) was mutated in MT‐2 (mutated sequence ACCAATAT). (B) Huh‐7.5 cells were mock‐infected or infected with HCV J6/JFH1 at a multiplicity of infection (MOI) of 1 and co‐transfected with the indicated WT, MT‐1 or MT‐2 LAMP‐2A promoter luciferase reporter plasmid. Gray bars: Mock‐infected; Black bars: HCV‐infected. LAMP‐2A promoter activity was measured by dual‐luciferase assays and normalized to Renilla luciferase activity. At 48 h post‐infection, luciferase activities were measured. Data represent mean ± SEM from three independent experiments. Statistical analysis was carried out using the Student's t‐test. **p < 0.01. (C, D) ChIP assays were performed in mock‐infected (C) or HCV‐infected (D). Huh‐7.5 cells were co‐transfected with pCAG‐FLAG‐NRF2 plasmid together with wild‐type or mutant (MT‐1) of LAMP‐2A promoter luciferase reporter. Chromatin immunoprecipitation (ChIP) assay was performed using an anti‐NRF2 antibody or normal rabbit IgG (as a negative control). Enrichment of the LAMP‐2A promoter region containing the putative NRF2‐binding site was analyzed by qPCR and expressed as fold enrichment relative to input chromatin. Data represent mean ± SEM from three independent experiments. Statistical analysis was carried out using the Student's t‐test. *p < 0.05; **p < 0.01.

3.5. HCV Infection Promotes Nuclear Accumulation and Activation of NRF2

NRF2 is activated through dissociation from its cytoplasmic repressor Kelch‐like ECH‐associated protein 1 (Keap1), and subsequent nuclear translocation [24, 33, 34, 35]. To determine whether HCV infection increases nuclear localization of NRF2, we performed immunofluorescence analysis using exogenous FLAG‐NRF2 and complemented this approach with biochemical fractionation of HCV‐infected cells. In mock‐infected cells, NRF2 was distributed in both the nucleus and cytoplasm (Figure 5A, left panel, Red signal). By contrast, HCV infection increased the proportion of NS5A‐positive cells showing predominantly nuclear NRF2 signals (Figure 5A, right panel, red signal). Consistent with this observation, cell fractionation assays showed no obvious change in cytoplasmic NRF2, but a clear increase in nuclear NRF2 in HCV‐infected cells (Figure 5B,C first panel, lane 2). Nuclear phosphorylated NRF2 (Ser‐40) were also increased, consistent with enhanced NRF2 activation. These results support the conclusion that HCV infection promotes nuclear accumulation and activation of NRF2.

4. Discussion

Hepatitis C virus (HCV) infection triggers oxidative stress, endoplasmic reticulum (ER) stress, and autophagy‐related signaling, and these responses are increasingly recognized as active determinants of viral persistence [36, 37, 38, 39, 40, 41, 42, 43]. NRF2 is a master regulator of antioxidant and cytoprotective transcriptional programs [44, 45]. Previous studies have shown that HCV proteins can modulate NRF2/ARE signaling, although the downstream lysosomal targets of this response remain incompletely defined [46, 47, 48]. In the present study, we identified LAMP‐2A as a transcriptional target linked to NRF2 signaling in HCV‐infected cells. This finding provides a mechanistic connection between virus‐associated stress responses and regulation of the CMA machinery.

CMA is a selective lysosomal degradation pathway distinct from macroautophagy and depends on LAMP‐2A as its rate‐limiting lysosomal receptor [9, 10, 49]. Changes in LAMP‐2A expression directly affect CMA activity and cellular stress adaptation [32, 50]. We previously demonstrated that HCV NS5A protein exploits HSC70‐dependent selective lysosomal degradation pathways [15, 16, 17, 18, 19]. Here, we extend those observations by showing that HCV increases LAMP‐2A expression through NRF2‐dependent transcriptional regulation. Together with the reduction in HCV RNA and viral proteins after LAMP‐2A knockdown, these findings indicate that LAMP‐2A contributes functionally to the HCV life cycle. A limitation of this study is that we did not directly measure CMA flux. Therefore, although the increase in LAMP‐2A expression is consistent with enhanced CMA‐related lysosomal capacity, our data do not by themselves prove increased CMA flux in HCV‐infected cells. Nevertheless, the identification of an NRF2‐responsive element in the LAMP‐2A promoter and the increased nuclear accumulation of NRF2 in infected cells provide a coherent mechanism for HCV‐induced LAMP‐2A upregulation (Figure 6).

Figure 6.

Proposed model of the NRF2‐LAMP‐2A regulatory axis in HCV‐infected cells. Based on the current data and prior literature, HCV infection promotes nuclear translocation of NRF2. Activated NRF2 directly binds to the antioxidant element (ARE) within the LAMP‐2A promoter and transcriptionally upregulates LAMP‐2A expression. Increased LAMP‐2A at the lysosomal membrane is proposed to enhance chaperone‐mediated autophagy (CMA)‐related lysosomal capacity. Through HSC70‐mediated recognition of substrate proteins bearing KFERQ‐like motifs, this pathway may facilitate selective lysosomal remodeling that supports efficient HCV replication.

Our candidate factor analysis suggests that LAMP‐2A transcription is controlled by multiple stress‐responsive pathways, but that NRF2 is the predominant direct regulator in this system. Although HIF‐1α overexpression increased promoter activity, mutation of the predicted HIF‐1α binding site did not suppress the reporter activity. These findings argue against direct binding of HIF‐1α to the site tested in this study and instead suggest that HIF‐1α may influence LAMP‐2A expression indirectly or through cooperative interactions with other stress‐responsive transcription factors. NFAT1 protein levels increased after HCV infection but did not augment promoter activity in Huh‐7.5 cells, implying that its contribution is context‐dependent. Thus, while several stress‐related transcription factors may be altered during HCV infection, the current data most strongly support direct transcriptional regulation of LAMP‐2A by NRF2.

Although multiple putative transcription factor‐binding motifs were identified in the LAMP‐2A promoter region, the presence of a predicted motif does not necessarily indicate functional transcriptional regulation. Our data support NRF2 as the principal direct regulator in this system, while the contribution of HIF‐1α appears limited and NFAT1 was not functionally validated in the promoter assay conditions used here.

Knockdown of LAMP‐2A reduced HCV RNA and protein levels, demonstrating that LAMP‐2A supports efficient viral replication. Our interpretation is that increased LAMP‐2A helps infected cells maintain a lysosomal environment favorable for replication‐associated stress adaptation. However, the present experiments do not identify the exact replication step affected or the specific CMA substrates responsible for this phenotype. Determining whether LAMP‐2A promotes replication by eliminating antiviral factors, remodeling metabolic regulators, or supporting proteostasis will require flux assays and substrate‐level analyses in the future studies.

HCV infection increased LAMP‐2A protein abundance in Huh‐7.5 cells (Figure 1C). Although a subtle difference in electrophoretic mobility was observed, the present study focused on the increase in overall LAMP‐2A abundance and did not directly examine the underlying post‐translational modification state. Because LAMP‐2A is a heavily glycosylated lysosomal membrane protein, the apparent mobility difference observed in HCV‐infected cells may reflect altered post‐translational modification and/or maturation state. Although this possibility was not directly addressed in the present study, it will be important to investigate whether HCV infection affects LAMP‐2A glycosylation or lysosomal processing in future work.

We aimed to assess the longer‐term effect of HCV infection on LAMP‐2A expression in Figure 1B and therefore LAMP‐2A mRNA levels were measured at days 2, 4, and 6 after infection and normalized to GAPDH relative to mock‐infected controls. In contrast, the experiment shown in Figure 2A was designed to evaluate the effect of shRNA‐mediated LAMP‐2A knockdown during an earlier time window (12–48 h), and the data were normalized within that experimental setting.

Our data in Figure 4C,D support NRF2 binding to the LAMP‐2A promoter and are consistent with HCV‐enhanced NRF2‐dependent transcription. Nevertheless, a direct quantitative comparison of NRF2 occupancy between HCV‐negative and HCV‐positive cells would further strengthen the conclusion regarding HCV‐induced promoter recruitment and remains an important subject for future study.

The apparent difference between the immunofluorescence and fractionation data in Figure 5A,B likely reflects the distinct nature of the assays. Immunofluorescence visualizes intracellular distribution in individual infected cells, whereas fractionation/immunoblotting provides biochemical information averaged across cell populations and may be influenced by both endogenous and ectopically expressed NRF2 species depending on the experimental setting. Thus, the two approaches collectively support HCV‐associated nuclear enrichment of NRF2 but are not expected to produce identical visual patterns.

Emerging evidence indicates that certain viruses actively exploit CMA to promote immune evasion and optimize intracellular conditions for viral replication. Viruses can selectively eliminate antiviral host factors via CMA, thereby maintaining host cell viability while attenuating innate immune signaling and establishing an environment favorable for persistent infection [51, 52]. For example, the NS2A protein of Zika virus has been shown to induce CMA‐dependent degradation of karyopherin subunit alpha 2 (KPNA2), a nuclear transport factor required for efficient interferon signaling. Loss of KPNA2 impairs STAT nuclear translocation and dampens type I interferon responses, thereby facilitating viral replication [53]. Similarly, Porcine reproductive and respiratory syndrome virus promotes CMA‐mediated degradation of TANK‐binding kinase 1 (TBK1), a central kinase in the RIG‐I‐MAVS signaling cascade, thereby suppressing IRF3 activation and type I interferon production [54]. Furthermore, recent studies have reported that LAMP‐2A‐mediated CMA promotes lysosomal degradation of key innate immune signaling molecules such as STING and TBK1, and that targeting LAMP‐2A enhances the efficacy of anti‐PD‐1 immune checkpoint blockade [55]. Collectively, these findings support a broad model in which viruses hijack CMA as a strategic immune evasion mechanism, inducing lysosomal degradation of key signaling intermediates to attenuate interferon‐mediating antiviral defenses and promote efficient replication. Further elucidation of virus‐CMA interactions will provide important insights into host‐pathogen dynamics and may uncover novel therapeutic targets aimed at restoring antiviral immunity.

We distinguish here between increased total protein abundance and functional activation. While Figure 3B demonstrates elevated levels of NRF2, HIF‐1α, and NFAT1 proteins, the activation‐related evidence for NRF2 is more directly supported by its nuclear accumulation and increased Ser40 phosphorylation in Figure 5. Additional early time‐point analyses would further clarify the temporal relationship between NRF2 activation and LAMP‐2A induction. Although the reporter assay in Figure 3B revealed differential effects among the tested transcription factors, direct validation of overexpression levels would further strengthen the interpretation of these findings. Therefore, the present data should be interpreted primarily as functional reporter assay results under the indicated transfection conditions.

In the current study, the main lines of evidence supporting the NRF2–LAMP‐2A axis are: (i) identification of an NRF2‐responsive element in the LAMP‐2A promoter, (ii) reduced promoter activity after mutation of this element, (iii) ChIP‐based detection of NRF2 binding to the promoter region, and (iv) increased nuclear accumulation and Ser40 phosphorylation of NRF2 during HCV infection. Direct depletion of NRF2 would provide complementary evidence for the NRF2 dependence of LAMP‐2A induction.

Our data also raise questions about the upstream pathways that activate NRF2 during HCV infection. Previous studies have linked viral infection to NRF2 activation through oxidative stress, reactive oxygen species, or ER stress‐PERK signaling [56, 57]. In the present study, we observed increased nuclear accumulation and Ser40 phosphorylation of NRF2, but we did not directly test the responsible upstream trigger. Moreover, our conclusions are based on the Huh‐7.5 hepatoma cell model and on assays that included exogenous FLAG‐NRF2. Additional studies using endogenous NRF2 perturbation, primary hepatocytes, or more physiologically relevant infection systems will be important to strengthen the generalizability of our findings. Even with these limitations, the present data define a transcriptional route by which HCV can increase LAMP‐2A expression and thereby potentially reinforce CMA‐associated host remodeling.

5. Conclusion

In conclusion, HCV increases LAMP‐2A expression at the promoter, mRNA, and protein levels in infected hepatoma cells. We identified NRF2 as the principal direct transcriptional regulator of this response. NRF2‐associated promoter occupancy was detected by ChIP, its binding site was required for basal and HCV‐responsive promoter activity, and HCV infection enhanced nuclear accumulation and Ser40 phosphorylation of NRF2. Although HIF‐1α affected the LAMP‐2A promoter activity in overexpression experiments, its contribution appears indirect under the conditions tested. Functionally depletion of LAMP‐2A reduced intracellular HCV RNA and virus protein levels, indicating that LAMP‐2A supports efficient viral replication. Collectively, these findings define an NRF2‐LAMP‐2A regulatory axis engaged during HCV infection and provide a framework for future studies examining how selective lysosomal degradation pathways support the HCV life cycle (Figure 6).

Author Contributions

Chieko Matsui, and Ikuo Shoji conceived and designed the experiments. Zihan Xu and Chieko Matsui carried out most of the experiments. M. A. S., F. P., D.L., and Takayuki Abe assisted with the construction and the data analysis. Zihan Xu, Chieko Matsui, and Ikuo Shoji wrote the manuscript.

Ethics Statement

This study did not include human samples or animal experiments that require approval from the Ethics Committee of the institution.

Conflicts of Interest

The authors declare no conflicts of interest.

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request. All data are presented in the main figures. Raw sequencing data, microscopy images, materials, and sequence information are available upon request. Correspondence and requests for materials should be addressed to Professor Ikuo Shoji.