Synergetic contributions of Seneca Valley virus 3 C and 3D proteins to induction of ferroptosis for viral replication

Abstract

Seneca Valley virus (SVV) infection leads to severe vesicular diseases in pigs, posing a significant threat to the global swine industry. Ferroptosis, a novel form of non-apoptotic cell death, is characterized by iron-dependent phospholipid peroxidation. However, the role of ferroptosis in SVV replication remains poorly understood. In this study, we demonstrate that SVV infection induces ferroptosis, as evidenced by lipid peroxidation, reactive oxygen species (ROS) accumulation, and glutathione (GSH) depletion. The GPX4 and nuclear receptor coactivator 4 (NCOA4)-mediated ferritinophagy are key contributors to ferroptosis induction. Furthermore, our findings reveal that the SVV 3 C proteinase (3C^pro) targets the GPX4 for degradation, thereby promoting ferroptosis. Simultaneously, the SVV 3D protein enhances the NCOA4-FTH1 interaction, leading to increased ferritin degradation and subsequent ferritinophagy. Notably, inhibition of ferroptosis significantly reduces SVV replication and its associated inflammatory effects. Collectively, these results elucidate the intricate molecular mechanisms underlying SVV-induced ferroptosis, highlighting the synergistic roles of 3C^pro and 3D in activating ferroptotic pathways and presenting potential targets for therapeutic intervention in SVV infections.

Supplementary Information

The online version contains supplementary material available at 10.1007/s00018-025-05951-y.

Introduction

Seneca Valley virus (SVV) was first considered as a contaminant in human fetal retinoblast cell cultures in the United States in 2002 [1, 2]. As an emerging pathogen, the first case of SVV infection was reported in China in 2015, followed by numerous cases globally [3–8]. SVV, an oncolytic RNA virus, causes vesicular diseases that are clinically indistinguishable from those of foot-and-mouth disease virus (FMDV), vesicular stomatitis virus (VSV), and swine vesicular disease virus (SVDV). Additionally, SVV shows promise as a therapeutic modality against human cancers [9, 10]. SVV is a non-enveloped, single-stranded, positive-sense RNA virus within the Picornaviridae family [1, 11]. Its genome, approximately 7.2 kb in length, encodes a large open reading frame (ORF) that is cleaved into four structural proteins (VP4, VP2, VP3, and VP1) forming the viral capsid, and eight nonstructural proteins (Lpro, 2 A, 2B, 2 C, 3 A, 3B, 3 C, and 3D) [1, 12]. The SVV 3 C protease (3C^pro) antagonizes host innate immune responses by targeting key molecules, including selective autophagy receptors SQSTM1/p62 and OPTN (optineurin) [13–15], and gasdermin D (GSDMD) [16]. 3C^pro also cleaves signal transducer and activator of transcription STAT2, histone deacetylase 4 (HDAC4) and TRIM32 inhibit the type I interferon signaling pathway [17–19]. In addition, 3C^pro cleaves key interferon-stimulated genes (ISGs) to antagonize the antiviral effect, such as mRNA decapping enzyme 1 A (DCP1A) [20].

Ferroptosis is an iron-dependent and non-apoptotic cell death characterized by lipid peroxidation [21]. Unlike apoptosis, necroptosis, or autophagic cell death, ferroptosis involves the excessive production of intracellular lipid ROS, leading to fatal cellar lipid peroxidation due to diminished cellular antioxidant capacity [22]. This process is marked by Fe²⁺ overload, GSH depletion, and mitochondrial shrinkage. Key regulators of ferroptosis include GPX4 (glutathione peroxidase 4), ACSL4 (Acyl-Coenzyme A synthetase long-chain family member 4), and SLC7A11 (solute carrier family 7 member 11, also known as xCT) [23, 24]. The GPX4-dependent system reduces lipid peroxidation, and its inactivation or inhibition leads to lipid ROS accumulation and the induction of ferroptosis [23, 25]. Viruses may trigger ferroptosis by subverting the GPX4-dependent pathway to promote replication.

Iron is an essential nutrient for both hosts and invading pathogens. Cellular iron is primarily stored in ferritin, an iron-binding protein composed of ferritin heavy chain (FTH1) and ferritin light chain (FTL) subunits. Ferroptosis is an autophagy-dependent cell death process [26], and ferritinophagy is mediated by the autophagic cargo receptor nuclear receptor coactivator 4 (NCOA4). NCOA4 facilitates the degradation of iron-bound ferritin via autophagy, releasing iron from ferritin [27–29]. Ferritinophagy induces ferroptosis by regulating intracellular iron homeostasis and ROS generation. The interaction between FTH1 and NCOA4 is necessary for functional ferritinophagy, and NCOA4 deletion completely inhibits ferritin degradation under iron depletion [27]. Transferrin receptor 1 (TFR1), a single membrane-spanning glycoprotein, mediates cellular iron uptake. Iron-loaded transferrin interacts with TFR1, delivering iron to cells [30, 31]. TFR1 also serves as a viral entry receptor [32].

Viruses can induce or inhibit ferroptosis to facilitate replication, pathogenesis, and evasion of host immune surveillance [33, 34]. For instance, herpes simplex virus 1 (HSV-1) and influenza A virus trigger ferroptosis through the nuclear factor-erythroid 2-related factor 2 (Nrf2)-Kelch-like ECH-associated protein 1 (Keap1) pathway, which respectively leads to viral encephalitis and nasal mucosal epithelial inflammation [35, 36]. Newcastle disease virus (NDV) induces ferroptosis via the p53-SLC7A11-GPX4 pathway by activating p53 [37]. Severe acute respiratory syndrome coronavirus 2(SARS-CoV-2) ORF3a causes the degradation of Nrf2, which leads to reduced cellular resistance to oxidative stress and facilitates ferroptosis [38]. Japanese encephalitis virus (JEV) induces neuronal ferroptosis by inhibiting GPX4 expression and enhancing lipid peroxidation via the yes-associated protein 1 (YAP1)/ACSL4 pathway [39]. However, it remains unclear whether SVV infection induces ferroptosis or ferritinophagy.

In this study, we investigated the ferroptosis pathway during SVV infection. Specifically, we investigate how SVV manipulates GPX4 axis and NCOA4-mediated ferritinophagy to induce ferroptosis, thereby facilitating viral replication. By understanding these mechanisms, we hope to uncover potential therapeutic targets that could disrupt SVV replication and offer new strategies for combating SVV-related diseases. The findings from this study not only advance our knowledge of SVV pathogenesis but also contribute to the broader understanding of virus-host interactions, particularly how viruses exploit cellular death pathways to their advantage.

Materials and methods

Cells, viruses, antibodies, and reagents

BHK-21 (ATCC, CCL-10) cells were cultured in Dulbecco’s modified Eagle’s medium (Invitrogen, California, USA) supplemented with 10% fetal bovine serum (Invitrogen). The cell culture was maintained at 37 °C in an incubator with 5% CO₂. The SVV strain CHhb17 that had been utilized in our prior research [40]. eGFP-tagged recombinant SVV (rSVV-eGFP) was kindly provided by Fuxiao Liu (Qingdao Agricultural University) [41]. Rabbit anti-GFP (50430-2-AP), and rabbit anti-TFR1 (10084-2-AP) were purchased from Proteintech (Wuhan, China). Rabbit anti-GPX4 (A1933), rabbit anti-SLC7A11 (A13685), and mouse anti-HA tag monoclonal antibody (A1933) were purchased from Abclonal (Wuhan, China). Rabbit anti-ferritin (ET1610-78) and rabbit anti-NCOA4 (PSH05-44) were purchased from Huabio (Huaan, China). LC3B (L7543) and FITC-conjugated goat anti-mouse IgG (H + L) (F0257; Sigma) were purchased from Sigma-Aldrich. SQSTM1/p62 (ab56416) was purchased from Abcam. HRP-conjugated goat anti-rabbit IgG (H + L) (1706515) and HRP-conjugated goat anti-mouse IgG (H + L) (1706516) were purchased from Bio-rad. The dilution factors of primary and secondary antibodies were 1:1,000 and 1:10,000, respectively. Alexa-568-conjugated goat anti-rabbit IgG (H + L) (11011) was purchased from Invitrogen. The mouse anti-VP1 monoclonal antibody was utilized in our previous studies [40]. Ferrostatin-1 (Fer-1, HY-100579), erastin (HY-15763), N-acetyl-L-cysteine (NAC, HY-B0215) were purchased from MedChemExpress (Shanghai, China). MG132 (S2619), Z-VAD-FMK (S7023), bafilomycin A1 (S1413), chloroquine (CQ, S6999), and 3-methyladenine (3-MA, S2767) were obtained from Selleck Chemicals (Shanghai, China).

Plasmids construction

The genes of TFR1, NCOA4, and FTH1 were amplified from PK-15 cells and then ligated into pCMV-HA vectors (Clontech, 631604) and p3×FLAG-CMV-10 vectors (Sigma, E4401) with the use of DNA assembly mix plus (D0204P, Lablead). The plasmids of GFP-tagged SVV structural and non-structural proteins, the single-point mutant plasmids GFP-3 C H48A and GFP-3 C C160A, as well as the double-mutant plasmid GFP-3 C [DM] (with double mutations of H48A and C160A), which were employed in our previous studies [15, 17]. Primers were listed in Table 1.

Table 1.

Primers used in this study

Primersa		Sequence (5’−3’)b
HA-NCOA4-EcoRI-F HA-NCOA4-KpnI-R Q-BHK-GPX4-F Q-BHK-GPX4-R Q-BHK-SLC7A11-F Q-BHK-SLC7A11-R Q-IL-6-F Q-IL-6-R Q-TNF-F Q-TNF-R Q-CCL20-F Q-CCL20-R Q-IL-1β-F Q-IL-1β-R Q-IL-18-F Q-IL-18-R	TGGCCATGGAGGCCCGAATTCGGATGAGTACCTCACAGGACCAGAAT GATCCCCGCGGCCGCGGTACCTCACATCTGTAGAGGAGTTCGATA CTAGGAGGAGGAAAGCTCGC TGTGACCGTCGATGTCCTTG GGTTACCTGCAGGGCAATGT TGCCCGTGTTCTGGAGTATG CGCAAGAGACTTCCATCCACT TGAAGTCTCCTCTCCGGACTT CAACCCTATCATCGGCTCCA TAAACCAGGTACAGCCCGTC GTCAGAAGCAGCAAGCAACTTT TTTGGATCAGCACACACGGAT ACAGAAATGCCTCGTGCTGT GTGGGCGTGTCACCTTTCAT TGGAGACCTGGAATCAGACGA ACAGAGAGGGTTACAGGCAGT

a F denotes forward PCR primer; R denotes reverse PCR primer

Detection of intracellular ROS levels

The detection of intracellular reactive oxygen species (ROS) was carried out using the DCFH-DA fluorescent probe (S0033S, Beyotime, Shanghai, China). BHK-21 cells were incubated with the DCFH-DA fluorescent probe for 30 min at 37 °C within a cell culture incubator. Subsequently, after being washed twice with phosphate-buffered saline (PBS), the BHK-21 cells were placed in fresh culture medium and imaged under a Nikon Al confocal microscope. For flow cytometry analysis, the washed cells were harvested by treating them with trypsin. Following centrifugation at 300 × g for 5 min, the cell pellets were resuspended in phosphate buffered saline (PBS) and then analyzed with a NovoCyte flow cytometer (Agilent, CA, USA).

Mitochondrial membrane potential (MMP) assay

The MMP was detected using JC-1 fluorescent probe (C2003S, Beyotime, Shanghai, China). BHK-21 cells were stained with JC-1 fluorescent probe (1:200) for 30 min at 37 °C in a cell culture incubator. After two washes with JC-1 washing buffer, BHK-21 cells were incubated with new culture medium and captured under a a Nikon Al confocal microscope. For flowcytometry analysis, washed cells were collected by trypsinization. After centrifugation at 300 × g for 5 min, the cell pellets were resuspended in PBS and analyzed using a NovoCyte flow cytometer.

Detection of mitochondrial ROS

BHK-21 cells were stained with mitoSOX red mitochondrial superoxide indicator (MCE, Shanghai, China) for 30 min at 37 °C in a cell culture incubator. After two washes with growth medium, cells were incubated with new culture medium and captured under a Nikon Al confocal microscope. Spectrofluorometer (Biotek) was used to examine mitoSOX at 510–580 nm wavelengths.

Lipid peroxidation assay

Lipid ROS was detected using BODIPY 581/591 C11 fluorescent probe (D3861, Invitrogen, Waltham, MA, USA). BHK-21 cells cultivated in 6-well plates were were incubated with 1 µM BODIPY 581/591 C11 at 37 °C for 30 min. After two washes with growth medium, cells were incubated with new culture medium and captured under a a Nikon Al confocal microscope. For flowcytometry analysis, washed cells were collected by trypsinization. After centrifugation at 300 × g for 5 min, the cell pellets were resuspended in PBS and analyzed using a NovoCyte flow cytometer.

Cell viability and LDH cytotoxicity assay

BHK-21 cells were seeded in 96-well plates, the viability of cells was examined using cell counting kit 8 according to the manufacturer’s instructions (CCK-8, C0038, Beyotime). The absorbance at 450 nm was measured using a plate reader (Bio-Rad, Hercules, CA, USA). The LDH cytotoxicity was determined using LDH cytotoxicity assay kit (40209ES76, yeasen) according to the manufacturer’s instructions. The absorbance at 490 nm was measured using a plate reader (Bio-Rad).

Measurement of MDA and GSH levels

The levels of malondialdehyde (MDA) and glutathione (GSH) were evaluated by employing specific assay kits. BHK-21 cells were plated in 6-well plates. The concentrations of MDA were determined using an MDA assay kit (S0131M, Beyotime) following the detailed instructions provided by the manufacturer. A microplate reader (Bio-Rad) was utilized to measure the absorbance at a wavelength of 532 nm. The content of GSH was analyzed with a reduced glutathione content assay kit (BC1175, Solarbio, China). The absorbance at 532 nm was also measured by means of a microplate reader (Bio-Rad).

Western blotting

Cell samples are lysed in lysis buffer containing protease inhibitor, removed cellular debris and boiled at 100 °C with 5 × SDS-PAGE loading buffer. The lysis products were separated from proteins by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), then proteins transferred from gels to a nitrocellulose membrane under ice bath conditions. Membranes were blocked with 5% skimmed milk in phosphate buffered saline (PBS) for 1 h at room temperature (RT). They were incubated with diluted primary antibodies at room temperature or at 4 °C for 12–16 h. Rabbit or mouse IgG antibodies conjugated with horseradish peroxidase (HRP) were used as secondary antibodies. Membranes were washed with PBST then visualized by enhanced ECL chemiluminescent substrate kit.

Co-immunoprecipitation (Co-IP)

BHK cells cultured in 2-cm dishes were transfected with the indicated plasmids. At 24 h post-transfection, the cells were lysed with lysis buffer containing protease inhibitors. Samples were centrifuged at 13,000 × g for 15 min at 4 °C to remove cellular debris. A portion of the cell lysate clarified supernatants were left as a control and the rest were mixed with the anti-HA magnetic beads. Then the mixture was incubated in a rolling incubator at 4 °C overnight. The beads were washed three times with lysis buffer, then boiled with 5 × SDS loading buffer. The proteins were analyzed with western blotting using the indicated antibodies.

Quantitative RT-PCR

RNA from the cells was extracted using the FastPure Cell/Tissue Total RNA Isolation Kit-BOX 2 (Vazyme, RC101-01). First-stand complementary DNA was reverse transcribed from 1 µg of total RNA using first-strand synthesis master mix (F0202, Lablead). Real-time PCR was performed on a Bio-rad CF96 Real-Time PCR System using Realab Green PCR mix. (R0202, Lablead) were normalised to β-actin as control. Relative gene expression was quantified by calculating 2^−△△CT (CT is the threshold cycle). Primers were listed in Table 1.

12 indirect Immunofluorescence (IFA)

BHK Cells were seeded in 24-well plates and incubate until density reaches 70%. Cell samples after treatment were fixed in 4% paraformaldehyde for 10 min, then permeabilized with 2% bovine serum albumin (BSA) in 0.5% Triton X-100 for 10 min, and blocked them using 2% BSA for 1 h. Then cells were incubated with the diluted primary antibody overnight at 4 °C, followed by incubation with Alexa Fluor 488 or Alexa Fluor 568 secondary antibody for 2 h at room temperature. Incubated with DAPI for 1 min and observed the color of the cells under the confocal laser scanning microscope.

RNA interference (RNAi)

The small interfering RNAs (siRNAs) were designed and synthesized by GenePharma (Suzhou, China): si-FTH1-110 (sense, 5’- CCUACGUCUAUCUGUCUAUTT-3’; antisense, 5’- AUAGACAGAUAGACGUAGGTT-3’), si-NCOA4-188 (sense, 5’-GCAUAAGUCGCCAUCUAGATT-3’; antisense, 5’- UCUAGAUGGCGACUUAUGCTT-3’), si-NCOA4-526 (sense, 5’-GCUCAUACUAACUCUUCAATT-3’; antisense, 5’- UUGAAGAGUUAGUAUGAGCTT-3’), si-TFR1-371 (sense, 5’-GGGCAGACCUCAAGAAACUTT-3’; antisense, 5’- AGUUUCUUGAGGUCUGCCCTT-3’), si-TFR1-1086 (sense, 5’-GGCAUCUUCACAGGAUAAATT-3’; antisense, 5’- UUUAUCCUGUGAAGAUGCCTT-3’), si-GPX4-287 (sense, 5’-GCUCCAUGCACGAAUUCUCTT-3’; antisense, 5’-GAGAAUUCGUGCAUGGAGCTT-3’), the siNC (sense, 5’-UUCUCCGAACGUGUCACGUTT-3’; antisense, 5’-ACGUGACACGUUCGGAGAATT-3’) served as a control. BHK-21 cells in 6-well plates were transfected with siRNAs using Lipofectamine RNAiMAX (13778150, Thermo Fisher). Cells were infected with SVV at 36 h post-transfection, the expression of proteins were subjected to Western blot.

TCID₅₀ assay

BHK-21 cells were infected with SVV at the multiplicity of infection (MOI) indicated. After incubation with SVV for 1 h at 37 °C, were washed three times with DMEM and supplemented with fresh culture medium containing 2% FBS. The cultured medium and cells were collected at the indicated times after SVV infection, then samples were titrated on BHK- 21 cells using limiting dilution assay by using TCID₅₀ assay [40].

Statistical analysis

Statistical analysis was processed using GraphPad Prism. Results were expressed as means ± standard deviations of triple independent tests. The error bars represent the standard deviation. ^∗P < 0.05, ^∗∗P < 0.01 were considered statistically significant.

Results

SVV infection induces ferroptosis

SVV infection induces host cell pyroptosis and apoptosis [16, 42, 43]. However, other forms of cell death have not been investigated during SVV infection. Therefore, we examined lipid peroxidation using the BODIPY 581/591 C11 fluorescent probe in conjunction with flow cytometry (Fig. 1A) and laser confocal microscopy (Fig. 1B). Erastin served as a positive control for the induction of ferroptosis. The results demonstrated that both SVV infection and erastin treatment significantly increased lipid ROS generation, as evidenced by the fluorescence shift from non-oxidized BODIPY 581/591 C11 (red) to oxidized BODIPY 581/591 C11 (green) (Fig. 1B). Notably, ferrostatin-1 (Fer-1) effectively inhibited SVV-induced lipid ROS production and SVV replication (Fig. 1A-D). Additionally, SVV infection-induced cell death, as assessed by the cell counting kit-8 (CCK-8) assay (Fig. 1E), and this cytotoxic effect was mitigated in the presence of Fer-1 (Fig. 1F, G). Intracellular lipid peroxidation and glutathione levels were quantitatively measured using malondialdehyde (MDA) and GSH assays, respectively. SVV infection and erastin treatment significantly decreased GSH levels, even in the presence of Fer-1 (Fig. 1H). Conversely, MDA and lactate dehydrogenase (LDH) levels were markedly elevated following SVV infection and erastin treatment; however, these increases were attenuated by Fer-1 (Fig. 1I, J). Collectively, these findings indicate that SVV induces cell death and lipid peroxidation in a dose- and time-dependent manner.

Fig. 1.

SVV infection induce ferroptosis (A, B) BHK-21 cells were infected with SVV (MOI = 5) and treated with erastin (10 µM) or Fer-1 (15 µM). At 12 hpi, intracellular lipid ROS was measured using C11-BODIPY 581/591 (10 µM) and examined by flow cytometry and confocal microscopy, respectively. (C) Relative fluorescence intensity in BHK-21 cells from (B). (D) BHK-21 cells were mock infected or infected with SVV (MOI = 5), then treated with Fer-1 (15 µM). At 12 hpi, SVV VP3 protein was stained by VP3 antibody. The statistical results percentage of SVV infected cells were using Image J. Error bars indicate mean ± SD from three independent infection experiments (**, P < 0.01). (E) BHK-21 cells were infected with SVV (MOI = 0.1, 1, and 5) and treated with erastin (10 µM) as a positive control, cell viability was measured using CCK-8 assay at 12 hpi. (F) BHK-21 cells were infected with SVV (MOI = 5) and treated with Fer-1 with different dosage (30 µM, 15 µM, 5 µM), cell viability was measured using CCK-8 assay at 12 hpi. (G) BHK-21 cells were infected with SVV (MOI = 5) and treated with DMSO or Fer-1 (15 µM), cell viability was measured using CCK-8 assay at 6 hpi and 12 hpi. (H-J) BHK-21 cells were infected with SVV (MOI = 5), and treated with erastin (10 µM) or Fer-1 (15 µM), the GSH at 12 hpi, MDA, and LDH level was measured using a GSSG/GSH quantification kit, and MDA assay kit at 0 hpi, 6 hpi and 12 hpi, respectively

SVV-induced ferroptosis is associated with increased intracellular ROS production

Intracellular ROS accumulation, a hallmark of ferroptosis, was detected using a DCFH-DA fluorescent probe through flow cytometry (Fig. 2A) and laser confocal microscopy (Fig. 2B). Significant ROS accumulation was observed in SVV-infected, erastin-treated, and Rosup-treated BHK-21 cells, indicating the induction of oxidative stress. This ROS production was notably reduced by treatment with N-acetyl-L-cysteine (NAC), a ROS scavenger, and Fer-1, a ferroptosis inhibitor (Fig. 2A-C). These findings suggest that ROS generation is crucial during ferroptosis. Further validation using the CellROX Deep Red probe confirmed that SVV infection led to increased ROS accumulation (Fig. 2D). Mitochondrial membrane potential (MMP) was assessed using the JC-1 fluorescent probe, revealing that MMP significantly decreased in SVV-infected, erastin-treated, and CCCP-treated cells, as indicated by a higher green fluorescence intensity of the JC-1 monomer (Fig. 2E-G). This decrease was mitigated by NAC or Fer-1 treatment (Fig. 2E-G). Additionally, elevated mitochondrial ROS was detected in SVV-infected and erastin-treated cells using the mitoSOX red mitochondrial superoxide indicator, which was subsequently inhibited by Fer-1 (Fig. 2H-J). These results demonstrate that SVV-induced ROS production, a critical factor in ferroptosis, is closely associated with mitochondrial dysfunction.

Fig. 2.

SVV infection induce ROS acculumation. (A) BHK-21 cells were infected with SVV (multiplicity of infection [MOI] = 5) at 12 hpi. Intracellular ROS was stained by DCFH-DA probe (10µM) and detected by confocal microscopy, in indicated group. Rosup as a positive control. erastin (10 µM), NAC (5 mM), Fer-1 (15 µM). (B) The statistical results of the relative ROS fluorescence from figure (A) using Image J. Error bars indicate mean ± SD from three independent infection experiments (*, P < 0.05; ***, P < 0.001; NS, not significant). (C) BHK-21 cells were mock infected or infected with SVV (MOI = 5), then treated with NAC (5 mM) and Fer-1 (15 µM). At 12 hpi, SVV VP3 protein was stained by VP3 antibody. The statistical results percentage of SVV infected cells were using Image J. Error bars indicate mean ± SD from three independent infection experiments (**, P < 0.01). (D) BHK-21 cells were infected with SVV (MOI = 5) at 12 hpi. Intracellular ROS was stained by CellROX Deep Red probe (10 µM) and detected by flow cytometry, erastin (10 µM). (E) Assessment of mitochondrial membrane potential (MMP) levels in BHK-21 cells using JC-1 fluorescent probe and detected by flow cytometry. erastin (10 µM), NAC (5 mM), Fer-1 (15 µM). (F) BHK-21 cells were mock infected or infected with SVV (MOI = 5), then treated with erastin (10 µM), NAC (5 mM), Fer-1 (15 µM), or CCCP (10 µM). At 12 hpt, MMP was measured using JC-1 fluorescent probe and detected by flow cytometry. (G) Relative aggregates/monomers fluorescence intensity ratio in BHK-21 cells from (F). (H) BHK-21 cells were mock infected or infected with SVV (MOI = 5), then treated with erastin. At 12 hpt, mitochondrial ROS was measured using mitoSOX red mitochondrial superoxide indicator (5 µM) and detected by flow cytometry. (I) BHK-21 cells were mock infected or infected with SVV (MOI = 5), then treated with erastin (10 µM) and Fer-1 (15 µM). At 12 hpt, mitochondrial ROS was measured using mitoSOX red mitochondrial superoxide indicator (5 µM) and examined by confocal microscopy. (J) Relative fluorescence intensity ratio in BHK-21 cells from (I)

SVV induces ferroptosis via inducing GPX4 degradation

Infection of SVV decreased the abundance of GPX4 and FTH1 in a time-dependent manner (Fig. 3A). As shown in Fig. 3A, TFR1 and NCOA4 expression were notably increased in SVV-infected cells. The mRNA transcriptional levels of GPX4 was also reduced remarkably, while TFR1 and NCOA4 were significantly elevated (Fig. 3B), which agreed with the protein level (Fig. 3A). SVV infection induced cell death at later stage (Fig. 3C). Inhibiting ferroptosis impairs SVV replication (Fig. 3D). In contrast, erastin treatment had the opposite effect (Fig. 3D). Ferroptosis, modulated by pathways such as autophagy, is shown to be impaired by Fer-1 treatment, which alleviates SVV-induced degradation of GPX4 and inhibits LC3-II conversion (Fig. 3E, F). Furthermore, erastin treatment enhances autophagosome formation and the expression of TFR1 and NCOA4 while downregulating GPX4 and FTH1 (Fig. 3G, H). Together, these results suggested that SVV induces ferroptosis via inducing GPX4 degradation to promote viral replicaiton.

Fig. 3.

SVV infection degraded GPX4 to induce ferroptosis (A) BHK-21 cells were infected with SVV (MOI = 5). The cell lysates were collected at 0, 3, 6, 9, 12, and 24 hpi and analyzed by immunoblotting with antibodies against SLC7A11, GPX4, NCOA4, TFR1, FTH1, VP1, and β-actin as an internal control. (B) The transcriptional expression level of SLC7A11, GPX4, TFR1, NCOA4, and FTH1 was analyzed using quantitative RT-PCR and normalized to β-actin mRNA. Error bars indicate mean ± SD from three independent infection experiments (*, P < 0.05; ***, P < 0.001; NS, not significant). (C) BHK-21 cells were infected with SVV (MOI = 5), the cell viability was measured using CCK-8 assay at 0, 3, 6, 9, 12, and 24 hpi. (D) BHK-21 cells were infected with SVV (MOI = 0.5) and treated with Fer-1 (15 µM) or erastin (10 µM). At 3, 6, 9, and 12 hpi, the cells were harvested and titrated using the TCID₅₀ assay. (E, G) BHK-21 cells were infected with SVV (MOI = 5) and treated with Fer-1 (15 µM) (E) and erastin (10 µM) (G), respectively. The cell lysates were collected at 12 hpi and analyzed by immunoblotting with indicated antibodies. (F, H) The ratios to β-actin from three independent experiments (E and G) are shown, respectively (*, P < 0.05; **, P < 0.01; ***, P < 0.001; ns, not significant). Image J was used to quantify the level of protein

NCOA4-mediated ferritin degradation facilitates SVV replication

NCOA4 is a cargo receptor for selective autophagic turnover of FTH1 during ferroptosis. Autophagy suppression significantly inhibits the NCOA4-mediated ferritinophagy signaling axis. The autophagy inhibitors, bafilomycin A1 (Baf A1) and CQ, blocked SVV-induced GPX4 and FTH1 degradation and inhibited TFR1 and NCOA4 expression (Fig. 4A, B, D), thereby reducing NCOA4-medicated ferritinophagy, indicating that autophagy contributes to SVV-induced ferroptosis. NCOA4 knockdown blocked SVV-induced LC3-II conversion and p62/SQSTM1 degradation (Fig. 4C) and prevented FTH1 degradation, leading to increased NCOA4 expression. TFR1, NCOA4, and FTH1 were substantially downregulated by siRNA-mediated knockdown (Fig. 4E, F). Silencing TFR1 and NCOA4 inhibited SVV propagation, while overexpression of TFR1 and NCOA4 promoted viral replication (Fig. 4G, H). Conversely, FTH1 silencing had the opposite effect on SVV replication (Fig. 4G, H). Overall, these findings suggest that SVV infection accelerates ferritin degradation by enhancing NCOA4-mediated ferritinophagy, which facilitates viral replication.

Fig. 4.

Blocking autophagy inbibit SVV-induced ferroptosis (A, B) BHK-21 cells were infected with SVV (MOI = 5) and treated with Baf A1 (200 nM) or CQ (40 µM). Immunoblot analyzed the expression of TFR1, NCOA4, FTH1, GPX4, p62, and LC3. (C) BHK-21 cells were transfected with siRNAs for 36 h, then infected with SVV (MOI = 5) for 12 h. Immunoblot analyzed the expression of TFR1, NCOA4, FTH1, p62, and LC3. (D) The ratios relative to β-actin from three independent experiments (A) are presented (*, P < 0.05; **, P < 0.01; ***, P < 0.001; ns, not significant). Image J was employed to quantify protein levels. (E) RT-qPCR assessment of the knockdown efficiency of TFR1, NCOA4, GPX4, and FTH1. BHK-21 cells were transfected with TFR1, NCOA4, GPX4, and FTH1 siRNAs at a concentration of 20 pmol, with siNC-transfected cells serving as negative controls. (F) Immunoblot analyzed the expression of TFR1, NCOA4, GPX4, and FTH1 after siRNA-medicated knockdown. (G) Immunoblot analyzed VP1 protein production in siRNAs, HA-TFR1, HA-NCOA4, and Flag-FTH1 transfected BHK-21 cells (MOI = 0.5). (H) Growth curves of SVV (MOI=0.5) after transfection with siRNAs, HA-TFR1, HA-NCOA4, HA-GPX4, and Flag-FTH1 in BHK-21 cells

SVV 3 C^pro targeted GPX4 axis for degradation

SVV 3C^pro triggers pyroptosis through the cleavage of gasdermin D (GSDMD) and causes host cell apoptosis via the cleavage of poly (ADP-ribose) polymerase (PARP) [16, 42, 43]. To identify the viral protein responsible for antagonizing the GPX4, SVV protein-expressing plasmids were co-transfected with HA-tagged GPX4. SVV 3C^pro induced the degradation of GPX4 in vitro (Fig. 5A-B). SVV 3C^pro plasmids with protease activity mutations (GFP-3 C-H48A, GFP-3 C-C160A, and GFP-3 C-DM) showed that 3C^pro-mediated degradation was protease activity-dependent (Fig. 5C-D). Z-VAD-FMK treatment significantly reduced GPX4 degradation (Fig. 5E-F). In contrast, inhibitors of the proteasome (MG132), lysosome (NH₄Cl), and autophagy (bafilomycin A1, chloroquine, 3-methyladenine) did not affect degradation (Fig. 5E-F), indicating that 3C^pro mediates degradation via the caspase pathway. The endogenous degradation of GPX4 was examined in the presence of inducers or inhibitors. SVV protein-expressing plasmids transfected into BHK-21 cells and treated with Fer-1 showed that SVV 3C^pro induced GPX4 degradation in vivo (Fig. 5G-H). These findings suggest that SVV 3C^pro degrades Nrf2, HO-1, and GPX4 in a caspase-dependent manner.

Fig. 5.

SVV 3 C^pro targets GPX4 for degradation (A) BHK-21 cells were cotransfected influenza hemagglutinin (HA)-tagged GPX4 with different green fluorescent protein (GFP)-tagged SVV protein expression plasmids for 24 h. Cell samples were subjected to immunoblotting with antibodies against HA, GFP, and β-actin. (B) Quantitative analysis of protein expression in (A) using Image J, where the ratio levels of HA-GPX4 bands were normalized against β-actin. (C) BHK-21 cells grown on 6-well plates were cotransfected single mutant or double mutants GFP-tagged 3C^pro, including 3 C-H48A, 3 C-C160A, and 3 C-DM (H48A-C160A) with HA-tagged GPX4 plasmids for 24 h. Cell samples were subjected to immunoblotting with antibodies against HA, GFP, and β-actin (D) Quantitative analysis of protein expression in (C) using Image J, where the ratio levels of HA-GPX4 bands were normalized against β-actin. (E) BHK-21 cells grown on 6-well plates were cotransfected GFP-3 C with HA-tagged GPX4 plasmids for 24 h, and GFP empty vector with HA-OPTN as a control. NH₄Cl (10 mM), MG132 (10 µM), Z-VAD-FMK (50 µM), Baf A1 (200 nM), CQ (40 µM), and 3-MA (25 mM), (50 µM) were added to cells at 12 h post transfection (hpt), dimethyl sulfoxide (DMSO) as a control. Cells were collected at 24 hpt and subjected to immunoblot with antibodies against HA, GFP, and β-actin. (F) Quantitative analysis of protein expression in (E) using Image J, where the ratio levels of HA-GPX4 bands were normalized against β-actin. (G) BHK-21 cells grown on 6-well plates were cotransfected GFP-tagged SVV protein expression plasmids for 24 h. Fer-1 was added to the cells at 12 hpt. Cells were collected at 24 hpt and subjected to immunoblot with antibodies against GPX4, GFP, and β-actin. (H) Quantitative analysis of protein expression in (G) using Image J, where the ratio levels of HA-GPX4 bands were normalized against β-actin. Data are presented as the mean ± standard deviation (SD) derived from three independent experiments. (*, P < 0.05; ***P < 0.001)

SVV 3D induces ferritinophagy via NCOA4-mediated FTH1 degradation

Ferritinophagy, the selective degradation of ferritin via NCOA4-mediated autophagy, leads to increased intracellular iron and can promote ferroptosis. We observed that SVV 3D protein upregulated NCOA4 (Fig. 6A). Confocal microscopy revealed that SVV 3D co-localized with HA-NCOA4 and showed a punctate distribution in the cytoplasm (Fig. 6B-C). Due to the ability of NCOA4 to facilitate ferritin delivery to lysosomes by direct interaction with FTH1, we examined the ability of NCOA4 to bind FTH1 in the presence of the 3D protein. NCOA4 and FTH1 formed punctate structures in the cytoplasm, and the 3D protein derived from larger condensates (Fig. 6D). These results suggest that 3D is sufficient for the formation of NCOA4-FTH1 condensates. However, 3D cultures promoted endogenous NCOA4 interaction with FTH1 (Fig. 6E), and SVV infection enhanced the NCOA4-FTH1 interaction (Figure. 6 F). Consistently, the amount of precipitated FTH1 considerably increased in the presence of the 3D protein (Fig. 6G). TAX1BP1 is optional for NCOA4-FTH1 condensate formation; however, it was necessary for macro and microautophagy [44]. We found that 3D promoted the NCOA4-TAX1BP1 interaction (Fig. 6H, Figure S1). These results indicated that 3D enhanced the NCOA4-FTH1 interaction and caused FTH1 degradation, resulting in greater sensitivity to ferroptosis.

Fig. 6.

SVV 3D induce ferritinophagy enhancing NCOA4-FTH1 interaction (A) BHK-21 cells were transfected GFP-tagged SVV protein expression plasmids for 24 h. The protein expression level of TFR1, NCOA4, and ACSL4 was analyzed using immunoblotting with antibodies against TFR1, NCOA4, ACSL4, and β-actin as an internal control. (B-D) BHK-21 cells were cotransfected indicated plasmids for 24 h, respectively. Cells were stained with HA antibody, Flag antibody, myc antibody, and DAPI (blue), then examined by confocal microscopy. (E-F) Endogenous NCOA4-FTH1 interaction was examined with Co-IP assay after GFP-3D transfection (E) or SVV infection (F). (G, H) BHK-21 cells were cotransfected indicated plasmids for 24 h, respectively. Cell samples were prepared for Co-IP analysis

Discussion

Ferroptosis, a novel form of programmed cell death (PCD), is characterized by the iron-dependent accumulation of lethal levels of peroxidized lipids [21]. Reportedly, viruses adopt various strategies to induce ferroptosis and promote replication [33, 35–39]. The Epstein-Barr virus (EBV) promotes primary B cell lipid peroxidation to ferroptosis for latency [45]. Newcastle disease virus (NDV) increases intracellular ROS and lipid peroxides in tumor cells, and NDV-induced ferritinophagy promotes ferroptosis via ferrous iron release [37]. Enterovirus, a virus belonging to the family Picornaviridae, induces ferroptosis via ACSL4 [46]. Notably, hepatitis A virus (HAV), which is within the family Picornaviridae, the viral 3C^pro induces ferroptosis dependent on its protease activity [47]. The present study shows that SVV infection induces ferroptosis via inducing GPX4 degradation, whereas the NCOA4-medicated ferritinophagy, viral proteins 3C^pro and 3D synergistically contribute to the activation of ferroptosis signaling pathways.

ROS are of great significance in programmed cell death (PCD), such as necroptosis, apoptosis, autophagy, and ferroptosis [48]. An over-accumulation of ROS attacks biomembranes and propagates lipid peroxidation chain reactions that induce PCD [48]. The current study revealed that SVV infection induced ferroptosis, as evidenced by an increase in oxidized C11-BODIPY signaling (Fig. 1A-C). Further studies indicated a decline in cell viability and GSH levels after SVV infection, which was restored in the presence of Fer-1 (Figure. 1E-H). Additionally, the MDA and LDH levels were upregulated following SVV infection (Figure. 1I, J). Enterovirus 71 (EV71) triggers neural cell death via enhancing ROS production and reducing the activity of the Nrf2/HO-1 pathway [49]. We investigated the generation of ROS following SVV infection. It was discovered that either SVV infection or erastin treatment notably led to the accumulation of ROS and lipid ROS, accompanied by a significant reduction in MMP (Fig. 2A-G). ROS production was inhibited with Fer-1 and NAC, indicating that ferroptosis participates in SVV-induced ROS production.

Ferroptosis is controlled by multiple metabolic events and signaling pathways, including glucose-regulated AMPK signaling, Ecadherin-NF2-Hippo-YAP signaling, hypoxia signaling, and p53 and BAP1 tumor suppressor functions [50]. Reportedly, JEV induces ferroptosis through the YAP1-ACSL4 pathway, and ferroptosis contributes to JEV-induced neuroinflammatory and neuronal damage [39]. HSV-1-induced Nrf2 degradation contributes to ferroptosis, and is associated with viral encephalitis [35]. SARS-CoV-2 ORF3a degrades Nrf2 by recruiting Keap1 promoted cells to ferroptosis [38]. Ferroptosis is involved in inflammation [51].The mRNA transcription of inflammatory factors (CCL20, TNF- α, IL-6, and IL-1β) was much more significant than SVV infection with erastin; however, it was impaired in the Fer-1 treatment group (Figure S2). In addition, SVV infection exploits DNA damage response (DDR) to facilitate viral replication, SVV-induced DDR triggered NF-κB signaling accompanied by upregulation of pro-inflammatory cytokines [52]. Our data demonstrated that SVV-induced inflammatory effects are associated with ferroptosis.

GPX4 is a critical negative regulator of ferroptosis. SVV 3C^pro simultaneously targets GPX4 for degradation (Fig. 5), whereas SVV 3D drives the NCOA4-FTH1 interaction and promotes NCOA4-mediated ferritin degradation (Fig. 6). This indicates that more than one viral protein is involved in ferroptosis: 3C^pro and 3D induced ferroptosis by targeting the GPX4 axis and NCOA4-FTH1 interaction, respectively. However, SVV 3C^pro induces pyroptosis by cleaving gasdermin D (GSDMD) and induces host cell apoptosis by cleaving poly (ADP-ribose) polymerase (PARP) [16, 42, 43]. Overall, these results suggested that SVV 3C^pro induces cell death in multiple ways.

Ferritinophagy is medicated by adaptor NCOA4 through autophagic degradation of ferritin, causing the release of iron from lysosome to cytosol [28]. Inhibition of FTH1 expression results in loss of ferritin’s ability to store iron, leading to the accumulation of Fe²⁺ and induction of ferroptosis. NCOA4-FTH1 interaction forms punctate structures, and the SVV 3D protein drives NCOA4-FTH1 to form large condensates and promotes their interaction (Fig. 6C-H). Mycobacterium tuberculosis (Mtb) hijacks NCOA4-mediated ferritinophagy to induce autophagic ferritin degradation and therefore promoted intracellular growth [53, 54]. While human parainfluenza virus type 2 (hPIV-2) V protein interferes with interaction between FTH1 and NCOA4 and subsequent ferritinophagy [55], indicating that invading pathogens use different strategies regulating ferritinophagy for their effcient survival. TFR1 is a specific ferroptosis marker. Inhibiton of autophagy substantially dampened SVV-induced GPX4 and FTH1 degradation, impaired TFR1 and NCOA4 expression (Fig. 4A, B), thereby inhibiting ferroptosis, indicating that autophagy contributes to SVV-induced ferroptosis. Downregulating NCOA4 expression blocks SVV to induce FTH1 dagradation and LC3-II consversion, thus inhibiting ferritinophagy (Fig. 4C). Importantly, NCOA4 knockdown strikingly inhibit inflammatory responses (Figure S2). CVB3 infection induces TFR1 expression and enhances cellular iron level to lead to iron overload [56]. Overall, autophagy plays a positive role in regulating SVV-induced ferroptosis. Ferritin assembles to form large condensates that exhibit liquid-like properties, NCOA4-ferritin condensate formation is driven by NCOA4-FTH1 interaction that is responsible for ferritin degradation, and TAX1BP1 is dispensable for the condensates formation but required for its recognition by macroautophagy and microautophagy [44]. Tax1 binding protein 1 (TAX1BP1) serves as a direct interactor of NCOA4 required for lysosomal ferritin flux [57]. TAX1BP1 is necessary for clearance of insoluble NCOA4 condensates and ferritin via recruitment of FIP200 [58]. The hemagglutinin of influenza A virus interacts with NCOA4 and TAX1BP1, facilitating the formation of ferritin-NCOA4 condensates and inducing ferritinophagy [59]. African swine fever virus A151R downregulates cGAS-STING-mediated IFN-β production by promoting lipid peroxidation through ferritinophagy-induced ferroptosis [60]. We found that SVV 3D promoted NCOA4-TAX1BP1 condensate formation (Fig. 6C, D). FTH1 knockdown increased Mtb growth, and NCOA4 knockdown decreased Mtb growth [53, 54]. In this study, we found that FTH1 knockdown enhanced SVV growth, and NCOA4 knockdown diminished SVV growth (Fig. 4H), indicating that SVV-induced ferritinophagy facilitates viral replication. SVV employs dual mechanisms to induce ferroptosis (GPX4 degradation and ferritinophagy), it indicated that ferroptosis is critical for SVV replication. Picornaviruses specifically manipulate ferroptotic pathways, such as enterovirus-A71 preferentially infects and replicates in human motor neurons, inducing neurodegeneration by ferroptosis [61], and complement components modulate ferroptosis in coxsackievirus B3 (CVB3)-induced viral myocarditis through interaction with transferrin receptor (TFRC) [62].

Overall, our study proposes a model in which SVV infection sensitizes cells to ferroptosis by inducing ROS accumulation, lipid peroxidation, and glutathione (GSH) depletion (Fig. 7). SVV 3C^pro and 3D synergistically target GPX4 for degradation and promote NCOA4-mediated ferritinophagy to induce ferroptosis. These findings provide further insight into SVV pathogenesis, leading to the development of a potential target for intervention in iron metabolism against SVV infection.

Fig. 7.

Proposed model for SVV-induced ferroptosis SVV infection induces intracellular and mitochondrial ROS accululation, lipid peroxidation, mitochondrial membrane potential (MMP) decreasing, and glutathione (GSH) depletion, thereby triggering ferroptosis. SVV 3C^pro targets GPX4 for degradation to induce ferroptosis. SVV 3D protein drives NCOA4-FTH1 and NCOA4-TAX1BP1 interaction, promoting NCOA4-mediated ferritin degradation, therefore resulting in ferritinophagy, and further strengthened ferroptosis

Supplementary Information

Below is the link to the electronic supplementary material.

Authors’ contributions

Conceptualization: Jiangwei Song.

Data curation: Jingjing Yang, Ruiyi Ma.

Formal analysis: Ruiyi Ma, Shijie Xie.

Funding acquisition: Jiangwei Song.

Investigation: Jingjing Yang, Ruiyi Ma.

Methodology: Shijie Xie, Jiayao Su.

Project administration: Jiangwei Song, Jingjing Yang, Jiayao Su.

Resources: Zijian Li, Ruiyi Ma.

Supervision: Jiangwei Song.

Validation: Shijie Xie, Zijian Li.

Writing–original draft: Jingjing Yang, Peipei Cheng.

Writing–review & editing: Jiangwei Song.

Funding

This work was supported by the National Natural Science Foundation of China (32372980), the Reform and Development of Institute of Animal Husbandry and Veterinary Medicine, Beijing Academy of Agriculture and Forestry Sciences (XMS202521 and XMS202503), the Outstanding Young Foundation of Beijing Academy of Agriculture and Forestry Sciences (YXQN202302).

Data availability

The data that support the findings of this study are available from the corresponding author upon reasonable request.

Declarations

Conflict of interest

The authors declare no potential conflict of interest.