Spring Viremia of Carp Virus N Protein Negatively Regulates IFN Induction through Autophagy-Lysosome–Dependent Degradation of STING

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The IFN system is the first line of host defense against viral invasion. On viral infection, the host pattern recognition receptors recognize viral nucleic acids and activate a series of signal transduction reactions, which eventually initiate IFN expression. Retinoic acid–inducible gene I (RIG-I)-like receptors (RLRs) are a common class of pattern recognition receptors and consist of three members: RIG-I, melanoma differentiation-associated gene 5 (MDA5), and laboratory of genetics and physiology 2 (1, 2). Once host cells are infected by an RNA virus, RIG-I or MDA5 senses the viral RNA and recruits the mitochondrial antiviral signaling protein (MAVS, also known as VISA, IPS-1, or Cardif) and then transmits signals to the downstream stimulator of IFN genes (STING, also known as MITA, ERIS, or MYPS) and TANK-binding kinase 1 (TBK1) (3, 4). Subsequently, IFN regulatory factor 3 (IRF3) and IRF7 become activated after phosphorylation by TBK1, then translocate into the nucleus to trigger IFN transcription (5). To date, orthologs of the human RLR signaling pathway have been identified in fish (6). For example, zebrafish MAVS is a critical component of the RLR pathway to induce IFN expression, and it has a powerful antiviral function (7). Grass carp TBK1, an upstream kinase of both IRF3 and IRF7, plays a crucial role in the cellular antiviral response (8). Crucian carp STING displays an antiviral function through inducing an IRF3/7-dependent IFN response (9). Overall, the functions of fish RLRs are well conserved.

However, to avoid eradication, viruses have evolved numerous strategies to escape the host IFN immune response. STING is a pivotal molecule that participates in the RLR signaling pathway (4, 10). During a viral infection, STING acts as a scaffold protein that interacts with MAVS and facilitates TBK1-mediated phosphorylation of IRF3/7, resulting in IFN induction (11). STING-knockout mice exhibit high susceptibility to lethal vesicular stomatitis virus infections (12). Simultaneously, it has been reported that fish STING is also involved in the RLR pathway, which plays a vital role in the activation of IFN production and inhibition of RNA and DNA virus replication (13, 14). Fish STING also contains four N-terminal transmembrane (TM) domains and a C-terminal cytosolic dimerization domain (13). In addition, the dominant-negative fish STING mutant (STING-CT, lacking the N-terminal TM domains) loses its IFN induction activity compared with wild-type STING (15). Hence many viruses choose STING as a target to inhibit the host IFN response. For example, STING can be cleaved by the dengue virus protease NS2B3 (16). Moreover, STING can be degraded by MGF-505-7R of the African swine fever virus (ASFV) through the autophagy pathway or NS2B3 of the zika virus via the ubiquitin–proteasome pathway (17, 18). Our laboratory previously reported that grass carp reovirus VP41 suppresses the fish IFN response by reducing STING phosphorylation (19). Despite this, STING is always targeted by mammalian viruses, and it is still unclear whether STING is attacked by other aquatic viruses.

Spring viremia of carp virus (SVCV), which belongs to the genus Vesiculovirus of the family Rhabdoviridae, has caused severe mortality in common carp (Cyprinus carpio) and several freshwater fish species (20). SVCV is a negative-sense single-stranded RNA virus that encodes five proteins in the following order (3′–5′): nucleoprotein (N protein), phosphoprotein (P protein), matrix protein, glycoprotein, and viral RNA-dependent RNA polymerase (21, 22). The specific functions of the viral proteins involved in SVCV replication and proliferation are not yet known. Based on the studies of rhabdoviruses, the N protein is a highly rich protein that interacts with viral RNA to form the nucleocapsid. The P protein is a component of the nucleocapsid that associates with the N protein and viral RNA-dependent RNA polymerase protein to mediate the transcription process; the matrix protein is involved in the assembly and budding formation of the viruses; and the glycoprotein on the outer surface of the virus induces receptor-mediated endocytosis (20, 23). To date, only a few studies exploring the SVCV pathogenic mechanisms have been conducted. For instance, SVCV uses autophagy to promote viral replication (24). In addition, SVCV infection induces host oxidative stress to accelerate tissue injury in the affected fish (25). Moreover, our laboratory has revealed a few immune evasion mechanisms of SVCV to modulate the host IFN response. The SVCV N protein degrades MAVS via the ubiquitin–proteasome pathway, and the P protein reduces TBK1-mediated IRF3 phosphorylation (26, 27). However, much information concerning SVCV immune evasion strategies is still unknown and requires further investigation.

In this study, we further investigated the immune evasion strategies of SVCV and revealed that the SVCV N protein interacts with STING and degrades it through an autophagy-lysosome–dependent pathway, thereby inhibiting IFN production and promoting virus replication. These data provide new information for understanding SVCV pathogenesis and reveal a novel strategy for SVCV prevention in the aquaculture industry.

Epithelioma papulosum cyprini (EPC) cells were maintained at 28°C in 5% CO₂ in medium 199 (Invitrogen) supplemented with 10% FBS. SVCV was propagated in EPC cells until a cytopathic effect was observed; then the cultured medium with cells was stored at −80°C and prepared for use.

The cDNA fragment of SVCV N protein (MZ343157.1) was obtained by RT-PCR using RNA of cells infected with SVCV and cloned into pcDNA3.1(+), pCMV-Tag2C, or pCMV-Myc vectors. The open reading frames (ORFs) of zebrafish MAVS (NM_001080584.2), TBK1 (NM_001044748.2), STING (NM_001278837.1), IRF3 (NM_001143904), ATG5 (NM_001009914.2), Beclin1 (AB266448.1), and ATG9b (NM_001320078.1) were cloned into pCMV-hemagglutinin (HA), pCMV-Myc, pCMV-Tag2C, and pCS2-mCherry vectors, respectively. The short hairpin RNA of N protein, Pimephales promelas Beclin1 (XM_039685426.1) and Pimephales promelas MAVS (XM_039672710.1) were designed by BLOCK-iT RNAi Designer and cloned into the pLKO.1-TRC Cloning vector. The ORF of N protein was also inserted into p-enhanced GFP-N3 and pCS2-mCherry vectors to obtain an N plasmid labeled with GFP or red fluorescent protein. In addition, the plasmids containing zebrafish IFNφ1pro-Luc, IFNφ3pro-Luc, and IFN-stimulated response element (ISRE)-Luc in the pGL3-Basic luciferase reporter vector (Promega) were constructed as described previously (28). The Renilla luciferase internal control vector (pRL-TK) was purchased from Promega. All constructs were verified by sequencing, and primers used in this study are listed in Supplemental Table I. Polyinosinic-polycytidylic acid [poly(I:C)] was purchased from Sigma-Aldrich and used at a final concentration of 1 μg/ml.

EPC cells were seeded in 24-well plates overnight, and the monolayer cells were transfected with pCMV-Tag2C or Tag2C-N; then at 24 h posttransfection (hpt), the cells were mock or transfected with poly(I:C). After 24 hpt, the cells were lysed for measuring luciferase activity by the dual-luciferase reporter assay system (Promega) according to the manufacturer’s instructions. As for RLR-induced IFN promoter activation assay, various plasmids were at a ratio of 10:10:10:1 [expression vectors of pcDNA3.1(+)-N: pcDNA3.1(+)-MAVS/STING/IRF3: IFNφ1pro/IFNφ3pro/ISRE-Luc: pRL-TK]. The empty vector pcDNA3.1(+) was used to ensure equivalent amounts of total DNA in each well.

Transient transfections were performed in EPC cells seeded in 6-well or 24-well plates by using FishTrans (MeiSenTe Biotechnology) according to the manufacturer’s protocol. For the antiviral assay, using 24-well plates, EPC cells were transfected with the indicated plasmids. At 24 hpt, cells were infected with SVCV at a multiplicity of infection (MOI = 0.001). After 48 or 72 h, supernatant aliquots were harvested for detection of virus titers; the cell monolayers were fixed by 4% paraformaldehyde (PFA) and stained with 1% crystal violet for visualizing the cytopathic effect. For virus titration, 100 μl of culture medium was collected at 48 h postinfection and used for plaque assay. The supernatants were subjected to 10-fold serial dilutions and then added (100 μl) onto a monolayer of EPC cells cultured in 96-well plates. After 48 or 72 h, the medium was removed and the cells were washed with PBS, fixed by 4% PFA, and stained with 1% crystal violet. The virus titer was expressed as 50% tissue culture-infective dose (TCID₅₀/ml).

Total RNAs were extracted by the TRIzol reagent (Invitrogen). First-strand cDNA was synthesized by using the PrimeScript RT reagent Kit with gDNA Eraser (Takara). Quantitative real-time PCR (qPCR) was performed with Fast SYBR green PCR Master Mix (Bio-Rad) on a CFX96 Real-Time System (Bio-Rad). PCR conditions were as follows: 95°C for 5 min and then 42 cycles of 95°C for 20 s, 60°C for 20 s, and 72°C for 20 s. The primers for qPCR are as listed in Supplemental Table I. β-actin gene was used as an internal control. The relative fold changes were calculated by comparison with the corresponding controls using the 2^−ΔΔCT (where CT is threshold cycle) method.

EPC cells seeded in 10-cm² dishes overnight were transfected with the total of 10 μg plasmids indicated in the figure legends. At 24 hpt, the medium was removed carefully, and the cell monolayer was washed with 10 ml ice-cold PBS. Then the cells were lysed in 1 ml of radioimmunoprecipitation lysis buffer (1% Nonidet P-40, 50 mM Tris–HCl [pH 7.5], 150 mM NaCl, 1 mM EDTA, 1 mM NaF, 1 mM sodium orthovanadate [Na₃VO₄], 1 mM PMSF, 0.25% sodium deoxycholate) containing protease inhibitor mixture (Sigma-Aldrich) at 4°C for 1 h on a rocker platform. The cellular debris was removed by centrifugation at 12,000 × g for 15 min at 4°C. The supernatant was transferred to a fresh tube and incubated with 15 μl anti–HA-agarose beads or anti-Flag affinity gel (Sigma-Aldrich) overnight at 4°C with constant agitation. These samples were further analyzed by immunoblotting (IB). Immunoprecipitated proteins were collected by centrifugation at 5000 × g for 1 min at 4°C, washed three times with lysis buffer, and resuspended in 100 μl 1 × SDS sample buffer. The immunoprecipitates and whole-cell lysates (WCLs) were analyzed by IB with the indicated Abs.

WCLs and immunoprecipitates were separated by electrophoresis on 10% SDS-PAGE and then transferred to polyvinylidene fluoride membrane. The membranes were blocked for 1 h at room temperature in TBST buffer (25 mM Tris–HCl, 150 mM NaCl, 0.1% Tween 20 [pH 7.5]) containing 5% nonfat dry milk. Then they were incubated with the primary Abs indicated on the figures at an appropriate dilution overnight at 4°C, washed three times with TBST, and then incubated with secondary Abs for 2 h at room temperature. After washing three times with TBST, the membranes were stained with Immobilon Western chemiluminescent HRP substrate (Millipore) and detected by using an ImageQuant LAS 4000 system (GE Healthcare). Abs were diluted as follows: anti–β-actin (Cell Signaling Technology) at 1:5000, anti-Flag/HA (Sigma-Aldrich) at 1:3000, anti-Myc (Santa Cruz Biotechnology) at 1:3000, and HRP-conjugated anti-rabbit IgG or anti-mouse IgG (Thermo Scientific) at 1:5000. The indicated Abs of SVCV proteins (includes N and P), EPC TBK1, and EPC STING at 1:1000 were prepared by our laboratory.

EPC cells were plated onto coverslips in six-well plates and transfected with the indicated plasmids for 24 h. Then the cells were washed twice with PBS and fixed with 4% PFA for 1 h. After being washed three times with PBS, the cells were treated with 1 μg/ml DAPI (Beyotime) in the dark at room temperature. Finally, the coverslips were washed and observed with a confocal microscope under a 63× oil immersion objective (SP8; Leica).

EPC cells were seeded in six-well plates and transfected with indicated plasmids for 24 h. Then cells were washed three times with PBS, trypsinized, and transferred to a 1.5-ml centrifuge tube. Cell precipitation was collected by centrifugation at 2000 × g for 5 min. The cell pellets were resuspended with 2.5% glutaraldehyde in 0.075 mol/l phosphate buffer (pH 7.4) for 4 h at 4°C for prefixation. Then the cells were washed three times with solution containing 0.075 mol/l phosphate and 0.19 mol/l sucrose for 15 min each and postfixed in 1% osmium tetroxide (O_sO₄) in 0.24 mol/l phosphate buffer (pH 7.4) for 2 h. Then the cells were dehydrated with a graded series of ethanol and acetone and gradually infiltrated with epoxy resin. Samples were continuously polymerized at 37°C overnight and then polymerized at 60°C for 48 h. Ultrathin sections (74 nm) were cut using a microtome (UC7; Leica) and mounted on copper slot grids. Sections were doubly stained with 3% uranyl acetate-lead citrate for 10 min and observed under transmission electron microscope (HT7700; Hitachi).

Luciferase and qPCR assay data are expressed as the mean ± SEM. The p values were calculated by Student t test. A p value <0.05 was considered statistically significant.

Our previous study demonstrated that the SVCV N protein degrades MAVS and inhibits IFN induction, ultimately leading to host immune response escape (27). But during host–virus coevolution, SVCV adopted various tactics to block host antiviral signaling. Because of the relatively limited SVCV genome capacity, it is possible that one encoded protein may perform multiple functions to enable immune escape. Thus, to further explore the other strategies used by SVCV to invade the host, the SVCV N protein was chosen for subsequent study. Four type I IFNs (IFNφ1–IFNφ4) have been identified in zebrafish, and only IFNφ1 and IFNφ3 can be induced by poly(I:C) (a mimic of viral RNA). Thus, the IFNφ1 promoter (IFNφ1pro) and IFNφ3pro activities were detected in a luciferase assay. As shown in (Fig. 1A and 1B, poly(I:C) stimulation remarkably induced IFNφ1pro and IFNφ3pro activities, whereas these inductions were significantly suppressed by overexpression of the N protein. ISRE is considered a transcription factor binding motif in the promoter regions of IFN and IFN-stimulated genes (ISGs), which facilitates gene transcription (29). Consistently, poly(I:C)-induced ISRE activation was also antagonized by the N protein (Fig. 1C). Meanwhile, overexpression of N protein also blocked SVCV-activated IFNφ1/3pro and ISRE activities (Fig. 1D–F). Moreover, at the mRNA level, the ifn, vig1, and isg15-1 transcripts in EPC cells were upregulated by poly(I:C) or SVCV stimulation, and a strong decline was observed in the N protein group (Fig. 1G–L). These data suggest that SVCV N protein acts as a negative regulator to impede host IFN production.

It has been reported that the fish RLR signaling pathway plays a vital role in the activation of IFN production (6, 30, 31). To probe whether the N protein affects the capacity of RLR-induced IFN expression, we employed RLR key molecular constructs and IFN promoters. First, the RLR key molecules significantly activated IFN expression, and overexpression of the N protein suppressed MAVS- and STING-mediated IFN activation remarkably, but slightly affected IRF3-induced IFNφ1pro activity (Fig. 2A). Second, IFNφ3pro and ISRE were also monitored. Similarly, the N protein dampened MAVS/STING-induced activation (Fig. 2B, 2C). In addition, these inhibitions were displayed in a dose-dependent manner (Fig. 2D–F). IRF3 is the downstream molecule of the RLR signaling pathway that induces IFN expression individually. Thus, these data indicate N protein interferes with the RLR signaling pathway by targeting MAVS/STING.

Given that the N protein negatively regulates RLR molecule-induced IFN activation, it was necessary to investigate whether the N protein associates with RLR molecules at the protein level. First, a coimmunoprecipitation assay was conducted in EPC cells cotransfected with N-Myc and MAVS/STING/IRF3 with HA-tag, respectively. The results showed that the anti-HA Ab-immunoprecipitated protein STING was recognized by the Myc-tagged N protein (Fig. 3A). In a reverse assay, the anti-Myc Ab–immunoprecipitated N protein was recognized by the HA-tagged STING protein (Fig. 3B), which meant that the N protein interacts with STING. Next, the subcellular localizations of the N protein with the RLRs were monitored in EPC cells. Confocal microscopy analysis revealed that the N protein–enhanced GFP signals were mainly distributed in the cytoplasm in a plaque-like aggregation and partially overlapped the red signals from MAVS/STING/IRF3 (Fig. 3C). Then, to determine which STING functional domain interacts with the N protein, we constructed two truncated STING mutants: STING-N (lacking the C terminus) and STING-C (lacking the N-terminal TM region) (Fig. 3D). Consistent with wild-type STING, STING-N bound with the N protein, whereas this interaction was abolished in the STING-C group (Fig. 3E). These data suggest that N protein is distributed in the cytoplasm and interacts with the N terminus of STING.

To further investigate the specific regulatory mechanism by which the N protein modulates STING, we determined the effect of the N protein on the RLR molecules at the protein level. Overexpression of the N protein reduced the abundance of MAVS and STING, while hardly affecting TBK1 expression (Fig. 4A). In addition, there was a marked decline in STING with increasing doses of the N protein (Fig. 4B). Moreover, endogenous STING was also substantially decreased with overexpression of the N protein (Fig. 4C). In contrast, the expression of endogenous TBK1 did not appear to change (Fig. 4D). Furthermore, we examined whether the N protein–induced reduction in STING expression occurred at the transcriptional level. Overexpression of the N protein had no obvious effect on sting transcripts (Fig. 4E), meaning that the N protein reduces STING expression at the protein level. Given that our previous study revealed SVCV N protein degrades MAVS through the ubiquitin–proteasome pathway (27), it is necessary to explore whether the effect of SVCV N protein on STING is independent of its effect on MAVS. As shown in (Fig. 4F and 4G, knockdown of MAVS did not affect the degradation of STING by N protein. Meanwhile, the inhibition of STING-induced IFN production by N protein did not change in the MAVS-knockdown group (Fig. 4H, 4I). Furthermore, the N protein–induced degradation of STING disappeared in the N protein knockdown group (Fig. 4J). Meanwhile, SVCV stimulation inhibits STING-induced IFN expression, but knockdown of N protein of SVCV restores STING-activated IFN production (Fig. 4K). Next, to explore the specific degradation mechanism for STING, several reagents, including MG132 (the ubiquitin–proteasome pathway inhibitor), 3-methyladenine (3-MA; the autophagy pathway inhibitor), and NH₄Cl (the late stage of autophagy/lysosomal pathway inhibitor), were used to treat the cells. The N protein–mediated degradation of STING was rescued by the autophagy-related inhibitors 3-MA and NH₄Cl, but not by MG132 (Fig. 4L). In addition, STING protein levels were gradually restored with increasing doses of 3-MA and NH₄Cl (Fig. 4M, 4N). These results suggest that the N protein induces STING degradation through the autophagy-lysosome pathway and independent of its effect on MAVS.

Because the earlier results show that the N protein degrades STING in an autophagic manner, we predicted that the N protein induces cellular autophagy. The detection of LC3 by IB or immunofluorescence has become a reliable method for monitoring autophagy and autophagy-related processes (32). Thus, the cellular lipidized LC3-II levels were detected in cells overexpressing N protein. Interestingly, LC3-II remained unchanged by different doses of the N protein (Fig. 5A). To exclude the possibility that the N protein–induced cellular autophagic process was completed before the samples were harvested and thus caused undetectable changes in LC3-II, we monitored the LC3-II level by treatment with 3-MA and NH₄Cl. Consistent with the earlier results, the LC3-II band remained unchanged, suggesting that the N protein alone cannot cause cellular autophagy (Fig. 5B, 5C). Similarly, individual overexpression of STING did not influence LC3-II under normal conditions or after reagent treatment, respectively (Fig. 5D–F). It is well known that the aggregation of LC3-GFP into punctate structures represents an increased autophagy level (32). Few punctate LC3-GFPs were observed in cells transfected with the N protein or STING (Fig. 5G). Meanwhile, transmission electron microscopy (TEM) was performed for ultrastructural analysis. The autophagosome-like vesicles were barely observed in individual overexpressed N protein or STING cells (Fig. 5H). Thus, we conjectured that STING was the specific target and essential for the N protein to promote autophagic degradation. To prove this hypothesis, the N protein and STING were coexpressed in cells and treated with 3-MA and NH₄Cl. As anticipated, LC3-II enhancement was observed in the NH₄Cl-treated groups (Fig. 5I). In addition, obvious green punctate LC3-GFP signals of accumulation and significant autophagosome-like vesicles with cytosolic contents were observed in cells cotransfected with the N protein and STING (Fig. 5J, 5K). To further explore the mechanism of N protein–mediated STING autophagic degradation, we detected the interactions of key autophagic system molecules with N protein. As exhibited in (Fig. 5L, EPC cells cotransfected with N-Flag and Beclin1/ATG5/ATG9b with Myc-tag, respectively. The results showed that only the anti-Myc Ab–immunoprecipitated protein Beclin1 was recognized by the Flag-tagged N protein. Next, short hairpin RNA against beclin1 was designed for subsequent assays, and the N protein–induced STING degradation was notably impaired by beclin1 knockdown (Fig. 5M). In addition, the inhibitory effect of N protein on STING-induced IFN production was significantly diminished in the presence of beclin1 knockdown (Fig. 5N, 5O). Collectively, these data demonstrate that coexpression of the N protein and STING is necessary to induce cellular autophagy to degrade STING, and Beclin1 is an essential factor in this process.

It has been reported that STING serves as a scaffold protein to recruit TBK1 and IRF3, thus facilitating IRF3 activation and leading to IFN production (5). This event also exists in fish (26). Therefore, the effect of the N protein on the STING–TBK1 interaction was explored, and the N protein dramatically impaired this association. Meanwhile, 3-MA treatment was employed to block the degradation of STING by N protein before coimmunoprecipitation assay and found that the interaction between STING and TBK1 was slightly changed when STING degradation was blocked. This means that the weakening of the interaction between TBK1 and STING was caused by N protein–mediated degradation of STING (Fig. 6A). Furthermore, we investigated whether the N protein influences STING-induced IFN production. At the mRNA level, STING induced the expression of ifn and other ISGs, but this activation was attenuated by overexpression of the N protein (Fig. 6B–E). These data suggest that the N protein impairs STING–TBK1 interaction and weakens STING-induced IFN response.

Fish STING is a powerful antiviral factor and plays a vital role in defending the host against DNA and RNA viruses (13, 14). Thus, it needs to be determined whether the N protein affects STING-induced cellular antiviral responses. As depicted in (Fig. 7A and 7B, overexpression of STING protected the cells from viral infection and reduced the viral titer 3162-fold (10^8.83 to 10^5.33 TCID₅₀/ml), as compared with that in the control group. In contrast, simultaneous overexpression of the N protein weakened the protective effect of STING on cells and increased the viral titer 17.39-fold (10^5.33–10^6.57 TCID₅₀/ml), as compared with that in the STING overexpression group. In addition, at both the mRNA and the protein level, overexpression of STING significantly suppressed viral gene expression, whereas the N protein disrupted this inhibition (Fig. 7C–G). These results suggest that the N protein has a negative regulatory effect on STING-mediated cellular antiviral responses.

As in mammals, the fish IFN system plays an important role in the defense against viruses. On viral infection, the host cells activate a series of signaling pathways to induce IFN expression and eliminate the viruses (33). Meanwhile, aquatic viruses have evolved a variety of strategies to resist the host IFN response to permit viral replication and proliferation. The aquatic virus SVCV causes high mortality in cultured fish industries; however, little is known about the pathogenic mechanisms by which SVCV modulates the host immune response. In this article, we report that the SVCV N protein interacts with STING and promotes its degradation through an autophagy-lysosome–dependent pathway, thereby inhibiting IFN induction. This finding broadens the understanding of SVCV pathogenesis.

It is well known that there are two main protein degradation mechanisms in eukaryotic cells: the ubiquitin–proteasome system and the autophagy-lysosome pathway. Generally, viruses also use these two degradation systems to modulate host proteins to achieve immune escape. For instance, pI215L of ASFV negatively regulates the cyclic GMP-AMP synthase–STING signaling pathway through recruiting RING-finger protein 138 to inhibit the k63-linked ubiquitination of TBK1. In addition, the A137R protein of ASFV inhibits IFN production via the autophagy-mediated lysosomal degradation of TBK1 (34, 35). Considering this information, a similar mechanism may exist in aquatic viruses. Our previous study demonstrated that the SVCV N protein degraded MAVS through the ubiquitin–proteasome system to block IFN production (27). In addition, in this study, the SVCV N protein degraded STING via the autophagy-lysosome pathway. This shows that aquatic viruses also use the host’s degradation system to attack their targets.

It has been reported that STING plays a vital role against infection by DNA and RNA viruses (12, 13). Consequently, viruses use multiple strategies to intercept STING signaling, including interaction disruption, cleavage, and posttranslational modification (16, 36, 37). There are multiple reasons for choosing STING as a target compared with other molecules. First, STING acts both as a DNA sensor by direct binding of cyclic dinucleotides translocating from the endoplasmic reticulum (ER) to the Golgi and as a scaffold protein to facilitate interactions between TBK1 and IRF3, resulting in IFN induction. Second, a recent study revealed that STING is also involved in regulating multiple cellular processes, including autophagy, cellular stress responses, and metabolism (38). Thus, it is effective for viruses to escape the host IFN response to disable STING function. In this study, the SVCV N protein interacted with the TM region of STING, which is required for STING to anchor to the ER. However, it is unknown whether this interaction affects the translocation of STING from the ER to the Golgi.

Autophagy is an evolutionarily conserved intracellular recycling process that maintains cellular homeostasis by sequestering and degrading cytoplasmic targets, including cellular organelles and microbes or their products. It has been demonstrated that autophagy is involved in diverse physiological processes, such as cell differentiation, cell death, and immune responses (39–42). On viral invasion, the host often activates autophagy to reduce infection by transporting the virus to the lysosome for degradation. However, some viruses have evolved strategies to subvert or hijack the autophagic process to escape host immune responses and benefit their own replication. For instance, the ORF3a of the coronavirus disease 2019 virus (severe acute respiratory syndrome coronavirus 2) interacts with and sequestrates vacuolar protein sorting 39, the component of homogeneous fusion and protein sorting, thereby preventing the interaction of the homogeneous fusion and protein sorting complex with autophagosome-localized syntaxin 17, thus inhibiting autophagic activity and achieving immune escape (43). Foot-and-mouth disease virus activates autophagy to facilitate viral trafficking and infection (44). As for SVCV, it can reportedly induce autophagy to promote its own replication (24). Our study revealed that the SVCV N protein uses the autophagy pathway to degrade STING and facilitate its amplification. However, further exploration is needed to determine whether there are other SVCV-encoded proteins that use autophagy to help SVCV achieve immune escape.

Taken together, this study reveals an important mechanism used by the SVCV N protein to suppress IFN production through degrading STING via the autophagy-lysosome–dependent pathway. These data further reveal the specific mechanisms of how SVCV uses autophagy to achieve immune escape and provide new ideas for the preventive treatment of SVCV.

We thank Fang Zhou (Analysis and Testing Center, Institute of Hydrobiology [IHB], Chinese Academy of Sciences [CAS]) for assistance with confocal microscopy analysis and Dr. Feng Xiong (China Zebrafish Resource Center, IHB, CAS) for assistance with qPCR analysis. We thank Yuan Xiao and Zhen-Fei Xing (Analysis and Testing Center, IHB, CAS) for assistance with TEM observation.

The authors have no financial conflicts of interest.