The alpha-coronavirus E protein inhibits the JAK-STAT pathway signaling by triggering STAT2 degradation through OPTN- and NBR1-mediated selective autophagy

ABSTRACT

The zoonotic transmission of coronaviruses continues to pose a considerable threat to humans. Swine acute diarrhea syndrome coronavirus (SADS-CoV), a bat coronavirus related to HKU2, causes severe economic losses in the pig industry and has the potential to trigger outbreaks in humans. However, our understanding of how SADS-CoV evades the host’s innate immunity remains limited, hindering effective responses to potential human outbreaks. In this study, we demonstrate that the SADS-CoV envelope protein (E) inhibits type I interferon (IFN-I) signaling by inducing the degradation of STAT2 via the macroautophagy/autophagy-lysosome pathway. Mechanistically, the E protein evades host innate immunity by promoting STAT2 degradation through autophagy, mediated by the NBR1 and OPTN receptors. Notably, ubiquitination of E protein is required for the autophagic degradation of STAT2. Additionally, lysine residue K61 of the E protein is crucial for its stable expression; however, it is not involved in its ubiquitination. In conclusion, our study reveals a novel mechanism by which the E protein disrupts IFN-I signaling by targeting STAT2 via autophagy, enhancing our understanding of SADS-CoV’s immune evasion strategies and providing potential drug targets for controlling viral infections.

Abbreviations: 3-MA: 3-methyladenine; ATG: autophagy related; BafA1: bafilomycin A₁; BSA: bovine serum albumin; CALCOCO2/NDP52: calcium binding and coiled-coil domain 2; CC: coiled-coil; CHX: cycloheximide; Co-IP: co-immunoprecipitation; DAPI: 4’,6-diamidino-2-phenylindole; DBD: DNA-binding domain; DMEM: Dulbecco’s Modified Eagle’s medium; DMSO: dimethyl sulfoxide; E, Envelope. FW: four-tryptophan; GAPDH: glyceraldehyde-3-phosphate dehydrogenase; HA: hemagglutinin; hpt: hours post-treatment; IF: indirect immunofluorescence; IFNB/IFN-β: interferon beta; IgG: immunoglobulin G; ISG: IFN-stimulated genes; ISRE: interferon-stimulated response element; MAP1LC3/LC3: microtubule associated protein 1 light chain 3; MOI: multiplicity of infection; NBR1: NBR1 autophagy cargo receptor; OPTN: optineurin; PBS: phosphate-buffered saline; PRRs: pattern recognition receptors; qPCR: quantitative polymerase chain reaction; SAR: selective autophagy receptor; SQSTM1/p62: sequestosome 1; STAT: signal transduction and activator of transcription; TBS-T: Tris-buffered saline with Tween 20; TCID50: 50% tissue culture infective dose; TOLLIP: toll interacting protein; Ub: ubiquitin; UBA: C-terminal ubiquitin-associated; VSV: vesicular stomatitis virus; WB: western blotting. WT: wild type.

Introduction

Over the past two decades, coronaviruses have posed considerable threats to human health, life, and economic development [1]. Swine acute diarrhea syndrome coronavirus (SADS-CoV), also known as porcine enteric alphacoronavirus (PEAV), is a novel, highly pathogenic α-CoV that primarily causes diarrhea and vomiting in piglets, with a mortality rate of nearly 90% in piglets younger than 5 days [2]. SADS-CoV was first identified in Guangdong province, China, in 2017 [3] and has since been reported in various regions of China in 2019, 2021, and 2023. It is suggested that SADS-CoV circulates sporadically in epidemic patterns within China [4–6]. Originating from HKU2, an α-CoV commonly found in bats, SADS-CoV crossed species through an unknown intermediate host [7]. The virus exhibits broad cell tropism and the potential for host transmission [8,9]. Notably, SADS-CoV has been shown to infect a wide range of primary human lung and intestinal cells, raising concerns about its ability to infect humans in vivo [8]. This increases the likelihood of future zoonotic disease transmission.

SADS-CoV is an enveloped, positive-sense, single-stranded RNA virus with a genome of approximately 27 kb, encoding 16 nonstructural proteins, four structural proteins, and three accessory proteins [3,10]. The envelope (E) protein, a small integral membrane protein, plays a key role in viral assembly, budding, formation, and pathogenesis [11–13]. Although the role of E in viral replication has been extensively studied [14], little is known about how E regulates the pathogenesis of α-coronaviruses.

Innate immunity is the host’s first line of defense against viral infection, and a virus’s ability to overcome type-I interferon (IFN-I) innate immune defenses is critical for establishing a successful infection [15,16]. Pattern recognition receptors detect pathogen-associated molecular patterns, leading to IFN-I production. IFN-I is a crucial component of innate immunity, produced by nearly all nucleated cell [17,18]. Once secreted, IFN-I binds to the IFN-I receptor (IFNAR) and triggers the expression of interferon-stimulated genes (ISGs) via the JAK (Janus kinase)-STAT (signal transducer and activator of transcription) signaling pathway [19]. STAT2, a key component of the JAK-STAT pathway, is recruited by IFNAR, phosphorylated by JAK1, and subsequently translocates to the nucleus with phosphorylated STAT1 (p-STAT1) and IRF9 to activate ISG transcription [18,20].

Macroautophagy, commonly referred to as autophagy, is an essential cellular homeostatic process that controls the quality and quantity of intracellular components by targeting cytoplasmic elements for degradation [21,22]. Autophagy plays a pivotal role in immune system development, supporting both innate and adaptive immune responses while directly combating intracellular microbes as a cell-autonomous defense mechanism [23,24]. It is also part of the host’s stress response and is frequently triggered by viral infections [25]. Coronaviruses often exploit components of the autophagy pathway to facilitate their replication [26]. Selective autophagy degrades specific targets such as damaged organelles, aggregating proteins, or invading bacteria, thereby critically participating in cellular quality control [27]. This process involves selective autophagy receptor (SAR) activity, which mediates the targeted degradation of intracellular components [28]. The concept of selective autophagy in mammalian cells emerged with the discovery of SQSTM1/p62 (sequestosome 1), a substrate and SAR that facilitates the degradation of ubiquitinated cargo via autophagy [29]. Ubiquitination has been proposed as a signal for selective autophagy, and several ubiquitin (Ub)-dependent SARs have been identified in recent years [30]. These include NBR1 (NBR1 autophagy cargo receptor), OPTN (optineurin), CALCOCO2/NDP52 (calcium binding and coiled-coil domain 2), and TOLLIP (toll interacting protein), which can bind both cargo and Ub, initiating pathways that trigger autophagy and membrane recruitment [27,31,32].

In this study, we demonstrated that the SADS-CoV envelope is highly immunosuppressive, promoting the selective autophagic degradation of STAT2, a critical protein in the IFN-I signaling pathway.

Results

SADS-CoV was sensitive to IFN-I pre-treatment

IFN-I plays a crucial role in limiting viral replication and regulating host antiviral immune responses. Due to the limited research on the sensitivity of SADS-CoV to IFN-I, we designed experiments to evaluate the effects of IFN-I pre-treatment and post-treatment on SADS-CoV (Figure 1A). Vesicular stomatitis virus (VSV), which is highly sensitive to IFN-I, was used as a control [33]. We used Vero E6 cells for these studies because they support robust replication of SADS-CoV and can respond to recombinant IFN despite failing to secrete IFN-I [34]. As shown in Figure 1B,C, high concentrations of IFNB/IFN-β in the pre-treatment group considerably reduced SADS-CoV replication in Vero cells and the infection levels in Vero, HeLa, and IPI-2I cells. In the post-treatment group, IFNB was far less effective in inhibiting the replication and infection levels of SADS-CoV and VSV-GFP than in the pre-treatment group (Figure 1B,C). VSV-GFP exhibited greater sensitivity to IFNB in both treatment groups. The antiviral effect of IFN is well known to be most effective when administered prior to infection [35]. Reduced efficacy in already infected cells is often attributed to insufficient time for ISG products to be transcribed and translated [36], as well as the virus’s damaging effects on cells. Indeed, many viruses have evolved specific mechanisms to evade IFN-I [37], often by synthesizing viral proteins that block IFN production or signaling pathways, as seen in our previous study [38]. In conclusion, SADS-CoV was sensitive to IFN-I pre-treatment; however, its sensitivity decreased after infection was established.

Figure 1.

Sensitivity analysis of SADS-CoV to IFN-I. Vero E6 cells were treated with varying concentrations of IFNB for 12 h and subsequently infected with SADS-CoV (MOI = 0.1) or VSV-GFP (MOI = 0.1) for 24 h. Alternatively, vero E6 cells were first infected with SADS-CoV (MOI = 0.1) or VSV-GFP (MOI = 0.1) for 24 h, followed by treatment with different concentrations of IFNB for 12 h. (A) schematic representation of IFN-I treatment timing addition and virus infection time points. (B) the effect of IFNB on SADS-CoV and VSV-GFP replication was assessed. SADS-CoV replication was evaluated through immunostaining for the viral nuclear protein; in contrast, VSV-GFP replication was analyzed via fluorescence microscopy after cells were fixed and stained with DAPI. Scale bar: 100 μm. (C) the ability of ifnb-treated SDS-CoV to infect different cell lines was determined. Supernatants were collected at 36 h post-infection for both groups, and SADS-CoV infection in various cell types was assessed by calculating the median tissue culture infectious dose (TCID50). The data are representative of three independent experiments (C). *p < 0.05, **p < 0.01, ***p < 0.001.

SADS-CoV antagonizes IFN-I signaling

Currently, few studies have investigated the immune evasion mechanisms of SADS-CoV, with most published research focusing on the upstream pathway of IFN-I [39,40]. Little is known about how SADS-CoV antagonizes the IFN-I-induced JAK-STAT pathway. To explore whether SADS-CoV interferes with IFN-I signaling, we examined interferon-stimulated response element (ISRE)-dependent gene expression in both mock-infected and virus-infected cells. As shown in Figure 2A, IFNB stimulation significantly increased ISRE promoter activity. However, in SADS-CoV-infected cells, IFNB-induced ISRE promoter activity was significantly reduced and inversely correlated with the level of viral infection. Quantitative polymerase chain reaction (qPCR) analysis further confirmed that the mRNA levels of ISG15, IFIT2/ISG54, and OAS1 in HeLa and IPI-2I cells, typically induced by IFNB, were markedly suppressed by SADS-CoV infection, with inhibition correlating negatively with the infection dose (Figure 2B,C).

Figure 2.

SADS-CoV antagonizes IFN-I signaling. (A) SADS-CoV inhibited the promoter activity of ifnb-activated ISRE in a dose-dependent manner. HeLa cells in 24-well plates were co-transfected with the ISRE promoter-driven firefly luciferase reporter plasmid ISRE-Luc (125 ng) and the renilla luciferase control plasmid phRluc-tk (25 ng), followed by infection with SADS-CoV at various MOI (0.01, 0.1, and 1). At 24 h post-infection, cells were treated with 1,000 U/mL of IFNB for 12 h and subjected to dual-luciferase reporter assays. (B and C) SADS-CoV inhibited ifnb-activated ISG transcription in HeLa and IPI-2I cells in a dose-dependent manner. Both cell lines were infected with SADS-CoV at different MOIs (0.01, 0.1, and 1) for 24 h, followed by treatment with 1,000 U/mL of IFNB for 12 h. RNA was extracted, and rt-qPCR was performed to evaluate ISG15, IFIT2, and OAS1 mRNA levels, with GAPDH as an internal control. (D and E) SADS-CoV infection inhibited the activation of p-STAT2, p-STAT1 and STAT2. Vero and IPI-2I cells were infected with either live or uv-inactivated SADS-CoV (MOI = 0.1) for 24 h, followed by IFNB treatment for 2 h. Protein expression was assessed by WB analysis. (F) identification of proteins inhibited by SADS-CoV in IFN-I signaling. HEK293T cells in 24-well plates were co-transfected with isre-luc (125 ng), hRluc-tk (25 ng), and either SADS-CoV protein or an empty vector (400 ng). After 24 h, cells were treated with 1,000 U/mL of IFNB for 12 h, followed by a dual-luciferase reporter assay. The inhibited protein is marked by a red “*.” the data are representative of three independent experiments (A, B, C and F). *p < 0.05, **p < 0.01, ***p < 0.001.

Next, we investigated the expression of p-STAT1 and p-STAT2 along with STAT1 and STAT2 in Vero E6 and IPI-2I cells to uncover the mechanism by which SADS-CoV antagonizes IFN-I signaling. To determine whether viral replication was required for this inhibitory effect, we used ultraviolet light to inactivate a group of SADS-CoV virions. As shown in Figure 2D,E and Figure S1A, B, STAT2 protein expression was significantly reduced in SADS-CoV-infected cells. The inhibition of p-STAT1 and p-STAT2 was also significant, with p-STAT1 appearing to be more substantially affected. Interestingly, SADS-CoV infection caused a slight increase in STAT1 protein levels, which may be related to a mechanism similar to that seen in porcine epidemic diarrhea virus, where p-STAT1 inhibition occurs through enhanced STAT1 acetylation via HDAC1 activity inhibition [41].

Finally, using a dual-luciferase reporter assay, we screened for SADS-CoV proteins that antagonize IFN-I signaling. The results showed that 9 proteins (nsp1, nsp2, nsp5, nsp7, nsp9, nsp12, nsp13, nsp16, E, M) significantly inhibited IFN-I-activated ISRE promoter activity (Figure 2F). Although western blotting (WB) failed to detect the expression of nsp6, nsp12, ns3, and M protein, their normal expression was confirmed using immunofluorescence assays (Figure S1C).

STAT2 is degraded by the SADS-CoV envelope protein via the autophagy pathway

Because SADS-CoV strongly inhibits STAT2 protein expression, we hypothesized that this inhibition contributes to its suppression of IFN-I signaling. To identify the specific proteins responsible for STAT2 degradation by SADS-CoV, we tested eight proteins that inhibit IFN-I signaling and found that the SADS-CoV E protein was the most effective in degrading STAT2 (marked with a red “*”) (Figure 3A). In eukaryotic cells, protein degradation is primarily controlled by three systems: the Ub-proteasome system, the autophagy-lysosome pathways, and caspase activation via apoptosis [42,43]. To determine the mechanism by which E protein induces STAT2 degradation, we treated E protein-transfected cells with several inhibitors: the proteasome inhibitor MG132, the autophagy inhibitor 3-methyladenine (3-MA), the autophagosome-lysosome fusion inhibitor bafilomycin A₁ (BafA1), the lysosome inhibitor NH₄Cl, and the caspase inhibitor Z-VAD-FMK. The results showed that E protein induced STAT2 degradation primarily through the autophagy-lysosome pathway (Figure 3B,C). The extent of STAT2 degradation was significantly and negatively correlated with the dose of transfected E protein (Figure 3D). Moreover, E protein promoted LC3 lipidation (Figure 3B–D). Additionally, STAT2 and SQSTM1/p62 protein levels decreased over time following E protein transfection (Figure 3E). Importantly, overexpression of STAT2 significantly reversed the E protein -mediated suppression of IFNB-activated ISRE promoter activity (Figure S2A). To further determine whether E protein degradation of STAT2 is dependent on autophagy, we evaluated the effect of E protein on STAT2 protein in cells with ATG5 (autophagy related 5) and ATG7 knockdown, as shown in Figure 3F,G E protein lost its ability to degrade STAT2 in cells depleted of ATG5 and ATG7. Furthermore, the inhibitory effect of E protein on IFN-I-activated ISRE promoter activity was attenuated in ATG5- and ATG7-depleted cells (Figure S2B).

Figure 3.

Autophagy – lysosome pathway associated with SADS-CoV E-induced STAT2 degradation. (A) SADS-CoV envelope is the primary protein responsible for degrading STAT2. HEK293T cells in 24-well plates were transfected with SADS-CoV Nsp2, Nsp5, Nsp7, Nsp12, Nsp13, Nsp16, E, M or an empty vector (400 ng). After 24 h, the cells were harvested, and the expression of the indicated proteins was analyzed by WB. (B and C) SADS-CoV envelope induces STAT2 degradation through the autophagic pathway. HEK293T cells in 12-well plates were transfected with the E protein, followed by treatment with DMSO or 3-MA (5 mm), MG132 (10 μM), Z-VAD (20 μM), BafA1 (100 nM), or NH₄Cl (50 mm) at 12 hpt. At 24 hpt the cells were then collected, and protein expression was evaluated via WB. (D) E protein degrades STAT2 in a dose-dependent manner and promotes LC3-I to LC3-II conversion. HEK293T cells in 12-well plates were transfected with varying amounts of E protein (250 ng, 500 ng, and 1 μg). After 24 hpt, the cells were harvested, and WB was performed to assess the expression of the designated proteins. (E) E protein degrades STAT2 and SQSTM1 over time. HEK293T cells in 12-well plates were transfected with E protein (1 μg), and cells were collected at 6, 12, 24, and 36 hpt for WB. (F and G) E lost the ability to degrade STAT2 in depleted cells of ATG5 and ATG7. HEK-293T cells were transfected with siRNA targeting ATG5 (siATG5, 20 mm), ATG7 (siATG7, 20 mm), or a negative control (siNC, 20 mm). At 24 hpt, E protein (1 μg) was transfected into the cells. After 24 h, the cells were harvested, and WB was performed using the indicated antibodies. (H) E protein induces GFP-LC3 puncta formation. HeLa cells grown on glass-bottom confocal dishes were co-transfected with GFP-LC3 (2 μg) and E protein (2 μg) or an empty vector. Rapamycin (50 nM) was used as a positive control at 12 hpt. At 24 hpt, cells were stained with anti-flag (red) and analyzed by confocal microscopy. Nuclei were stained with DAPI (blue) (scale bar: 10 μm). (I) E protein exhibits complete autophagic flux. HeLa cells on glass-bottom confocal dishes were co-transfected with GFP-LC3 and E-BFP or an empty vector. At 24 hpt, cells were treated with LysoTracker red (50 nm) for 1 h and analyzed by confocal microscopy (scale bar: 10 μm). The data are representative of three independent experiments (B, C, D, E, F, and G). *p < 0.05, **p < 0.01, ***p < 0.001.

Previous studies have shown that SADS-CoV induces autophagy [44,45]. Our results further demonstrated that the E protein degrades STAT2 via the autophagy-lysosome pathway and promotes MAP1LC3/LC3 (microtubule associated protein 1 light chain 3) lipidation. To confirm that E protein induces complete autophagy, we used indirect immunofluorescence to examine the effects of the E protein on LC3 and lysosomes. As shown in Figure 3H,I the E protein strongly increased the number of GFP-LC3-positive autophagosomes, colocalized with LC3-positive puncta, and promoted GFP-LC3 fusion with lysosomes. In addition, BafA1 treatment led to the accumulation of E protein in the cells (Figure S2C), and E protein-transfected cells induced more LC3-II than non-transfected cells (Figure S2D).

In conclusion, these data suggest that the SADS-CoV E protein induces complete autophagy and antagonizes IFN-I signaling by promoting the autolysosome degradation of STAT2.

The E protein primarily associates with the DNA-binding domain (DBD) of STAT2 and mediates its entry into lysosomes

To further investigate the mechanism behind STAT2 degradation by the E protein via autophagy, we conducted co-immunoprecipitation (co-IP) and confocal microscopy were used to determine the association between E protein and STAT2. As shown in Figure 4A,B both exogenous and endogenous STAT2 were found to associate with the E protein. Confocal microscopy revealed colocalization of STAT2 with the E protein (Figure 4C). STAT2 consists of six conserved functional domains: the N-terminal domain, coiled-coil (CC) domain, DBD, linker domain, SRC-homology 2 domain, and transcriptional activation domain [46]. To identify the primary STAT2 domain associating with the E protein, we generated truncated STAT2 mutants (Figure 4D) and analyzed their association with the E protein. WB and co-IP assays demonstrated that STAT2 (1–480 aa) and wild-type STAT2 (STAT2-WT) strongly bound to the E protein; however, STAT2 (1–320 aa) and STAT2 (480–851 aa) weakly bound, suggesting that the STAT2 DBD is the key domain responsible for associating with the E protein (Figure 4E). Given that STAT2 degradation by the E protein is associated with LC3 lipidation, we investigated whether the E protein promotes the colocalization of STAT2 with LC3-II. The results are shown in Figure 4F. Additionally, we observed that the E protein facilitated STAT2 colocalization with LAMP1 (Figure 4G). Together, these findings suggest that the E protein associats with the STAT2 DBD and promoted STAT2 entry into autophagosomes and lysosomes.

Figure 4.

Association between E protein and STAT2, and mediation of STAT2 entry into lysosomes. (A) E protein associates with exogenous STAT2. E protein (5 μg) was co-transfected with STAT2 (5 μg) into HEK293T cells in 100-mm dishes. At 24 hpt, cells were harvested, and co-ip was performed on cell lysates using the indicated antibodies, followed by protein immunoblotting. (B) E protein associates with endogenous STAT2. E protein (10 μg/well) was transfected into HEK293T cells in 100-mm dishes. At 24 hpt, cell lysates were subjected to co-ip with the indicated antibodies, followed by protein immunoblotting. (C) E proteins associate strongly with the STAT2 DBD domain. E protein (5 μg) and STAT2 truncated mutants (1–320 aa, 5 μg; 1–480 aa, 5 μg; and 480–851 aa, 5 μg) were co-transfected into HEK293T cells in 100-mm culture dishes. At 24 hpt, cells were harvested, and cell lysates underwent co-ip with the indicated antibodies, followed by protein immunoblotting. (D) Schematic representation of STAT2 truncated mutants. (E) E protein colocalizes with STAT2. At 24 hpt, E protein (1.5 μg) was transfected alone or co-transfected with STAT2 (1.5 μg) into HeLa cells in glass-bottom dishes. The subcellular localization of the indicated proteins was analyzed by confocal microscopy. Nuclei were stained with DAPI (scale bar: 10 μm). (F) E protein promotes punctate aggregation of STAT2 with LC3. At 24 hpt, GFP-LC3 (1 μg), STAT2 (1 μg), or E protein (1 μg) were co-transfected into HeLa cells in glass-bottom dishes. The subcellular localization of the indicated proteins was analyzed by confocal microscopy (scale bar: 10 μm). (G) E protein promotes STAT2 colocalization with lysosomes. HeLa cells were transfected with GFP-STAT2 along with either E protein or an empty vector in glass-bottom dishes. After 24 hpt, cells were stained with anti-LAMP1 (red), and the subcellular localization of the indicated proteins was analyzed by confocal microscopy (scale bar: 10 μm).

NBR1 and OPTN are autophagy receptors involved in E protein-mediated STAT2 degradation

Selective autophagy regulates the abundance of specific cellular components through SARs, which target protein complexes, aggregates, and organelles for lysosomal degradation [47]. To identify the SARs responsible for E protein-mediated STAT2 degradation, we assessed the interactions among STAT2, SAR, and E protein. Co-IP assays showed that the E protein associated with NBR1, OPTN, CALCOCO2, and TOLLIP (Figure 5A), with the strongest binding to OPTN and TOLLIP. Further analysis of STAT2’s association with SARs in the presence of the E protein revealed that STAT2 associated with NBR1, OPTN, and SQSTM1 (Figure S3A). However, the E protein specifically promoted the association of STAT2 with NBR1 and OPTN, but not SQSTM1 (Figure 5B). This was also confirmed by the results of laser confocal, where E protein promoted the punctate colocalization of NBR1 and OPTN with STAT2 but not with SQSTM1 (Figure S3B–D).

Figure 5.

NBR1 and OPTN are key cargo receptors mediating E protein-induced autophagic degradation of STAT2. (A and B) the indicated proteins were transfected into HEK293T cells in 100-mm dishes. At 24 hpt, cells were harvested, and cell lysates underwent co-ip using the specified antibodies, followed by protein immunoblotting. (A) E protein associates with NBR1, OPTN, CALCOCO2, SQSTM1, and TOLLIP. E protein (5 μg) was transfected with NBR1 (5 μg), CALCOCO2 (5 μg), OPTN (5 μg), SQSTM1 (5 μg), or TOLLIP (5 μg), respectively. (B) E protein promotes STAT2 binding to OPTN and NBR1. STAT2 (5 μg) or E protein (5 μg) were transfected with NBR1 (5 μg), OPTN (5 μg), or SQSTM1 (5 μg). (C) E protein promotes punctate colocalization of NBR1 and OPTN with LC3. GFP-LC3 (1 μg) was co-transfected with NBR1 (1 μg), OPTN (1 μg), or E protein (1 μg) into HeLa cells in glass-bottom dishes. At 24 hpt, cells were stained with anti-flag (red) and anti-ha (blue). The subcellular localization of the indicated proteins was analyzed via confocal microscopy (scale bar: 10 μm). (D) E protein promotes the binding of NBR1 and OPTN to lysosomes. GFP-NBR1 (1 μg) and GFP-OPTN (1 μg) alone or in combination with E protein (1 μg) were transfected into HeLa cells in glass-bottom dishes. At 24 hpt, cells were stained with anti-LAMP1 (red) and anti-ha (blue). The subcellular localization of the indicated proteins was analyzed using confocal microscopy (scale bar: 10 μm). (E) Depletion of NBR1 and OPTN impairs the ability of E protein to degrade STAT2. HEK-293T cells were transfected with siRNA targeting NBR1 (siNBR1, 20 mm), OPTN (siOPTN, 20 mm), or a negative control (siNC, 20 mm). At 24 hpt, E protein (1 μg) was transfected into the cells. After 24 h, the cells were harvested, and WB was performed using the indicated antibodies. (F) Complementation of NBR1 and OPTN restored the ability of E protein to degrade STAT2. HEK-293T cells were transfected with siRNA targeting NBR1 (siNBR1, 20 mm), OPTN (siOPTN, 20 mm). At 24 hpt, E (0.5 μg) and NBR1 (0.5 μg), OPTN (0.5 μg) were co-transfected into the cells. After 24 h, the cells were harvested, and WB was performed using the indicated antibodies. (G) Depletion of NBR1 or OPTN impaired the ability of the E protein to antagonize IFN-I signaling. HEK-293T cells were transfected with siRNA targeting NBR1 (siNBR1, 20 mm), OPTN (siOPTN, 20 mm), or a negative control (siNC, 20 mm). At 24 hpt, E protein (400 μg) or an empty vector (400 ng) was co-transfected with isre-luc (125 ng), hRluc-tk (25 ng). After 24 h, cells were treated with 1,000 U/mL of IFNB for 12 h, followed by a dual-luciferase reporter assay. The data are representative of three independent experiments (E, F, and G). *p < 0.05, **p < 0.01, ***p < 0.001.

Selective autophagy differs from bulk autophagy in its use of SARs, which attach to cargo and use the LC3-interacting region (LIR) motif to bind mammalian Atg8 homologs on the phagophore membrane [29]. The cargo is then recognized, attached to the phagophore, and degraded via autophagy, a process requiring large amounts of Atg8-family proteins [28]. Confocal microscopy demonstrated that the E protein considerably increased the colocalization of NBR1 and OPTN with LC3, forming punctate structures in the cytoplasm (Figure 5C). Subsequently, we used confocal microscopy to examine whether the E protein promoted the incorporation of NBR1 and OPTN into lysosomes. As expected, the E protein facilitated the punctate colocalization of NBR1 and OPTN with lysosomes (Figure 5D). These findings suggest that during E protein-mediated STAT2 degradation, NBR1 and OPTN act as SARs, accompanying the E protein and STAT2 into lysosomes for degradation.

Next, we investigated whether NBR1 or OPTN is necessary for E protein-induced STAT2 degradation. Knockdown of NBR1 and OPTN via siRNA had no direct effect on STAT2 protein expression (Figure S3E). However, depletion of NBR1 and OPTN impaired the E protein’s ability to degrade STAT2, and co-depletion of both NBR1 and OPTN restored STAT2 expression more effectively than depletion of either one alone. Moreover, E protein decreased the expression levels of OPTN and NBR1 in cells (Figure 5E). However, after knockdown of NBR1 and OPTN individually or even together, E protein still significantly degraded STAT2 protein level. Suggesting that there are other SAR-independent mechanisms that contribute the effect. Moreover, exogenous NBR1 and OPTN could not resist the siRNA-mediated silencing (Figure S3F). Complementation of NBR1 or OPTN could reactivate the ability of E protein to degrade STAT2 (Figure 5F). Importantly, in cells depleted of either NBR1 or OPTN or both NBR1 and OPTN, E protein significantly reduced the inhibitory effect on IFN-I-activated ISRE promoter activity (Figure 5G).

In summary, these results indicate that NBR1 and OPTN are important cargo receptors that mediate E protein-induced STAT2 autophagic degradation.

Associations between the CC, four-tryptophan (FW), and C-terminal Ub-associated (UBA) domains of NBR1 and the CC and Ub-Binding domains of OPTN with the E protein

To further understand how NBR1 and OPTN contribute to STAT2 degradation by the E protein, we investigated which domains of NBR1 and OPTN associate with the E protein. NBR1 contains several functional domains, including a Phox-Bem1 domain, zinc finger domain, LIR, UBA domains, FW domains, two CC domains, and an amphipathic helix domain. OPTN also contains multiple domains, including two CC domains, a leucine zipper domain, LIR, a Ub-binding domain, and a zinc finger domain. The truncated NBR1 construct is shown in Figure 6A.

Figure 6.

Functional domains of NBR1 and OPTN involved in E protein association. (A and D) Schematic representation of NBR1 and OPTN truncated mutants. (B and E) NBR1 truncated mutants N1 (1–337 aa), N2 (281–472 aa), and N4 (693–957 aa) and OPTN truncated mutants O3 (210–512 aa) and O4 (454–577 aa) colocalized with E protein. HeLa cells in glass-bottom dishes were transfected with 1 μg of truncated NBR1 or OPTN mutants, either alone or co-transfected with 1 μg of E protein. At 24 hpt, cells were stained with anti-flag (green) and anti-ha (red) antibodies, and nuclei were stained with DAPI. Subcellular localization of the indicated proteins was analyzed by confocal microscopy (scale bar: 10 μm). (C and F) NBR1 truncated mutants N1 (1–337 aa), N2 (281–472 aa), and N4 (693–957 aa) and OPTN truncated mutants O3 (210–512 aa) and O4 (454–577 aa) associate with the E protein. Full-length and truncated mutants of NBR1 (5 μg) or OPTN (5 μg) were co-transfected with E protein (5 μg) into HEK293T cells in 10-mm cell culture dishes. At 24 hpt, cells were harvested, and cell lysates underwent co-ip using the indicated antibodies, followed by protein immunoblotting.

Confocal microscopy revealed that the E protein colocalized with NBR1 domains N1 (1–337 aa), N2 (281–472 aa), and N4 (693–957 aa) in punctate patterns (Figure 6B). This suggests that the E protein primarily associated with the CC, FW, and UBA domains of NBR1. Interestingly, the two truncated NBR1 mutants showed altered subcellular localization. Specifically, N1 (1–337 aa) localized to the cell membrane; however, upon association with the E protein, its localization shifted to a punctate distribution in the cytoplasm and nucleus (Figure 6B). Similarly, N2 (281–472 aa), which typically resides in the nucleus, exited the nucleus and colocalized with the E protein in the cytoplasm (Figure 6B). Co-IP assays confirmed that the E protein associated with N1 (1–337 aa), N2 (281–472 aa), and N4 (693–957 aa) but not with N3 (497–732 aa) (Figure 6C).

We then constructed truncated OPTN mutants (Figure 6D). Both confocal microscopy and co-IP assays showed that the E protein associated with O3 (210–512 aa) and O4 (454–577 aa) (Figure 6E,F). Interestingly, unlike OPTN-WT, all truncated OPTN mutants exhibited varying degrees of nuclear localization (Figure 6E).

In conclusion, this study identified the key domains of NBR1 and OPTN that associated with the E protein. Notably, the E protein was found to associated with the Ub-binding domains of NBR1 and OPTN, suggesting that its recruitment to these proteins may be related to its ubiquitination.

E protein undergoes linear ubiquitination modification via non-lysine residues

In the selective autophagy pathway, NBR1 and OPTN typically recognize ubiquitinated substrates and deliver them to autophagosomes for selective degradation [30]. To explore whether the E protein undergoes ubiquitination, we conducted immunoprecipitation (IP) and confocal microscopy experiments. The results demonstrated that the E protein was polyubiquitinated, but not monoubiquitinated (Figure 7A). Additionally, E protein showed strong colocalization with Ub (Figure 7B).

Figure 7.

Linear ubiquitination of the E protein. (A) Polyubiquitination of the E protein. HEK293T cells were co-transfected with E protein (5 μg) and UB (5 μg) in 10-mm culture dishes. At 24 hpt, cells were harvested, and cell lysates underwent co-ip with the indicated antibodies, followed by WB. (B) HeLa cells were transfected with UB (1.5 μg) either alone or co-transfected with E protein (1.5 μg) in glass-bottom dishes. At 24 hpt, cells were stained with anti-flag (green) and anti-ha (red), and nuclei were stained with DAPI. Subcellular localization of the indicated proteins was analyzed using confocal microscopy (scale bar: 10 μm). (C) E protein associates with ub single-lysine mutants. HEK293T cells were co-transfected with E protein (5 μg) and UB (5 μg) or single lysine site retention UB (K6), UB (K11), UB (K27), UB (K29), UB (K33), UB (K48), or UB (K63) (5 μg each) into 10-mm cell culture dishes. At 24 hpt, cells were harvested, and cell lysates underwent co-ip with the indicated antibodies, followed by protein immunoblotting. (D) Identification of ubiquitination sites on the E protein. HEK293T cells were co-transfected with E protein lysine mutants, where individual or collective lysine residues were mutated to arginine (5 μg each), along with UB (5 μg) in 10-mm cell culture dishes. At 24 hpt, cells were harvested, and co-ip of cell lysates was performed using the indicated antibodies, followed by protein immunoblotting. (E) Linear ubiquitination of the E protein and its mutants. HEK293T cells were co-transfected with E protein (5 μg), its mutants E^K61R (5 μg) and E^{K4R,K34R,K61R} (5 μg), along with UB-K0 (5 μg) in 10-mm cell culture dishes. At 24 hpt, cells were harvested, and co-ip of cell lysates was performed using the specified antibodies, followed by WB.

To further investigate, we co-transfected E protein with either WT Ub or six single-lysine mutant Ub plasmids (Ub [K6], Ub [K11], Ub [K27], Ub [K29], Ub [K33], Ub [K48], and Ub [K63], which contain arginine substitutions at all lysine [K] residues except the corresponding lysine site) (Figure 7C). The E protein associated with all mutant Ub variants. Subsequent experiments revealed that E protein also associated with Ub mutants where single lysines were mutated to arginine (Ub^K6R, Ub^K11R, Ub^K27R, Ub^K29R, Ub^K33R, Ub^K48R, and Ub^K63R) (Figure S4A), suggesting that the E protein was polyubiquitinated via multiple Ub chains or underwent linear ubiquitination through the M1 Ub chain. Importantly, the E protein did not increase the ubiquitination of NBR1, OPTN, or STAT2 (Figure S4B and C).

To determine if the E protein’s selective autophagy is dependent on its ubiquitination, we mutated all lysine residues of the E protein (K4, K34, and K61) to arginine. Interestingly, mutating lysine 61 (K61) considerably reduced E protein’s expression; however, it did not appear to affect its ubiquitination, as the level of E^K61R ubiquitination decreased proportionally to its reduced protein expression (Figure 7D, Figure S5D). Remarkably, the E protein remained ubiquitinated even when all lysine residues were mutated (Figure S5E). This led us to speculate that the E protein undergoes lysine-independent linear ubiquitination. As expected, the E protein exhibited linear ubiquitination regardless of the lysine mutations (Figure 7E).

Together, these findings suggest that E protein undergoes lysine-independent linear ubiquitination.

Ubiquitination of the E protein is required for STAT2 degradation

We next examined whether the E protein mutations affected its ability to degrade STAT2. Interestingly, despite the reduced expression of the mutant E protein, it retained the ability to degrade STAT2 and continued to facilitate STAT2 degradation through the autophagic pathway (Figure 8A). To investigate whether the autophagic degradation of STAT2 by the E protein is related to its ubiquitination, cells were treated with PYR-41, an E1 inhibitor that blocks over 90% of E1 activity [48]. The results showed that PYR-41 considerably reduced E protein ubiquitination (Figure 8B) and reversed the E protein’s ability to degrade STAT2 while restoring the expression of E, NBR1, and OPTN (Figure 8C). Additionally, Ub overexpression markedly decreased the half-life of the E protein in cells treated with cycloheximide (CHX) (Figure 8D). These findings indicate that E protein ubiquitination is essential for STAT2 degradation and further suggest that the K61 residue of the E protein stabilizes its expression independent of ubiquitination.

Figure 8.

E protein degradation of STAT2 is associated with its ubiquitination modification. (A) EK61R degrades STAT2 via autophagy. E protein and its mutants (1 μg) were transfected into HEK293T cells in 12-well plates. Cells were treated with or without 3-MA (5 mm) at 12 hpt and harvested at 24 hpt for WB using the indicated antibodies. (B) PYR-41 inhibits ubiquitination of E protein. HEK293T cells were co-transfected with E protein (5 μg) and its mutants E^K61R (5 μg) and E^{K4R,K34R,K61R} (5 μg), along with UB (5 μg), in a 10-mm cell culture dish. At 6 hpt, cells were treated with PYR-41 (50 μm). At 24 hpt, cells were harvested. Cell lysates underwent co-ip using the indicated antibodies and were analyzed via protein immunoblotting. (C) PYR-41 impairs the ability of E protein to degrade STAT2. E protein (1 μg) and its mutants E^K61R (1 μg) and E^{K4R,K34R,K61R} (1 μg) were transfected into HEK293T cells in 12-well plates. Cells were treated with or without PYR-41 (50 μm) at 6 hpt and harvested at 24 hpt. Protein immunoblotting was performed using the indicated antibodies. (D) UB reduced the half-life of E protein. E protein (1 μg) and UB (1 μg) or an empty vector were co-transfected into HEK293T cells in 12-well plates. At 24 hpt, cells were treated with CHX (100 μg/ml) for varying durations. The data are representative of three independent experiments (A and C). ***p < 0.001.

Discussion

Coronaviruses have evolved multiple immune escape pathways to suppress innate immunity against IFN-I, thereby promoting their replication [34,41,49]. As a novel type of coronavirus, the immune escape and pathogenic mechanism of SADS-CoV are not yet fully understood. Our data reveal that SADS-CoV antagonizes IFN-I signaling by degrading STAT2, a critical protein in the IFN-I antiviral response [50]. Various viruses have been shown to target STAT2 to disrupt the IFN-I-mediated innate antiviral response and promote viral replication; however, the mechanisms used by different viruses vary. For example, the measles virus V protein inhibits STAT2 association with IRF9 by binding with the C-terminal domain of STAT2 [51]. Additionally, porcine delta coronavirus nsp5 cleaves STAT2 at glutamine 685 (Q685) and Q758, impairing its function [52]. Zika virus NS5 also mediates STAT2 degradation via a proteasome-dependent pathway [53]. In this study, we demonstrated a novel mechanism for STAT2 degradation: the SADS-CoV Envelope induces selective autophagy of STAT2 via the dual autophagy receptors NBR1 and OPTN (Figure 9).

Figure 9.

Following ubiquitination, the SADS-CoV E protein is recruited by NBR1 and OPTN and degraded together with STAT2 via the autolysosome pathway, antagonizing IFN-I signaling.

We conducted an indiscriminate screening of SADS-CoV proteins that inhibit IFN-I signaling and identified multiple immunosuppressive E proteins, including Nsp1 and Nsp5, which have already been characterized for their antagonism of IFN-I signaling [54,55]. However, the specific mechanisms behind the immunosuppressive effects of these proteins remain unclear. Our findings indicate that STAT2 degradation by the SADS-CoV E protein via the autophagic pathway strongly inhibits IFN-I signaling. The E protein induces the conversion of LC3-I to LC3-II, activates autophagic flow, and ultimately recruits STAT2 to lysosomes for degradation by associating with STAT2. Most studies of the E protein in coronaviruses focus on its role in viral assembly [56]. Here, we highlight the autophagy-inducing and immunosuppressive functions of the alphacoronavirus E protein, offering a new perspective on the functional diversity of alphacoronavirus envelope proteins.

Selective autophagy plays a dual role in antiviral innate immune responses [22]. Xenophagy, a cellular defense mechanism, specifically recognizes intracellular pathogens and targets them for autophagic degradation [57]. In the later stages of infection, autophagy facilitates antigen processing, triggering an adaptive immune response [58]. However, viruses can exploit a SAR as a means of immune evasion, using it to degrade host proteins and inhibit innate immunity. For instance, avibirnavirus VP3 induces TRAF6 autophagy in an SQSTM1-dependent manner to evade host innate immunity [59]. Similarly, the α-herpesvirus tegument UL21 ubiquitinates CGAS, recruiting TOLLIP to mediate CGAS degradation via selective autophagy, thereby suppressing innate immunity [60]. β-coronavirus SADS-CoV-2 NSP13 inhibits IFN-I production by selectively degrading TBK1 through an SQSTM1-dependent autophagy pathway [61]. Although STAT2 is crucial in IFN-I signaling and has been implicated in cancer development [62], its association with SARs has not been reported. In this study, we identified several STAT2-associating SARs, including SQSTM1, NBR1, and OPTN, which may play critical roles in regulating STAT2 stability. Our results suggest that the SADS-CoV envelope recruits STAT2 via NBR1 and OPTN, promoting its autophagic degradation and inhibiting IFN-I-mediated innate immunity. This highlights the involvement of NBR1 and OPTN in regulating STAT2 expression.

Ubiquitination is essential for selective autophagy [47], where protein marked with Ub signals are recognized by Ub-binding receptors and delivered to proteasomes or autophagosomes for degradation [30,63]. We found that the E protein binds strongly to Ub, and its ubiquitination occurs through non-lysine residues. Lysine is typically considered the primary site for substrate ubiquitination [64]. However, recent studies have shown that other residues, including cysteine, serine, threonine, and N-terminal free amino groups, can serve as ubiquitination sites [65]. For example, the Drosophila protein Bruce targets the apoptosis inhibitor rpr (reaper) for ubiquitination through a non-lysine pathway [66]. Our results suggest that E protein ubiquitination via a non-lysine pathway is necessary for STAT2 degradation, and the K61 residue stabilizes E protein expression independently of ubiquitination. We speculate that this stabilization may involve acetylation, a common cellular mechanism that increases protein stability by competing with ubiquitination [67].

Although our data did not identify the specific Ub chain responsible for STAT2 polyubiquitination, and further studies are needed to clarify the role of the E protein in this process, our findings underscore the critical role of ubiquitination in STAT2 degradation mediated by the E protein. These results may provide insights for future strategies aimed at preventing and controlling SADS-CoV infections.

In conclusion, our study elucidates the mechanism by which the SADS-CoV E protein antagonizes IFN-I signaling by degrading STAT2. Following ubiquitination, the E protein associates with STAT2 and is recruited by NBR1 and OPTN for degradation via the autophagy – lysosomal pathway. Additionally, the K61 residue is essential for stabilizing the E protein.

Materials and methods

Cells and viruses

The HEK293T, Vero, HeLa, and IPI-2I cell lines were obtained from the Department of Infectious Diseases at the College of Veterinary Medicine, South China Agricultural University. HEK293T, Vero, HeLa, and IPI-2I cells were cultured in Dulbecco’s Modified Eagle’s medium (DMEM; Gibco 11,965,092) supplemented with 10% fetal bovine serum and 1% antibodies (penicillin, streptomycin, and amphotericin B). The SADS-CoV (PEAV) strain GDS04 was generously provided by Yongchang Cao from Sun Yat-sen University and was propagated in Vero cells in DMEM containing 0.3% trypsin (Gibco 15,050,065).

Antibodies and reagents

Our laboratory provided a murine polyclonal antibody specific to the SADS-CoV Nuclear protein. GFP (66002–1-IG) was obtained from Proteintech. Murine monoclonal antibodies against HA (M20003) and FLAG (M20008) were purchased from Abmart, and GAPDH (HC301–01) was sourced from Full Gold. Rabbit monoclonal antibodies against HA (3724), p-STAT1 (9167), and p-STAT2 (88410) were acquired from Cell Signaling Technology, and ATG7 (ET1610–53) was obtained from Hua Bioscience. LC3B2 (EPR18709) was purchased from Abcam. Rabbit polyclonal antibodies against STAT2 (16674–1-AP), STAT1 (10144–2-AP), NBR1 (16004–1-AP), HA (51064–2-AP), Flag (20543–1-AP), and OPTN (16004–1-AP) were sourced from Proteintech.

The secondary antibodies used included IRDye® 800CW Goat (polyclonal) Anti-Rabbit/Mouse IgG (H+L) from LI-COR (D30322–1/25), Goat anti-Rabbit IgG (H+L) Cross-Adsorbed Secondary Antibody, Alexa Fluor 555 (A-31570) from Thermo Fisher Scientific, and Alexa Fluor 488-labeled Goat Anti-Mouse IgG (H+L) (A0428), Alexa Fluor 647-labeled Goat anti-Rabbit IgG (H+L) (A0468), and Alexa Fluor 555-labeled Donkey Anti-Mouse IgG (H+L) (A0460) from Beyotime. Murine (A7031) and rabbit (A7016) IgGs were obtained from Beyotime Biotechnology.

DMSO (HY-Y0320), MG132 (HY-13259), 3-MA (HY-19312), BafA1 (HY-100558), NH₄Cl (HY-Y1269), CHX (HY-12320), and PYR-41 (HY-13296) were purchased from MedChemExpress. Z-VAD-FMK (C1202), a dual-luciferase reporter gene detection kit (RG028), 4’,6-diamidino-2-phenylindole (DAPI; C1002), PMSF (ST507), and cell lysis buffer for WB and IP (P0013) were obtained from Beyotime. Protein A/G Agarose gels (sc-2003) were sourced from Santa Cruz Biotechnology. Recombinant human IFNB (300-02BC) was purchased from PeproTech. Lipofectamine 3000 transfection reagent (L3000150), Lipofectamine RNAiMAX transfection reagent (13778150), and TRIzol™ (15596026CN) were acquired from Thermo Fisher Scientific, and PEI transfection reagent was obtained from FUSHENBIO. HiScript II Reverse Transcriptase (RL201) and AceQ qPCR SYBR Green Master Mix (Q121) were sourced from Vazyme.

Plasmid

The pGL3-ISRE-Luc and pRL-TK luciferase vectors were generously donated by Professor Changxu Song from the South China Agricultural University. FLAG-tagged constructs of NBR1, OPTN, CALCOCO2, SQSTM1, TOLLIP, and truncated mutants of NBR1, OPTN, and STAT2 were created through seamless cloning into the pCDNA3.1-FLAG-N vector (MIAOLINGBIO, P0794). The FLAG-tagged E protein was cloned into the pCAGGS-FLAG-N vector. FLAG-tagged mutants E^K4R, E^K34R, E^K61R, and E^K4,34,61R were synthesized using the pCAGGS-FLAG-N vector obtained from Sangon Bioengineering Co., Ltd. GFP-tagged vectors for STAT2, NBR1, OPTN, and SQSTM1 were constructed in the pEGFP-C1 vector using seamless cloning. The blue fluorescent E protein was cloned into the pmTagBFP2 vector (MIAOLINGBIO, P24146). Ha-tagged SDS-CoV protein, STAT2-FLAG, GFP-LC3, HA-tagged Ub single-site retained, and single-site mutant plasmids are stored in our laboratory. pRK5-Ub-HA-K0 was purchased from MIAOLINGBIO (P51112).

Generation of KD cells

The ATG5, ATG7, NBR1, and OPTN siRNAs were synthesized and designed by Ribo. All primer- and siRNA-targeted gene sequences are listed in Tables S1 and S2, respectively. KD cells were obtained using reverse transfection. Dilute 20 mm siRNA in 200 µl Opti Medium without serum (Gibco 31,985,070) in the well of the tissue culture plate, then add 3 µl Lipofectamine™ RNAiMAX to each well. Mix gently and incubate for 15 min at room temperature. Dilute cells in complete growth medium without antibiotics. (cell density should be 30–50% confluent 24 h after plating). To each well, add 1 ml of the diluted cells. Incubate the cells 24–72 h at 37°C in a CO₂ incubator until ready to assay for gene knockdown.

Luciferase assay

Cells were transiently transfected with the pGL3-ISRE-Luc and pRL-TK luciferase vectors, along with the indicated plasmids. After 24 h of transient transfection, the cells were stimulated with or without IFNB (1,000 U/mL) for 12 h, and luciferase activity was measured using a dual-luciferase assay system (Promega, E5311) according to the manufacturer’s instructions.

Virus infection and titration

All viral stock solutions were produced in Vero cells. The maintenance medium for virus propagation in Vero, HeLa, and IPI-2I cells consisted of serum-free DMEM supplemented with 0.3%, 0.05%, and 0.1% trypsin, respectively. Cells were seeded into 96-well plates, and the virus solution was diluted from 10⁻¹ to 10⁻⁸ using DMEM as the maintenance medium for the different cell types before being added to the cells in the 96-well plates. The cells were then cultured in a viral maintenance medium for 48 h, and cytopathic effects were recorded. Viral titers were calculated using the Reed – Muench method.

WB and IP

Cells were lysed in cell lysis buffer supplemented with 1% PMSF for WB and IP. Protein sample buffer (5×) was added, and the samples were boiled. For WB analysis, the protein samples were electrophoresed on 10% SDS-PAGE gels and transferred to polyvinylidene difluoride membranes (Beyotime, P0965). Following the transfer, the membranes were blocked with 5% skim milk powder in Tris-buffered saline (Beyotime, ST667) with Tween 20 (Beyotime, ST825; TBS-T). The primary antibody was diluted in TBS-T containing 2% bovine serum albumin (BSA; Beyotime, ST023), with dilution ratios specific to each antibody. The secondary antibody used was IRDye® 800CW Goat (polyclonal) Anti-Rabbit/Mouse IgG (H+L), diluted at 1:15,000.

For co-IP, the cells were seeded in 10-mm cell culture dishes and transfected with the indicated plasmids. After a specific treatment time (hours post-transfection [hpt]), the cells were lysed in cell lysis buffer for WB and IP (Beyotime, P0013). Protein A/G conjugated with IP antibodies, or IgG was incubated with the protein lysates for 2 to 4 h at 4°C. Following incubation, protein A/G was washed three times with phosphate-buffered saline (biosharp, BL601A) with Tween 20, and SDS protein loading buffer was added. The samples were boiled for 10 min for subsequent SDS-PAGE and WB analysis.

Confocal microscopy

HeLa cells were seeded into confocal dishes and transfected with the specified plasmids. After 24 h, the cells were stimulated with or without IFNB (1,000 U/mL) for 2 h. The cells were then fixed with 4% paraformaldehyde for 10 minutes and permeabilized with phosphate-buffered saline containing 0.3% Triton X-100 (Beyotime, ST1723). Following three washes with PBS, the cells were incubated with 2% BSA at 37°C for 1 h. Anti-STAT2 and STAT1 antibodies were applied at a dilution of 1:200; in contrast, anti-HA and anti-FLAG antibodies were used at a dilution of 1:1,000. The samples were incubated with primary antibodies overnight at 4°C and with secondary antibodies for 1 h at room temperature. Subsequently, DAPI was used to stain cells for 15 minutes. Fluorescence images were acquired using a confocal laser scanning microscope (Leica TCS SP8).

RNA extraction and real-time qPCR

Total RNA was extracted from the samples using TRIzol reagent, and RNA was reverse-transcribed using HiScript II Reverse Transcriptase. The resulting cDNA was utilized for qPCR with the AceQ qPCR SYBR Green Master Mix using the Bio-Rad, CFX96. The abundance of individual mRNA transcripts in each sample was measured in triplicates, and gene expression levels were normalized to GAPDH or ACTB/β-actin. The qPCT primers used are listed in Table S3.

Statistical analyses

All data represent at least two independent experiments. Data were analyzed using a two-tailed Student’s t-test via GraphPad Prism v.8 (San Diego, California, USA). P-values of < 0.05 were considered statistically significant.

Funding Statement

This research was funded by the National Key Research and Development Program of China [2022YFD1800804], the 16th Batch of Special Grants (Center) from the China Postdoctoral Science Foundation [2023T160232], the 74th Batch of General support from the China Postdoctoral Foundation [2023M741220)], and the China Agriculture Research System of MOF and MARA [CARS-35].

Disclosure statement

No potential conflict of interest was reported by the author(s).

Data availability statement

All data included in this study are available from the corresponding author upon request.

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/15548627.2025.2479671