Targeting liquid–liquid phase separation of SARS-CoV-2 nucleocapsid protein promotes innate antiviral immunity by elevating MAVS activity

Wang, Shuai; Dai, Tong; Qin, Ziran; Pan, Ting; Chu, Feng; Lou, Lingfeng; Zhang, Long; Yang, Bing; Huang, Huizhe; Lu, Huasong; Zhou, Fangfang

doi:10.1038/s41556-021-00710-0

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Published: 08 July 2021

Targeting liquid–liquid phase separation of SARS-CoV-2 nucleocapsid protein promotes innate antiviral immunity by elevating MAVS activity

Shuai Wang¹^na1,
Tong Dai¹^na1,
Ziran Qin¹^na1,
Ting Pan²^na1,
Feng Chu¹,
Lingfeng Lou¹,
Long Zhang ORCID: orcid.org/0000-0001-8139-0474³,
Bing Yang^3,4,
Huizhe Huang⁵,
Huasong Lu³ &
…
Fangfang Zhou ORCID: orcid.org/0000-0003-1870-7584¹

Nature Cell Biology volume 23, pages 718–732 (2021)Cite this article

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Abstract

Patients with Coronavirus disease 2019 exhibit low expression of interferon-stimulated genes, contributing to a limited antiviral response. Uncovering the underlying mechanism of innate immune suppression and rescuing the innate antiviral response remain urgent issues in the current pandemic. Here we identified that the dimerization domain of the SARS-CoV-2 nucleocapsid protein (SARS2-NP) is required for SARS2-NP to undergo liquid–liquid phase separation with RNA, which inhibits Lys63-linked poly-ubiquitination and aggregation of MAVS and thereby suppresses the innate antiviral immune response. Mice infected with an RNA virus carrying SARS2-NP exhibited reduced innate immunity, an increased viral load and high morbidity. Notably, we identified SARS2-NP acetylation at Lys375 by host acetyltransferase and reported frequently occurring acetylation-mimicking mutations of Lys375, all of which impaired SARS2-NP liquid–liquid phase separation with RNA. Importantly, a peptide targeting the dimerization domain was screened out to disrupt the SARS2-NP liquid–liquid phase separation and demonstrated to inhibit SARS-CoV-2 replication and rescue innate antiviral immunity both in vitro and in vivo.

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Main

Coronavirus disease 2019 (COVID-19), caused by the novel severe acute respiratory syndrome (SARS) coronavirus 2 (SARS-CoV-2), emerged and spread rapidly worldwide, posing a major threat to global public health and safety¹. The SARS-CoV-2 genomic RNA is harboured within a viral particle consisting of four structural proteins—the crown-like spike (S) glycoprotein, the membrane (M) protein, the ion channel envelope (E) protein and the nucleocapsid protein (NP; SARS2-NP). The S protein binds to the human ACE2 receptor and is responsible for viral entry into host cells. The M protein facilitates viral assembly in the endoplasmic reticulum and the NP assembles with viral RNA to form a helical ribonucleoprotein complex called the nucleocapsid^2,3.

Although the precise interaction between the innate immune system and SARS-CoV-2 is yet to be fully elucidated, it has been suggested that the innate immune response plays a crucial role in determining the severity of COVID-19 (refs. ^4,5,6). During coronavirus infection, the viral components and replication intermediates serve as pathogen-associated molecular patterns recognized by host germline-encoded pattern recognition receptors, including retinoic acid-inducible gene I (RIG-I)-like receptor and members of the Toll-like receptor family^7,8,9,10. The activation of innate immune receptors promotes the production of type I interferons (IFN-I; including IFN-β and IFN-α), which mediates the antiviral response through the upregulation of IFN-stimulated genes (ISGs)^{11,12,13,14,15}.

The NP of coronaviruses associates with the large positive-strand RNA viral genome and the carboxy (C)-terminal endodomain of the M protein, which is critical for coronavirus replication¹⁶. SARS2-NP possesses self-assembly properties, with its dimerization domain (DD) being responsible for homodimer formation as well as C-terminal-mediated tetramer formation¹⁷. However, whether SARS2-NP possesses a pathogenic mechanism other than the packaging of viral RNA and whether the self-assembly properties of SARS2-NP could be targeted for COVID-19 prevention and therapy remain unknown. Here we reveal the mechanisms of NP-mediated innate immune evasion, NP modification and mutation, which together permit the optimization of therapeutic strategies and antiviral drug against COVID-19.

Results

The SARS2-NP DD is required for LLPS and viral RNA association

SARS2-NP contains three predicted intrinsically disordered regions (IDRs), an amino (N)-terminal RNA-binding domain and a C-terminal DD^17,18,19 (Extended Data Fig. 1a). Given that proteins with IDRs tend to undergo liquid–liquid phase separation (LLPS)^20,21, we tested whether NP has such a property. Purified green fluorescent protein (GFP)-conjugated NP (Extended Data Fig. 1b) formed microsized droplets in solutions (Fig. 1a) and more numerous and larger droplets were observed at low salt concentrations, weak acidic conditions (pH 4.5–7.5) and high temperatures (Fig. 1a–c). The droplets dissolved when the NaCl concentration was increased, suggesting that electrostatic interactions are needed (Fig. 1a). SARS2-NP droplet formation was inhibited by 5% 1,6-hexanediol (Hex)—a compound that disrupts weak hydrophobic interactions²² (Extended Data Fig. 1c)—and completely disrupted by heat or proteinase K (Extended Data Fig. 1d,e). The droplet size of NPs labelled with AF488 increased with time (Fig. 1d), indicating that the puncta were not artificially formed by the GFP tag. A fluorescence recovery after photobleaching (FRAP) assay revealed rapid fluorescence recovery after bleaching (Fig. 1e), indicating efficient diffusion within the SARS2-NP droplets.

**Fig. 1: The DD is required for LLPS of SARS2-NP and RNA.**

To evaluate whether SARS2-NP undergoes LLPS in cells, we transfected HeLa cells with GFP–SARS2-NP and observed that GFP–SARS2-NP formed discrete puncta in the cytoplasm (Fig. 1f,g). In line with the in vitro results, the GFP–SARS2-NP-enriched droplets in cells readily underwent fusion (Fig. 1f) and recovered quickly following FRAP (Fig. 1g). Because RNA is required for viral ribonucleoprotein assembly, we evaluated whether RNA is involved in GFP–SARS2-NP droplet formation by co-transfecting Cy5-labelled SARS-CoV-2 open reading frame (ORF) RNA with GFP–SARS2-NP. The GFP–SARS2-NP-enriched puncta were found to co-localize with Cy5–RNA fluorescence (Fig. 1h), and fusion of GFP–SARS2-NP and Cy3–RNA droplets was observed (Fig. 1i).

To determine which domain is required for NP LLPS, a series of truncated SARS2-NPs labelled with GFP were bacterially expressed and purified (Extended Data Fig. 1b). Although deletion of either the IDR or RNA-binding domain barely affected SARS2-NP-droplet formation, the DD-depleted mutant (d_DD) exhibited no or very low LLPS (Fig. 1j). In HeLa cells, GFP-labelled SARS2-NP d_DD was evenly distributed in the cytoplasm, similar to the localization of wild-type (WT) GFP–SARS2-NP after Hex treatment (Fig. 1k). Co-immunoprecipitation revealed that DD deletion prevented SARS2-NP self-association (Fig. 1l). Semi-denaturing detergent agarose gel electrophoresis (SDD–AGE) analysis of the cytosolic fraction revealed a smear of sodium dodecyl sulfate (SDS)-resistant high-molecular-weight SARS2-NP, but not d_DD (Extended Data Fig. 1f,g), in the absence but not in the presence of β-mercaptoethanol (Extended Data Fig. 1f). In sucrose gradients, SARS2-NP WT, but not d_DD, sedimented as a much larger assembly than predicted by its molecular weight (Extended Data Fig. 1h). A proximity ligation assay (PLA)^23,24 showed that the Flag– and Myc–SARS2-NP together yielded PLA signals in the cytoplasm, which were detected for the DD-deficient mutant (Fig. 1m). Thus, the DD is required for SARS2-NP condensation in vitro and in vivo.

The NP is highly basic (pI of about 10) and contains multiple RNA-binding sites^25,26. Its DD can form a tightly linked dimer with a large RNA-binding groove^17,25,27. To evaluate the binding affinity between SARS2-NP and viral RNA, we employed surface plasmon resonance (SPR), where the RNA–SARS2-NP binding affinity was analysed by flowing recombinant SARS2-NP over the RNA-captured sensor chip. As expected, the SARS2-NP exhibited a high binding capacity for viral RNA, which was substantially reduced in the DD-depleted mutant (Fig. 1n), indicating that the DD is required for SARS2-NP to bind viral RNA.

NP-induced innate immune evasion relies on the DD

To explore the function of NP in the innate antiviral response, constructs encoding SARS-NP or SARS2-NP along with IFN-β and PRD I–III (contains the IRF3-binding site of the Ifnb1 promoter) reporter plasmids were transfected into HEK293T cells, followed by Sendai virus (SeV) stimulation. Exogenous expression of both NPs resulted in significant downregulation of IFN-β and PRD I–III promoter activity (Fig. 2a). Overexpression of SARS-NP and SARS2-NP also resulted in reduced expression of IFNB1 messenger RNA and downstream ISGs following induction by SeV (Fig. 2b). Consistent with these results, transfection with both NPs inhibited the 5′-triphosphorylated RNA (5′-ppp RNA)- or poly(I:C)-induced expression of IFNB1 mRNA (Fig. 2c). Furthermore, a slightly stronger inhibitory effect on IFN-β signalling was exhibited by SARS2-NP than by SARS-NP (Fig. 2a–c).

**Fig. 2: SARS2-NP-mediated dampening of IFN-β signalling relies on the DD.**

Deletion of the SARS2-NP DD almost completely abolished the inhibitory effect of SARS2-NP on the expression of IFNB1, CCL5 and CXCL10 mRNA as well as on IFN-β promoter activity (Fig. 2d and Extended Data Fig. 2a,b). The same results were obtained using human lung carcinoma A549 cells (Extended Data Fig. 2c). In line with these findings, overexpression of SARS2-NP d_DD had no obvious effect on IFNB1 mRNA expression induced by poly(I:C) and 5′-ppp RNA (Fig. 2e and Extended Data Fig. 2d). Enzyme-linked immunosorbent assay (ELISA) analysis indicated that the ectopic expression of SARS2-NP WT in RAW264.7 and A549 cells significantly repressed SeV-induced IFN-β secretion, whereas the ectopic expression of SARS2-NP d_DD had no effect (Fig. 2f). After challenge with vesicular stomatitis virus (VSV), an RNA virus, Ifnb1 expression was inhibited and the VSV-specific mRNA, protein and titres (determined as plaque-forming units, p.f.u.) were increased by WT NP but not by NP d_DD (Fig. 2g,h and Extended Data Fig. 2e,f). Accordingly, the replication of VSV expressing GFP (GFP–VSV) was increased by the ectopic expression of WT NP, whereas NP d_DD had no effect on viral replication (Fig. 2i). In addition, ectopic expression of WT NP, but not NP d_DD, maximized the cytopathic effect induced by SeV infection (Fig. 2j and Extended Data Fig. 2g). Thus, SARS2-NP negatively regulates IFN-β signalling and DD is critical for these inhibitory effects.

The DD mediates NP-induced innate immune evasion in vivo

To further elucidate the role of SARS2-NP in the regulation of innate antiviral immunity in mice, we genetically engineered recombinant VSVs by employing the virulence-attenuated VSV backbone harbouring an M51R substitution in the matrix (M) protein^28,29. We inserted the coding sequence of control GFP, SARS2-NP WT or SARS2-NP d_DD into the VSV backbone between the VSV glycoprotein (G protein) and the polymerase protein (L protein)³⁰, hereafter denoted as VSV, NP–VSV and NP d_DD–VSV, respectively. This enabled us to perform viral infections in mice systematically and observe the effects of NPs on virulence and antiviral immunity (Fig. 3a). All three viruses displayed similar replication kinetics in plaque assays in Vero E6 cells. However, the NP–VSV replicated more efficiently than the control VSV and NP d_DD–VSV in HEK293T cells (Fig. 3b). Because Vero E6 cells lack an IFN response, the observation described above implied that NP-mediated innate immunosuppression probably contributes to viral proliferation.

**Fig. 3: The DD is essential for the SARS2-NP-mediated suppression of innate antiviral responses in vivo.**

We first infected primary peritoneal macrophages with recombinant VSVs at a multiplicity of infection (m.o.i.) of 0.1. After stimulation, Ifnb1 expression was inhibited and the VSV-specific mRNA and VSV titres were increased in cells infected with NP–VSV but not in those infected with NP d_DD–VSV (Fig. 3c). To investigate the function of NP in vivo, mice were challenged with the recombinant VSVs. The levels of Ifnb1 mRNA were significantly lower in the spleen, liver and lungs of the NP–VSV-infected mice than in those of the mice infected with control VSV or NP d_DD–VSV (Fig. 3d). The serum concentration of IFN-β was also lower in NP–VSV-infected mice than control VSV- and NP d_DD–VSV-infected mice (Fig. 3e). In contrast, the VSV-specific mRNA and protein expression as well as the VSV titres were all higher in the lungs, liver and spleen of the NP–VSV-infected mice than the control VSV- and NP d_DD–VSV-infected mice (Fig. 3f–h). Haematoxylin and eosin (H&E) staining of the lungs after infection revealed higher immune-cell infiltration as well as severe tissue injury in the NP–VSV-infected mice than in their control VSV- and NP d_DD–VSV-infected counterparts (Fig. 3i). Moreover, the mice that were infected with recombinant VSV harbouring SARS2-NP WT exhibited substantially higher sensitivity to the virus relative to the mice that were challenged with control VSV. However, such a difference was not observed in the mice that were infected with NP d_DD–VSV (Fig. 3j). These findings indicate that the DD mediates the NP-induced suppression of innate antiviral immunity and contributes to viral replication in vivo.

SARS2-NP negatively regulates MAVS–IRF3 activation and is enriched in MAVS droplets

Next, we investigated whether NP inhibited IFN-β signalling by evaluating virus-induced phosphorylation of TBK1 and IRF3 as well as IRF3 dimerization and its nuclear accumulation^31,32. Surprisingly, all of these molecular events were inhibited by ectopic expression of SARS2-NP WT but not SARS2-NP d_DD (Fig. 4a, left). Similar results were found when we analysed SeV-induced signalling events in A549 cells (Fig. 4a, right). Moreover, cells expressing SARS2-NP WT, but not d_DD, exhibited severely impaired IRF3 nuclear translocation after stimulation with SeV (Fig. 4b,c). Thus, SARS2-NP might target critical IFN-β signalling components upstream of TBK1–IRF3.

**Fig. 4: SARS2-NP negatively regulates TBK1–IRF3 activation and is enriched in MAVS droplets.**

To determine which signalling protein could be targeted by SARS2-NP, expression plasmids encoding a series of essential antiviral signalling proteins were co-expressed with SARS2-NP. We found that ectopically expressed SARS2-NP specifically interacted with MAVS (Fig. 4d). Endogenous MAVS immunoprecipitated together with SARS2-NP in cells infected with SARS-CoV-2 (Fig. 4e). MAVS is comprised of a CARD (amino acids 1–90), a Pro-rich (amino acids 91–172) and a transmembrane (amino acids 513–540) domain (Extended Data Fig. 3a). To determine which MAVS domain mediates its association with SARS2-NP, a series of truncated MAVS variants were constructed for interaction assays with SARS2-NP. Deletion of the critical N terminus of MAVS—removal of either the CARD domain or the Pro-rich region—impeded the MAVS–SARS2-NP interaction (Fig. 4f). Following infection with RNA virus, MAVS serves as a central antiviral signalling hub via Lys63-linked poly-ubiquitination by the E3 ligase TRIM31 and aggregates to stimulate downstream signalling cascades. Interestingly, the CARD domain of RIG-I interacts with the CARD domains of MAVS³³, whereas TRIM31 interacts with MAVS through the Pro-rich domain of MAVS³⁴. We therefore tested whether SARS2-NP regulates the interaction of MAVS with RIG-I and TRIM31. We indeed observed endogenous TRIM31 and RIG-I associated with endogenous MAVS after infection with SeV in both HEK293T and A549 cells. Furthermore, the infection-induced associations were impeded by SARS2-NP WT, whereas SARS2-NP d_DD had no effect (Fig. 4g). Thus, SARS2-NP suppresses innate immune signalling at the level of MAVS.

MAVS was reported to form clusters in vivo and prion-like aggregates in vitro³³; this also suggests that MAVS might undergo LLPS and thus benefit antiviral signalling transduction^33,35. In cells, MAVS clusters were found to be disrupted into segments by SARS2-NP WT, but not by SARS2-NP d_DD, and those segments could still partially co-condensate with SARS2-NP WT (Extended Data Fig. 3b). Purified GFP–MAVS formed microsized droplets spontaneously in vitro, and the droplets were larger at higher protein concentrations (Fig. 4h). We next evaluated the potential relationship between SARS2-NP LLPS and its inhibitory regulation of MAVS. Notably, when the MAVS protein was present in excess (above the diagonal), GFP–MAVS was enriched in the droplets (Fig. 4h). In contrast, when SARS2-NP was in excess (below the diagonal), the MAVS–MAVS enrichment pattern was displaced by MAVS–SARS2-NP and SARS2-NP–SARS2-NP enrichment patterns, in which mCherry–SARS2-NP exhibited an opposite pattern of enrichment (Fig. 4h). In contrast, SARS2-NP d_DD barely affected the MAVS droplet formation (Extended Data Fig. 3c). Thus, SARS2-NP negatively regulates MAVS–IRF3 signalling cascades by binding MAVS and interfering with its phase separation.

SARS2-NP inhibits Lys63-linked poly-ubiquitination and aggregation of MAVS

Due to the inhibitory effect of SARS2-NP on the association between MAVS and TRIM31, we observed that ectopic SARS2-NP expression repressed MAVS poly-ubiquitination with WT and Lys63-linked ubiquitin (Fig. 5a). Using tandem ubiquitin-binding entities (TUBE), we pulled-down endogenous K63-ubiquitin-conjugated MAVS from SeV-stimulated HEK293T and A549 cells, which was severely impeded by SARS2-NP WT but not by SARS2-NP d_DD (Fig. 5b,c).

**Fig. 5: SARS2-NP inhibits the Lys63-linked poly-ubiquitination and aggregation of MAVS.**

MAVS aggregation relies on Lys63-linked poly-ubiquitination induced by virus infection that also recruits TBK1 and IKKε for phosphorylation and activation^34,36. Consistent with impaired ubiquitination of MAVS, we found that the aggregation of MAVS—either transfected or at endogenous levels induced by SeV—can be suppressed by the ectopic expression of SARS2-NP WT but not SARS2-NP d_DD (Fig. 5d–f). In sucrose-gradient ultracentrifugation, SeV infection strongly shifted MAVS from the low-molecular-weight fractions to the high-molecular-weight fractions, and this shift was severely impaired in cells with ectopic SARS2-NP WT expression but not in those with ectopic expression of SARS2-NP d_DD (Fig. 5g). Consistent with these results, SARS2-NP WT, but not SARS2-NP d_DD, strongly interfered with the association between co-expressed MAVS and TBK1/IKKε (Fig. 5h). At endogenous levels, SeV-induced MAVS–TBK1–IKKε complex formation in both HEK293T and A549 cells was disrupted by SARS2-NP WT but not by SARS2-NP d_DD (Fig. 5i). Together, these results indicate that SARS2-NP strongly inhibits MAVS activation in cells.

SARS2-NP acetylation at Lys375 abrogates its LLPS and the NP-mediated suppression of MAVS signalling

These findings raised the question of how hosts manage to mount a defence against viral infections in the presence of high amounts of NP. We therefore investigated whether NP itself is regulated in the host. In fact, we noticed that a few mice infected with NP–VSV seemed to have mild pathological symptoms compared with others and exhibited only slight weight loss. Thus, we extracted NP from the tissue of these mice for mass spectrometry analysis. We observed that SARS2-NP can be highly acetylated in vivo at Lys375 (Fig. 6a), a motif that is adjacent to the DD and conserved between SARS-CoV-NP and Middle East respiratory syndrome coronavirus (MERS-CoV)-NP (Fig. 6b). Using an acetyl-Lys antibody, we confirmed that abundant NP can be acetylated in host cells, and Lys375 was the most critical residue for NP acetylation (Fig. 6c). Co-expression of CREB-binding protein (CBP), but not the other acetyltransferases tested, catalysed NP acetylation at Lys375 (Extended Data Fig. 4a,b). Furthermore, knockdown of endogenous CBP with independent small interfering RNA (siRNA) significantly inhibited NP acetylation (Extended Data Fig. 4c), indicating that host CBP mediates NP acetylation at Lys375. To consolidate that NP was acetylated at Lys375, we generated an antibody specific to Lys375-acetylated NP and found that cells treated with either the sirtuin inhibitor nicotinamide (NAM) or the HDAC inhibitor trichostatin A showed increased Lys375 acetylation of NP (Extended Data Fig. 4d), revealing that both sirtuins and HDACs might be involved in deacetylating the NP. Among the eleven HDACs (HDAC1–11) and seven sirtuins (SIRT1–7) tested, multiple enzymes—including HDAC6, -7 and -9, and Sirt1, -3, -4 and -5—were found to effectively deacetylate NP (Extended Data Fig. 4e).

**Fig. 6: Acetylation of SARS2-NP at Lys375 reduces LLPS and its ability to bind RNA.**

We next generated a mutant by replacing Lys with Gln, thus mimicking the acetylation state of Lys. Although SARS2-NP WT exhibited efficiency in the RNA-binding assay, a K375Q substitution strongly inhibited NP activity (Fig. 6d). Moreover, a K375R replacement resulted in a greater than fivefold increase in RNA-binding activity compared with the WT control (Fig. 6d). In line with its role in regulating Lys375 acetylation, gain or loss of CBP expression significantly enhanced or reduced the RNA-binding affinity of SARS2-NP, respectively (Extended Data Fig. 4f,g). Compared with SARS2-NP WT, the K375Q mutant exhibited severely reduced PLA signals both at the basal and virus-induced levels (Fig. 6e). Moreover, the K375Q mutant partially lost activity towards Lys63-linked poly-ubiquitination and aggregation of MAVS, whereas K375R exhibited enhanced ability (Fig. 6f). Consistent with this, the K375Q mutant was less efficient in suppressing IFN-β promoter activity and IFNB1 mRNA production as well as in enhancing GFP–VSV replication than SARS2-NP WT, whereas these activities were significantly enhanced in the K375R mutant (Fig. 6g and Extended Data Fig. 4h,i). Therefore, the NP-mediated attenuation of MAVS activity and antiviral signalling could be partially rescued by Lys375 acetylation.

To verify the effect of acetylation on NP function, we employed genetic code expansion orthogonal systems^37,38 to incorporate acetyl-Lys into a recombinant SARS2-NP to create a fully Lys375-acetylated NP (SARS2-NP^Lys375-Ac). In this system, a pBAD/Myc–His vector carrying the ORF for SARS2-NP with an amber codon at the desired site for Lys375 was co-transformed with a plasmid harbouring aminoacyl-transfer RNA synthetase (pSupAR-tRNA-chRS42) into the Escherichia coli strain DH10B. The E. coli were subsequently treated with NAM and N^ε-acetyl-Lys and induced with l-arabinose to express fully N^ε-acetylated Lys375 NP (SARS2-NP^Lys375-Ac; Fig. 6h, left); SARS2-NP WT in the same vector were purified as non-acetylated control (SARS2-NP^Non-Ac; Fig. 6h, left).

As expected, antibodies specific to Lys375-acetylated NP can detect SARS2-NP^Lys375-Ac but not SARS2-NP^Non-Ac (Fig. 6h, right). In SPR assays, SARS2-NP^Lys375-Ac was much less efficient than SARS2-NP^Non-Ac in binding to viral RNA in vitro (Fig. 6i), indicating that Lys375-acetylated NP is indeed less active. To test whether acetylation alters phase separation of NP in vitro, we incubated fluorescently labelled NP with viral RNA. SARS2-NP^Non-Ac and RNA rapidly formed micrometre-sized liquid droplets and the small liquid droplets quickly fused into larger ones, accompanied by increased fluorescence intensity and a larger equivalent diameter. In contrast, SARS2-NP^Lys375-Ac and RNA slowly formed smaller droplets with much less liquid (Fig. 6j). FRAP experiments showed that following bleaching, SARS2-NP^Non-Ac binding to RNA was recovered more efficiently than that of SARS2-NP^Lys375-Ac (Fig. 6k). Thus, NP acetylation at Lys375 abrogates its LLPS.

K375N, K375E and K375Q variants that mimic acetylation exhibit reduced LLPS and ability to suppress innate antiviral immunity

RNA viruses have very high mutation rates, enhancing evolvability and altering their virulence³⁹. On 26 May 2021, 1,096,918 SARS-CoV-2 sequences had already been reported and deposited (https://bigd.big.ac.cn/ncov)^40,41. Analysis of the corresponding NPs revealed that several variants, including K375Q, and more frequently reported as K375N and K375E, existed for the SARS2-NP (Fig. 7a). Mutation of Lys to Gln, Asn and particularly Glu reduces its positive charge (Gln and Asn are neutral, and Glu has a negative charge), which to a certain extent mimics acetylation of the corresponding Lys residue (Fig. 7a). Considering that Lys375 is adjacent to the DD, a key region responsible for the NP–RNA association, NP LLPS and the NP-mediated blockage of RIG-I-like-receptor signalling, acetylation and acetyl-mimicking mutations at this site probably affect the electrostatic interactions between the DD and RNA, thereby interfering with SARS2-NP function. Thus, the K375N and K375E variants probably attenuate NP-mediated innate immune suppression, in turn reducing virulence. We sought to experimentally confirm this hypothesis.

**Fig. 7: K375N and K375E variants that mimic SARS2-NP acetylation exhibit lower RNA affinity, LLPS and immunosuppression.**

As expected, the purified K375N and K375E mutants seemed to exhibit reduced binding affinity for viral RNA in SPR and oligo pull-down experiments in vitro, during which a more obvious reduction was observed for K375E (Fig. 7b,c). This was consistent with the idea that K-to-E mutations resulted in a more dramatic charge reduction. To test whether these viral mutations induce changes in the phase separation of SARS2-NP, we incubated fluorescently labelled K375N and K375E with viral RNA and compared the micrometre-sized liquid droplet formation with that of SARS2-NP WT. Both the K375N and K375E mutants seemed to have compromised LLPS (Fig. 7d). All of the Lys375 variants of SARS2-NP—that is, K375N, K375E and K375Q—were less effective in suppressing MAVS droplet formation when they were incubated in vitro (Extended Data Fig. 5).

In cells, PLA revealed much weaker signals for K375N and K375E when compared with WT SARS2-NP (Fig. 7e), which is indicative of the impaired dimerization and self-association of SARS2-NP. An SDD–AGE assay confirmed that the K375N and K375E variants were also less efficient at forming detergent-resistant condensates (Extended Data Fig. 6a). In line with this, both variants partially lost their ability to inhibit the SeV-induced Lys63-linked poly-ubiquitination and aggregation of MAVS (Fig. 7f,g). Furthermore, a significantly reduced ability to suppress IFN-β promoter activity (Extended Data Fig. 6b) and IFNB1 mRNA expression (Extended Data Fig. 6c) was observed for the mutants in addition to a reduced ability to upregulate VSV–GFP replication (Extended Data Fig. 6d). To evaluate the functional consequence of SARS2-NP mutations in vivo, we engineered recombinant VSVs expressing the K375N, K375E and K375Q variants (hereafter, NP–VSV K375N, NP–VSV K375E and NP–VSV K375Q, respectively). Compared with mice infected with NP–VSV, expression of Ifnb1 mRNA was significantly higher in the spleen, liver, and lungs of the mice infected with NP–VSV K375N, NP–VSV K375E and NP–VSV K375Q (Fig. 7h). The concentration of IFN-β in the sera of the mice infected with NP–VSV K375N, NP–VSV K375E or NP–VSV K375Q was also higher (Fig. 7i). The levels of VSV-specific mRNA and the VSV titres were lower in the spleen, liver and lungs of the mice infected with the K375N, K375E and K375Q NP–VSV variants than the NP–VSV-infected mice (Fig. 7j,k). Mice that were infected with the three mutant VSVs had lower mortality rates than the NP–VSV-infected mice (Fig. 7l).

To further determine the role of SARS2-NP mutations at Lys375 in SARS-CoV-2 infection, Caco-2 cells were transfected to express SARS2-NP WT, K375N, K375E or K375Q and subsequently challenged with SARS-CoV-2. As expected, exogenous expression of SARS2-NP WT sharply impeded the expression of IFNB1 mRNA and enhanced the replication of SARS-CoV-2 genomic RNA, whereas the Lys375 mutants poorly inhibited IFN-β production, resulting in a lower SARS-CoV-2 genomic RNA load (Fig. 7m). Together, modifications and mutations of the critical Lys375 residue of SARS2-NP could suppress its function, enhancing the innate antiviral response of the host.

Interfering peptide NIP-V targeting the DD disrupts SARS2-NP LLPS and thus enhances the innate antiviral response both in vitro and in vivo

Our findings revealed the indispensable role of the DD of SARS2-NP for SARS2-NP function and immunosuppression. This prompted us to develop a strategy to block the DD. We noticed that protein domains containing natural l-peptides can be mimicked using d-amino acids with a retro-reversed sequence⁴². Peptide modified with such d-retroinverso (DRI) isoforms can yield new chemical properties, potentially enhancing the peptide potency^43,44,45. Several DRI-modified peptides were well-tolerated and therapeutically effective in a double-blinded, randomized, placebo-controlled phase IIb trial^46,47,48 and a phase I trial for the systemic treatment of solid tumours⁴⁹. This existing evidence provided the rationale for designing a well-tolerated and therapeutically effective anti-SARS2-NP peptide with a DRI conformation. Furthermore, recent reports determined the structure of the SARS2-NP DD dimer^17,19, describing that the predominant dimerization interface of each DD subunit included two β strands and η1, α3, α4 and α5 helices, located in five regions of the DD. To prevent SARS2-NP-mediated innate immune suppression, we designed and synthesised multiple interfering peptides with a DRI conformation, named NP interfering peptide (NIP) I–V, all of which were designed to inhibit SARS2-NP dimer formation by blocking different DD interaction regions (Fig. 8a,b). To facilitate the cellular uptake of NIPs, peptides were designed to fuse with HIV-TAT (Fig. 8a), a basic and hydrophilic sequence that allows for the energy-independent cellular uptake of cargo through transient pore formation^43,50.

**Fig. 8: The NIP-V peptide targeting the DD disrupts SARS2-NP LLPS and thus elevates the innate antiviral response in vitro and in vivo.**

In a phase separation assay, NIP-V was screened out to disrupt droplet formation in vitro (Fig. 8c and Extended Data Fig. 7a). NIP-V treatment hampered the formation of NP-enriched liquid-phase droplets in living cells (Fig. 8d and Extended Data Fig. 7b), thereby establishing the inhibitory role of NIP-V on SARS2-NP LLPS in vivo. Compared with the control, cells treated with NIP-V showed no detectable NP self-association (Extended Data Fig. 7c). We subsequently examined whether this interfering peptide could counteract the inhibitory effect of NP on IFN-β signalling. As expected, treatment with NIP-V significantly enhanced the expression of Ifnb1 and Isg56 while reducing the levels of VSV-specific mRNA (Fig. 8e). Higher NIP-V concentrations were found to inhibit SARS2-NP condensation in cells (Fig. 8f) and thus relieve the NP-mediated suppression of MAVS aggregation (Fig. 8g), indicating that this peptide could probably rescue MAVS activity. Quantitative PCR (qPCR) analysis revealed that NP overexpression blocked IFNB1 mRNA expression induced by RIG-IN and MAVS but not by IRF3-5D. This effect could be partially rescued by NIP-V (Extended Data Fig. 7d), confirming the innate immunity-enhancing effect of this peptide.

To determine the function of NIP-V in vivo, mice that had been pre-injected with or without NIP-V were infected with NP–VSV. The expression of Ifnb1 and Isgs mRNA was significantly higher in the spleen, liver and lungs of the NIP-V-treated mice compared with the PBS-treated mice (Extended Data Fig. 8a,b). An ELISA assay revealed that the concentration of IFN-β was also significantly higher in the serum of the NIP-V-treated mice (Extended Data Fig. 8c). Consistent with these results, NIP-V treatment considerably reduced the VSV-specific mRNA levels, VSV-specific protein VSV-G expression and NP–VSV titres in the lungs, liver and spleen of mice (Extended Data Fig. 8d–f). Notably, the mice that were pretreated with NIP-V seemed to be less sensitive to NP–VSV infection than those pretreated with PBS (Extended Data Fig. 8g). Staining with H&E confirmed that there was less damage in the lungs of the NIP-V-treated mice (Extended Data Fig. 8h). Importantly, to determine whether blocking NP could inhibit SARS-CoV-2 replication, we treated SARS-CoV-2-infected CaCo-2 cells with NIP-V. We observed a gradual dose-dependent elevation of IFNB1 mRNA expression following NIP-V treatment, which consequently suppressed active viral replication (as assessed by the levels of the SARS-CoV-2 subgenomic RNA4-encoding E gene) and viral titres (Fig. 8h). To further evaluate the function of NIP-V in the context of SARS-CoV-2 infection, human ACE2 (hACE2) transgenic mice were pre-injected with NIP-V or PBS and then intranasally inoculated with SARS-CoV-2 at 50% tissue culture infectious dose (TCID₅₀) of 1 × 10⁵. Compared with the PBS-treated mice, SARS-CoV-2-induced Ifnb1 and Isg56 mRNA expression in the spleen, liver and lungs were higher in the NIP-V-treated mice (Fig. 8i and Extended Data Fig. 8i). NIP-V treatment also potentiated the IFN-β induction in serum of SARS-CoV-2-infected mice (Fig. 8j). Consistent with these results, the levels of SARS-CoV-2 genomic RNA in the spleen, liver and lungs were drastically reduced in the NIP-V-treated mice (Fig. 8k). Staining with H&E indicated that SARS-CoV-2 infection led to severe lung damage, characterized by thickened alveolar septa and inflammatory-cell infiltration in the PBS-treated hACE2 transgenic mice, whereas only minor lung damage was observed in the mice pretreated with NIP-V (Fig. 8l). Consistent with this, the expression of the S (Fig. 8m) and N (Extended Data Fig. 8j) proteins in the lung were remarkably reduced in the NIP-V-treated mice compared with the PBS-treated mice. Together, the results from the in vitro and in vivo analyses demonstrated that NIP-V could disrupt SARS2-NP LLPS and thus alleviate the NP-mediated suppression of the innate antiviral responses (Fig. 8n).

Discussion

The IFN-I response has been demonstrated to protect the host against SARS-CoV and MERS-CoV in various animal models, including mice, marmosets and macaques^51,52,53. The addition of exogenous IFN-I to cell cultures efficiently inhibits the proliferation of SARS-CoV, MERS-CoV and SARS-CoV-2 (refs. ^{51,54,55,56,57,58,59}). To survive in the host, viruses employ various immune evasion mechanisms to counteract innate immune sensing and IFN-I production. Compared with other respiratory RNA viruses, SARS-CoV-2 exhibits an increased immunosuppressive ability and induces low levels IFN-I production both in vitro and in vivo^54,60. In the serum of patients with COVID-19, IFN-I expression could not be detected by commonly used assays, despite efficient replication of SARS-CoV-2 in human lung tissue^1,54,61,62, which suggests that SARS-CoV-2 harbours a highly efficient mechanism for the inhibition of innate antiviral immunity. However, although several SARS-CoV-2 viral proteins were shown to be implied in immune evasion⁶³, the precise evasion mechanism remains elusive. In this study we demonstrated that the introduction of SARS2-NP, but not SARS2-NP d_DD, leads to a compromised MAVS signalling response, increased viral proliferation and higher mortality in mice. This finding is in agreement with that of a previous report stating that the virulence and clinical severity of coronaviruses are associated with immune evasion⁶⁴. It is known that human coronavirus 229E induces high levels of IFN and thus exhibits mild virulence⁶⁵, whereas SARS-CoV and MERS-CoV utilize multiple mechanisms to repress the IFN response, contributing to their high lethality^66,67,68.

While our manuscript was in consideration, other groups reported that SARS2-NP undergoes LLPS with RNA^{26,39,69,70,71,72,73,74}, which contributes to virion assembly as well as viral transcription and replication^59,60,61. Our findings indicate that SARS2-NP LLPS is associated with its ability to inhibit IFN-I signalling. Notably, we observed SARS2-NP acetylation and deacetylation at Lys375 by host acetyltransferases and deacetylases, which in turn modulated NP self-condensation and NP-regulated MAVS activity. Importantly, we found that the patient-derived SARS-CoV-2 virus was frequently mutated at SARS2-NP Lys375—including the K375N, K375E and K375Q variants, all of which had a reduced positive charge and thus mimicked acetylation. Furthermore, SARS2-NP harbouring these mutations exhibited lower levels of RNA binding, reduced LLPS and reduced ability to suppress innate antiviral immunity.

In this study we designed interfering peptides against all interaction structures within the DD of SARS2-NP. One of the peptides, NIP-V, was screened out as a potent in vitro inhibitor of SARS2-NP LLPS, rescuing MAVS activity and the antiviral immune response in vivo. Thus, the current findings highlight SARS2-NP LLPS as a promising therapeutic target for rescuing innate antiviral immunity during SARS-CoV-2 infection. We believe these insights may help curb the ongoing pandemic as well as any future outbreaks.

Methods

Mice

Mice were maintained under specific-pathogen-free conditions in the animal facility of Soochow University. The hACE2 transgenic mice were purchased from Shanghai Model Organisms Center, Inc. Animal experiments involving infectious virus were performed in the physical containment level 2 laboratory of Soochow University. The Institutional Committee for Animal Welfare of Soochow University approved and oversaw this study. Authentic SARS-CoV-2 challenge studies were approved by the Ethics Committee of ZSSOM of Sun Yat-sen University on Laboratory Animal Care. All of the authentic SARS-CoV-2-related experiments were conducted in a biosafety-level-3 facility. All of the animal studies were conducted in accordance with the Soochow University guidelines or in compliance with the guidelines and regulations of the Laboratory Monitoring Committee of Guangdong Province of China.

Cells and reagents

HEK293T, HeLa, A549, U2OS, Vero E6 and Caco-2 cells obtained from the American Type Culture Collection were cultured at 37 °C under 5% CO₂ in DMEM medium supplemented with 10% fetal bovine serum, 100 U ml⁻¹ penicillin and 100 μg ml⁻¹ streptomycin. Peritoneal macrophages were harvested from mice 4 d after injection of thioglycolate (BD) and cultured in DMEM supplemented with 5% fetal bovine serum. All cells were negative for mycoplasma. Poly(I:C) and 5′-ppp RNA were purchased from Invivogen. MG132 was purchased from SelleckChem.

Plasmids and transfection

Expression plasmids for RIG-I, MAVS, TRAF3, TBK1, IKKɛ and IRF3, and plasmids for the IFNB1-Luc and ISRE-Luc transcriptional reporters were described previously⁷⁵. SARS-CoV NP and SARS2-NP were synthesised by GENEWIZ, China. Sequences encoding the NPs were amplified by standard PCR and cloned into the pcDNA3.1-Myc and pLV-Flag vectors. Point mutations were generated by site-directed mutagenesis with KOD plus polymerase (Toyobo) and the plasmids encoding the WT protein were used as the template. All constructs were confirmed by DNA sequencing. Standard polyethylenimine was used for transient transfection of plasmids and siRNA into HEK293T cells. Plasmids and RNA mimics were transfected into macrophages using Geneporter 2 transfection reagent (Genlantis).

Transcription reporter assay

HEK293T cells were seeded into 48-well plates, transfected with plasmids using polyethylenimine, and subsequently (mock) treated for various times—as indicated in the figures—and harvested for the measurement of luciferase activity. The internal transfection control Renilla was used to normalize the luciferase activities to control for differences in transfection efficiency.

Real-time qPCR with reverse transcription

Total RNA was prepared using the RNAiso Plus kit (Takara). A total of 1 μg RNA was reverse-transcribed using a PrimeScript RT reagent kit (Perfect Real Time; Takara). Real-time PCR was conducted using TB Green premix Ex Taq (Tli RNaseH Plus; Takara) using a StepOne Plus real-time PCR system (Applied Bioscience). Quantitation of all target gene expression was normalized to the control gene Gapdh for mouse genes and GAPDH for human genes.

The qPCR primers were as follows: murine Ifnb1 forward, 5′-TCCTGCTGTGCTTCTCCACCACA-3′; murine Ifnb1 reverse, 5′-AAGTCCGCCCTGTAGGTGAGGTT-3′; murine Cxcl10 forward, 5′-ATCATCCCTGCGAGCCTATCCT-3′; murine Cxcl10 reverse, 5′-GACCTTTTTTGGCTAAACGCTTTC-3′; murine Ccl5 forward, 5′-TCACCATATGGCTCGGACACCAC-3′; murine Ccl5 reverse, 5′-TTGGCACACACTTGGCGGTTC-3′; murine Isg56 forward, 5′-AAGACAAGGCAATCACCCTCTACT-3′; murine Isg56 reverse, 5′-GTCTTTCAGCCACTTTCTCCAAA-3′; murine Gapdh forward, 5′-GGCCTTCCGTGTTCCTACC-3′; murine Gapdh reverse, 5′-AGCCCAAGATGCCCTTCAGT-3′; VSV forward, 5′-ACGGCGTACTTCCAGATGG-3′; VSV reverse, 5′-CTCGGTTCAAGATCCAGGT-3′; human IFNB1 forward, 5′-CCAACAAGTGTCTCCTCCAAAT-3′; human IFNB1 reverse, 5′-AATCTCCTCAGGGATGTCAAAGT-3′; human GAPDH forward, 5′-AGGGCTGCTTTTAACTCTGGT-3′; human GAPDH reverse, 5′-CCCCACTTGATTTTGGAGGGA-3′. human ISG56 forward, 5′-GCTTTCAAATCCCTTCCGCTAT-3′; human ISG56 reverse, 5′-CTTGGCCCGTTCATAATTTTTTC-3′; human CXCL10 forward, 5′-TTTGCTGCCTTATCTTTCTGACT-3′; human CXCL10 reverse, 5′-ATTGTAGCAATGATCTCAACACG-3′; SARS2 E forward, 5′-CTTTCGTGGTATTCTTGCTAGTT-3′ and SARS2 E reverse, 5′-CACGTTAACAATATTGCAGCA-3′.

Immunoprecipitation and immunoblot analyses

Cells were lysed with 1 ml lysis buffer (20 mM Tris–HCl, pH 7.4, 2 mM EDTA, 25 mM NaF and 1% Triton X-100) containing protease inhibitors (Sigma) for 10 min at 4 °C. Immunoprecipitation was performed with anti-Flag M2 beads (Sigma, A2220) for 1 h at 4 °C or with various antibodies (see below) and protein A–sepharose (GE Healthcare Bio-Sciences AB) for 3 h at 4 °C. Thereafter, the precipitates were washed three times with washing buffer (50 mM Tris–HCl, pH 8.0, 150 mM NaCl, 1% Nonidet P-40, 0.5% sodium deoxycholate and 0.1% SDS) and the immunocomplexes were eluted with sample buffer containing 1% SDS for 5 min at 95 °C. Immunoblot analysis was performed with specific antibodies and secondary anti-mouse or anti-rabbit antibodies conjugated to horseradish peroxidase (HRP). Visualization was achieved with chemiluminescence. The antibodies used for immunoprecipitation (IP), immunoblotting (IB) and immunofluorescence (IF) were as follows: mouse monoclonal antibody to SARS2-NP (ABclonal, A20142; 1:2,000 for IB), rabbit monoclonal antibody to MAVS (Cell Signaling, 4983 and 3993; 1:1,000 for IB), anti-acetyl-Lys (Abcam, ab80178; 1:2,000 for IB), MAVS antibody (provided by F. Hou; 1:200 for IP), rabbit monoclonal antibody to IRF3 (Cell Signaling, D614C; 1:1,000 for IB and 1:200 for IF), rabbit monoclonal antibody to phospho-IRF3(Ser 396) (Cell Signaling, 4D4G; 1:1,000 for IB), rabbit monoclonal antibody to TBK1 (Cell Signaling, D1B4; 1:1,000 for IB), rabbit monoclonal antibody to phospho-TBK1(Ser172) (Cell Signaling, D52C2; 1:1,000 for IB), rabbit monoclonal antibody to IKKɛ (Cell Signaling, D20G4; 1:1,000 for IB), rabbit monoclonal anti-phospho-IKKɛ (Ser172) (Cell Signaling, 8766; 1:1,000 for IB), mouse monoclonal anti-Flag (M2) (Sigma-Aldrich, F3165; 1:2,000 for IB and 1:200 for IF), rabbit polyclonal anti-HA (Y11) (Santa Cruz Biotechnology, sc-805; 1:2,000 for IB), rabbit monoclonal anti-HA (12CA5) (made in-house; 1:2,000 for IB), rabbit polyclonal anti-Myc (A-14) (Santa Cruz Biotechnology, sc-789; 1:2,000 for IB), rabbit polyclonal anti-CBP (Santa Cruz Biotechnology, sc-583; 1:1,000 for IB), mouse monoclonal anti-Myc (9E10) (Santa Cruz Biotechnology, sc-40; 1:2,000 for IB), mouse monoclonal anti-actin (Sigma-Aldrich, AC-15; 1:2,000 for IB), anti-VSV-G (ABGENT, AP1016a; 1:1,000 for IB), HRP-mouse anti-biotin (Invitrogen, 03-3720; 1:1,000), protein A–HRP (Sigma-Aldrich, GENA9120; 1:10,000 for IB) and HRP-conjugated secondary antibodies (Cell Signaling, 7076 (anti-mouse IgG) or 7074 (anti-rabbit IgG); 1:10,000 for IB); the polyclonal antibody to acetylated Lys375 of SARS2-NP was generated by immunizing rabbits with Cys-KDKKK(Kac)ADETQAL peptide (1:2,000 for IB and 1:200 for IP). Fluorescein AffiniPure goat anti-mouse IgG (H + L) (115-095-146), rhodamine (TRITC) AffiniPure goat anti-mouse IgG (H + L) (115-025-146), fluorescein AffiniPure goat anti-rabbit IgG (H + L) (111-095-003) and rhodamine (TRITC) AffiniPure goat anti-rabbit IgG (H + L) (111-025-003) were from Jackson ImmunoResearch.

SDD–AGE

The SDD–AGE protocol was previously described³⁶. Briefly, cells were lysed in Buffer A (10 mM Tris–HCl, pH 7.5, 10 mM KCl, 1.5 mM MgCl₂, 0.25 M d-mannitol and protease inhibitor cocktail) by repeated douncing, and the homogenized cells were centrifuged at 700g for 10 min at 4 °C. The supernatant was transferred and centrifuged at 10,000g for 30 min at 4 °C to form a pellet, which was the intact crude mitochondrion (P5). The crude mitochondria (P5) were resuspended in 1×sample buffer (0.5×TBE, 10% glycerol, 2% SDS and 0.0025% bromophenol blue) and loaded onto a vertical 1.5% agarose gel. After electrophoresis in the running buffer (1×TBE and 0.1% SDS) for 35 min with a constant voltage of 100 V at 4 °C, the proteins were transferred to an Immobilon membrane (Millipore) for immunoblotting.

Protein fractions in sucrose gradient

Protein-fraction isolation on a sucrose gradient was previously described³⁶. Briefly, crude mitochondria (P5) isolated as described in the ‘SDD–AGE’ section were resuspended in Buffer B (20 mM HEPES (pH 7.4), 10% glycerol, 0.5 mM EGTA and 0.25 M d-mannitol) and loaded on top of a centrifuge tube containing 1 ml of 50% sucrose in PBS on the bottom layer and 1 ml of 40% sucrose in PBS on the top layer. After centrifugation at 100,000g for 30 min, the mitochondria enriched at the interface of the two layers were collected and solubilized in PBS containing 1% n-dodecyl-β-d-maltosid. The mitochondrial lysate was loaded onto a sucrose gradient (10–60%) and centrifuged at 160,000g for 2 h. Seven fractions of equal volume were taken from the top to bottom of the tube and loaded on the SDS–PAGE gel.

TUBE1 assay

The isolation of endogenous K63-linked polyubiquitinated substrates by TUBE1 was previously described⁷⁶. Equilibrated agarose–TUBEs were incubated with cell lysis buffer for 4 h at 4 °C. The cell lysate was clarified by centrifugation for 10 min at approximately 12,000g (4 °C). The agarose–TUBEs were then washed with TBST and the resin was treated with 0.2 M glycine–HCl, pH 2.5 on a rocking platform for at least one 1 h at 4 °C. The resin was then collected by high-speed centrifugation (13,000g) for 5 min and the supernatant was recovered.

In vivo ubiquitination assay

Cells were washed with PBS and lysed in two pellet volumes of RIPA buffer (20 mM NAP, pH 7.4, 150 mM NaCl, 1% Triton X-100, 0.5% sodium deoxycholate and 1% SDS) supplemented with protease inhibitors and 10 mM N-ethylmaleimide. The lysates were sonicated, boiled at 95 °C for 5 min, diluted with RIPA buffer containing 0.1% SDS and then centrifuged at 4 °C (16,000g for 15 min). The supernatant was incubated with specific antibody and protein A–sepharose for 3 h at 4 °C. After extensive washing, the bound proteins were eluted with 2×SDS sample buffer and separated on an SDS–PAGE gel, followed by western blotting.

Native PAGE

Cells were lysed in RIPA lysis buffer (50 mM Tris–HCl, pH 7.4, 150 mM NaCl, 1% NP-40 and 100 μM PMSF) and 15 μg supernatant of the TCL was used for native PAGE after centrifugation; 7.5% native PAGE gels were made without SDS. The gels were pre-run with 25 mM Tris and 192 mM glycine, pH 8.4, with 1% deoxycholate in a cathode chamber for 30 min at 40 mA. Samples in native sample buffer (10 μg protein, 62.5 mM Tris–HCl, pH 6.8, 15% glycerol and 1% deoxycholate) were size-fractionated by electrophoresis for 60 min at 25 mA and transferred to PVDF membranes for immunoblot analysis.

Purification of Lys375-specific fully acetylated SARS2-NP recombinant proteins

The K375 acetylated SARS2-NP recombinant proteins were generated according to a previous report³⁸. After optimizing the constructs and methods, the E. coli DH10B strain was transformed with the plasmid aminoacyl-tRNA synthetase pSupAR-tRNA-chRS42 and pBAD/Myc–His vector carrying the ORF of SARS2-NP with an amber codon at the desired site for Lys375. The cells were first cultured overnight in LB medium supplemented with 100 μg ml⁻¹ ampicillin and 50 mg ml⁻¹ chloramphenicol (Amp⁺ Cm⁺) at 37 °C. A 10-ml volume of the overnight bacterial culture was then subcultured in 200 ml LB Amp⁺ Cm⁺. When the optical density at 600 nm (OD₆₀₀) reached 0.6–0.7, 20 mM NAM and 100 mM acetyl-Lys were added, and 30 min later, when the OD₆₀₀ reached 0.8–1.0, protein expression was induced at 30 °C for 18–20 h by adding 2 mM l-arabinose. The cells were harvested after induction and washed with ice-cold PBS containing 20 mM NAM; the proteins were purified with HisTrap FF (GE Healthcare, 17-5319-01) according to the manufacturer’s protocol.

Immunofluorescence and confocal microscopy

HeLa or U2OS cells cultured on glass coverslips were transfected with plasmids or treated with various stimuli as indicated in the figures. The cells were then washed with PBS, fixed with 4% paraformaldehyde in PBS for 20 min, permeabilized with 0.2% Triton X-100 and blocked with 3% BSA. The cells were then stained with the appropriate antibodies (as listed in the ‘Immunoprecipitation and immunoblot analyses’ section and indicated in the figures), followed by incubation with fluorescent-dye-conjugated secondary antibodies. The nuclei were counterstained with DAPI (Sigma-Aldrich). The tissue paraffin sections were blocked with 3% BSA and 0.1% Tween-20 in PBS for 1 h after rehydration and antigen retrieval, and the sections were then stained with the indicated antibodies in 3% BSA, followed by incubation with fluorescent-dye-conjugated secondary antibodies in 3% BSA. The nuclei were counterstained with DAPI in 3% BSA.

In vitro phase separation assay

Purified proteins were diluted to various concentrations in buffer containing 20 mM Tris–HCl pH 5.5 and 1 mM dithiothreitol with the indicated salt concentrations. Protein solution (5 μl) was loaded onto a glass slide, covered with a coverslip and imaged using an LSM880 confocal microscope system (Zeiss). Phase separation of recombinant EGFP–SARS2-NP with Cy3-labelled RNA was performed in physiological buffer (20 mM Tris–HCl, pH 7.0, 15 mM NaCl, 135 mM KCl, 5 mM phosphate, 1.5 mM MgCl₂ and 1 mg ml⁻¹ BSA). The mixture was transferred to 96-well plates (Corning) coated with 20 mg ml⁻¹ BSA (Sigma) and imaged.

The sequence of the Cy3- or Cy5-labelled SARS-CoV-2 RNA was as follows: 5′-CACUCGCUAUGUCGAUAACAACUUCUGUGGCCCUGAUGGCUACCCUCUUGAGUGCAUUAAAGA-3′.

In vitro and in vivo FRAP assays

The FRAP experiments were performed on a Zeiss LSM880 confocal microscope system. For FRAP of SARS2-NP, spots of approximately 2 μm in diameter in droplets of approximately 10 μm were photobleached with 20% laser power for 1 s using 488 nm lasers. Cellular FRAP was performed on a Zeiss LSM880 confocal microscope system at 37 °C in a live-cell-imaging chamber and the SARS2-NP was fully or partially photobleached with 20% laser power for 2 s using a 488 nm laser. Time-lapse images were acquired over 1 min after bleaching. Images were processed by ImageJ and FRAP data were fitted to a single exponential model using GraphPad Prism 8.

SPR

Biotin-labelled RNA was immobilized to a SA sensor chip using a Biacore T200 system (GE Healthcare). The running buffer contained 10 mM HEPES, pH 7.4, 150 mM NaCl, 3 mM EDTA and 0.05% P20. Serial dilutions of purified recombinant SARS2-NP were injected at a range of concentrations as indicated in the figure legends. The resulting data were fit to a 1:1 binding model using the Biacore Evaluation Software (GE Healthcare).

The biotin-labelled SARS-CoV-2 RNA had the following sequence: 5′-CACUCGCUAUGUCGAUAACAACUUCUGUGGCCCUGAUGGCUACCCUCUUGAGUGCAUUAAAGA-3′.

Oligo pull-down assay

To assay RNA binding to SARS2-NP in cell lysates, HEK293T cells were transfected with the indicated plasmids. After 36 h, the HEK293T cells were washed twice, collected and lysed in buffer containing 50 mM Tris–HCl, 150 mM NaCl, 1% NP-40, 5 mM EDTA, protease inhibitor cocktail and phosphatase inhibitor cocktail (Roche) for 30 min. The cell debris was pelleted by centrifugation at 16,000g for 15 min at 4 °C. Biotin-labelled RNA (2 μg) was first immobilized onto 15 μl of streptavidin sepharose beads. The beads were washed three times with lysis buffer (described above) to remove unconjugated ligands and then added to pre-cleared lysates to perform the standard pull-down assay.

Interfering peptides

The NIPs were as follows: NIP-I, H-FNGQTQEPGRpprrrqrrkkrg-OH (molecular weight (M_w), 2,706; lowercase font, HIV-TAT sequence); NIP-II, H-IRSMGFFASASPAFQAIQPWpprrrqrrkkrg-OH (M_w, 3,785); NIP-III, H-LKIAGTYTLWTGSPTVEMGIpprrrqrrkkrg-OH (M_w, 3,711); NIP-IV, H-SDASSMSQQLQKSFDDLpprrrqrrkkrg-OH (M_w, 3,460); and NIP-V, H-IHKNLLIVQDKFpprrrqrrkkrg-OH (M_w, 3,041).

The interfering peptides were manufactured by GL Biochem at >99% purity and stored at −20 °C in powder aliquots of 1 mg until use to avoid freeze–thaw artefacts. For in vitro experiments, the immunoprecipitates were dissolved in PBS to generate a 25 mM stock. For in vivo experiments, the immunoprecipitates were dissolved in PBS to generate a 5 mg ml⁻¹ stock solution, which was kept on ice until injection. Before injection, the solution was brought to room temperature.

Viral infection and plaque assay

To construct the recombinant VSV carrying GFP, SARS2-NP WT or SARS2-NP d_DD, the coding sequence of the GFP, SARS2-NP WT or SARS2-NP d_DD was inserted into VSV backbones between the VSV glycoprotein (G protein) and polymerase protein (L protein) using NheI and XhoI restriction enzyme sites³⁰. The VSV backbone harbours the M51R substitution in the matrix (M) gene, attenuating the virulence of original strain^28,29. The recombinant virus (GFP–VSV, NP–VSV or NP–VSV d_DD) was then recovered in Vero E6 cells³⁰. Mouse macrophages or other cells (2 × 10⁵) were plated 24 h before infection. The cells were infected with VSV (m.o.i. of 0.1) or SeV (100 haemagglutination units (HAU) ml⁻¹) for varying times as indicated in the figures. VSV plaque assay and VSV replication were determined by a standard TCID₅₀ assay on permissive Vero E6 cell monolayers in 96-well plates with a tenfold dilution series of the samples. After 1 h of infection, the plates were incubated for 48 h. The medium was then removed, and the cells were fixed with 4% paraformaldehyde for 15 min and stained with 1% crystal violet for 30 min before plaque counting.

Viral infection in vivo

For in vivo viral infection studies, eight-week-old mice were infected with VSV (5 × 10⁸ p.f.u. per mouse) by i.p. injection for 24 h. The eight-week-old hACE2 transgenic mice were intranasally inoculated with SARS-CoV-2 strain HB-01 at a dosage of 1 × 10⁵ TCID₅₀ for 16 and 24 h. We collected blood from the orbital sinus for ELISA and the lungs, spleen and liver from each mouse for analysis of RNA, protein and viral titres. To measure the VSV titres in the lungs, spleen and liver, snap-frozen tissues were weighed and homogenized three times (5 s each) in MEM. After homogenization, the suspensions were centrifuged at 1,620g for 30 min and the supernatants were used for plaque assays on monolayers of Vero E6 cells seeded in 96-well plates with a tenfold dilution series of the samples. For the survival experiments, the mice were monitored for survival after infection with VSV.

ELISA

The concentrations of IFN-β in culture supernatants and serum were measured using ELISA kits (R&D Systems).

Lung histology

Lungs from control or virus-infected mice were dissected, fixed in 10% phosphate-buffered formalin, embedded in paraffin, sectioned, stained with H&E solution and examined by light microscopy for histological changes.

Immunohistochemical staining and evaluation

Primary antibodies to Spike (Abcam, ab273433; 1:1,000) and Spike (ABclonal, A20022; 1:1,000) were used for immunohistochemical staining of formalin-fixed paraffin-embedded microarrays of lung tissues from each group of mice, according to previously described staining protocols⁷⁷. The quantification of staining was expressed as an H score. The H score was determined using the formula 3 × the percentage of strongly stained cells + 2 × the percentage of moderately stained cells + the percentage of weakly stained cells, yielding a range of 0–300.

Statistics and reproducibility

Statistical analyses were performed using a two-tailed unpaired Student’s t-test or as indicated in the figure legends. All of the assays were performed three times with similar results. The exact value of n, representing the number of mice in the experiments, is indicated in the figure legends. Statistical differences with a P value of 0.05 or less were considered significant. There was no exclusion of data points or mice. No randomization or blinding were used.

Reporting Summary

Further information on research design is available in the Nature Research Reporting Summary linked to this article.

Data availability

Source data are provided with this paper. All other raw data supporting the findings of this study are available from the corresponding author on reasonable request.

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Acknowledgements

This work was supported by a special programme from the Ministry of Science and Technology of China (grant no. 2016YFA0502500 to L.Z.), the Chinese National Natural Science Funds (grant nos U20A201376, 82041009, 31871405, 91753139, 31925013, 31671457, 31571460, 31870902, 32070907, 31871405, 31771619, 92053114, and 32070632), National Postdoctoral Program for Innovative Talent (grant no. BX2021208), Jiangsu National Science Foundation (grant nos BK20180043 and 19KJA550003), Shenzhen Science and Technology Program (grant no. JSGG20200225150431472), and special COVID-19 program of the Sino-German Center for Research Promotion (grant no. C-0023).

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These authors contributed equally: Shuai Wang, Tong Dai, Ziran Qin, Ting Pan.

Authors and Affiliations

Institutes of Biology and Medical Science, Soochow University, Suzhou, China
Shuai Wang, Tong Dai, Ziran Qin, Feng Chu, Lingfeng Lou & Fangfang Zhou
Center for Infection and Immunity Studies, School of Medicine, Sun Yat-sen University, Shenzhen, China
Ting Pan
MOE Laboratory of Biosystems Homeostasis and Protection and Innovation Center for Cell Signaling Network, Life Sciences Institute, Zhejiang University, Hangzhou, China
Long Zhang, Bing Yang & Huasong Lu
Department of Pharmaceutical Chemistry and the Cardiovascular Research Institute, University of California, San Francisco, CA, USA
Bing Yang
Faculty of Basic Medical Sciences, Chonqing Medical University, Chongqing, China
Huizhe Huang

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Shuai Wang
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Tong Dai
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Ziran Qin
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Ting Pan
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Feng Chu
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Lingfeng Lou
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Long Zhang
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Bing Yang
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Huizhe Huang
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Huasong Lu
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Fangfang Zhou
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Contributions

S.W., T.D. and Z.Q. designed the experiments and analysed the data. S.W., T.D., Z.Q., F.C., L.L. and T.P. performed the experiments. B.Y. performed the mass spectrometry analysis. H.H., H.L. and L.Z. provided valuable discussion. T.D. and F.Z. wrote the manuscript.

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Correspondence to Fangfang Zhou.

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Competing interests

The peptide NIP-V are subject to patent applications (patent no. 202110466103.7; China; F.Z., T.D., S.W., and L.Z.) and may be shared with research organizations for research and educational purpose only under an material transfer agreement to be discussed in good faith with the recipient; such a material transfer agreement may restrict the recipient to make any modifications to these materials. All of the other authors declare no competing interests.

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Peer review information Nature Cell Biology thanks Luke O’Neill and the other, anonymous, reviewer(s) for their contribution to the peer review of this work. Peer reviewer reports are available.

a-b, qPCR analysis of Ifnb1, Isg56 (a) and Cxcl10 (b) mRNA in the spleen, liver and lung of mice (n = 5 mice per group) injected with PBS (i.p.) or NIP-V (5 mg/kg, i.p.) for 2 h and then infected with NP–VSV (5 × 10⁸ PFU per mouse, i.p.) for 24 h. c, ELISA determination of IFN-β level in serum from mice as in a. d-e, qPCR analysis of VSV RNA (d) and plaque assay of VSV (e) in the spleen, liver, and lung of infected mice as in a. f, IB analysis of VSV-G in the spleen (top), liver (middle) and lung (bottom) of infected mice selected randomly as in a. n = 3 mice per group. g, Survival curves of mice (n = 10 mice per group) administrated with PBS or NIP-V (5 mg/kg, i.p.) for 2 h and then infected with NP–VSV (1 × 10⁹ PFU per mouse, i.p.). h, Microscopy of hematoxylin-and-eosin-stained lung sections from mice treated with PBS or VSV as in a. Scale bar, 500 µm (top) or 100 µm (bottom). Images represent five independent samples with a similar pattern. i, qPCR of Isg56 mRNA in the spleen, liver and lung (right) of hACE2 transgenic mice (n = 6 mice per control uninfected groups, n = 8 mice per infected groups) pretreated with PBS or NIP-V (25 mg/kg, i.p.) for 2 h and then given intranasal inoculation with SARS-CoV-2 (1×10⁵ TCID50 per mouse) for another 16 or 24 h. j, Immunofluorescence assay of the SARS2-NP antigen (green) of lung sections from hACE2 transgenic mice pretreated with PBS or NIP-V (25 mg/kg, i.p.) for 2 h and then intranasally inoculated with SARS-CoV-2 (1×10⁵ TCID50 per mouse) for 24 h. The nuclei were stained with DAPI (blue). Scale bar, 500 µm. n = 8 independent samples. Data are presented as mean ± s.d. (a-e, i, j (right)). Statistical analysis was performed using two-tailed Student’s t-test (a-e, i, j (right)) or log-rank test (g). Source data are available online.

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Wang, S., Dai, T., Qin, Z. et al. Targeting liquid–liquid phase separation of SARS-CoV-2 nucleocapsid protein promotes innate antiviral immunity by elevating MAVS activity. Nat Cell Biol 23, 718–732 (2021). https://doi.org/10.1038/s41556-021-00710-0

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Received: 17 December 2020
Accepted: 07 June 2021
Published: 08 July 2021
Issue Date: July 2021
DOI: https://doi.org/10.1038/s41556-021-00710-0

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Abstract

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Main

Results

The SARS2-NP DD is required for LLPS and viral RNA association

NP-induced innate immune evasion relies on the DD

The DD mediates NP-induced innate immune evasion in vivo

SARS2-NP negatively regulates MAVS–IRF3 activation and is enriched in MAVS droplets

SARS2-NP inhibits Lys63-linked poly-ubiquitination and aggregation of MAVS

SARS2-NP acetylation at Lys375 abrogates its LLPS and the NP-mediated suppression of MAVS signalling

K375N, K375E and K375Q variants that mimic acetylation exhibit reduced LLPS and ability to suppress innate antiviral immunity

Interfering peptide NIP-V targeting the DD disrupts SARS2-NP LLPS and thus enhances the innate antiviral response both in vitro and in vivo

Discussion

Methods

Mice

Cells and reagents

Plasmids and transfection

Transcription reporter assay

Real-time qPCR with reverse transcription

Immunoprecipitation and immunoblot analyses

SDD–AGE

Protein fractions in sucrose gradient

TUBE1 assay

In vivo ubiquitination assay

Native PAGE

Purification of Lys375-specific fully acetylated SARS2-NP recombinant proteins

Immunofluorescence and confocal microscopy

In vitro phase separation assay

In vitro and in vivo FRAP assays

SPR

Oligo pull-down assay

Interfering peptides

Viral infection and plaque assay

Viral infection in vivo

ELISA

Lung histology

Immunohistochemical staining and evaluation

Statistics and reproducibility

Reporting Summary

Data availability

References

Acknowledgements

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Authors and Affiliations

Contributions

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Supplementary information

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