Suppressor tRNA-mediated conditionally replicating live vaccine effectively protects pigs against Seneca Valley virus challenge

Abstract

Seneca Valley virus (SVV) causes porcine idiopathic vesicular disease, severely endangering the economic benefits of the swine industry. Artificial control of viral replication via the introduction of premature termination codons (PTCs) into the viral genome using suppressor tRNAs (sup-tRNAs) is a promising strategy for developing efficient vaccines. In this study, optimization of the SVV 3D gene for the incorporation of multiple PTCs resulted in a progeny virus harboring three PTCs with high replicative capacity and genetic stability. In mouse and pig models, compared with inactivated vaccines, the SVV-PTC vaccination elicited high antibody titers at 14 days post-primary immunization and induced a stronger Th1-biased cellular immune response. We further validated the safety and protective efficacy of the SVV-PTC vaccine candidate in swine models, confirming its complete attenuation in vivo, absence of horizontal transmission, and protective efficacy comparable to that of the commercial inactivated SVV vaccine. In conclusion, the development of live but replication-incompetent attenuated vaccines using sup-tRNA is a promising prevention and control strategy, providing a new insight for the development of novel SVV vaccines.

Introduction

Seneca Valley virus (SVV) belongs to the family Picornaviridae and is the sole member of the genus Senecavirus. It possesses a typical picornavirus “L-4-3-4” genome layout. The SVV genome is approximately 7.2 kb in length, containing a unique open reading frame (ORF) that encodes a large polyprotein (2181 amino acids). This polyprotein is cleaved to produce 12 mature proteins (5′-L-VP4-VP2-VP3-VP1-2A-2B-2C-3A-3B-3C-3D-3′), and the closest relatives of SVV within this family are members of the genus Cardiovirus¹. SVV can induce idiopathic vesicular disease in pigs, with clinical manifestations including vesicles on the snouts and coronary bands, sometimes accompanied by lameness, anorexia, lethargy, and transient fever. The mortality rate of neonatal piglets (mainly those aged 7 days) increases by 30–70%^2–4. Since 2015, Seneca Valley virus (SVV) has been reported in multiple countries, including Brazil, the United States, China, Thailand, Colombia, Vietnam, Chile, and others^3,5–10. Recent research reports indicate that SVV induces severe vesicles clinically observable on the lips, oral cavity, and hoof nails of infected buffaloes^11,12. Hence, the development of safe and highly effective vaccines is of great significance to prevent the transmission and potential pandemic of SVV.

Currently, various types of vaccines have been developed for SVV, including inactivated vaccines (commercially available), nanoparticle vaccines, synthetic peptide vaccines, virus-like particle (VLP) vaccines, pseudorabies virus (PRV)-vectored recombinant live-vector vaccines, and live attenuated vaccines^13–18. These vaccines exhibit favorable protective efficacy, but some limitations to their immunogenicity still exist. The inactivated vaccines fail to induce rapid, robust immune responses after primary immunization¹⁴, while live attenuated vaccines carry safety risks¹⁹. With the advancement of synthetic biology, a growing number of studies have integrated genetic code expansion (GCE) technology with viral reverse genetics, giving rise to a novel type of viral vaccine: live but replication-incompetent virus vaccines^20–22. GCE technology enables site-specific incorporation of unnatural amino acids (UAAs) into nascent polypeptide chains, realizing precise in vitro regulation of viral replication^23,24. It has been successfully used to produce replication-incompetent influenza viruses²⁰. Nevertheless, the broader implementation of this methodology is hindered by suboptimal UAA incorporation rates and a dependence on extensive, stochastic mutagenesis-lacking predictive rational design-for target site identification^20,25,26. Additionally, nonsense suppressor tRNA (sup-tRNA or ACE-tRNA) achieves sense readthrough of PTCs by cross-reacting with endogenous aminoacyl-tRNA synthetases (aaRSs) to charge the correct amino acid²⁷, and is currently being explored for developing novel pseudorabies virus (PRV) vaccines²¹.

In this study, we compared the packaging efficiencies of GCE and sup-tRNA technologies for SVV-PTC production. Based on the comparison results, sup-tRNA technology was ultimately selected for the construction and preparation of the SVV-PTC strain. Although the PTC strategy has been successfully applied to rescue several viruses, our study not only evaluated the immunogenicity of the SVV-PTC vaccine in a mouse model but also further validated its in vivo safety, immunogenicity, and challenge protection efficacy in the natural host swine, demonstrating its favorable protective efficacy and safety in vivo. Collectively, our findings demonstrate that leveraging sup-tRNA technology to engineer replication-deficient viruses holds extensive translational promise for vaccine design. Furthermore, this research establishes a robust empirical foundation to guide the development of next-generation prophylactic strategies.

Results

Sup-tRNA exhibits higher PTC readthrough efficiency than GCE

Currently, the production of PTC-containing viruses mainly relies on two strategies: genetic code expansion technology and suppressor tRNA-mediated readthrough systems^20–22. This study aimed to generate SVV-PTC viruses using these two methods and compare their packaging efficiencies.

For construction of the GCE system, we employed an orthogonal translation system consisting of the pyrrolysyl-tRNA synthetase/tRNAᴄᴜᴀ pair (PylRS/tRNAᴄᴜᴀ) and the orthogonal unnatural amino acid (UAA) N(ε)-Boc-L-lysine (BOCK). In cells expressing PylRS/tRNAᴄᴜᴀ, BOCK supplementation allowed readthrough of viral genome PTC sites, thereby promoting synthesis of intact viral proteins and virus production. Without BOCK, translation aborted prematurely, precluding formation of functional viral proteins and infectious viral particles (Fig. 1A).

Fig. 1. Comparison of the PTC read-through activity of GCE and suppressor tRNA.

A Schematic of SVV-PTC generation using GCE technology, created with BioGDP.com. B Schematic diagram illustrating the mechanism of SVV-PTC production based on suppressor tRNA, created with BioGDP.com. C Engineering of a natural tyrosine-charging tRNA into a suppressor tRNA capable of recognizing the UAG termination codon. D The pCMV-8tRNA vector (pCMV-Tem3-8tRNA-chFSYRS-I267A-L349Y-final) and pLKO.1-2sup-tRNAy vector used in transient transfection. E pCMV-8tRNA was cotransfected with the mCherry or mCherryY43* in HEK293T cells. At 24 h post-transfection, fluorescence microscopy was used to evaluate the suppression efficacy. Scale bar: 200 μm. F Suppression efficiency of pLKO.1-2sup-tRNAy analyzed by fluorescence microscopy in transfected HEK293T cells. Scale bar: 200 μm. G pCMV-8tRNA was cotransfected with SVV 3D mutant plasmids (Y333, Y343, and 2Y*) into HEK293T. After transfection, UAA was added (+) or omitted (−), and cells were harvested at 24 h for Western blot analysis. H Quantitative analysis of the protein level of 3D in (G). I pCMV-8tRNA was cotransfected with SVV-eGFP mutants (3D-Y333*, 3D-Y343* and 3D-2Y*) into HEK293T cells, followed by the addition of UAA. EGFP expression was observed under a fluorescence microscope at 48 h post‑transfection. Scale bar: 200 μm. J pLKO.1-2×sup-tRNAy and SVV 3D mutant plasmids were co-transfected into HEK293T cells, and lysates were harvested for Western blot analysis. K Quantitative analysis of the protein level of 3D in (J). L pLKO.1-2×sup-tRNAy and SVV-eGFP mutants were cotransfected into HEK293T cells, and eGFP expression was observed. Scale bar: 200 μm. Error bars denote SEM. Significance was determined by one-way ANOVA. Exact p values are shown (n = 3).

In contrast, since suppressor tRNAs are derived from natural tRNA mutations, their readthrough of PTC sites does not require exogenous aminoacyl-tRNA synthetases or UAAs²⁸. Given that Tyr-tRNAᴄᴜᴀ (sup-tRNAy) efficiently reads through nonsense mutations²⁹, it was selected for SVV-PTC virus production. In cells harboring sup-tRNAy, during translation at the PTC site, sup-tRNAy competes with the eukaryotic release factor 1 (eRF1) to recognize the premature termination codon (TAG) and inserts tyrosine, allowing translation to proceed, achieving nonsense mutation readthrough, and ultimately producing infectious viral particles²⁷. Due to the presence of PTC sites in the viral genome, SVV-PTC can only complete a single round of infection in wild-type cells (Fig. 1B, C).

To evaluate the feasibility of PTC readthrough by the GCE system and sup-tRNAy, we constructed a reporter plasmid mCherryY43ᵀᴬᴳ (mCherryY43*) by site-directed mutagenesis. Subsequently, this plasmid was cotransfected into cells with the PylRS/tRNAᴄᴜᴀ expression vector, and BOCK was added to the culture medium. Fluorescence observation confirmed successful expression of mCherryY43* at position 43, which was dependent on the presence of PylRS, pyrrolysyl-tRNA, and BOCK, with the amber codon substitution being effectively achieved (Fig. 1D, E). After cotransfection of the mCherryY43* plasmid with the sup-tRNAy expression vector into cells, distinct red fluorescence was observed under fluorescence microscopy, while no fluorescence was detected in the control group (Fig. 1D, F). These findings confirm that both GCE technology and the sup-tRNAy system effectively enable readthrough of PTC sites.

To assess the read-through capability of the two technologies on the SVV 3D protein, two tyrosine codons (Tyr333 and Tyr343) were randomly selected in the gene encoding the SVV 3D protein and substituted with amber codons (TAG) by site-directed mutagenesis. The 3D single mutants 3D-Y333TAG (Y333*), 3D-Y343TAG (Y343*), and the double mutant 3D-Y333TAG-Y343TAG (2Y*) were co-transfected into HEK293T cells with the pCMV-Tem3-8tRNA-chFSYRS-I267A-L349Y-final (pCMV-8tRNA) or pLKO.1-2sup-tRNAy vector, respectively. Western blot analysis showed that, compared with sup-tRNA-mediated readthrough, GCE resulted in a greater amount of truncated, unreadthrough protein (Fig. 1G, J; Supplementary Fig. 1). Densitometric analysis further indicated that sup‑tRNAy exhibited better readthrough efficiency. (Fig. 1H, K). Furthermore, eGFP-tagged SVV PTC mutant plasmids (pSKII-SVV-eGFP-3D-Y333*, Y343*, and 2Y*) were cotransfected with pCMV-8tRNA or pLKO.1-2sup-tRNAy into cells for PTC virus packaging. Results demonstrated that eGFP expression was barely detectable in the GCE system. In contrast, in 293 T cells expressing sup-tRNAy, SVV with a single PTC site exhibited high read-through efficiency, while no eGFP expression was observed for SVV carrying the double PTC sites (Fig. 1I, L). Thus, suppressor tRNAy exhibits greater potential than GCE in the rescue of SVV-PTC.

Efficient packaging of SVV‑PTC mediated by sup‑tRNAy

Based on comparisons between the two technologies, we employed sup-tRNA to produce SVV-PTC. To improve the production efficiency of SVV-PTC, we constructed the vector pPB-CMV-mCherryY43*-IRES-Puro-8tRNAy (PB8) for SVV-PTC production (Supplementary Fig. 2A–C). We found that the PTC readthrough efficiency and viral rescue efficiency gradually increased with elevated sup-tRNAy copy numbers (Supplementary Fig. 2D–F). The SVV 3D gene was selected for PTC insertion, as upstream genes can be normally transcribed and translated post-infection to enhance host antiviral response. Therefore, we individually mutated all tyrosine codons (21 in total) in the 3D gene to the TAG stop codon by site-directed mutagenesis in both the 3D expression vector and the full-length infectious clone pSKII-SVV-eGFP.

Following cotransfection of PB8 with each 3D mutant, Western blot showed that all 3D mutant proteins could be read through to produce full length products. (Fig. 2A–D; Supplementary Figs. 4-5). We further evaluated the packaging efficiency of SVV-PTC using PB8. All PTC mutants displayed strong eGFP fluorescence and induced obvious cytopathic effect (CPE) in HEK293T cells (Fig. 2E). Viral copy numbers of the SVV-PTC variants were then determined by absolute quantitative PCR. The packaging efficiency varied among different mutants: the Y117 and Y394 mutants exhibited rescue efficiency similar to WT virus, and even the Y333* mutant with the lowest efficiency maintained a viral titer above 2×10⁸ VP/mL (Fig. 2F; Supplementary Fig. 6).

Fig. 2. Construction and examination of SVV-PTC mutants.

A–D Twenty-one TAG mutants were constructed by replacing Y codons with TAG in pCAGGS-HA-3D; each was cotransfected with PB8 or empty vector into HEK293T cells, and cells harvested at 24 hpt for Western blot. E Twenty-one TAG mutants were generated by replacing tyrosine (Y) codons with TAG at all Y residues in the 3D gene of the SVV-eGFP plasmid; each mutant was co-transfected with PB8 into HEK293T cells, and eGFP fluorescence expression was detected at 48 h post-transfection. Scale bar: 200 μm. F Quantitative detection of the copy numbers of SVV-eGFP-3D mutants was performed using qPCR (n = 3). Error bars denote SEM.

To further examine whether sup-tRNAy also enables efficient packaging of PTCs in other viral proteins, we mutated tyrosine residues in L, VP4, and 3C to TAG in the infectious clone and cotransfected the constructs with PB8 into HEK293T cells. Consistent with previous results, extensive eGFP fluorescence and clear CPE were observed at 48 h post-transfection (Supplementary Fig. 7).

Next, we applied the GCE system to rescue the 21 SVV-PTC mutants. Only two mutants (Y65 and Y73) showed weak eGFP expression, while no obvious fluorescence was detected in the others (Supplementary Fig. 8). In conclusion, these data confirm the efficiency and applicability of sup-tRNAy for SVV-PTC production.

Incorporation of multiple PTCs in the 3D gene enhances the genetic stability of SVV-PTC

Amber codons may revert to sense codons during the replication and passage of PTC viruses²⁰. To generate genetically stable PTC viruses, serial passage analysis was performed on all constructed mutant viruses in this study. Viruses obtained by co-transfection were designated as passage 0 (P0) and used to infect HEK293T cells expressing PB8 and wild-type HEK293T cells, respectively. Reversion mutations were determined by monitoring the fluorescent signals induced by viral infection. The results demonstrated that SVV mutants carrying PTC sites in the L, VP4, and 3 C proteins exhibited distinct fluorescent aggregation in wild-type HEK293T cells (Supplementary Fig. 9A). Among the 21 SVV mutants with PTC sites in the 3D protein, most exhibited fluorescence expression at varying levels, while only a few mutants (Y77, Y343, and Y444) displayed a marked reduction in fluorescent signals. (Fig. 3A, B; Supplementary Fig. 9B).

Fig. 3. Characterization of the in vitro stability of the SVV-PTC viruses.

A Each of the 21 SVV-3D-PTC viruses (P0) was used to infect HEK293T cells with or without PB8 pre-transfection; eGFP expression was examined at 48 h post-infection, and three stable PTC sites were screened out. Scale bar: 200 μm. B The three selected amino acid residues (Y77, Y343, and Y444) were labeled within the tertiary structure of the 3D protein. C Double and triple PTC mutants (3D-Y77*/Y343*, 3D-Y77*/Y444*, 3D-Y343*/Y444*, and 3D-Y77*/Y343*/Y444*) were generated using the SVV-eGFP plasmid. HEK293T cells were cotransfected with the respective mutant plasmids and PB8, followed by fluorescence observation at 48 h post-transfection and collection of the cell culture supernatants. Scale bar: 200 μm. D Viral copy numbers of mutants were quantified by qPCR (n = 3). Error bars denote SEM. E P0 double and triple PTC mutant viruses were serially passaged to P3 in HEK293T cells with or without PB8 cotransfection, and eGFP expression was monitored throughout passaging. Scale bar: 200 μm. F The eGFP was excised from the plasmid SVV-eGFP-3D-Y77*/Y343*/Y444*, and the resulting plasmid was cotransfected with PB8. The virus was then serially passaged up to P3. Wild-type HEK293T cells were infected with the P1 and P3 virus, and the CPE was observed at 48 h post-infection. Scale bar: 200 μm. G P1 and P3 SVV-PTC were subsequently sequenced. H Comparisons of the morphologies of the PTC viruses and the wild-type SVV by transmission electron microscopy. Scale bar: 100 nm.

Given the substantial impact of different PTC sites on viral genetic stability, we introduced multiple PTC sites into the 3D gene to reduce the reversion mutation frequency. Based on the results of the preliminary screening, three relatively stable sites (Y77, Y343, Y444) were selected to construct double and triple mutants, namely SVV-eGFP-3D-Y77*/Y343*, SVV-eGFP-3D-Y77*/Y444*, SVV-eGFP-3D-Y343*/Y444*, and SVV-eGFP-3D-Y77*/Y343*/Y444*. P0 PTC viruses were successfully rescued by cotransfecting the above mutant plasmids with PB8 into cells. Obvious CPE and extensive eGFP fluorescent expression were observed for both double and triple mutant viruses after co-transfection (Fig. 3C). The qPCR results showed that, compared with the virus containing two PTC sites, the titer of the triple mutant virus was not markedly reduced (Fig. 3D). Therefore, the PB8 vector we constructed can be used for subsequent PTC-virus production.

Subsequent serial passage experiments confirmed that multiple PTC sites significantly enhanced viral genetic stability: no eGFP expression was detected in wild-type HEK293T cells at P3 (Fig. 3E). Based on the above results, SVV-3D-Y77*/Y343*/Y444* (designated as SVV-PTC-3Y*) was selected as the candidate vaccine strain and the eGFP reporter gene was knocked out. When SVV-PTC-3Y* was passaged to P3 and then used to infect wild-type HEK293T cells, no CPE was observed (Fig. 3F; Supplementary Fig. 10A). DNA sequencing verification of the P3 virus confirmed the absence of reversion mutations at the PTC sites (Fig. 3G; Supplementary Fig. 10B). These results demonstrated that SVV-PTC-3Y* possessed high genetic stability under in vitro culture conditions. The SVV-PTC-3Y* was then purified, and transmission electron microscopy analysis showed no morphological differences between SVV-PTC and SVV-WT (Fig. 3H).

SVV-PTC-3Y* elicits early high-titer neutralizing antibodies in mice

We evaluated the potential of SVV-PTC-3Y* as a vaccine candidate in mice (Fig. 4A). Virus neutralization assay results demonstrated that SVV-PTC-3Y* induced high levels of neutralizing antibodies in mice. Neutralizing antibody titers in the IP and IM groups did not differ significantly from those in the inactivated vaccine group at 42 dpi, reaching approximately 1:4096 (P > 0.9999, P = 0.4146). Notably, at 7 dpi, the neutralizing antibody titers in the SVV-PTC-3Y* IP and IM groups reached ~1:26 and ~1:104, respectively, whereas no neutralizing antibody was detected in the inactivated vaccine group. On day 21 after the first immunization, the neutralizing antibody titers of the SVV-PTC-3Y* IP and IM groups reached ~1:512, which were significantly higher than those of the inactivated vaccine group (~1:128) (P = 0.0023; Fig. 4B).

Fig. 4. The SVV-PTC immunization induced an efficient antibody response in mice.

A Schematic of BALB/c mice experiment plan, created with BioGDP.com. B Detection of serum neutralizing antibody titers against SVV LNSY01-2017 strain at different time points. C–E Mouse serum IgG, IgG1, and IgG2a antibody responses to the SVV at different time points by indirect ELISA. Data in (B–E) are presented as mean ± SD (n = 5). One-way analysis of variance (ANOVA) with Newman-Keuls multiple comparisons test was performed. Exact p values are shown.

Detection of SVV-specific IgG antibodies in serum revealed that all immunized groups exhibited high IgG antibody levels except the PBS control group (Fig. 4C). At 42 dpi, the IgG antibody levels in the IP and IM groups were significantly higher than those in the inactivated vaccine group (P = 0.0481, P < 0.0001). Notably, at 14 days following the primary immunization, the SVV-PTC-3Y* IP and IM groups also exhibited significantly higher IgG responses compared with the inactivated vaccine group; the average OD₄₅₀ values were 1.056 in the IP group, 1.336 in the IM group, and 0.284 in the PBS control group (P < 0.0001).

The detection of SVV-specific IgG1 and IgG2a subtype antibodies in serum revealed that both the SVV-PTC-3Y* and inactivated vaccine groups had significantly higher antibody levels than the PBS control group (P < 0.0001) (Fig. 4D, E). The inactivated vaccine group had considerably higher IgG1 antibody levels than IgG2a, indicating that it induced a Th2-biased humoral immune response. In contrast, the SVV-PTC-3Y* groups had substantially higher IgG2a antibody levels than IgG1, particularly during the early stage of immunity, and their IgG2a antibody levels were significantly higher than those in the inactivated vaccine group (P < 0.0001), suggesting that SVV-PTC-3Y* preferentially induces a Th1-type immune response.

In summary, both SVV-PTC-3Y* and the inactivated vaccine can elicit robust humoral immune responses in mice. Compared with the inactivated vaccine, SVV-PTC-3Y* can induce high titers of neutralizing antibodies after the first immunization, showing an advantage of faster immune response. Among the different immunization routes, both intraperitoneal injection and intramuscular injection are superior to intranasal vaccination.

SVV-PTC-3Y* elicits effective Th1-biased cellular immune responses in mice

To quantify the intensity of cellular immunity, we detected the proliferation stimulation index (SI) of splenic lymphocytes on day 42 post immunization. The results showed that the SI values in the inactivated vaccine group (P = 0.0018), SVV-PTC-3Y* IP group (P < 0.0001), SVV-PTC-3Y* IM group (P = 0.0048), and SVV-PTC-3Y* IN group (P = 0.0312) were significantly higher than those in the PBS control group. Furthermore, the SI values in the SVV-PTC-3Y* IP group were significantly higher than those in the inactivated vaccine group (P < 0.0017; Fig. 5A). To investigate the effects of SVV-PTC-3Y* on SVV-specific cellular immune responses, mouse splenic lymphocytes were stimulated with inactivated SVV (0.1 MOI), and the proportions of CD3+ , CD4α+ , and CD8α+ T lymphocytes were quantified by flow cytometry at 42 dpi (Fig. 5B, C; Supplementary Fig. 11). The percentages of CD4α+ T cells within CD3+ T cells in both the inactivated vaccine group and all SVV-PTC-3Y* immunized groups were significantly higher than that in the PBS control group (P < 0.0001). Compared with the inactivated vaccine group, the SVV-PTC-3Y* IP group (P = 0.0148) and IN group (P = 0.0042) showed significantly increased percentages of CD4α+ T cells, whereas no significant difference was observed in the SVV-PTC-3Y* IM group (P = 0.9998). Regarding CD8α+ T cells, the percentage within CD3+ T cells in the inactivated vaccine group was not significantly different from that in the PBS group (P = 0.9177). In contrast, the SVV-PTC-3Y* IP group (P = 0.0077) and IM group (P < 0.0001) both exhibited significantly higher percentages of CD8α+ T cells compared with the PBS group.

Fig. 5. The SVV-PTC immunization drives a Th1-biased cellular immune response in mice.

A Proliferative responses of splenic lymphocytes in mice at 42 dpi. B, C The proportions of CD4α+ and CD8α+ cells within CD3+ T lymphocytes were detected by flow cytometry. D–H Detection of cytokine levels in lymphocyte-stimulated supernatants of SVV-PTC-immunized mice at 42 dpi. Concentrations of IL-2 (D), IL-4 (E), IL-10 (F), IFN-γ (G), and TNF-α (H) were measured by commercial ELISA kits. Data in (A-H) are presented as mean ± SD (n = 5). One-way ANOVA with Newman–Keuls test was performed. Exact p values are shown.

To evaluate the characteristics of the T cell immune response induced by the vaccine, we measured the secretion levels of key cytokines in mouse splenic lymphocytes by double-antibody sandwich ELISA, including IL-2, IFN-γ, TNF-α, IL-4, and IL-10 (Fig. 5D–H). Among the SVV‑PTC groups, the IM group showed a more considerable increase in cytokine levels compared with the other groups; thus, we focused our analysis on the SVV-PTC-3Y* IM group. The IM group elicited a mixed cytokine response, characterized by elevated levels of Th1‑type cytokines (IL‑2, IFN‑γ, TNF‑α), the Th2‑type cytokine IL‑4, and IL‑10. Notably, the induction of Th1‑type cytokines was predominant, and the secretion level of IFN‑γ (~620 pg/mL) was markedly higher than that of IL‑4 (~94 pg/mL), indicating a Th1‑biased immune response. This was consistent with the humoral immune results showing higher IgG2a antibody levels than IgG1. Furthermore, the secretion level of IL‑10 was significantly increased (P < 0.0001; Fig. 5F), which may contribute to immunoregulation and prevent excessive inflammation. These results demonstrated that the SVV-PTC-3Y* induced a balanced cellular immune response characterized by Th1 dominance and moderate immunoregulation. In contrast, the inactivated vaccine group showed a mild Th2‑skewed T cell response, as IFN‑γ (~164 pg/mL) was only slightly higher than IL‑4 ( ~142 pg/mL), with no obvious Th1 polarization. In summary, SVV-PTC-3Y* elicits effective Th1-biased cellular immune responses in mice.

SVV-PTC-3Y* exhibits no horizontal transmission and favorable safety in swine

To evaluate the in vivo safety of SVV-PTC, blood samples were collected at 1, 3, 5, and 7 days post‑immunization (dpi) with SVV-PTC, followed by amplification of the 3D gene of SVV-PTC virus in the blood. SVV 3D gene was successfully amplified from the blood samples at 1 and 3 dpi, and sequencing results confirmed that no reversion mutations occurred in SVV-PTC (Fig. 6A, B; Supplementary Fig. 12, 13A). In addition, after immunization with SVV-PTC, continuous observation for 14 days revealed no blister-like pathological changes (Supplementary Fig. 13B).

Fig. 6. Safety and horizontal transmission characteristics of SVV-PTC in swine.

A, B PCR amplification and sequencing of the 3D gene of SVV-PTC in blood samples at 1, 3, 5, and 7 dpi. C–H Horizontal transmission of wild-type virus and SVV-PTC among pigs (n = 3), as reflected by viral copy numbers in nasal swabs, pharyngeal swabs, and fecal swabs. Data plotted for individual pig and overlaid with mean ± SD. The horizontal dashed line shows the baseline viral load prior to challenge. I–K Dynamic changes in viral copy numbers of wild-type virus and SVV-PTC in nasal swabs, pharyngeal swabs, and anal swabs from infected pigs (n = 3) at different time points post-infection. Error bars denote SD. Significance was determined by one-way ANOVA. Exact p values are shown.

To evaluate the horizontal transmission characteristics of SVV-PTC, unvaccinated pigs were cohoused with pigs inoculated with either SVV-PTC or SVV-WT. In both groups, no significant differences in nasal swab viral copy numbers were observed between the cohoused unvaccinated pigs and their baseline levels at 0 days post-contact (dpc) (Fig. 6C, F). Analysis of pharyngeal swab viral dynamics revealed that in the SVV-WT group, viral loads in inoculated pigs gradually decreased from 0 to 4 dpc and then increased thereafter. Meanwhile, viral loads in cohoused unvaccinated pigs showed a sustained increase from 0 dpc and became significantly higher than those in inoculated pigs by 4 dpc (P < 0.0001). By 9 dpc, no significant difference in pharyngeal viral loads was observed between inoculated and cohoused pigs (P = 0.0784) (Fig. 6D). These dynamic changes indicate that SVV-WT can be transmitted horizontally through pharyngeal secretions. In contrast, in the SVV-PTC group, although inoculated pigs showed significantly higher pharyngeal viral copy numbers than cohoused pigs, the viral loads in cohoused pigs remained not significantly different from baseline (0 dpc) throughout the observation period (Fig. 6G). Furthermore, at 9 dpc, substantial virus shedding was detected in fecal swabs of pigs cohoused with SVV-WT, whereas no such shedding was observed in pigs cohoused with SVV-PTC (Fig. 6E, H). These results demonstrate that, unlike the SVV-WT cohabitation group, none of the contact pigs in the SVV-PTC cohabitation group exhibited a significant increase in viral copy numbers, confirming that SVV-PTC lacks the ability for horizontal transmission in the population. In addition, comparison of viral loads in nasal, pharyngeal, and anal swabs from pigs intramuscularly inoculated with SVV-WT or SVV-PTC showed that the SVV-PTC group maintained low viral loads throughout the observation period (Fig. 6I–K), further supporting its replication deficiency in vivo and favorable safety profile. Collectively, these data demonstrate that SVV-PTC exhibits good safety in pigs and does not possess horizontal transmission characteristics.

SVV-PTC-3Y* rapidly induces robust humoral immune responses in swine

We established a rational experimental strategy to assess protective immune efficacy in pigs (Fig. 7A). Serum samples were collected at 0, 14, 28, and 42 dpi to determine the neutralizing antibody titers (Fig. 7B). The results showed that the neutralizing antibody titers in the SVV-PTC group increased significantly at 14 dpi (P < 0.0001), with a mean titer of ~1:74., which was significantly higher than that in the inactivated vaccine group (~1:11) (P < 0.0001). The titers in both the SVV-PTC and inactivated vaccine groups rose markedly after the booster immunization, reaching a mean of ~1:512 in the SVV-PTC group and ~1:256. These findings demonstrated that SVV-PTC elicited a rapid and potent humoral immune response, with significantly higher neutralizing antibody levels than the inactivated vaccine as early as 14 days after primary immunization (P < 0.0001).

Fig. 7. The SVV-PTC vaccine induces robust immune responses in swine.

A Schematic illustration of the experimental design for swine immunization and challenge protection, created with BioGDP.com. B Serum neutralizing antibody titers against the SVV LNSY01-2017 strain in swine were determined at 0, 14, 28, and 42 dpi. C SVV-specific IgG antibody levels at different time points were measured by indirect ELISA using purified SVV, with absorbance determined at 450 nm (OD₄₅₀). D The proliferation of PBMCs from each group was detected following stimulation with UV-inactivated SVV. E, F The proportions of CD4α+ and CD8α+ cells within CD3+ T lymphocytes were detected by flow cytometry after stimulation with inactivated SVV. G–K After stimulation of PBMCs with inactivated SVV, cell culture supernatants were harvested for detection of the expression levels of cytokines IL‑2, IL‑4, IL‑10, IFN‑γ, and TNF‑α. Data are presented as mean ± SD (n = 5). Significance was determined by one-way ANOVA. Exact p values are shown.

Meanwhile, we determined the capacity of SVV-PTC to induce SVV-specific IgG antibody titers in swine (Fig. 7C). Interestingly, both SVV-PTC and the inactivated vaccine rapidly elicited high titers of SVV-specific IgG antibodies at 14 dpi, which were sustained thereafter. We speculated that non-neutralizing IgG might dominate the total IgG produced at the early stage of immunization, while neutralizing IgG became the major component after the booster immunization, which could be attributed to the immunological and biological characteristics of swine. In addition, statistical analysis revealed no significant difference in the SVV-specific IgG antibody titers induced by SVV-PTC and the inactivated vaccine group at 14 and 28 dpi (P = 0.5127, P = 0.0542), whereas the titers induced by SVV-PTC were significantly higher than those by the inactivated vaccine at 42 dpi (P = 0.0313).

SVV-PTC-3Y* induces a robust Th1-biased cellular immune response in swine

The determination of lymphocyte proliferative response is crucial for evaluating cellular immune responses. PBMCs from each group were stimulated with inactivated SVV, followed by measurement of OD₄₅₀ values and calculation of the stimulation index (SI) (Fig. 7D). The SI values of both the SVV-PTC vaccine group and the inactivated vaccine group were significantly higher than that of the PBS control group (P < 0.0001), with no significant difference observed between these two groups (P = 0.1851). These findings indicated that both the SVV-PTC vaccine and the inactivated vaccine could effectively enhance PBMCs proliferation through specific stimulation.

The percentages of CD3+ , CD4α+ , and CD8α+ T lymphocytes in PBMCs were determined by flow cytometry (Fig. 7E, F). The results showed that the percentages of CD4α+ T cells within CD3+ T cells in both the inactivated vaccine group (P = 0.0002) and the SVV-PTC group (P < 0.0001) were significantly higher than that in the PBS control group. Moreover, the percentage in the SVV-PTC group was significantly higher than that in the inactivated vaccine group (P = 0.0046). Similarly, the percentages of CD8α+ T cells within CD3+ T cells in both the inactivated vaccine group (P = 0.0104) and the SVV-PTC group (P = 0.0004) were significantly higher than that in the PBS control group, and the SVV-PTC group again showed a significantly higher percentage compared with the inactivated vaccine group (P = 0.0185).

The secretion levels of cytokines including IL-2, IL-4, IL-10, IFN-γ, and TNF-α in PBMCs at 42 dpi were quantified using commercial double-antibody sandwich ELISA kits (Fig. 7G–K). Compared with the PBS control group, SVV-PTC induced significantly elevated secretion levels of IL-2 ( ~842 pg/mL) (P = 0.0004), IFN‑γ (~1532 pg/mL) (P < 0.0001), and TNF‑α (~452 pg/mL) (P < 0.0001), which are representative Th1‑type cytokines. Although the secretion level of IL‑4 (5546 pg/mL) was also increased, the immune response was dominated by Th1‑type cytokines, indicating a Th1‑biased cellular immune response. In contrast, the inactivated vaccine induced a remarkably high secretion level of IL‑4 ( ~9106 pg/mL), accompanied by moderate increases in IL‑2 ( ~281 pg/mL), IFN‑γ (~145 pg/mL), and TNF‑α (~222 pg/mL), suggesting a Th2‑skewed immune profile. Regarding the IFN‑γ/IL‑4 ratio, the ratio in the SVV‑PTC group was 18‑fold higher than that in the inactivated vaccine group and 30‑fold higher than that in the PBS control group. Although the absolute concentration of IL‑4 in the SVV‑PTC group was relatively high, the relative IFN‑γ/IL‑4 ratio and the fold changes clearly demonstrate that SVV‑PTC induced a distinct Th1‑biased immune response. In addition, the secretion levels of IL‑10 were significantly elevated in both vaccine groups (P = 0.0009, P < 0.0001), especially in SVV-PTC, which may contribute to immunoregulation and prevent excessive inflammatory responses. Collectively, these results indicated that SVV-PTC, as a live but replication-incompetent vaccine, could induce a robust Th1‑biased cellular immunity compared with the inactivated vaccine.

SVV-PTC-3Y* confers complete protection against SVV challenge in swine

In this study, the in vivo protective efficacy of the SVV-PTC vaccine was evaluated using an established swine challenge model¹³. Pigs were continuously monitored for 14 days, with disease severity assessed by vesicular lesion sampling, imaging documentation, and clinical scoring after challenge. Clinical symptom scoring criteria followed³⁰: 0 points (no visible vesicular lesions), 1 point (1 mild lesion), 2 points (2 mild lesions), 3 points (2 severe lesions), and 4 points ( ≥ 3 severe lesions). No vesicular lesions were observed in any of the experimental pigs from the SVV-PTC vaccine group and the inactivated vaccine group (Fig. 8A, B). In contrast, all pigs in the PBS control group developed typical vesicular lesions at 1-3 days post-challenge, with disease severity peaking at 4-9 days post-challenge and subsequent symptom remission (Fig. 8C, D). Furthermore, duplex PCR assays targeting SVV and FMDV were performed on the collected vesicular samples. All tested samples yielded specific SVV amplicons, whereas no FMDV-specific bands were detected (Fig. 8E; Supplementary Fig. 14).

Fig. 8. Evaluation of clinical vesicular lesion scores and identification in swine challenged with SVV LNSY01–2017.

A–C Clinical vesicular lesions were scored and statistically analyzed in each group after challenge. D Photographs of clinical symptoms in individual pigs from the PBS control group after challenge. E Detection of SVV and FMDV in swine vesicular lesions using specific duplex PCR.

Discussion

Despite the availability of commercial inactivated vaccine against SVV, the vaccine inherently suffers from limitations such as relatively weak immunogenicity and reliance on adjuvants. Thus, developing live replication-controllable vaccines holds significant importance. Based on the sup-tRNA technology, our study successfully constructed a safe and effective replication-defective virus, SVV-PTC. Both GCE technology and sup-tRNA can be used to generate viruses containing PTCs, but the mechanisms are fundamentally different: GCE technology achieves readthrough by incorporating UAAs at PTC sites, while sup-tRNA mediates the insertion of natural amino acids^20,22.

Genetic code expansion platforms are widely used in virological studies, enabling the functionalization of virus-like particles, the optimization of viral vectors for gene therapy, and the investigation of essential viral properties^31,32. Despite their versatility, using GCE to engineer viruses with premature termination codons often encounters challenges related to the incorporation of unnatural amino acids (UAAs). Specifically, targeting UAAs to specific PTC locations can lead to issues such as failed viral rescue, reduced production yields, or compromised structural stability. Adding to these technical difficulties is the lack of reliable, predictive criteria for identifying the most effective PTC insertion sites^20,22. Conversely, sup-tRNAs facilitate the incorporation of the native amino acid at the targeted PTC locus, thereby preserving wild-type viral architecture and functionality. From a methodological standpoint, the propagation of PTC-harboring viruses via this system relies solely on the establishment of sup-tRNA-overexpressing cell lines; this circumvents the burdensome requirements of exogenous aminoacyl-tRNA synthetase co-expression and continuous UAA supplementation. Additionally, because sup-tRNAs exhibit negligible readthrough at canonical termination codons, they avert deleterious disruptions to host cellular homeostasis²⁹. However, the cell line we established could not efficiently support SVV-PTC replication, and virus production relied mainly on cotransfection, which limits production efficiency. Future work will employ a transposon system to generate a cell line with high sup-tRNAy expression. Consequently, relative to traditional GCE platforms, the sup-tRNA strategy represents a significantly more streamlined and efficacious paradigm for the engineering and rescue of PTC viruses.

For the development of SVV-PTC, ensuring its safety and stability is a core prerequisite. During viral passaging, it was found that a single PTC site is prone to mutation into a sense codon, and the stability of different PTC sites varies significantly, requiring extensive mutation screening to obtain stable sites. Previous studies have confirmed that increasing PTC sites can significantly improve the stability of PTC-viruses²⁰. However, it should be noted that an increase in the number of PTC sites is accompanied by a decrease in viral rescue efficiency. Therefore, there is an urgent need to construct cell lines with high-efficiency sup-tRNA expression³³. Currently reported strategies to improve PTC virus rescue efficiency include increasing the copy number of sup-tRNA, using strong promoters to drive expression, and knocking down or engineering release factors^34–36.

Interestingly, although both SVV and FMDV belong to the Picornaviridae, SVV-PTC-3Y* showed superior genetic stability compared to a previously reported PTC-containing FMDV, which remained unstable even with two amber codons³³. This discrepancy may be explained by several factors. First, FMDV has higher genetic diversity due to multiple serotypes, a broad host range, and high recombination rates^37,38, whereas SVV has only one serotype and a narrow host range (mainly pigs), limiting its evolutionary plasticity. Second, we screened 21 tyrosine residues in the 3D gene and combined three stable sites (Y77, Y343, Y444) to raise the genetic barrier against reversion. Third, sup-tRNAy system enables efficient readthrough without exogenous components, reducing selective pressure for reversion. In contrast, the GCE system used in the previous FMDV study has lower rescue efficiency and imposes stronger survival pressure that may drive reversion. Collectively, these features account for the favorable genetic stability of SVV-PTC-3Y*.

As a novel type of replication-defective live vaccine, SVV-PTC occupies a unique position between conventional inactivated vaccines and live attenuated vaccines. Because it retains the complete integrity of both the viral capsid and nucleic acid payload, the vaccine preserves an initial infectious capacity equivalent to that of wild-type strains. However, downstream viral proliferation is strictly contingent upon the presence of sup-tRNA. As a result, vaccination initiates efficient host cell entry followed by an abortive replication cycle in vivo, thereby striking an optimal balance between robust immunogenicity and stringent safety²⁰. In this study, we found that SVV-PTC exhibits distinct immunological characteristics, including a rapid increase in neutralizing antibody levels, a Th1-biased immune response, and a higher intensity of cellular immunity. The rapid elevation of neutralizing antibody levels induced by SVV-PTC may be attributed to its retention of the intact capsid structure and surface antigens of wild-type SVV, which can be directly recognized and rapidly activated by B cells without relying on adjuvants. Meanwhile, since we selected the PTC site in the 3D protein, SVV-PTC can complete a single round of infection in host cells. During the replication process, viral nucleic acid is released and sensed by intracellular pattern recognition receptors (PRRs), such as RIG-I and MDA5, to trigger innate immunity³⁹. This activation initiates the type I interferon signaling pathway, which rapidly recruits and activates follicular helper T cells (Tfh), thereby promoting B cell proliferation, differentiation, and antibody affinity maturation⁴⁰. Relevant studies have shown that a single immunization with the influenza virus mRNA-LNP vaccine can induce a protective antibody response within 10–14 dpi⁴¹, which shares a similar mechanism with SVV-PTC. AZD1222 (ChAdOx1 nCoV-19) is a replication-deficient simian adenovirus–vectored vaccine. Following vaccination, PBMCs isolated from recipients were stimulated with SARS-CoV-2 spike peptide pools, revealing a significant increase in the proportion of CD4+ T cells, along with markedly elevated levels of Th1-type cytokines (IL-2, IFN-γ, and TNF-α). These findings demonstrate that the vaccine induces a Th1-biased T cell response⁴², which is consistent with the T cell immunity profile elicited by the SVV-PTC vaccine. Furthermore, in a mouse model, antigen-specific IgG2a antibody titers were significantly higher than IgG1, providing additional evidence for the presence of Th1-type helper activity. However, we did not evaluate vaccine‑induced the long‑term durability of protection in this study. PTC-harboring viruses are inherently avirulent in vivo. Moreover, during co-infection scenarios, recombination between PTC mutants and wild-type strains actively attenuates the infectivity of the wild-type virus in vivo and in vitro²⁰. This study also confirmed that pigs-inoculated with SVV-PTC remained completely asymptomatic and exhibited no evidence of horizontal viral transmission. Collectively, these attributes highlight the substantial biosafety advantages and translational promise of the sup-tRNA-dependent SVV-PTC platform.

Notably, immune responses differed between mouse and pig models. In pigs, IgG levels peaked at day 14 and remained comparable to those of the inactivated vaccine group at days 14 and 28. In mice, however, SVV‑PTC induced significantly higher IgG levels early after immunization, with the peak occurring after the booster. Moreover, IL‑4 levels were markedly higher in pigs than in mice. These differences may arise from inherent interspecies variations in the composition and response patterns of the immune system, including differences in antigen‑presenting cell types and lymphocyte subset distribution, which may drive faster inflammatory responses in pigs, facilitating rapid antibody production. Despite these differences, SVV‑PTC consistently induced rapid neutralizing antibodies, enhanced cellular immunity, and a Th1‑biased response in both species.

In summary, we delineate an advanced methodological framework for engineering infectious virions by sup-tRNA technology. By enabling stringent control over the viral replication, our study establishes a robust empirical foundation for the design of next-generation SVV prophylactics.

Methods

Ethics statement

All animal experiments complied with ethical regulations for animal testing and research and were conducted in accordance with protocols approved by the Laboratory Animal Management Committee of Huazhong Agricultural University and the Scientific Ethics Committee of Huazhong Agricultural University (Hubei, China) (Ethics numbers: HZAUMO-2026-0034 for mice, YF-XJ-2025-023 for swine). All animal experiments were performed in compliance with the 2016 Chinese National Standard for laboratory animals and other relevant regulations of the Animal Welfare Act.

Viruses and cells

In this study, SVV (HB-CH-2016 strain) was isolated and identified by our laboratory from vesicular lesion tissue samples of piglets with Porcine Idiopathic Vesicular Disease (PIVD)⁴³. The SVV LNSY01-2017 (MH064435) challenge model, preserved and established in our laboratory, was utilized to perform the viral challenge. The commercial inactivated SVV vaccine was selected as the positive control, provided by Wuhan Keqian Biology Co., Ltd. Human embryonic kidney 293T cells (HEK293T, ATCC CRL-11268) and baby hamster kidney cells (BHK-21, ATCC CCL-10) were both cultured in Dulbecco’s Modified Eagle Medium (DMEM, Sigma, D6429) supplemented with 10% fetal bovine serum (FBS, Cell-Box, AUS-02S-02-S).

Plasmid construction

The SVV plasmid rescue systems (pSKII-SVV-HB-eGFP and pSKII-CMV-SVV-HB) were preserved in our laboratory. Mutant plasmids (pSKII-SVV-HB-eGFP-L-TAG, pSKII-SVV-HB-eGFP-VP4-TAG, pSKII-SVV-HB-eGFP-3C-TAG, and pSKII-SVV-HB-eGFP-3D-TAG) containing amber codons within the open reading frame of the wild-type plasmid pSKII-SVV-HB-eGFP were generated by site-directed mutagenesis (Vazyme, Nanjing, China). The plasmid pCAGGS-HA-3D was also preserved in our laboratory. An amber codon was introduced into the 3D sequence to construct the mutant plasmid pCAGGS-HA-3D-TAG. All mutant plasmids were verified by gene sequencing (Sangon Biotech, Wuhan).

The pyrrolysyl-tRNA synthetase/tRNA_CUA pair (PylRS/tRNA_CUA) for site-specific incorporation of the orthogonal unnatural amino acid N(e)-Boc-L-lysine (BOCK) was used as previously reported⁴⁴. This plasmid system was kindly provided by Professor Yanjun Liu from the National Key Laboratory of Crop Genetic Improvement, Huazhong Agricultural University. Plasmids pLKO.1-2×sup-tRNAy and pLVX-mCherryY43* were kindly provided by Professor Wenchun Fan from the Institute of Life Sciences, Zhejiang University.

The PB transposon system was purchased from VectorBuilder (Guangzhou, China). The mCherryY43* gene driven by the CMV promoter was cloned into the PB transposon vector to obtain the pPB-CMV-mCherryY43*-IRSE-Puro plasmid. Two copies of the U6-tRNAy sequence were amplified from plasmid pLKO.1-2×sup-tRNAy, and the 7SK promoter was amplified from pCMV-Tem3-8tRNA-chFSYRS-I267A-L349Y-final. Subsequently, two copies of the 7SK-tRNAy sequence were obtained by overlap extension PCR. The pPB-CMV-mCherryY43*-IRSE-Puro plasmid was digested with BamHI, and then the 2×U6-tRNAy and 2×7SK-tRNAy sequences were sequentially cloned into the vector using homologous recombination enzyme (Vazyme, Nanjing, China) to finally obtain the plasmid containing 8 copies of tRNAy, namely pPB-CMV-mCherryY43*-IRSE-Puro-[2×U6-tRNAy-2×7SK-tRNAy]2 (abbreviated as PB8), which was used for subsequent rescue of PTC viruses. All plasmids used for transfection were amplified using the Plasmid Mini Kit (Omega) according to the manufacturer’s instructions.

Generation of replication-incompetent SVV harboring amber codon(s) in their genome

For the preparation of SVV-PTC, HEK293T cells were seeded into 6-well plates at a density of 2 × 10⁵ cells per well 24 h before transfection. Subsequently, 1.5 μg each of the mutant plasmids from the virus rescue system and PB8 plasmid were mixed and transfected into the cells using JetPRIME transfection reagent (Polyplus Transfection, PT-114-15) according to the manufacturer’s instructions. After 4–6 h, the medium was replaced with DMEM supplemented with 2% FBS. The cells were further incubated at 37 °C in 5% CO₂ until more than 90% CPE was observed. The supernatant containing the virus was collected and centrifuged at 1000 × g for 10 min to remove residual cells.

For the generation of SVV-PTC using GCE technology, an almost identical procedure was performed with the following modifications: the PB8 plasmid was replaced with pCMV-Tem3-8tRNA-chFSYRS-I267A-L349Y-final, and the medium was supplemented with 1 mM N(e)-Boc-L-lysine (BOCK) after transfection. For subsequent animal experiments, the obtained SVV-PTC viruses were concentrated using an ultra-high-speed refrigerated centrifuge.

Genetic stability evaluation of SVV-PTC viruses in vitro

HEK293T cells were transfected with the PB8 plasmid when the cell confluency reached 70–80%. At 24 h post-transfection, the cells were infected with SVV-PTC at a multiplicity of infection (MOI) of 0.1. When more than 90% CPE was observed, the supernatant was collected for the next round of infection, and this procedure was repeated more than 3 times. A parallel control experiment was conducted by infecting HEK293T cells not transfected with the PB8 plasmid with SVV-PTC. The dependence of the virus on tRNAy was determined by observing the occurrence of CPE and the expression of the viral fluorescent protein eGFP.

After each passage, total cellular RNA was extracted using TRIzol reagent. Subsequently, cDNA was synthesized using reverse transcriptase (Yeasen Biotechnology) according to the manufacturer’s instructions. PCR amplification was performed with Phanta Max Super-Fidelity DNA Polymerase (Vazyme, China) in a 50 μL reaction system containing specific primers for the SVV 3D fragment. The obtained PCR products were subjected to gene sequencing to analyze whether mutations occurred during viral passage.

Purification and morphological observation of wild-type and replication incompetent Seneca Valley virus particles

Wild-type SVV was propagated by infection of BHK-21 cells at an MOI of 0.01, and SVV-PTC was prepared following the aforementioned protocol. The cell culture supernatants were collected and clarified by centrifugation at 8000 rpm for 20 min at 4 °C. The clarified supernatants were then concentrated by ultracentrifugation at 28000 rpm for 3 h at 4 °C using an SW32 rotor. Subsequently, the pelleted viruses were resuspended in 1 mL of PBS, and subjected to a 20–65% sucrose gradient ultracentrifugation at 38,000 × rpm for 4 h at 4 °C with an SW41 rotor. The virus-containing bands were harvested, diluted with PBS, and the sucrose was removed by an additional ultracentrifugation at 38,000 × rpm for 4 h at 4 °C using an SW41 rotor. The resulting virus pellets were resuspended in 500 μl of PBS. The purified viruses were either used immediately or aliquoted, flash-frozen, and stored at −80 °C until subsequent use. The morphology of the purified viruses was observed under a transmission electron microscope (TEM) by negative staining.

Western blotting

At 24 h post-transfection, HEK293T cells were lysed with NP40 lysis buffer on ice for 30 min. Cell debris was removed by centrifugation, and the protein concentration was quantified using the BCA assay. The protein samples were mixed with 5×loading buffer and boiled at 95 °C for 10 min in a dry bath incubator. Proteins were separated by 10% SDS-PAGE and then transferred onto PVDF membranes via wet transfer.

The membranes were blocked with PBST buffer containing 5% nonfat milk at room temperature for 2 h, followed by incubation with primary antibodies at 4 °C overnight. The following primary antibodies were diluted in antibody dilution buffer: anti-HA antibody (1:5000, 66006-2-Ig, Proteintech), α-Tubulin antibody (1:5000, 66031-1-Ig, Proteintech), and GAPDH antibody (1:5000, 60004-1-Ig, Proteintech). After washing the membranes three times with PBST, they were incubated with HRP-conjugated goat anti-mouse IgG (1:5000) at room temperature for 1 h. Following extensive washing, protein bands were visualized using an enhanced chemiluminescent (ECL) detection kit (Uelandy).

Mice immunity experiment schedule

Four-week-old female SPF BALB/c mice were purchased from the Experimental Animal Research Center of Huazhong Agricultural University. Mice were housed in individually ventilated cages (IVC) with sterile wood chip bedding, maintained at 22 ± 2 °C with a 12/12 h light/dark cycle, and provided with autoclaved food and water ad libitum. All mice were acclimatized to the facility before the start of experiments. For the immunogenicity study, 25 mice were randomly divided into 5 groups (n = 5 per group) by body weight stratification: SVV‑PTC (intraperitoneal, IP) group, SVV‑PTC (intramuscular, IM) group, SVV‑PTC (intranasal, IN) group, SVV inactivated vaccine group, and PBS control group. Blinding was implemented in this study: animal allocation was performed by an investigator who was not involved in subsequent experiments, and the personnel performing immunizations, blood collection, and data detection were unaware of the group assignments. Each mouse was immunized with 10⁹ VP of SVV‑PTC by the indicated route. The commercial SVV inactivated vaccine was administered intramuscularly at 100 μL per mouse. A booster immunization with the same dose was given on day 21 post‑primary immunization. Serum samples were collected from the orbital vein before immunization (day 0) and on days 7, 14, 21, 28, 35, and 42 dpi. On day 42 post‑immunization, mice were euthanized by carbon dioxide (CO₂) inhalation in a gradual fill chamber (20% chamber volume/min), followed by cervical dislocation as a secondary method to ensure death. Spleens were then harvested to isolate splenic lymphocytes for cellular immune assessment. All procedures were performed at the same time of day, the order of treatments was randomized, and cages were placed on the same rack (Supplementary Fig. 15).

Swine immunity and challenge experiment schedule

Thirteen‑week‑old SVV‑negative Landrace gilts (50–60 kg) were purchased from Guangxi Yangxiang Co., Ltd. Pigs were housed in individual pens in a biosafety level‑2 (BSL‑2) facility with controlled temperature (22 ± 3 °C). They were fed a commercial diet twice daily and had free access to water. All pigs were acclimatized before the study.

To evaluate the immunogenicity and challenge protection of the SVV‑PTC vaccine in pigs, 15 pigs were randomly allocated into three groups (n = 5 per group) by body weight stratification: SVV‑PTC group, inactivated vaccine group, and PBS control group. Blinding was implemented in this study: animal allocation was performed by an investigator not involved in subsequent experiments, and the personnel performing immunizations, blood collection, and data detection were unaware of the group assignments. Pigs were immunized intramuscularly on day 0 and day 21 with 2 mL of SVV‑PTC (10¹⁰ VP/mL), 2 mL of commercial SVV inactivated vaccine, or 2 mL of PBS, respectively. For humoral immunity assessment, blood samples were collected from the anterior vena cava at 0, 14, 28, and 42 dpi. At 42 dpi, PBMCs were isolated for cellular immune evaluation. At 43 dpi, all pigs were challenged: each pig received 6 mL of homologous virulent strain SVV LNSY01‑2017 (10¹⁰ TCID₅₀/mL) by combined intranasal spray (4 mL) and intramuscular injection (2 mL). Clinical symptoms were monitored and recorded for 14 days post‑challenge. Tissue samples from pigs with typical SVV vesicular lesions were collected for duplex PCR detection of SVV and FMDV. At the conclusion of the experiment, all pigs were humanely euthanized by intravenous injection of an overdose of sodium pentobarbital ( ≥ 90 mg/kg) via the ear vein, followed by exsanguination, in accordance with the Chinese National Standard GB/T 39760‑2021 “Laboratory animal—Guidelines for euthanasia”. Procedures were all performed at a fixed time of day, with the treatment order randomized. In addition, pigs were housed in individual pens under identical conditions (Supplementary Fig. 15).

Horizontal transmission assay of SVV-PTC

For assessing the horizontal transmission characteristics of SVV‑PTC, pigs (n = 3 per inoculated group) were housed in two separate pens. One day before inoculation, baseline swabs were collected. Pigs were then intramuscularly inoculated with 10¹⁰ VP/mL SVV‑PTC or SVV‑WT. One day after inoculation, 3 negative pigs were introduced into each pen as contact pigs. All pigs were kept together for the duration of the experiment. Nasal, pharyngeal, and fecal swabs were collected at 0, 2, 4, 6, and 9 dpc for viral copy number detection. Blinding was implemented in this study: animal allocation was performed by an investigator not involved in subsequent experiments, and the personnel performing immunizations, blood collection, and data detection were unaware of the group assignments. The background threshold was set as the mean viral load detected in all pigs prior to inoculation or contact (0 dpc). At the end of the transmission study, pigs were euthanized as described above. A fixed time of day was used for all procedures, the order of treatments was randomized, and all pens were located in the same room with uniform environmental conditions (Supplementary Fig. 15).

RNA extraction and real-time reverse transcriptase PCR (RT-qPCR)

Total RNA was extracted from SVV-PTC-infected cells and from nasal, pharyngeal, rectal swabs and blood samples of pigs using Trizol reagent. The primers (SVV 3D-F: 5′-AGAATTTGGAAGCCATGCTCT-3′; SVV 3D-R: 5′- GAGCCAACATAGATACA-GATTGC-3′) and TaqMan probe (5′-FAM-TTCAAACCAGGAACACTACTCGAG-TAMRA-3′) targeting the SVV 3D gene were synthesized¹⁴. SVV RNA was detected using the HiScript II One Step RT-PCR Kit (Vazyme, P611-01) following the manufacturer’s instructions. A 10-fold serial dilution series (from 10⁻¹ to 10⁻⁸) of plasmid pMD18-T-3D was prepared to generate a standard curve for quantification of viral genome copy numbers in the samples. For the determination of viral titers of viruses rescued from cells, viral RNA was extracted and then reverse-transcribed using the HiScript II Q RT SuperMix for qPCR (Vazyme, R223) in a two-step protocol, followed by absolute quantitative PCR to determine the viral copy numbers. Viral titers are expressed as VP/mL, defined as one viral particle corresponding to one genome copy.

Enzyme-linked immunosorbent assay (ELISA)

To determine SVV-specific IgG, IgG1, and IgG2a antibody levels in mice or swine, an indirect ELISA was employed¹³. Purified wild-type virus was diluted in 0.05 mol/L carbonate-bicarbonate buffer (pH 9.6), and 96-well microtiter plates were coated with 100 μL per well overnight at 4 °C. Plates were washed with PBS between each step. Wells were blocked with 100 μL of PBS-0.1% Tween 20 containing 2% BSA at 37 °C for 2 h. Serum samples (diluted 1:500, 100 μL/well) were added and incubated overnight at 4 °C, followed by incubation with HRP-conjugated goat anti-mouse IgG (1:5000), IgG1 (1:10000), IgG2a (1:10000), or HRP-conjugated goat anti-swine IgG (1:5000) (AB Clone, China) at 37 °C for 1 h. After the addition of tetramethyl benzidine (TMB; Solarbio, Beijing, China) and incubation at 37 °C for 15 min, the reaction was terminated with stop solution (Solarbio, Beijing, China). Absorbance was measured at 450 nm using a multimode microplate reader (Tecan Spark 10 M).

Virus neutralization (VN) assay

The neutralization assay was performed as described previously¹⁴ in 96-well cell culture plates. Serum samples were inactivated by heating at 56 °C for 30 min and subjected to two-fold serial dilution in DMEM supplemented with antibiotics (100 μg/mL streptomycin, 100 U/mL penicillin). Diluted sera were mixed with an equal volume of SVV (SVV LNSY01-2017, 200 TCID₅₀), and the mixture was incubated at 37 °C for 90 min. BHK-21 cell suspension in 2% DMEM was then added, with incubation continued for 72 h. Neutralizing antibody titers were calculated as the log₂ reciprocal of the highest serum dilution that inhibited 50% of SVV replication.

Lymphocyte proliferation

T lymphocyte proliferation was assessed at 42 dpi using a CCK-8 kit (Sigma, USA) as described previously⁴⁵. Briefly, lymphocytes were adjusted to 1 × 10⁶ cells/mL and seeded into 96-well plates with RPMI-1640 medium supplemented with 20% FBS. Inactivated SVV strain SVV LNSY01-2017 (0.1 MOI) was added to stimulate proliferation. Triplicate wells were used per group, with incubation at 37 °C for 72 h. Next, 10 μL of CCK-8 reagent was added to each well and incubated at 37 °C for 4 h. Absorbance at 450 nm was measured using a microplate reader. The stimulation index (SI) was calculated as follows: SI = (OD value of the immune group—OD value of the blank control group)/ (OD value of the negative control group—OD value of the blank control group).

Flow cytometry

CD3+ , CD4α+ and CD8α+ T lymphocyte percentages in lymphocytes were measured by flow cytometry as described⁴⁶. Lymphocytes were adjusted to 1 × 10⁸ cells/mL in 20% FBS-supplemented RPMI-1640, stimulated with inactivated SVV strain LNSY01-2017 (0.1MOI) for 4–6 h, then centrifuged at 1500 rpm. the cells were then collected and labeled with FITC goat anti-mouse CD3, PerCP-Cyanine 5.5 goat anti-mouse CD4α, and APC goat anti-mouse CD8α flow cytometric antibody (Proteintech, China). In the swine immunization assessment experiment, Alexa Fluor 647 mouse anti-pig CD3, FITC mouse anti-pig CD4α, and PE mouse anti-pig CD8α (Becton, Dickinson and Company, USA) were used. Antibody staining was performed following the manufacturer’s instructions. Both negative and positive control groups were included during the experiment. The negative control consisted of unstained cells, which were used to set the autofluorescence baseline. The positive controls consisted of CD3+ , CD4α+ , and CD8α+ single‑stained cells, which were used to adjust fluorescence compensation and define the position of the positive gates. For the detection of CD4α+ and CD8α + T cells, the gating strategy was as follows: lymphocytes were first gated on an FSC‑A/SSC‑A dot plot. CD3+ T cells were then identified from this lymphocyte gate. Finally, the percentages of CD4α+ and CD8α+ cells were calculated within the CD3+ gate. Data were collected using a flow cytometer (Beckman Coulter, Pasadena, CA, USA) and analyzed using CytExpert software (version 2.2) (Beckman Coulter, Pasadena, CA, USA).

Detection of cytokine assays

Cytokine detection was performed as previously described⁴⁷. Lymphocytes were adjusted to a concentration of 1 × 10⁶ cells/mL in RPMI-1640 medium supplemented with 20% fetal bovine serum, then stimulated with inactivated SVV (0.1 MOI) for 72 h before supernatant collection. Commercial Double-antibody sandwich ELISA kits (NeoBioscience, Wuhan, China) were used to quantify the secretion levels of IL-2, IL-4, IL-10, IFN-γ, and TNF-α in the lymphocyte supernatants.

Statistical analysis

All data were expressed as the mean ± standard deviation (SD) or mean ± standard error of the mean (SEM). One-way analysis of variance (ANOVA) with Newman-Keuls multiple comparisons test was used for single-factor group comparisons by GraphPad Prism 8.0.2. A P value < 0.05 was considered statistically significant.

Author contributions

P.Q. conceived the project, X.H.C., N.C., M.T.L., L.X.T., P. H., J.D., L.A.M., C.Z., C.W.L., Z.H.H. and J.Y.Z. performed the experiments. X.H.C. and X.M.L. analyzed the data. X.H.C. and N.C. wrote the paper. P.Q. and X.M.L. participated in the discussion and interpretation of the results. All authors have read, edited, and agreed to the published version of the manuscript.

Data availability

The datasets generated and/or analyzed during the current study are not publicly available because all relevant data are fully presented within this published article and its supplementary information files, but they are available from the corresponding author on reasonable request.

Competing interests

The authors declare no competing interests.

Supplementary information

The online version contains supplementary material available at https://doi.org/10.1038/s41541-026-01507-8.