Anatid herpesvirus 1 UL51 protein is palmitoylated by DHHC9 and DHHC18 for viral replication and virulence
Xiaolan Liu, Ruihao Zhang, Mingshu Wang, Anchun Cheng, Wei Zhang, Qiao Yang, Xumin Ou, Di Sun, Yu He, Bin Tian, Zhen Wu, Shaqiu Zhang, Juan Huang, Ying Wu, Yanling Yu, Ling Zhang, XinXin Zhao, Dekang Zhu, Shun Chen, Renyong Jia, Mafeng Liu
TL;DR
A viral protein called UL51 is modified through palmitoylation, which is crucial for its role in viral replication and virulence.
Contribution
This study identifies palmitoylation of UL51 by host enzymes DHHC9 and DHHC18 as a novel mechanism for viral pathogenesis.
Findings
Deletion of UL51 significantly reduces viral replication and virulence in ducks.
Palmitoylation of UL51 is essential for its interaction with UL10 and its localization to the Golgi apparatus.
Disruption of UL51 palmitoylation impairs secondary envelopment and infectious virus production.
Abstract
The tegument protein UL51 is conserved among herpesviruses, but the mechanisms underlying its role in viral pathogenesis remain unclear. In this study, using Anatid herpesvirus 1 (AnHV-1) as a model, we show that deletion of UL51 markedly attenuates viral replication and virulence in ducks. Immunoprecipitation coupled with LC–MS/MS identified interactions between pUL51 and multiple viral proteins, among which the association with pUL10 depends on pUL51 palmitoylation. This modification promotes pUL10 stability and its localization to the Golgi apparatus. Further analysis revealed that the palmitoylation of pUL51 is mediated by the host palmitoyltransferases DHHC9 and DHHC18 and is required to maintain pUL51 stability by preventing ubiquitin–proteasome-mediated degradation. Moreover, the deletion of pUL51 led to the accumulation of incompletely enveloped viral particles in the cytoplasm,…
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Fig 12- —http://dx.doi.org/10.13039/501100001809National Natural Science Foundation of China
- —China Agriculture Research System of MOF and MARA
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Taxonomy
TopicsHerpesvirus Infections and Treatments · Viral-associated cancers and disorders · Cytomegalovirus and herpesvirus research
Introduction
The Herpesviridae family is a large group of enveloped DNA viruses that infect a wide range of animals, including nearly all vertebrates and some invertebrates [1]. This family is divided into three major subfamilies: Alpha-, Beta-, and Gammaherpesvirinae. Herpes simplex virus 1 (HSV-1), Pseudorabies virus (PRV), Anatid herpesvirus 1 (AnHV-1), and varicella-zoster virus (VZV) are alphaherpesviruses. Herpesvirus virions are approximately spherical, with diameters ranging from 120 to 200 nm. The complete viral particle consists primarily of four layers, which, from inner to outer, consist of the double-stranded DNA core, capsid, tegument, and envelope [2]. The protein layer between the capsid and envelope is the tegument, a structure unique to herpesviruses [3] that serves multiple functions. The pUL51 protein and its homologs are highly conserved across the alphaherpesvirus subfamily, underscoring their fundamental role in the viral life cycle. In HSV-1 and PRV, pUL51 has been implicated in secondary envelopment [4], virion assembly [5], and cell-to-cell spread [6,7], thereby supporting efficient viral replication. In HSV-1-infected cells, pUL51 undergoes posttranslational modifications such as phosphorylation and palmitoylation, with palmitoylation being particularly critical for virus–cell membrane interactions [8,9]. Interestingly, the role of HSV-1 pUL51 in promoting cell-to-cell spread appears to be cell type dependent and does not always correlate with virion release [6]. Additionally, studies have shown that pUL51 interacts with other viral proteins, including pUL7, pUL14, and glycoprotein E (gE), to facilitate secondary envelopment and cell-to-cell spread [6,7,10]. The function of the N-terminus of pUL51 is similar to that of members of the endosomal sorting complex required for transport (ESCRT) protein family, which participate in viral assembly [5,11,12]. Homologs of pUL51 are also present in beta- and gammaherpesviruses. For instance, deletion of pUL71 (the pUL51 homolog) in human cytomegalovirus (HCMV) disrupts viral assembly compartment formation and secondary envelopment, leading to severely impaired replication [13,14]. Similarly, in Kaposi’s sarcoma-associated herpesvirus (KSHV), the pUL51 homolog pORF55 is essential for the production of infectious virions [15]. Collectively, these studies highlight pUL51 as a multifunctional protein that plays a conserved and critical role in the herpesvirus life cycle.
Palmitoylation is a critical lipid-based posttranslational modification (PTM) that involves the covalent attachment of medium-chain fatty acids, primarily palmitic and stearic acids, to specific residues of target proteins [16]. On the basis of the type of linkage, protein palmitoylation can be classified into three forms: S-palmitoylation, N-palmitoylation, and O-palmitoylation [17,18]. N-palmitoylation occurs at specific lysine residues via an amide bond, whereas O-palmitoylation involves the attachment of a palmitoyl group to serine residues through an ester bond. S-palmitoylation, which involves the formation of a thioester bond with cysteine residues, is a reversible lipid modification [19] and plays essential roles in regulating protein subcellular localization, trafficking, stability, membrane association, and signal transduction [20,21]. For instance, palmitoylation enhances the membrane fusion capacity of bovine foamy virus (BFV) envelope glycoproteins and facilitates the release of subviral particles (SVPs), thereby increasing BFV infectivity [22]. This modification also mediates autolysis of the hepatitis C virus (HCV) NS2 protein and the recruitment of viral envelope proteins to viral assembly sites, promoting HCV RNA replication and infectious particle assembly [23]. Additionally, palmitoylation is involved in the intracellular transport of the influenza B virus NB protein [24] and the binding of the hepatitis E virus ORF3 protein [25] and Sindbis virus TF protein to the plasma membrane [26] and promotes viral budding.
Protein palmitoylation is catalyzed by palmitoyl acyltransferases (PATs), which contain a highly conserved Asp-His-His-Cys (DHHC) zinc finger domain at their catalytic site and are therefore also known as DHHC proteins [27]. DHHC proteins catalyze protein palmitoylation through a two-step reaction. The palmitoyl group is transferred from palmitoyl-CoA to the enzyme itself, forming an acyl-enzyme intermediate, after which the palmitoyl group is transferred to a specific cysteine residue on the substrate protein [28,29]. Although viral proteins themselves do not possess PAT activity, viruses have evolved mechanisms to exploit host PAT systems to modify their own proteins [30,31]. For example, Chikungunya virus utilizes host DHHC2 and DHHC19 to modify its nonstructural protein 1 (NSP1) [32], whereas herpes simplex virus type 1 (HSV-1) uses DHHC3 to palmitoylate its membrane protein UL20 [33,34]. Human immunodeficiency virus type 1 (HIV-1) achieves palmitoylation of its Tat protein via DHHC20 [35]. Although the palmitoylation of the HSV-1 UL51 protein has been demonstrated, the functional significance of this modification in herpesvirus infection and its regulatory mechanisms remain unclear. In this study, we used Anatid herpesvirus 1 (AnHV-1), an alphaherpesvirus, as a model to investigate the role of pUL51 during infection. We observed that pUL51 promotes viral replication and virulence both in vitro and in vivo and that its palmitoylation is critical for increasing replication efficiency in vitro and pathogenicity in animals. We further observed that pUL51 undergoes palmitoylation at its N-terminal cysteine residue (C9). Substitution of this cysteine with alanine or pharmacological inhibition of palmitoylation with 2-bromopalmitate (2-BP) abrogated palmitoylation, leading to reduced pUL51 expression and disrupted Golgi localization. Finally, we determined that pUL51 utilizes the host palmitoyltransferases DHHC9 and DHHC18 to promote its own palmitoylation and stabilize protein expression, thereby promoting viral replication and increasing virulence.
Results
Construction and identification of recombinant viruses
To investigate the role of pUL51 in viral infection, we utilized an infectious clone platform containing the genome of a Chinese virulent strain (CHv) of AnHV-1. Using a label-free two-step Red recombination strategy, we generated infectious clone plasmids for a UL51 gene-deleted strain (AnHV-1‑ΔUL51) and a corresponding revertant strain (AnHV-1‑ΔUL51‑Rev) (Fig 1A). To generate recombinant viruses related to the UL51 gene, the positive infectious clone plasmids AnHV-1-ΔUL51 and AnHV-1-ΔUL51-Rev were transfected into duck embryo fibroblasts (DEFs), and the cellular status was monitored regularly. The results revealed fluorescent foci at 72 hours post-transfection (Fig 1B). These foci expanded over time and were accompanied by the appearance of lesions in DEFs under white light (P0). The rescued virus was subsequently passaged into fresh DEFs, where it produced fluorescence and corresponding cytopathic effects (P3). Over time, the inverted duplication of AnHV-1 genomic sequences inside the mini-F element underwent homologous recombination triggered by the recombinases in DEFs for the markerless excision of vector sequences from the AnHV-1 genome, resulting in the gradual disappearance of fluorescence. By passage 10 (P10), nonfluorescent AnHV-1-ΔUL51 deletion and UL51-revertant strains were successfully obtained. Deletion and subsequent restoration of the UL51 gene were confirmed by PCR analysis (Fig 1C), using DNA from DEFs as a negative control. Amplification of the UL51 locus yielded a product of approximately 549 bp in AnHV-1-ΔUL51 compared with a 1,308 bp product in both the parental virus (AnHV-1-CHv50) and the revertant strain (AnHV-1-ΔUL51-Rev). This difference in size of approximately 759 bp indicates the complete deletion of the UL51 coding region in the mutant virus. Subsequent sequencing results also confirmed the deletion of the UL51 gene in AnHV-1-ΔUL51. As expected, the expression of pUL51 was not detected in AnHV-1-ΔUL51-infected cells by Western blot analysis or IFA (Fig 1D and 1E), and the absence of pUL51 did not affect the expression of its adjoining proteins, pUL50 and pUL52 (Fig 1D). These results confirmed the successful generation of AnHV-1-ΔUL51, which was then used for subsequent experiments.
Construction and identification of recombinant viruses.(A) Schematic diagram of the construction of the infectious AnHV-1-△UL51 clone, (I) genomic structure of AnHV-1-CHv50, (II) first red recombination replacing the UL51 gene with the kanamycin gene, (III) product after the first red recombination, (IV) second red recombination where the I-Sec I restriction site is recognized by recombinase to remove the Kan fragment, and (IV) the genomic structure of AnHV-1-△UL51. (B) Rescue and purification of AnHV-1-△UL51. (C) PCR identification of the recombinant virus. (D) Western blot analysis of UL51 gene-deficient viral protein expression. (E) IFA identification of the recombinant virus, representative images from three independent replicate experiments, scale bar, 40 μm.
Deletion of pUL51 reduces the incidence and mortality of viral infection
An in vivo experiment was subsequently performed to assess the effect of UL51 on viral pathogenesis, following the experimental schematic outlined in Fig 2A. In group 1, 14-day-old ducks (n = 40) were inoculated with an equivalent dose of either the parental virus (AnHV-1-CHv50), the revertant virus (AnHV-1-ΔUL51-Rev), or the UL51-deficient virus (AnHV-1-ΔUL51) or mock-infected with minimal essential medium (MEM). Body temperature, weight change, and survival were monitored daily for 10 days post-infection. As shown in Fig 2B–2D, ducks infected with AnHV-1-ΔUL51 survived the entire observation period and maintained normal body temperature and body weight, a phenotype indistinguishable from that of the mock-infected MEM control group. In contrast, infection with AnHV-1-CHv50 and AnHV-1-ΔUL51-Rev viruses induced severe disease, characterized by hyperthermia, significant weight loss, and progressive lethality, culminating in final mortality rates of 60% and 70%, respectively. On the basis of the peak mortality observed between days 2 and 7 postinoculation in group 1, we next further evaluated the impact of UL51 deletion on AnHV-1 virulence in vivo. Twenty-four 14-day-old ducklings were inoculated with either MEM or 10⁷ TCID₅₀ of AnHV-1-CHv50, AnHV-1-ΔUL51-Rev, or AnHV-1-ΔUL51. Tissues were collected at 5 days post-inoculation for histopathological analysis and viral load quantification. No mortality was observed in ducklings from the MEM or AnHV-1‑ΔUL51 groups, and no gross lesions were detected upon necropsy. In contrast, infection with either the revertant or parental virus caused mortality and was associated with significant pathological alterations (indicated by the black arrow in the diagram), including hemorrhage and/or necrosis in the heart, liver, and spleen; thymic atrophy; and thinning and hemorrhage of the duodenal and cecal walls (Fig 2E). Liver, spleen, cecum, and thymus tissues from each group were collected and processed for hematoxylin and eosin (HE) staining. Histological analysis revealed no significant degeneration or necrosis in the MEM or AnHV-1-ΔUL51 groups. The liver exhibited preserved architecture with well-organized hepatic cords. The spleen structure was intact, the intestinal cecal villi appeared finger-like and neatly arranged, and the thymus demonstrated clear corticomedullary demarcation and uniform thymocyte distribution. However, the AnHV-1-ΔUL51-Rev and AnHV-1-CHv50 groups exhibited significant pathological alterations in multiple organs, including hepatic steatosis, cellular necrosis, and loss of normal tissue architecture. In the thymus, the focal areas displayed erythrocyte-filled reticular spaces with an absence of normal cellular structure. The spleen showed blurred cellular margins and a reticulated pattern resulting from the loss of parenchymal cells. Cecal villi were disrupted and sloughed, with no intact cellular morphology observed (Fig 2F). We used quantitative PCR to measure the viral load of AnHV-1 in various tissues to evaluate its replication in ducks (Fig 2G). The results revealed that viral loads in the heart, liver, spleen, glandular stomach, thymus, bursa of Fabricius, duodenum, and cecum of ducks infected with AnHV-1-ΔUL51 were significantly lower than those in ducks infected with either AnHV-1-CHv50 or AnHV-1-ΔUL51-Rev. Specifically, the viral loads in tissues of ducks infected with the ΔUL51-deficient virus ranged from 10^3.57^ to 10^4.73^ copies/1 g, whereas those in tissues from ducks infected with the parental virus or the revertant strain ranged between 10^6.24^ and 10^10.08^ copies/1 g. These findings indicate that deletion of the UL51 gene significantly impaired the replication capacity of AnHV-1 in vivo. Together, these results demonstrate that the absence of UL51 reduces the virulence of anatid herpesvirus 1.
*Deletion of pUL51 reduces the incidence and mortality of viral infection.(A) Experimental design for assessing the effects of UL51 deletion on AnHV-1 pathogenicity in vivo. Fourteen-day-old ducklings were intramuscularly inoculated with 10⁷ TCID₅₀ of AnHV-1-ΔUL51, AnHV-1-ΔUL51-Rev, or AnHV-1-CHv50. Ducklings injected with MEM served as mock-infected controls. Ducklings in Group 1 (n = 10 per group) were monitored for 10 days post-infection for changes in body temperature, body weight, and survival. Ducklings in Group 2 (n = 6 per group) were euthanized at 5 days post-infection for histopathological assessment of organ lesions and viral load quantification. The schematic images were hand-drawn by the authors and edited manually (licensed under CC BY 4.0). The nonoriginal icon within these figures was sourced from https://openclipart.org/detail/326143. (B–D) Body temperature, body weight, and survival rate of ducks infected with AnHV-1-ΔUL51 over a 10-day period. (E) Macroscopic pathological changes in organs from AnHV-1-ΔUL51-infected ducks at 5 days post-infection. These images were taken by the authors of this manuscript and edited manually (licensed under CC BY 4.0). (F) Histopathological analysis of various tissues collected from AnHV-1-ΔUL51-infected ducks on day 5 post infection (hematoxylin and eosin staining; original magnification, 100×). (G) Viral DNA copies in the tissues and organs of ducks infected with AnHV-1-ΔUL51 at 5 days post-infection. Data are expressed as mean ± SD from three independent biological replicates. Statistical analysis was performed using two-way ANOVA (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ***p < 0.0001).
pUL51 promotes viral replication in vitro
To investigate the mechanism by which pUL51 deletion attenuates AnHV-1 virulence, we examined the in vitro replication characteristics of AnHV-1-ΔUL51 by assessing viral growth kinetics. DEFs were infected with AnHV-1-CHv50, AnHV-1-ΔUL51-Rev, and AnHV-1-ΔUL51 at a multiplicity of infection (MOI) of 0.01, and cell samples were collected at 24, 48, 60, and 72 hours post-infection. The viral titer of AnHV-1-ΔUL51 remained consistently lower than those of the parental and revertant viruses, with the most pronounced difference observed at 48 hours post-infection (approximately 9-fold, Fig 3A). These results indicate that AnHV-1-∆UL51 produces progeny virus less efficiently than the parental and revertant viruses do, suggesting that deletion of UL51 impairs viral replication in vitro. To explore the influence of the UL51 gene on the steps of the viral life cycle, we first investigated the adsorption (Fig 3B) and invasion (Fig 3C) abilities of the UL51 gene deletion virus on the host cell surface. Next, we examined the impact of the UL51 gene on viral genomic DNA replication by infecting DEFs with three viruses at identical copy numbers and incubating them for 6 h. Infected cells were collected at 7, 9, 11, and 13 hours post-infection. Viral genomic DNA was extracted to assess the impact of the UL51 gene on viral genomic DNA replication. The results revealed no significant differences in viral DNA copy numbers among the three strains (Fig 3D), indicating that the deletion of the UL51 gene did not affect the replication of viral genomic DNA. These findings suggest that pUL51 is dispensable for the early stages of viral infection, including adsorption, invasion, and genomic replication.
*Multistep growth kinetics of recombinant viruses and the role of pUL51 in the viral life cycle.(A) Multistep growth curves of AnHV-1-ΔUL51. Duck embryo fibroblasts (DEFs) were infected with AnHV-1-ΔUL51, AnHV-1-ΔUL51-Rev, or AnHV-1-CHv50 at an MOI of 0.01. Total cells and supernatants were harvested at the indicated time points, and viral titers were determined. (B) Effect of pUL51 on viral adsorption. DEFs were infected with a labeled virus at an MOI of 1 for 2 hours 4°C, after which the number of viral DNA copies was quantified to evaluate the adsorption efficiency. (C) Effect of pUL51 on viral invasion. Following the adsorption period, the cells were incubated at 37°C for 2 hours. Samples were collected, and viral entry efficiency was assessed via quantitative PCR. (D) Effect of pUL51 on viral genomic DNA replication. (E) Effect of pUL51 on viral release. DEFs were infected with viruses. After 16 hours, the medium was replaced with fresh maintenance medium, and both the cells and the supernatants were harvested at 1, 2, 3, and 4 hours. The ratio of extracellular to intracellular viral titers was used to assess viral release efficiency. (F) Effect of pUL51 on cell-to-cell spread. Plaque morphology was visualized by crystal violet staining. The diameters of 30 representative plaques per group were measured using ImageJ software; the results are summarized in the right panel. Data are expressed as mean ± SD from three independent biological replicates. Statistical analysis was performed using one-way/two-way ANOVA (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ***p < 0.0001).
We subsequently examined the release of mature viral particles following infection with three viral strains and observed a gradual increase in the number of infectious viral particles over time (Fig 3E). To quantify the efficiency of viral release, we defined the supernatant titer as extracellular particles and the cell-associated titer as intracellular particles and then calculated the release efficiency as the ratio of the extracellular to the intracellular titer at each time point. Compared with the parental and revertant viruses, the UL51-deficient virus showed significantly reduced release efficiency at all time points examined (Fig 3E). These findings indicate that pUL51 is required for efficient progeny virus release, suggesting that it functions at a late stage of replication to promote viral release. Furthermore, the deletion of UL51 significantly altered plaque morphology. To quantify these differences, at least 30 randomly selected plaques from cells infected with each virus were analyzed statistically. When the plaque size of AnHV-1-CHv50 was used as the reference, the average plaque size of AnHV-1-ΔUL51 was only 30.7%, representing a 69.3% reduction compared with that of AnHV-1-CHv50 (Fig 3F). This pronounced plaque-size phenotype is consistent with the observed defect in particle release and likely reflects impaired cell-to-cell spread. In conclusion, pUL51 is not required for viral adsorption, invasion, or genomic DNA replication but facilitates the release of mature virions, suggesting that pUL51 functions primarily during the late stages of AnHV-1 replication.
Identification of pUL51-interacting viral proteins
pUL51 has been demonstrated to promote viral replication in vitro and pathogenicity in vivo. To elucidate the molecular mechanisms of pUL51 in AnHV-1 infection, we used LC‒MS/MS technology to preliminarily screen viral proteins that interact with pUL51. First, pUL51 was transiently expressed via plasmid transfection, followed by AnHV-1 infection of DEFs and immunoprecipitation (IP) enrichment (Fig 4A). Silver staining revealed a Flag-tagged pUL51 band in the experimental group, along with additional specific bands compared with those in the IgG control group (Fig 4B). LC‒MS/MS analysis of the affinity-purified samples revealed four specific viral interaction partners of pUL51: pUL27, pUL30, pUL38, and pUL10 (Fig 4C). To further validate these interactions, we cotransfected DEFs with pCAGGS-UL51-3xFlag and each of the following vectors: pCAGGS-UL27-3xHA, pCAGGS-UL30-3xHA, pCAGGS-UL38-3xHA, and pCAGGS-UL10-3xHA, followed by Co-IP experiments. The results revealed that pUL51 successfully pulled down pUL27, pUL30, pUL38, and pUL10, indicating that pUL51 interacts with all four proteins (Fig 4D). To further validate these interactions during viral infection, DEFs were infected with AnHV-1 at an MOI of 1. Cells were collected 36 hours post-infection and subjected to immunoprecipitation using a pUL51-specific rabbit polyclonal antibody and rabbit IgG as a control. The results demonstrated that pUL10, pUL27, pUL38 and pUL30 were detectable in the pUL51 immunoprecipitates under viral infection conditions (Fig 4E). Indirect immunofluorescence assays were used to analyze the colocalization of pUL51 with its interacting partners. While pUL30 and pUL38 did not significantly colocalize with pUL51, a clear colocalization signal was observed for pUL27 and pUL10 with pUL51 in specific subcellular compartments (Fig 4F), indicating a strong functional association between pUL27, pUL10, and pUL51.
Identification of viral proteins that interact with pUL51.(A) Schematic of the immunoprecipitation‒mass spectrometry (IP‒MS) workflow used to identify AnHV-1 pUL51-interacting proteins. Cells were transfected with a plasmid encoding pUL51-Flag. At 24 h posttransfection, the cells were infected with AnHV-1 at an MOI of 1 for another 24 h. The cell lysates were then subjected to immunoprecipitation using anti-Flag or control mouse IgG antibodies, followed by silver staining and Western blot analysis. (B) Silver staining and Western blot identification of pUL51-Flag IP-enriched samples. (C) Mass spectrometry results for pUL51-Flag. The raw mass spectrometry data results can be found at https://doi.org/10.6084/m9.figshare.31229161. (D) Co-IP experiments detecting interactions between pUL51 and pUL30, pUL38, pUL27, and pUL10 in DEFs. DEFs were cotransfected with 2 μg of pUL51-encoding plasmid and 2 μg of either pUL30, pUL38, pUL27, or pUL10-encoding plasmids. Samples were collected 36 h posttransfection for Co-IP analysis. (E) Coimmunoprecipitation experiments were performed to detect interactions between pUL51 and pUL30, pUL38, pUL27, and pUL10 during AnHV-1 infection. DEFs were infected with AnHV-1 at an MOI of 1. Cells were collected 36 hours post-infection and subjected to immunoprecipitation using a pUL51-specific rabbit polyclonal antibody and rabbit IgG as a control. Western blot analysis was performed using rabbit polyclonal antibodies specific to pUL51, pUL10, pUL27, and pUL30. DEFs were used as a control. (F) The colocalization of pUL51 with pUL30, pUL38, pUL27, and pUL10 in DEFs was detected by IFA. 0.5 μg of the pUL51-encoding plasmid was cotransfected with 0.5 μg each of pUL30, pUL38, pUL27, and pUL10 into DEFs. Samples were collected 24 h posttransfection for IFA analysis, representative images from three independent replicate experiments. Scale bar: 20 μm.
pUL51 stabilizes pUL10 expression and colocalizes with it in the Golgi apparatus
pUL51 interacts with both pUL27 and pUL10 and colocalizes with each other. To investigate the effect of pUL51 on the expression of these two proteins, we cotransfected cells with pUL51 and either pUL27 or pUL10. While pUL51 did not alter pUL27 expression (Fig 5A), it markedly increased pUL10 expression (Fig 5B). We next asked whether this effect is evident during viral infection. Comparisons of cells infected with AnHV-1-CHv50 and AnHV-1-ΔUL51 at an equal MOI revealed that the loss of pUL51 specifically reduced pUL10 expression without affecting the levels of other viral proteins (Fig 5C), providing further evidence that pUL51 increases pUL10 expression. To determine whether pUL51 regulates protein expression by affecting protein stability, we performed the experiments in the presence of the protein synthesis inhibitor cycloheximide (CHX). We found that pUL27 degradation remained largely unchanged regardless of the presence of pUL51 (Fig 5D). In contrast, the presence of pUL51 significantly delayed the degradation of pUL10, maintaining its protein stability (Fig 5E). These findings indicate that pUL51 specifically maintains the stability of pUL10, preventing its degradation. To determine the subcellular localization of pUL51 and pUL10, we performed immunofluorescence staining. Transiently expressed pUL51 colocalized with the Golgi marker GM130, whereas pUL10 was uniformly distributed in the cytoplasm. When the cells were cotransfected, distinct colocalization was observed, suggesting potential colocalization of both proteins at the Golgi apparatus (Fig 5F). Since both antibodies used to detect pUL51 and pUL10 were rabbit polyclonal antibodies, to exclude potential interference from antibody cross-reactivity, we used the recombinant virus AnHV-1-UL10_HA_ to label pUL10 with an HA tag during infection and used a rabbit anti-pUL51 antibody to detect endogenous pUL51. The results confirmed that under viral infection conditions, both pUL51 and pUL10 colocalized with GM130, and colocalization was also observed between pUL51 and pUL10 (Fig 5G), indicating that pUL51 and pUL10 colocalize at the Golgi apparatus during viral infection.
pUL51 increases the expression of pUL10 and colocalizes with it in the Golgi apparatus.(A) Transient expression of pUL51 and pUL27. DEFs were transfected with 1 μg of either empty pCAGGS vector or pCAGGS-UL51–3 × Flag, together with 1 μg of pCAGGS-UL27–3 × HA plasmid. Cells were harvested at 36 h posttransfection for Western blot analysis. (B) Transient expression of pUL51 and pUL10. DEFs were transfected with 1 μg of empty pCAGGS or pCAGGS-UL51–3 × Flag, along with 1 μg of pCAGGS-UL10–3 × HA plasmid. Lysates were collected at 36 h posttransfection for Western blot analysis. (C) Western blot analysis of pUL10 expression in cells infected with the UL51-deficient virus. DEFs were infected with AnHV-1-ΔUL51 or AnHV-1-CHv50 at an MOI of 0.1 and harvested at 18, 36, and 48 h post infection for Western blot. (D) DEF cells were cotransfected with the following plasmid combinations: (i) 0.5 μg of empty pCAGGS vector plus 0.5 μg of pCAGGS-UL27–3 × HA or (ii) 0.5 μg of pCAGGS-UL51–3 × Flag plus 0.5 μg of pCAGGS-UL27–3 × HA. Twenty-four hours post-transfection, the cells were treated with cycloheximide (CHX) and harvested at 0, 3, and 6 hours after CHX addition for Western blot analysis. (E) DEF cells were cotransfected with the following plasmid combinations: (i) 0.5 μg of empty pCAGGS vector plus 0.5 μg of pCAGGS-UL10–3 × HA or (ii) 0.5 μg of pCAGGS-UL51–3 × Flag plus 0.5 μg of pCAGGS-UL10–3 × HA. Twenty-four hours post-transfection, the cells were treated with cycloheximide (CHX) and harvested at 0, 3, 6, and 9 hours after CHX addition for Western blot analysis. (F) Colocalization of pUL51 and pUL10 with the Golgi marker GM130 in transfected DEFs. Cells were transfected with plasmids encoding UL51–3 × Flag and UL10–3 × HA. At 24 h posttransfection, immunofluorescence staining was performed using antibodies against GM130. Nuclei were stained with DAPI, representative images from three independent replicate experiments. Scale bar: 20 μm. Scale bar: 20 μm. (G) Colocalization of pUL51 and pUL10 with GM130 during AnHV-1 infection. DEFs were infected with AnHV-1-UL10HA. The proteins were labeled with anti-pUL51 (rabbit polyclonal) and anti-HA (mouse monoclonal) antibodies. The Golgi apparatus was stained with anti-GM130. The nuclei were counterstained with DAPI, representative images from three independent replicate experiments. Scale bar: 20 μm.
The C9 residue of pUL51 is critical for its interaction with pUL10
To identify the key interaction site between pUL51 and pUL10, we generated mutations based on the conserved domains of pUL51 (Fig 6A). Coimmunoprecipitation (Co-IP) assays revealed that the 1–161 amino acid region of pUL51 mediates binding to pUL10 (Fig 6B and 6C). Further stepwise deletion mutations of the N-terminal 1–161 amino acids revealed that deletion of the first 9 amino acids (UL51Δ1–9aa) already abolished the interaction with pUL10 (Fig 6D). Sequence alignment analysis revealed that the 9th cysteine (Cys) residue is highly conserved across pUL51 and its homologous proteins in various herpesviruses (Fig 6E). To assess the role of this site in protein interactions, we generated a mutation by substituting the ninth cysteine with alanine (C9A). AlphaFold predicted structures for both pUL51 and pUL51/C9A, revealing structural alterations in pUL51 following the C9A mutation (S1A Fig). Molecular docking simulations further revealed a decrease in the number of interaction sites between the pUL51 C9A mutant and pUL10 compared with the wild-type protein (S1B and S1C Fig). Coimmunoprecipitation (Co-IP) assays confirmed that the C9A mutation disrupted the pUL51–pUL10 interaction (Fig 6F). To determine whether this site also influences pUL51 interactions with other viral proteins (pUL27, pUL30, and pUL38), we performed Co‑IP experiments to compare wild‑type pUL51 and the pUL51/C9A mutant. Both proteins were able to bind to these viral partners (Fig 6G–6I), indicating that the C9A mutation does not affect pUL51 interactions with pUL27, pUL30, or pUL38. Previous studies have shown that pUL51 maintains the stability of pUL10. We further examined the impact of the C9A mutation on this function and found that pUL10 degradation accelerated in pUL51/C9A compared with that in wild-type pUL51 (Fig 6J), indicating that the stabilizing effect of pUL51 on pUL10 is weakened or lost because of this mutation. Notably, the C9A mutant of pUL51 still colocalized with pUL10 but no longer specifically colocalized with the Golgi apparatus (Fig 6K). Together, these results demonstrate that the cysteine residue at position 9 of pUL51 is critical for its interaction with pUL10.
The C9 residue of pUL51 is critical for its interaction with pUL10.(A) Schematic representation of pUL51 mutant plasmids. (B) Interaction between the N-terminal fragment of pUL51 (amino acids 1–161) and pUL10. DEFs were transfected with 2 μg of pCAGGS-UL511–161-3 × Flag and 2 μg of pCAGGS-UL10-3 × HA. Coimmunoprecipitation (Co-IP) was performed at 36 h post transfection. (C) Interaction between the C-terminal fragment of pUL51 (amino acids 162–252) and pUL10. DEFs were transfected with 2 μg of pCAGGS-UL51162–252-3 × Flag and 2 μg of pCAGGS-UL10-3 × HA. Cells were harvested at 36 h posttransfection for Co-IP analysis. (D) Interaction between truncated pUL51 and pUL10. DEFs were transfected with 2 μg of a plasmid encoding a truncated pUL51 fragment and 2 μg of pCAGGS-UL10-3 × HA. Samples were collected at 36 h posttransfection for Co-IP. (E) Sequence analysis of the pUL51 C9 site. Site-directed mutagenesis was performed to substitute cysteine with alanine at position 9 (highlighted in red). Multiple sequence alignment of the N-terminal 20 amino acids of pUL51 from eight herpesviruses: HSV-1 (GenBank: NC_001806.2), VZV (GenBank: NC_001348.1), PRV (GenBank: MZ219273.1), BoHV-1 (GenBank: MH791338.1), AnHV-1 (GenBank: JQ647509.1), HCMV (GenBank: AH013698.2), EBV (GenBank: NC_009334.1), and KSHV (GenBank: NC_009333.1). Alignments were performed using MEGA 7.0. (F) Interaction between pUL51/C9A and pUL10. DEFs were transfected with 2 μg of plasmid encoding either wild-type pUL51 or pUL51/C9A, together with 2 μg of pCAGGS-UL10-3 × HA. Co-IP analysis was conducted 36 h posttransfection. (G) Interaction between pUL51/C9A and pUL27. DEFs were transfected with 2 μg of plasmid encoding either wild-type pUL51 or pUL51/C9A, together with 2 μg of pCAGGS-UL27-3 × HA. Co-IP analysis was conducted 36 h posttransfection. (H) Interaction between pUL51/C9A and pUL38. DEFs were transfected with 2 μg of plasmid encoding either wild-type pUL51 or pUL51/C9A, together with 2 μg of pCAGGS-UL38-3 × HA. Co-IP analysis was conducted 36 h posttransfection. (I) Interaction between pUL51/C9A and pUL30. DEFs were transfected with 2 μg of plasmid encoding either wild-type pUL51 or pUL51/C9A, together with 2 μg of pCAGGS-UL30-3 × HA. Co-IP analysis was conducted 36 h posttransfection. (J) Effect of pUL51/C9A on pUL10 stability. DEF cells were transfected with 0.5 μg of plasmid encoding wild-type pUL51 or pUL51/C9A, and 0.5 μg of pCAGGS-UL10-3 × HA plasmid was concurrently transfected. Twenty-four hours post-transfection, the cells were treated with cycloheximide (CHX) and harvested at 0, 3, and 6 hours after CHX addition for Western blot analysis. (K) DEFs were transfected with plasmids expressing pUL51 or pUL51/C9A, along with 0.5 μg of pCAGGS-UL10-3 × HA plasmid. Twenty-four hours post-transfection, the cells were fixed and analyzed by immunofluorescence to assess protein colocalization. The nuclei were counterstained with DAPI, representative images from three independent replicate experiments. Scale bar: 20 μm.
The C9 residue is crucial for the palmitoylation and Golgi localization of pUL51
The pUL51 protein of AnHV-1, as well as those of other herpesviruses, contains a conserved cysteine residue at position 9 (Fig 6E), which has been reported to undergo palmitoylation. To determine whether AnHV-1 pUL51 is also palmitoylated and whether the C9 site is required for this modification, we substituted the cysteine at position 9 with alanine (C9A) using site-directed mutagenesis, generating the plasmid pCAGGs-UL51_C9A_-3xFlag. Flag-tagged wild-type pUL51 and mutant pUL51/C9A were expressed in DEFs, and palmitoylation of proteins was detected by immunoprecipitation and acyl-biotin exchange. Lysates from cells expressing pUL51-Flag or pUL51/C9A-Flag were subjected to immunoprecipitation using an anti-Flag antibody. The immunoprecipitated samples were then treated with N-ethylmaleimide (NEM) to block free thiol groups. Hydroxylamine (HAM) was subsequently applied to cleave thioester linkages, followed by conjugation with biotin-BMCC (1-biotinamido-4-[4’-(maleimidomethyl) cyclohexanecarboxamido] butane). Biotin incorporation was ultimately detected by Western blot analysis using horseradish peroxidase (HRP)-conjugated streptavidin.
pUL51-Flag was expressed in DEFs, which were subsequently treated with the protein palmitoylation inhibitor 2-bromopalmitate (2-BP) to inhibit protein palmitoylation. No band corresponding to palmitoylated pUL51-Flag was present in cells treated with 2-BP (Fig 7A, pUL51-Flag + 2-BP) compared with the mock-treated sample (Fig 7A, pUL51-Flag), confirming that pUL51-Flag was palmitoylated. Moreover, in HAM-treated samples (+HAM), a palmitoylated band for wild-type pUL51-Flag was detected (Fig 7B, pUL51-Flag), whereas no such band was observed in the untreated control group (-HAM). Conversely, the pUL51/C9A-Flag mutant exhibited no detectable palmitoylation signal, even after HAM treatment (Fig 7B, pUL51/C9A-Flag). These results demonstrate that AnHV-1 pUL51 is palmitoylated and that the conserved cysteine residue at position 9 (C9) is essential for this lipid modification. Next, to investigate the impact of palmitoylation on the biological function of AnHV-1 pUL51, we used the palmitoylation inhibitor 2-bromopalmitate (2-BP). To determine the appropriate working concentration of 2-BP, we assessed its effects on DEF activity using a CCK-8 assay kit. A concentration of 30 μM, which did not significantly affect cell viability, was selected for all subsequent experiments (S2A Fig). The results of the cytotoxicity assessment for the remaining inhibitors used (including MG132, CQ, and CHX) are shown in S2B-S2D Fig. As shown in S2E and S2F Fig, the cells were treated with 30 μM 2-BP during AnHV-1 infection. We found that inhibition of palmitoylation had no effect on viral adsorption or invasion (S2G and S2H Fig) but significantly reduced viral replication and release (S2I and S2J Fig), suggesting that palmitoylation promotes AnHV-1 replication. Palmitoylation is known to regulate the subcellular localization of proteins. To determine whether this modification influences the Golgi localization of pUL51, we performed immunofluorescence staining. The results revealed that transiently expressed wild-type pUL51 colocalized with the Golgi marker protein GM130. However, this colocalization was abolished following treatment with the palmitoylation inhibitor 2-BP. Similarly, the pUL51 C9A mutant failed to colocalize with GM130 (Fig 7C). These results demonstrate that palmitoylation at the C9 residue is essential for the correct Golgi localization of pUL51.
C9 is crucial for the palmitoylation and Golgi localization of pUL51.(A) Detection of palmitoylation modification of pUL51-Flag using the palmitoylation inhibitor 2-BP. (B) Measurement of pUL51 palmitoylation. DEFs were transfected with plasmids expressing pUL51-Flag or pUL51/C9A-Flag. At 36 h post transfection, the cell lysates were immunoprecipitated using anti-Flag magnetic beads and subjected to an ABE assay in the presence or absence of hydroxylamine (HAM) treatment, followed by Western blot analysis. (C) Subcellular localization of pUL51 and pUL51/C9A. DEFs were transfected with plasmids expressing pUL51-Flag or pUL51/C9A-Flag for 24 h and then treated with or without 30 μM 2-bromopalmitate (2-BP). Localization was assessed by immunofluorescence using an antibody against the Golgi marker GM130. The nuclei were counterstained with DAPI, representative images from three independent replicate experiments. Scale bar: 20 μm.
pUL51 palmitoylation at residue C9 is essential for protein stability
Palmitoylation plays a role in regulating protein stability. To investigate its effect on pUL51 protein stability, we exogenously expressed pUL51-Flag in DEFs and treated them with varying doses of the palmitoylation inhibitor 2-BP. The results revealed that pUL51-Flag expression decreased in a dose-dependent manner with increasing 2-BP concentration (Fig 8A), suggesting that inhibition of palmitoylation may promote pUL51 degradation. To further assess the degradation kinetics of pUL51, we performed a cycloheximide (CHX) chase assay to determine its protein half-life. As shown in Fig 8B and 8C, the apparent half-life of wild-type pUL51 was approximately 8 hours. In contrast, upon inhibition of palmitoylation with 2-BP, the half-life of pUL51-Flag was shortened to approximately 4 hours. A similar reduction in stability was observed for the palmitoylation-deficient mutant pUL51/C9A-Flag, which also exhibited a half-life of approximately 4 hours. These results demonstrate that the impairment of palmitoylation significantly accelerates the degradation of pUL51.
Palmitoylation at the C9 site maintains pUL51 stability by affecting the ubiquitin–proteasome degradation pathway.(A) DEFs were transfected with a pUL51-Flag plasmid for 24 h and then treated with increasing concentrations of 2-BP for 6 h. Protein expression levels were analyzed by Western blot. (B) DEFs transfected with pUL51-Flag for 12 h were treated with 30 μM 2-BP or DMSO for 12 h, followed by incubation with 100 mg/mL cycloheximide (CHX). Lysates were collected at the indicated time points and analyzed by Western blot. (C) DEFs expressing pUL51-Flag or pUL51/C9A-Flag were treated with CHX (100 mg/mL). Lysates were harvested at the indicated times and subjected to Western blot analysis. (D) DEFs expressing pUL51-Flag were treated for 6 h with 30 μM 2-BP alone or in combination with 50 μM MG132. DEFs expressing pUL51-Flag were treated for 6 h with 30 μM 2-BP alone or in combination with 50 μM chloroquine (CQ). Protein levels were assessed by Western blot. (E) DEFs were cotransfected with plasmids encoding pUL51-Flag or pUL51/C9A-Flag together with Ub-HA. After 36 h, the lysates were immunoprecipitated and analyzed by Western blot. (F) DEFs expressing pUL51/C9A-Flag were treated with or without 50 μM MG132 for 6 h, followed by CHX (100 mg/mL). Lysates were collected at the indicated times and analyzed by Western blot. The band intensity of pUL51-Flag was quantified using ImageJ software.
Protein degradation is mediated primarily by the ubiquitin‒proteasome system and the autophagy‒lysosome pathway. To elucidate the pathway responsible for pUL51 degradation upon palmitoylation inhibition, cells expressing pUL51-Flag were cotreated with 2-BP and either the proteasome inhibitor MG132 or the lysosome inhibitor chloroquine (CQ). Notably, inhibition of the proteasome with MG132 rescued the 2-BP-induced degradation of pUL51-Flag (Fig 8D), whereas CQ treatment had no significant effect. To directly examine ubiquitination levels, we cotransfected DEFs with a ubiquitin-expressing plasmid (Ub-HA) along with either pUL51-Flag or the palmitoylation-deficient pUL51/C9A-Flag mutant. Immunoprecipitation assays revealed that compared with its wild-type counterpart, the pUL51/C9A-Flag mutant exhibited markedly increased ubiquitination (Fig 8E). Finally, to confirm the dependence on the proteasome for accelerated degradation, we performed cycloheximide (CHX) chase assays on cells expressing the pUL51/C9A-Flag mutant in the presence or absence of MG132. The inhibition of the proteasome pathway significantly delayed the degradation of the mutant protein and prolonged its half-life (Fig 8F).
Duck DHHC9 and DHHC18 stabilize the pUL51 protein through palmitoylation
Protein palmitoylation is catalyzed by a family of DHHC domain-containing palmitoyltransferases. In this study, we cloned nine duck-derived DHHC genes and preliminarily screened their interactions with pUL51. Co-IP assays demonstrated that all nine palmitoyltransferases interact with pUL51 (Fig 9A). Further immunofluorescence staining revealed that DHHC2, DHHC9, DHHC14, and DHHC18 colocalized with pUL51 in the cytoplasm (Fig 9B). To investigate whether these enzymes affect pUL51 protein expression, we cotransfected cells with increasing amounts of DHHC2, DHHC9, DHHC14, or DHHC18 plasmids together with pUL51. Western blot analysis revealed that none of these palmitoyltransferases significantly increased pUL51 protein expression (Fig 9C–9F), suggesting that the interaction between palmitoyltransferases and pUL51 is not dose dependent. We hypothesized that these DHHC enzymes may regulate pUL51 stability via palmitoylation. To test this hypothesis, we examined the half-life of pUL51 in CHX-treated DEFs upon overexpression of each of the four DHHCs. The overexpression of DHHC9 and DHHC18 significantly extended the half-life of pUL51 and promoted its accumulation, whereas DHHC2 and DHHC14 did not have a significant stabilizing effect (Fig 9G–9J). To investigate the subcellular localization of DHHC9 and DHHC18, we examined their colocalization with the Golgi marker GM130, given that protein palmitoylation typically occurs in the endoplasmic reticulum or Golgi apparatus. Our results revealed that both DHHC9 and DHHC18 strongly colocalized with GM130 (Fig 9K). Furthermore, to directly determine whether DHHC9 and DHHC18 mediate pUL51 palmitoylation, we cotransfected cells with DHHC9-Flag or DHHC18-Flag along with pUL51-HA and measured palmitoylation levels using acyl-biotin exchange assays. We found that DHHC9 and DHHC18 significantly promoted the palmitoylation of pUL51-HA (Fig 9L). Next, we synthesized short hairpin RNA (shRNA) expression plasmids targeting DHHC9 and DHHC18 to knockdown their gene expression (S2K and S2L Fig). From these, we selected the effective targets shRNA-DHHC9–687 and shRNA-DHHC18–1019 for subsequent functional studies. We subsequently expressed pUL51 in cells in which DHHC9 or DHHC18 was knocked down and assessed its palmitoylation levels via acyl-biotin exchange assays. Compared with those in the shRNA-NC control group, pUL51 palmitoylation signals were significantly reduced following DHHC9 or DHHC18 knockdown (Fig 9M), and viral replication was also impaired (S2M Fig). In summary, the results of this study demonstrate that pUL51 interacts with multiple palmitoyltransferases, among which DHHC9 and DHHC18 promote the palmitoylation of pUL51, thereby increasing its stability. These findings provide crucial evidence of the regulatory mechanisms of pUL51 in viral replication.
The palmitoyltransferases DHHC9 and DHHC18 promote the palmitoylation of pUL51.(A) DEFs were cotransfected with plasmids encoding pUL51-HA and Flag-tagged DHHC proteins for 36 hours. The cell lysates were subjected to immunoprecipitation followed by Western blot analysis. (B) DEFs were cotransfected with pUL51-HA and DHHC-Flag plasmids for 24 hours and then fixed and immunostained. The colocalization of the pUL51-HA and DHHC-Flag proteins was observed using confocal microscopy. Scale bar: 20 μm. (C–F) DEFs were transfected with different concentrations of plasmids encoding DHHC2, DHHC9, DHHC14, or DHHC18, along with 1 μg of the pCAGGS-UL51-3xHA plasmid. Samples were harvested 36 hours post-transfection for Western blot analysis of protein expression. (G–J) DEFs were cotransfected with plasmids encoding DHHC2, DHHC9, DHHC14, or DHHC18 together with 1 μg of the pCAGGS-UL51-3xHA plasmid for 24 hours. Cycloheximide (CHX, 100 mg/mL) was then added, and cell lysates were collected at 0, 2, 4, 6, and 8 hours post-treatment for Western blot analysis. The band intensity of pUL51-HA was quantified using ImageJ software. (K) Subcellular localization of DHHC9 and DHHC18. DEFs were transfected with plasmids expressing DHHC9-Flag and DHHC18-Flag for 24 h. Localization was assessed by immunofluorescence using an antibody against the Golgi marker GM130. The nuclei were counterstained with DAPI, representative images from three independent replicate experiments. Scale bar: 20 μm. (L) DEFs were cotransfected with plasmids encoding DHHC9 or DHHC18 along with the pUL51-HA expression plasmid for 36 hours. Cell lysates were immunoprecipitated using mouse anti-HA magnetic beads. The samples were analyzed by Western blot to detect palmitoylation with or without hydroxylamine (HAM) treatment. (M) DEFs were cotransfected for 36 hours with a pUL51-Flag plasmid along with shRNA plasmids (shRNA-NC, shRNA-DHHC9-687, or shRNA-DHHC18-1019). Cell lysates were immunoprecipitated using mouse anti-Flag magnetic beads. The samples were analyzed by Western blot to detect palmitoylation with or without hydroxylamine (HAM) treatment.
Inhibition of pUL51 palmitoylation by C9A mutation impairs AnHV-1 replication in vitro
To investigate the role of pUL51 palmitoylation in AnHV-1 replication, we generated a recombinant AnHV-1 mutant (AnHV-1-UL51_C9A_) in which the cysteine at position 9 of pUL51 was substituted with alanine (S3A Fig). Both the palmitoylation-site mutant virus and its revertant, AnHV-1-UL51_C9A_ Rev, were successfully rescued. Sequencing and Western blot analyses confirmed the introduction of the pUL51/C9A mutation in AnHV-1-UL51_C9A_ and the successful expression of pUL51 in the revertant virus (S3B and S3C Fig). To further analyze the impact of the mutations on viral replication, DEFs were infected with AnHV-1-UL51_C9A_, AnHV-1-UL51_C9A_ Rev, AnHV-1-CHv50, and 2-BP-treated AnHV-1-CHv50 at the same MOIs. Viral replication kinetics were assessed using a 50% tissue culture infectious dose (TCID₅₀) assay. The results revealed that between 12 and 72 hours post-infection, AnHV-1-UL51_C9A_ viral titers were significantly lower than those of the parental virus and the revertant virus. Similarly, compared with no treatment, AnHV-1-CHv50,2-BP-treated AnHV-1-CHv50 resulted in lower replication levels (Fig 10A and 10B). These findings indicate that inhibiting pUL51 palmitoylation impairs AnHV-1 replication in vitro. Furthermore, analysis of lesion size in DEFs revealed that plaques formed by 2-BP-treated AnHV-1-CHv50 and AnHV-1-UL51_C9A_ were 2-fold and 3-fold smaller, respectively, than those produced by untreated AnHV-1-CHv50 and AnHV-1-UL51_C9A_ Rev (Fig 10C), reinforcing the conclusion that pUL51 palmitoylation is critical for efficient viral replication. To confirm the critical role of the C9 site in palmitoylation and subcellular localization, we assessed the palmitoylation status of pUL51 in DEFs infected with AnHV-1-UL51_C9A_ using Western blot analysis. We found that the palmitoylated band disappeared for the mutant virus, whereas this modification remained detectable for the wild-type virus (Fig 10D). Immunofluorescence staining further demonstrated that pUL51 no longer colocalized with GM130 in AnHV-1-UL51_C9A_-infected cells (Fig 10E). Together, these data demonstrate that cysteine 9 is essential for pUL51 palmitoylation and its targeting to the Golgi apparatus.
*Effect of pUL51/C9A mutation on AnHV-1 replication.(A) Multistep growth kinetics of AnHV-1-UL51C9A. DEFs were infected with AnHV-1-UL51C9A, AnHV-1-UL51C9A and AnHV-1-CHv50 at an MOI of 0.01. Total cell samples were collected at the indicated time points, and viral titers were determined. (B) Statistical analysis of viral titers at each time point. (C) Effect of the C9A mutation on viral lesion size. Viral spread was assessed by crystal violet staining. Thirty representative plaques were measured per group. Plaque diameters were quantified using ImageJ software. (D) DEFs were infected with AnHV-1-CHv50 or AnHV-1-UL51C9A at an MOI of 0.1 for 36 h. The cell lysates were immunoprecipitated using a rabbit anti-UL51 polyclonal antibody and analyzed by Western blot to detect pUL51 palmitoylation with or without hydroxylamine (HAM) treatment. (E) DEFs were infected with AnHV-1-CHv50, AnHV-1-UL51C9A, or AnHV-1-UL51C9A Rev at an MOI of 0.1 for 36 h and then fixed and immunostained with an antibody against GM130 (a Golgi marker). The nuclei were counterstained with DAPI. Scale bar: 20 μm. Data are expressed as mean ± SD from three independent biological replicates. Statistical analysis was performed using two-way ANOVA (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ***p < 0.0001).
Palmitoylation of pUL51 is essential for secondary envelopment and efficient virion production
To investigate the role of pUL51 in viral assembly, we evaluated the formation of mature virions in DEFs by transmission electron microscopy (TEM). No significant differences in the organization of the nucleus or viral replication centers were observed between wild-type and mutant infections. In stark contrast, significant differences between the parental virus and UL51-deficient viruses emerged in the cytoplasm. The most noteworthy differences are the presence of mature cytoplasmic virions (Fig 11A-11D), and the appearance of multivesicular bodies (MVBs) (Fig 11E-11H). In cells infected with the parental or revertant viruses, cytoplasmic viral particles at various stages of envelope formation, including numerous mature particles with intact envelopes, were observed. Notably, these mature particles were frequently enclosed within multivesicular bodies (MVBs) (Fig 11E, yellow triangles). Quantitative analysis revealed that in parental virus-infected cells, 27.87% of cytoplasmic viral particles had intact envelopes, and 31.08% were released extracellularly. In contrast, in AnHV-1-ΔUL51-infected cells, 54.76% of cytoplasmic particles exhibited incomplete secondary envelopment, only 10.2% possessed intact envelopes, and the proportion of released particles was drastically reduced to 1.02%. To determine whether pUL51 palmitoylation is involved in this process, we analyzed cells infected with the palmitoylation-site mutant AnHV-1-UL51_C9A_. Its phenotype resembled that of the deletion mutant, with 43.28% of cytoplasmic particles showing incomplete envelopes (Table 1 and Fig 11I). Although MVBs were present in all infection groups, significantly fewer viral particles were localized within MVBs in cells infected with AnHV-1-ΔUL51 or AnHV-1-UL51_C9A_ (Fig 11E-11H), indicating that pUL51 is required for the efficient recruitment of viral components into MVBs. In summary, pUL51 is essential for the formation of the secondary viral envelope, the production of intact viral particles, and their release, with its palmitoylation playing a crucial role. Furthermore, pUL51 likely facilitates viral assembly and maturation by directing viral components into MVBs.
Table 1: Effects of mutations in UL51 on the distribution of viral particles in AnHV-1-infected DEFs.
Electron microscopy analysis of AnHV-1-CHv50-, AnHV-1-ΔUL51-, AnHV-1-UL51C9A- and AnHV-1-UL51C9A Rev-infected DEFs.(A–D) Electron microscopy images of DEFs infected with AnHV-1-CHv50, AnHV-1-ΔUL51, AnHV-1-UL51C9A and AnHV-1-UL51C9A Rev are presented. (D–G) Close-up image of the enlarged area showing virions in the cytoplasm of AnHV-1-CHv50-, AnHV-1-ΔUL51-, AnHV-1-UL51C9A- and AnHV-1-UL51C9A Rev-infected cells. Red solid arrows indicate viral particles located within vesicles or those that are mature. Red hollow arrows point to viral particles with incomplete envelopes. The black hollow arrows highlight viral particles lacking a secondary envelope. Yellow triangles indicate MVB. The nucleus (N) and cytoplasm (C) are marked. Bars, 1 μm (A-D) and 500 nm (E-H). (I) A total of 200 to 300 viral particles were counted across five cells. The figure shows the percentage of viral particles at the morphogenesis stage of infection.
Inhibiting pUL51 palmitoylation by C9A mutation attenuates AnHV-1 virulence in ducks
To investigate the effect of inhibiting pUL51 palmitoylation on the pathogenicity of AnHV-1 in vivo, 14-day-old ducks were infected with serial doses (10⁷, 10⁶, and 10⁵ TCID₅₀) of the mutant AnHV-1-UL51_C9A_, its revertant AnHV-1-UL51_C9A_ Rev, or the parental AnHV-1-CHv50. Body temperature, weight changes, and mortality were recorded daily for 10 consecutive days. The results revealed that all the ducks in the mock-infected control group survived and maintained normal body temperatures and body weights. Among ducks infected with AnHV-1-UL51_C9A_, only those inoculated with the highest dose (10⁷ TCID₅₀) had increased body temperatures and exhibited a survival rate of 80%. In contrast, ducks infected with 10⁶ or 10⁵ TCID₅₀ of AnHV-1-UL51_C9A_ showed no signs of fever or reduced body weight, and all survived. Ducks in the AnHV-1-UL51_C9A_ Rev and AnHV-1-CHv50 infection groups exhibited sustained elevated body temperatures (>42.5°C) starting at 2 days post-infection, reduced body weights, and dose-dependent mortality. Specifically, survival rates in the AnHV-1-UL51_C9A_ Rev group at doses of 10⁷, 10⁶, and 10⁵ TCID₅₀ doses were 30%, 30%, and 40%, respectively, whereas those in the AnHV-1-CHv50 group were 0%, 0%, and 30%, respectively (S4A-S4C Fig). These results indicate that the palmitoylation defect in pUL51 significantly reduces the pathogenicity of AnHV-1 in ducks. To further evaluate the impact of the pUL51/C9A mutation on viral replication and virulence in vivo, ducklings were infected with 10⁶ TCID₅₀ of the respective viruses. Necropsies were conducted at 3, 5, and 10 days post-infection to assess organ pathology and tissue viral loads. No mortality occurred in either the MEM or AnHV-1-UL51_C9A_ groups, and ducks in these groups exhibited no symptoms. In contrast, ducks infected with AnHV-1-UL51_C9A_ Rev or AnHV-1-CHv50 exhibited characteristic clinical signs, including lacrimation, periorbital wetness, and ocular discharge (Fig 12A). Gross pathological examination revealed severe lesions—including hemorrhages in multiple organs, thymic atrophy, and detachment of the intestinal mucosa—in ducks infected with AnHV-1-UL51_C9A_ Rev or AnHV-1-CHv50 (black arrow in the diagram). In contrast, no significant pathological changes were observed in the MEM or AnHV-1-UL51_C9A_ groups (Fig 12B). Quantitative PCR (qPCR) analysis of tissue viral loads revealed that compared with the AnHV-1-UL51_C9A_ Rev and AnHV-1-CHv50 groups, ducks infected with AnHV-1-UL51_C9A_ had significantly lower viral loads in the heart, liver, spleen, glandular stomach, thymus, bursa of Fabricius, duodenum, and cecum at all time points examined (Fig 12C). These results indicate that the pUL51/C9A mutation impairs AnHV-1 replication in vivo.
*Effect of the pUL51/C9A mutation on AnHV-1 virulence.(A) Clinical signs in the heads of ducklings were observed following infection with different viruses. (B) Histopathological examination of tissues from 14-day-old ducks intramuscularly inoculated with 1 mL (10⁶ TCID₅₀) of AnHV-1-UL51C9A, AnHV-1-UL51C9A Rev, AnHV-1-CHv50, or MEM. Necropsy was performed on day 5 post infection to assess lesions in various tissues. These images were taken by the authors of this manuscript and edited manually (licensed under CC BY 4.0). (C) Replication kinetics of the pUL51/C9A mutant virus in ducks. 14-day-old ducks (n = 9) were intramuscularly inoculated with 1 mL (10⁶ TCID₅₀) of AnHV-1-UL51C9A, AnHV-1-UL51C9A Rev, or AnHV-1-CHv50. Heart, liver, spleen, gizzard, bursa of Fabricius, duodenum, cecum, and thymus tissues were collected on days 3, 5, and 10 post infection. Viral DNA copy numbers were quantified by quantitative PCR. Data are expressed as mean ± SD from three independent biological replicates. Statistical analysis was performed using two-way ANOVA (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ***p < 0.0001).
Discussion
The UL51 gene encodes pUL51, the tegument protein of alphaherpesvirus. Although homologous proteins are present in all herpesviruses, their pathogenic mechanisms vary across different viruses. For example, pUL51 influences the pathogenicity of HSV-1 during intracranial infection in mice, with phosphorylation at Ser184 being critical for high virulence following ocular infection [36]. Moreover, the fusion of a TAP tag to the C-terminus of UL51 impairs the pathogenic function of MDV in vivo [37]. Conversely, the ORF7 protein of VZV has been identified as a neurovirulence factor that plays a key role in determining the neurotropism of VZV [38]. In this study, we investigated the role of pUL51 in the virulence and pathogenicity of viruses. We found that the UL51-deficient strain exhibited a significantly reduced replication capacity in ducklings and failed to induce pathogenicity, confirming that UL51 is an important virulence gene in AnHV-1. To define the role of pUL51 in AnHV-1 pathogenesis, we performed a functional analysis of a pUL51 deletion mutant. Following pUL51 deletion, AnHV-1 retained the ability to replicate in DEFs, indicating that pUL51 is nonessential for viral growth within cells. pUL51 does not affect viral adsorption, invasion, or genomic DNA replication, but it promotes the formation of the secondary viral envelope, the production of intact viral particles, and their release. Thus, pUL51 contributes to efficient viral replication in vitro. Although pUL51 is not essential for viral replication, its critical role in cell-to-cell spread (CCS) has been demonstrated in HSV-1, PRV, and BoHV-1. In this study, compared with the parental and revertant viruses, the deletion of UL51 reduced AnHV-1 plaque size by approximately 69%, suggesting that pUL51 promotes AnHV-1 cell-to-cell spread. Although reduced plaque size does not exclusively indicate cell-to-cell spread, this phenotype, combined with the conserved role of pUL51 in cell-to-cell spread in other herpesviruses, strongly supports its involvement in this process. This function is typically associated with the conserved N-terminal YXXΦ motif, which is also present in AnHV-1 pUL51 [39]. Notably, the YXXΦ motif is widely found in various viral proteins, including the cytoplasmic domains of gE and gB [40], suggesting possible functional redundancy in mediating viral cell-to-cell spread.
Virus‒host and virus‒virus protein interactions are essential for successful viral infection, highlighting the importance of elucidating these interaction networks to identify potential antiviral targets. Previous studies have indicated that pUL51 and its homologs are involved in the secondary envelopment of herpesviruses, although the precise underlying molecular mechanisms remain incompletely understood. In HSV-1, pUL51 interacts with several viral proteins, including pUL7, pUL14, and gE [7,10]. Notably, pUL51 forms a complex with pUL7 that is critical for viral assembly [7]. Additionally, its interaction with pUL14 contributes to the regulation of secondary envelopment in HSV-1 [41]. To characterize the protein interaction network of pUL51 during infection, we performed immunoprecipitation followed by liquid chromatography‒tandem mass spectrometry (LC‒MS/MS) using AnHV-1-infected DEFs. This analysis revealed four novel viral protein interactors of pUL51: pUL10, pUL27, pUL30, and pUL38. Among these genes, pUL10 encodes the envelope glycoprotein gM, which is highly conserved among herpesviruses and plays a critical role in membrane fusion [42], viral assembly, cell-to-cell spread [43], and immune evasion [44,45]. This study revealed that the interaction between pUL51 and pUL10 depends on the palmitoylation of pUL51. Acyl-biotin exchange (ABE) experiments confirmed that cysteine 9 (C9) is the key palmitoylation site. When this site was mutated (pUL51/C9A), pUL51 lost its Golgi localization, failed to interact with pUL10, and could not prevent pUL10 degradation, indicating that pUL51 anchors to specific membrane structures (such as the Golgi apparatus) via palmitoylation, potentially providing a stable platform for pUL10. We propose that pUL51 maintains pUL10 stability by promoting its correct folding and oligomerization [46], thereby ensuring an adequate supply of this critical envelope glycoprotein. The interactions of pUL51 with pUL27 (gB), pUL30, and pUL38 are independent of its palmitoylation, as the C9A mutant retains binding to all three proteins. Notably, pUL51 directly binds to gB, the key glycoprotein for viral adsorption, membrane fusion, and cell-to-cell spread [47–49]. Given the reduced plaque size observed upon UL51 deletion or C9A mutation, we hypothesize that this interaction may enhance the function of gB in intercellular transmission. pUL30 (DNA polymerase) [50] and pUL38 (a nucleocytoplasmic shuttling protein) [51] are localized primarily in the nucleus or perinuclear region and are not significantly colocalized with cytoplasmic pUL51, suggesting that their interactions are transient. We hypothesize that during late replication, pUL51 may recruit these nuclear proteins to cytoplasmic assembly sites, ensuring their targeted delivery to the viral assembly site rather than cytoplasmic diffusion. The specific structural basis for the interactions between pUL51 and pUL27, pUL30, and pUL38, as well as their precise dynamics throughout the infection cycle, remain to be further investigated.
Transmission electron microscopy revealed that deletion of pUL51 in AnHV-1 resulted in abnormal accumulation of immature viral particles in the cytoplasm, accompanied by a reduction in the number of viral particles within the vesicles. Similarly, following infection with the pUL51 C9A mutant virus, the cytoplasm predominantly contained virions with incomplete envelope coverage. These findings suggest that loss of pUL51 palmitoylation impairs secondary envelope formation by affecting the targeted transport of both pUL51 itself and pUL10 to the Golgi apparatus, ultimately hindering viral particle maturation and release. Furthermore, transmission electron microscopy revealed a notable observation: cells infected with either the parental or the revertant virus contained numerous viral particles enclosed within multivesicular body (MVB)-like structures. MVBs are large vesicles formed by the fusion of endosomes and other secretory vesicles. MVBs have important functions in cells: the fusion of MVBs with lysosomes results in the degradation of MVB contents; MVBs serve as platforms for intracellular signaling, and MVB trafficking to and fusion at the cell surface results in the secretion of exosomes that may influence intracellular signaling [52,53]. The fate of viral particles localized to MVBs is not yet fully understood. While the fusion of MVBs with lysosomes could represent a terminal degradative pathway for the virus, MVBs may also serve as vehicles for transport out of the cell via exocytosis [54,55]. The hijacking of MVBs by viruses constitutes an important route of entry and egress for enveloped viruses and has been shown to be a mechanism through which viruses remodel exosomes to alter intercellular communication and inflammation [54,56–58]. Within the herpesvirus family, ESCRT components and MVBs are involved in the maturation and egress of herpes simplex virus 1 (HSV-1) [59,60]. Specifically, ESCRT-III promotes HSV-1 primary envelopment by mediating scission during HSV-1 budding through the INM [61], and secondary envelopment of HSV-1 requires VPS-4 [62], a component of the ESCRT-III machinery essential for generating intraluminal vesicles within MVBs [60,63]. Similarly, HCMV and human herpesvirus 6 (HHV-6) virions have been observed within MVBs [64], and HCMV replication depends on the ESCRT proteins VPS-4 and CHIMP-1 [65]. MVBs and the ESCRT machinery are associated with numerous cellular processes, such as signaling, membrane and lipid turnover, and cellular proliferation and polarization [66]. These studies collectively underscore the central importance of MVBs in virus envelopment and egress. In this study, compared with cells infected with the parental virus, ΔUL51-infected cells exhibited a marked reduction in multivesicular body (MVB) structures or displayed morphologically abnormal MVBs, which also contained fewer encapsidated viral particles. These observations suggest that pUL51 may play a role in regulating the host endosomal sorting and transport pathway. It has been reported that the N-terminal region of the UL51 protein can form filamentous structures in vitro, adopting a conformation similar to that of the ESCRT-III component CHMP4B and assembling into filaments reminiscent of ESCRT-III polymers [5]. In HCMV, residues 300–325 of pUL71, the homologous protein of pUL51, directly bind to the MIT domain of VPS4A. This motif plays a crucial role in recruiting VPS4A to the HCMV assembly compartment [12]. Our study demonstrated that pUL51 is localized to the Golgi apparatus through palmitoylation, which serves as a primary source of lipids and proteins for multivesicular body (MVB) membranes. Additionally, pUL51 interacts with and regulates the transport of multiple viral proteins, facilitating their enrichment on MVB membranes. We hypothesize that pUL51 may promote or modify MVB biogenesis, provide a membrane platform for viral assembly, or function as a molecular adapter that recruits immature viral particles or other components from the cytoplasm into MVBs.
S-Palmitoylation is a reversible, enzyme-catalyzed posttranslational modification that involves the formation of a thioester bond between a palmitoyl group and a cysteine residue. This modification plays critical roles in regulating protein trafficking, subcellular localization, protein stability, and protein–protein interactions [21,67]. We first investigated the effect of palmitoylation on the stability of pUL51. Treatment with the broad-spectrum palmitoylation inhibitor 2-bromopalmitate (2-BP) reduced the expression stability of pUL51 and shortened its half-life. Similarly, the half-life of the exogenously expressed palmitoylation-deficient mutant pUL51/C9A was significantly shorter than that of wild-type pUL51. These results indicate that palmitoylation at the C9 site is critical for maintaining pUL51 stability because it prevents its premature degradation, thereby ensuring normal protein function. Previous studies have reported that pUL51 in other herpesviruses is localized to the Golgi apparatus. Here, AnHV-1 pUL51 was also localized to the Golgi, whereas the C9A mutant lost this localization, demonstrating that palmitoylation is essential for its ability to target the Golgi. Palmitoylation has also been reported for pUL51 in HSV-1 and KSHV [8,15], and pUL51 and its homologs localize to the Golgi in multiple herpesviruses, including HSV-1, PRV, VZV, HCMV, and KSHV [4,14,68]. These findings suggest that palmitoylation is a highly conserved posttranslational modification of herpesvirus pUL51 proteins and plays a crucial role in their Golgi localization. Therefore, further investigations into the mechanism underlying pUL51 palmitoylation are warranted.
Protein S-palmitoylation relies on the palmitoyltransferase family in host cells for catalysis [69]. To this end, we cloned a series of duck-derived DHHC enzymes and determined that DHHC9 and DHHC18 are capable of stabilizing pUL51 expression and promoting palmitoylation of this protein. The palmitoylation of viral proteins typically occurs in the endoplasmic reticulum or Golgi apparatus [30,70]. The Golgi localization of pUL51, its colocalization with DHHC9 and DHHC18, is therefore highly significant. In our experiments, we observed that multiple palmoyltransferases interact with AnHV-1 pUL51. The overexpression or knockdown of DHHC9 or DHHC18 resulted in varying degrees of increase or decrease in the palmitoylation band of pUL51, indicating that DHHC9 and DHHC18 are the primary enzymes that mediate pUL51 palmitoylation. Although other DHHC enzymes interact with pUL51, they fail to provide effective functional compensation. Further screening among interacting DHHC enzymes revealed that DHHC9 and DHHC18 not only colocalize with pUL51 in the Golgi apparatus but also maintain its stability and promote its palmitoylation. These two enzymes synergistically support the functional activity of pUL51. DHHC enzymes play diverse regulatory roles in both host and viral proteins; for instance, DHHC9 has been reported to participate in the palmitoylation of the SARS-CoV-2 Spike protein, interacting with it in the endoplasmic reticulum or Golgi apparatus to promote palmitoylation, thereby promoting viral membrane fusion and infectivity [71]. DHHC18 catalyzes the palmitoylation of cGAS, inducing conformational changes that inhibit the binding of cGAS to DNA and its dimerization process [72]. These observations demonstrate that DHHC enzymes can modulate a wide range of host and viral proteins, highlighting their potential as targets for antiviral strategies. Additionally, the palmitoylation of some host proteins is increased by the synergistic action of multiple DHHC enzymes. For example, more than half of the DHHC family members can increase IFITM3 palmitoylation, with DHHC3, DHHC7, DHHC15, and DHHC20 exerting the strongest effects [73]. In another case, DHHC3, DHHC7, and DHHC17 act together to palmitoylate SNAP25, thereby promoting its membrane localization [74]. This cooperative modification pattern is also observed for viral proteins. The NSP1 protein of Chikungunya virus is palmitoylated by DHHC2 and DHHC19 [32], while HSV-1 UL20 and HIV-1 Tat are modified by DHHC3 and DHHC20, respectively [34,35]. The precise mechanism by which protein palmitoylation affects viral infection remains elusive. Studies have indicated that depalmitoylation inhibits the secretion of infectious viral particles and significantly reduces viral infectivity, suggesting that palmitoylation plays a critical role in viral assembly and infection [75,76]. In this study, treatment with the inhibitor 2-bromopalmitate (2-BP) reduced viral replication and decreased plaque size by approximately 50%. Furthermore, infection with the pUL51/C9A mutant virus AnHV-1-UL51_C9A_ resulted in a further decrease in replication capacity, with plaque size reduced by approximately 75%. In vivo infection of ducklings with AnHV-1-UL51_C9A_ demonstrated that this mutation attenuated AnHV-1 pathogenicity by inhibiting pUL51 palmitoylation. These findings indicate that palmitoylation at the C9 residue of pUL51 is critical for both the in vitro replication efficiency and the in vivo virulence of AnHV-1. Thus, this study confirms that palmitoylation at the C9 residue affects viral replication capacity in vitro and in vivo and provides a molecular mechanism underlying the reduced virulence of AnHV-1 caused by pUL51 deletion. Furthermore, this work is the first systematic elucidation of the palmitoylation mechanism and function of pUL51 in α-herpesviruses.
In summary, the UL51 gene is a viral pathogenicity gene. Palmitoylation mediated by the palmitoyltransferases DHHC9 and DHHC18 increases the pathogenicity of the virus. This modification not only influences the interaction between pUL51 and the viral envelope glycoprotein pUL10 but also contributes to maintaining the stability of pUL51 and its targeted localization to the Golgi apparatus. Collectively, these functions facilitate viral replication, secondary envelopment, infectious virion production, and cell-to-cell spread, ultimately enhancing viral proliferation and pathogenicity in vivo.
Materials and methods
Ethical statement
All animal experiments were approved by the Animal Welfare Committee of Sichuan Agricultural University (Protocol Permit Number: 20240638) and were performed in accordance with relevant guidelines and regulations.
Cells and viruses
Nine-day-old nonimmune-fertilized duck embryos and 14-day-old healthy ducklings were purchased in Ya ‘an city, Sichuan Province. All the experimental ducks were free of AnHV-1, and AnHV-1 antibodies were negative.
Duck embryo fibroblasts (DEFs) were prepared from 9-day-old duck embryos. Following removal of the head and internal organs, the embryos were minced and digested with 0.25% trypsin. The resulting cell suspension was then centrifuged at 4,500 × g for 5 min, and the pellet was resuspended and cultured in Dulbecco’s modified Eagle’s medium (DMEM; Gibco, Shanghai, China) supplemented with 10% newborn calf serum (NBCS; Gibco, Shanghai, China). The cells were maintained at 37°C in a 5% CO₂ atmosphere.
Antibodies and other reagents
The commercial antibodies used in this study included mouse anti-Flag monoclonal antibody (1:5,000; Cat: M185-3S) and mouse anti-HA monoclonal antibody (1:5,000; Cat: M132-3), which were purchased from Medical & Biological Laboratories (USA); mouse anti-GOLGA2/GM130 monoclonal antibody (1:250; Cat: 66662–1-Ig) and rabbit anti-β-actin monoclonal antibody (1:5,000; Cat: 20536–1-AP) were purchased from Proteintech; and rabbit anti-Flag monoclonal antibody (1:5,000; Cat: AE092), rabbit anti-HA monoclonal antibody (1:5,000; Cat: AE105), HRP-labeled goat anti-rabbit IgG (H + L) (1:5,000; Cat: AS014), and HRP-labeled goat anti-mouse IgG (H + L) (1:5,000; Cat: AS003) were purchased from ABclonal Technology. Alexa Fluor 568-labeled goat anti-mouse IgG antibody (1:1,000; Cat: A11004) and Alexa Fluor 488-labeled goat anti-rabbit IgG antibody (1:1,000; Cat: A11008) were purchased from Thermo Fisher Scientific (USA). Mouse IgG (Cat A7028) and the Rapid Silver Stain Kit (Cat P0017S) were obtained from Beyotime Biotechnology. Protein A/G magnetic beads (Cat HY-K0202) were obtained from MedChemExpress. Rabbit polyclonal antibodies against the AnHV-1 proteins UL51, UL50, UL52, ICP8, UL10, UL23, UL48, UL47, UL27 and UL30 were generated and aliquoted in our laboratory. Methylcellulose (Cat M6385) and 2-bromopalmitate (2-BP, Cat 21604-1G) were obtained from Sigma‒Aldrich. Cycloheximide (CHX, Cat S7418) was purchased from Selleck Chemicals. All the compounds were dissolved in 0.1% DMSO for experimental use. A palmitoylation-specific immunoblot kit with HRP detection (Cat AM10314) was acquired from AIMSMASS.
Strains and viruses
The 50th generation cell passage virus Anatid herpesvirus 1 Chinese virulent strain (AnHV-1-CHv) (GenBank: NO. JQ647509.1) and its bacterial artificial chromosome infectious clone pAnHV-1-CHv50 and strain pEP-Kan-S of AnHV-1 were preserved and provided by the Institute of Animal Medicine Immunology, Sichuan Agricultural University.
Plasmids
The recombinant plasmids pCAGGS-UL10-3xHA, pCAGGS-UL27-3xHA, pCAGGS-UL30-3xHA, pCAGGS-UL38-3xHA, pCAGGS, and Ub-HA were maintained and provided by the Institute of Animal Medicine Immunology, Sichuan Agricultural University. UL51 and UL51 mutant gene fragments were amplified from the AnHV-1-CHv50 strain and cloned and inserted into the pCAGGS vector linearized with EcoR I and Kpn I restriction enzymes. The DHHC gene fragment was obtained by RT‒PCR using total RNA extracted from DEFs as the template, and the resulting cDNA was subsequently cloned and inserted into the pCAGGS vector. All the primers used in this study were synthesized by Tsingke Biotechnology Co., Ltd., and their sequences are provided in S1 Table.
Construction and rescue of recombinant viruses
In this study, we used a bacterial artificial chromosome (BAC)-based gene editing platform for recombinant anatid herpesvirus 1. Using the scarless recombination system in Escherichia coli ΔMiniF-GS1783, we constructed the AnHV-1-△UL51 and AnHV-1-△UL51-Rev viruses [34], as shown in Fig 1A. To generate the infectious clone of AnHV-1-△UL51, a ΔUL51-Kan fragment was amplified with the primers ΔUL51-F/R using the pEP-Kan-S plasmid as a template. This fragment contained a 40 bp upstream homologous arm (derived from the left side of the UL51 gene) and a 40 bp downstream homologous arm (derived from both the left and right sides of the UL51 gene). The kanamycin (Kan) resistance cassette included an I-SceI endonuclease site. The Kan resistance gene replaced the UL51 gene during the first homologous recombination. Subsequently, in vivo cleavage at the I-SceI site and a second homologous recombination cleavage using the 40 bp repeat sequence removed the Kan resistance gene, yielding the infectious cloning plasmid pAnHV-1-△UL51. Their primer sequences are provided in S1 Table. The PCR products containing homologous arms with the UL51 gene or UL51_C9A_ gene, the I-SceI site, the Kan resistance gene, and a 40 bp repeat sequence were electrotransferred into E. coli GS1783 harboring pAnHV-1-ΔUL51. pAnHV-1-ΔUL51-Rev and pAnHV-1-UL51_C9A_ were constructed following the same procedure described above. The recombinant plasmid was verified by PCR amplification using the primers UL51-JD-F and UL51-JD-R and subsequently confirmed by DNA sequencing. To obtain deletion and recovery viruses, we extracted recombinant positive plasmids and transfected DEFs with Hief Trans Liposomal Transfection Reagent (40802ES03; Yeasen, Shanghai, China). After incubation at 37°C for 6 h, the supernatant was discarded, and the medium was replaced with fresh DMEM containing 1% NBCS until many green spots were produced. The cell supernatant was collected and inoculated into new DEF monolayer cells until the green fluorescent spot disappeared and obvious cytopathic changes were observed. The recombinant viruses were ultimately identified by PCR and Western blot analysis.
Virus multistep growth curve
The AnHV-1-CHv50, AnHV-1-∆UL51, and AnHV-1-∆UL51-Rev viruses were inoculated at an MOI of 0.01 onto monolayer cultures of DEFs in 12-well plates. The supernatants containing the virus were harvested at 24, 48, 60, and 72 hours post-infection. Following three freeze‒thaw cycles, 100 µL aliquots from each time point were subjected to tenfold serial dilutions, ranging from 10^-1^ to 10^-8^. Each dilution was inoculated into 96-well plates (100 µL per well), with eight replicates per dilution. After inoculation, the cells were incubated at 37°C under 5% CO₂ for 5–7 days. Cytopathic effects (CPEs) were monitored and recorded daily under a microscope. The 50% tissue culture infectious dose (TCID_50_) was calculated using the Reed–Muench method, and a multistep viral growth curve was generated.
Real-time fluorescence quantitative PCR
Viral genomic DNA was extracted using the Viral Genomic DNA Extraction Kit (Cat: DP304–03; TIANGEN, Beijing, China). The viral DNA copy number was quantified by quantitative PCR (qPCR) targeting the AnHV-1 UL30 gene using a standard curve previously established in our laboratory. The qPCR assays were performed with Premix Ex Taq (Probe qPCR) (Cat: RR390, Takara, Japan) on a Roche LightCycler 96 instrument under the following cycling conditions: initial denaturation at 95°C for 30 sec, followed by 40 cycles of 95°C for 5 sec and 60°C for 30 sec. The copy numbers were determined on the basis of the standard curve. All reactions were run in triplicate. Statistical analyses were performed using GraphPad Prism version 9.5. The sequences of the primers and probes used are provided in S1 Table.
Detection of virus adsorption, invasion, replication, release and plaque size
(1) Adsorption.
Single-layer DEFs were precooled at 4°C for 2 h, after which the cell fragments were removed by washing with PBS that was precooled to 4°C. Cells were infected with a viral dose of 10⁷ copies/0.1 mL (AnHV-1-CHv50, AnHV-1-ΔUL51, AnHV-1-ΔUL51-Rev). Following infection, the cells were incubated at 4°C for 2 hours and then washed five times with prechilled PBS. Cell samples were collected for real-time quantitative PCR to determine viral DNA copy numbers.
(2) Invasion.
Single-layer DEFs were precooled at 4°C for 2 hours and then were washed with precooled PBS to remove cellular debris. The cells were infected with viruses (AnHV-1-CHv50, AnHV-1-∆UL51-Rev, and AnHV-1-∆UL51) at a dosage of 10⁷ copies/0.1 mL. After incubation at 4°C for 2 hours, the supernatant was discarded, and the unadsorbed viruses were removed by washing with precooled PBS. The cells were further incubated at 37°C for 2 hours and washed again with precooled PBS. Genomic DNA was extracted from the collected cell samples, and the number of AnHV-1 genome copies was quantified via qPCR.
(3) Genome replication.
Single-layer DEFs were infected with viruses (AnHV-1-CHv50, AnHV-1-∆UL51-Rev, and AnHV-1-∆UL51) at a dose of 10⁷ copies/0.1 mL and incubated at 37 °C for 6 hours. The supernatant was then discarded and replaced with cell culture maintenance medium. The cells were cultured in a 5% CO₂ incubator at 37°C. Cell samples were collected at 7, 9, 11, and 13 hours after medium replacement. Genomic DNA was extracted, and the number of AnHV-1 genome copies was quantified via qPCR.
(4) Release.
Single-layer DEFs were infected with viruses (AnHV-1-CHv50, AnHV-1-∆UL51-Rev, and AnHV-1-∆UL51) at an MOI of 1 and incubated at 37 °C for 16 hours. The supernatant was then discarded and replaced with cell culture maintenance medium. The cells were cultured in a 5% CO₂ incubator at 37°C. The supernatant and corresponding cell samples were collected at 1, 2, 3, and 4 hours after medium replacement, and viral titers were determined using the TCID₅₀ assay.
(5) Virus plaque size.
Single-layer DEFs were infected with viruses (AnHV-1-CHv50, AnHV-1-∆UL51-Rev, and AnHV-1-∆UL51) at an MOI of 0.0001 and incubated at 37 °C for 2 hours. The supernatant was then discarded and replaced with cell culture maintenance medium containing 1% methylcellulose. The cells were cultured in a 5% CO₂ incubator at 37°C for 5 days. The medium was subsequently discarded, and the cells were washed several times with PBS. Each well was fixed with 1 mL of precooled 4% paraformaldehyde at room temperature for 15 minutes. After the residual paraformaldehyde was removed with PBS, the cells were stained with 0.5% crystal violet solution for 5 minutes. Plates were gently rinsed with tap water to remove excess stain, and plaques were imaged and quantified for size and number. The average plaque size was quantified using ImageJ software. The lesion size of the deficient virus was then calculated and expressed as a percentage relative to that of the parental virus, which was normalized to 100%.
Electron microscopy analysis of recombinant viruses
All the samples to be examined were subjected to transmission electron microscopy as previously described [77]. DEFs were infected with AnHV-1-CHv50, AnHV-1-UL51_C9A_-Rev, AnHV-1-UL51_C9A_, and AnHV-1-∆UL51 at an MOI of 5. At 18 hours post-infection, the cells were harvested by gentle scraping, pelleted via centrifugation at 3,500 × g for 10 minutes, and fixed overnight at 4°C in 2.5% glutaraldehyde. All the samples were sent to Chengdu Lilai Biotechnology Co., Ltd., for analysis by transmission electron microscopy (JEM-1400FLASH, Tokyo, Japan).
Pathogenicity assay
Sixty-four healthy day-old Cherry Valley ducklings were obtained from a breeding facility at Sichuan Agricultural University (Sichuan, China). Prior to the experiment, all the ducklings were confirmed to be free of AnHV-1 infection and tested negative for AnHV-1 antibodies. To evaluate the role of the UL51 gene in AnHV-1 replication and viral pathogenicity in vivo, forty 14-day-old ducklings were randomly allocated into four groups (n = 10 per group). Each group was inoculated intramuscularly with 10⁷ TCID₅₀ of AnHV-1-CHv50, AnHV-1-ΔUL51-Rev, AnHV-1-ΔUL51, or an equivalent volume of MEM (mock infection). Body temperature, body weight, and mortality were recorded daily over a 10-day period. In another experiment, 24 14-day-old ducklings were randomly divided into 4 groups (6 ducklings per group) and injected with the same dose of AnHV-1-CHv50, AnHV-1-ΔUL51-Rev, AnHV-1-ΔUL51 virus or MEM and sacrificed 5 days post-infection. The tissues of each duckling were collected to analyze the lesions and viral loads.
Preparation and analysis of mass spectrometry samples
To identify proteins that interact with pUL51, we performed immunoprecipitation coupled with mass spectrometry analysis. DEFs were cultured at 37°C and 5% CO₂ until they were fully confluent. The cells were then transfected with the pCAGGS-UL51-Flag plasmid. At 24 hours post-transfection, the cells were infected with AnHV-1-CHv50 at an MOI of 1. After 36 hours of infection, the cells were collected, and the supernatant was discarded. The cells were lysed using immunoprecipitation-compatible lysis buffer (Beyotime) and incubated on ice for 30 min, followed by centrifugation at 12,000 × g for 10 min at 4°C. The supernatant was divided equally into two portions. One portion was incubated with mouse anti-Flag monoclonal antibody at a 1:100 dilution, while the other was incubated with mouse IgG (as a negative control) at the same dilution; both were incubated in a shaking incubator at 4°C for 12 hours. Protein A/G magnetic beads were subsequently added at a 1:10 ratio and incubated at 37°C for 1 hour. After five washes with PBS, the immunoprecipitated complexes were resuspended in 80 μL of buffer. A portion of the eluate was analyzed by Western blot to verify the enrichment of target proteins. Differential protein bands were visualized using a rapid silver staining kit. The remaining samples were subjected to mass spectrometry analysis by Oebiotech (Shanghai, China). The raw MS data were processed using Proteome Discoverer software (version 2.5) to identify potential pUL51-interacting proteins.
Coimmunoprecipitation (Co-IP) Analysis
36 hours after plasmid transfection, the cells were lysed using immunoprecipitation lysis buffer, incubated on ice for 30 minutes, and centrifuged at 12,000 × g for 10 minutes at 4°C. The lysates were incubated with mouse anti-HA or anti-Flag monoclonal antibodies at a 1:100 dilution with constant rotation at 4°C for 12 hours. Protein A/G magnetic beads were then added at a 1:10 bead-to-lysate ratio and incubated at 37 °C for 1 hour. After five washes with PBS, the supernatant was discarded, and the beads were resuspended in 40 µL of PBS. Finally, 10 µL of 5 × protein loading buffer was added to the eluted samples for subsequent Western blot analysis.
Western blot
Following viral infection or plasmid transfection, the cells were washed with PBS and lysed using RIPA lysis buffer (Beyotime) supplemented with 1% PMSF (Beyotime). The proteins were separated by SDS‒PAGE and transferred onto PVDF membranes. The membranes were blocked with 5% nonfat milk at 37°C for 3 hours, followed by incubation with primary antibody at 4°C overnight. After being washed three times with Tris-buffered saline containing Tween 20 (TBST), the membranes were incubated with HRP-conjugated secondary antibodies (anti-rabbit IgG or anti-mouse IgG) at 37°C for 1 hour. The membranes were then washed three times with TBST. The protein bands were visualized using an enhanced chemiluminescence (ECL) hypersensitive detection kit (Cat: 36208ES76; Yeasen, China) on a ChemiDoc MP imaging system (Bio-Rad).
Indirect immunofluorescence assay
DEFs were cultured on coverslips and transfected for 24 h. Subsequently, the cells were washed three times with PBS and fixed with 4% paraformaldehyde overnight at 4°C. After fixation, the cells were permeabilized with 0.25% Triton X-100 at 4°C for 30 min and blocked with 5% bovine serum albumin (BSA; Cat: B2064; Sigma‒Aldrich) at 37°C for 3 h. The cells were then incubated with the appropriate primary antibody for 3 h at room temperature, followed by washing and incubation with Alexa Fluor 568-conjugated goat anti-mouse IgG (1:1,000) and Alexa Fluor 488-conjugated goat anti-rabbit IgG (1:1,000) secondary antibodies for 1 h at room temperature. After additional washing, the cell nuclei were stained with 4′,6-diamidino-2-phenylindole (DAPI; Cat: 10236276001; Roche) for 15 min at room temperature. Finally, the coverslips were mounted using 50% glycerol in PBS, and images were acquired with a laser scanning confocal microscope (FV4000; Olympus, Japan).
Cell viability assay
When DEFs reached 80% confluence in 96-well plates, they were treated with different concentrations of the palmitoylation inhibitor 2-BP. Cell viability was evaluated using the Cell Counting Kit-8 (CCK-8; Cat: C0037, Beyotime) following the manufacturer’s protocol. Briefly, CCK-8 solution was added to each well, and the plates were incubated at 37°C for 1 hour. The absorbance at 450 nm was then measured using a microplate reader.
Immunoprecipitation and acyl-biotin exchange assay
pUL51 palmitoylation was determined by immunoprecipitation and acyl-biotin exchange (IP-ABE) assays. Briefly, DEFs were transfected with plasmids expressing pUL51-Flag or pUL51/C9A-Flag for 36 hours. The samples were processed according to the manufacturer’s protocol for the Specific Palmitoylation Immunoblot Kit (Cat: AM10314; AIMSMASS, China), followed by Western blot analysis. Biotinylated palmitoylated proteins were detected using HRP-conjugated streptavidin, while immunoprecipitated UL51-Flag and UL51/C9A-Flag proteins were detected using a rabbit anti-Flag monoclonal antibody.
CHX chase assay
To examine the effect of the palmitoylation inhibitor 2-BP on the stability of pUL51, DEFs were transfected with a plasmid expressing pUL51-Flag. At 12 hours post-transfection, the cells were treated with 30 μM 2-BP. After 12 hours of incubation at 37°C, the culture medium was replaced with fresh medium containing 100 mg/mL cycloheximide (CHX). The cells were further incubated at 37°C, and the lysates were collected at 0, 0.5, 1, 1.5, 2, and 4 hours after CHX addition. Protein expression was analyzed by Western blot. To analyze the effect of the C9A mutation on pUL51 stability, DEFs were transfected with plasmids expressing pUL51-Flag and pUL51/C9A-Flag. At 24 hours post-transfection, the medium was replaced with medium containing 100 mg/mL CHX. The cells were incubated at 37°C, and the lysates were harvested at 0, 2, 4, 6, 8, and 10 hours after CHX treatment. Protein expression was assessed by Western blot. The band intensity of UL51-Flag was quantified using ImageJ software and normalized to that of β-actin. Protein decay curves were generated, and the half-life was calculated using GraphPad Prism version 9.5.
Knockdown of DHHC9 and DHHC18
On the basis of the sequences of DHHC9 (GenBank: XM_038184538.2) and DHHC18 (GenBank: XM_038167258.2), we designed and synthesized a short hairpin RNA expression plasmid (S1 Table), which was synthesized by GenePharma. Short hairpin RNA (shRNA) expression plasmids, along with DHHC9 and DHHC18 eukaryotic expression plasmids, were transfected into DEFs. The protein expression levels of DHHC9 and DHHC18 were detected by Western blot analysis at 36 hours post-transfection.
Statistical analysis
In this paper, the data are expressed as the group mean ± standard deviation (SD) and were analyzed using GraphPad Prism 9.5 via t tests or one-way/two-way ANOVA. Statistical significance thresholds were uniformly set at *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001, with ns denoting nonsignificance (P > 0.05).
Supporting information
S1 FigPredicted structures and molecular docking analysis of pUL51 and pUL51/C9A with pUL10.(A) AlphaFold-predicted structures of wild-type pUL51 (green) and the mutant pUL51/C9A (blue). The site of the C9A mutation is indicated in red. (B) Molecular docking model of wild-type pUL51 (green) with pUL10 (pink). (C) Molecular docking model of the mutant pUL51/C9A (blue) with pUL10 (pink). All the structures were visualized using PyMOL software.(TIF)
S2 FigPalmitoylation modifications affect multiple stages of the AnHV-1 life cycle.(A) Viability of DEFs treated with increasing concentrations of the palmitoylation inhibitor 2-bromopalmitate (2-BP), as determined by a CCK-8 assay after 12 hours. (B) DEF viability after treatment with different concentrations of MG132, as measured by the CCK-8 assay after 6 hours. (C) DEF viability after treatment with different concentrations of CQ, as measured by the CCK-8 assay after 12 hours. (D) DEF viability after treatment with different concentrations of CHX, as measured by a CCK-8 assay after 12 hours. (E) Schematic diagram of the experimental design for assessing the effects of palmitoylation on AnHV-1 adsorption and invasion. (F) Schematic diagram of the experimental design for evaluating the role of palmitoylation in AnHV-1 replication and release. (G, H) Viral adsorption assay: DEFs were precooled to 4°C for 2 h, after which the medium was replaced with fresh DMEM containing 2-BP, DMSO, or AnHV-1 (MOI of 1). After incubation at 4°C for 2 h, the cells were washed three times with ice-cold PBS. For the viral invasion assay, the medium was replaced with 2-BP or DMSO immediately after the adsorption period. The cells were incubated at 37°C for 2 h and then were washed three times with PBS. Viral DNA copy numbers were quantified by qPCR. (I) Replication assay: DEFs were infected with AnHV-1 at an MOI of 1. At 6 h post infection, the medium was replaced with DMEM containing 30 μM 2-BP or DMSO. Cells were harvested at 6, 12, 24, and 36 h postinfection, and viral DNA copies were quantified by qPCR. (J) Release assay: DEFs were infected with AnHV-1 at an MOI of 1 and incubated at 37°C for 16 h. The medium was then replaced with DMEM containing 30 μM 2-bromopalmitate (2-BP) or DMSO. The cells and supernatants were collected at 0.5, 1, 2, and 3 h posttreatment. The ratio of extracellular to intracellular viral titers was used to assess viral release efficiency. (M) To evaluate the knockdown efficiency of DHHC9-targeting shRNAs, DEFs were cotransfected for 36 hours with a DHHC9–3 × Flag expression plasmid alongside shRNA plasmids (shRNA-NC, shRNA-DHHC9–687, shRNA-DHHC9–892, or shRNA-DHHC9–1121). Protein expression levels were then analyzed by Western blot using an anti-Flag antibody. (L) To evaluate the knockdown efficiency of DHHC18-targeting shRNAs, DEFs were cotransfected for 36 hours with a DHHC18–3 × Flag expression plasmid alongside shRNA plasmids (shRNA-NC, shRNA-DHHC18–583, shRNA-DHHC18–1019, or shRNA-DHHC18–1197). Protein expression levels were then analyzed by Western blot using an anti-Flag antibody. (M) DEFs were transfected for 24 hours with shRNA plasmids (shRNA-NC, shRNA-DHHC18–1019, or shRNA-DHHC9–687). The cells were subsequently infected with AnHV-1-CHv50 at an MOI of 1. At the indicated times post infection, the cells were harvested, and viral DNA copies were quantified. The data are presented as the mean ± SD. Significance was determined by one-way or two-way ANOVA (ns, not significant; *p < 0.05; **p < 0.01; ***p < 0.001; ****p < 0.0001). The results were repeated in three independent experiments with three replicates each.(TIF)
S3 FigConstruction and identification of the AnHV-1 pUL51 palmitoylation mutant virus.(A) Schematic representation of the pUL51 palmitoylation-deficient mutant virus AnHV-1-UL51_C9A_. (B) Sequencing results of AnHV-1-UL51_C9A_ and AnHV-1-UL51_C9A_ Rev. (C) Western blot analysis of pUL51 expression. DEFs were infected with AnHV-1-UL51_C9A_, AnHV-1-UL51_C9A_ Rev, or AnHV-1-CHv50 at an MOI of 1 for 36 hours. Mock-infected DEFs were used as a control.(TIF)
S4 FigEffects of pUL51 palmitoylation-modified mutant viruses on the survival rate, body temperature, and body weight of ducks.14-day-old ducks (n = 10 per group) were intramuscularly inoculated with 1 mL of serial dilutions (10⁵ to 10⁷ TCID₅₀) of AnHV-1-UL51_C9A_, AnHV-1-UL51_C9A_ Rev, AnHV-1-CHv50, or MEM (mock control). (A) Survival was monitored daily for 10 days post-inoculation. Survival rates are shown. (B) Body temperature was measured daily over the 10-day period. (C) Body weight was recorded daily for 10 days.(TIF)
S1 TableSequences of primers used in this study for constructing eukaryotic expression plasmids and AnHV-1 recombinant viruses.(DOCX)
S2 TableRaw data.(XLSX)
S1 Raw ImageAll the Figure’s WB original data.(PDF)
S2 Raw ImageRaw data from other images.(PDF)
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