Proteasomal processing of the viral replicase ORF1 facilitates HEV-induced liver fibrosis

Significance

Chronic hepatitis E virus (HEV) infections, particularly those of genotype 3 (G3), frequently lead to liver fibrosis and cirrhosis, yet the precise mechanisms remain elusive. This study aimed to uncover key pathogenic factors and virus–host interactions responsible for HEV-induced liver damage. We identified a “proteasome-produced” viral protein in human and rodent specimens and elucidated its role in HEV-induced liver pathogenesis by potentiating fibrogenic TGF-beta/SMAD signaling. This finding advances our understanding of virus–host interactions in liver diseases, highlighting complex interplays between viral proteins and host signaling pathways and suggesting a potential therapeutic target, emphasizing the importance of host–virus interactions in managing chronic viral diseases.

Abstract

Chronic infections with hepatitis E virus (HEV), especially those of genotype 3 (G3), frequently lead to liver fibrosis and cirrhosis in patients. However, the causation and mechanism of liver fibrosis triggered by chronic HEV infection remain poorly understood. Here, we found that the viral multiple-domain replicase (ORF1) undergoes unique ubiquitin–proteasomal processing leading to formation of the HEV-Derived SMAD Activator (HDSA), a viral polypeptide lacking putative helicase and RNA polymerase domains. The HDSA is stable, non-HSP90-bound, localizes to the nucleus, and is abundant in G3 HEV-infected hepatocytes of various origins. Markedly, the HDSA in hepatocytes potentiates the fibrogenic TGF-β/SMAD pathway by forming compact complexes with SMAD3 to facilitate its promoter binding and coactivator recruitment, leading to significant fibrosis in HEV-susceptible gerbils. Virus infection–induced liver fibrosis in HEV-susceptible gerbils could be prevented by mutating the residues P989C, A990C, and A991C (PAA-3C) within ORF1, which are required for proteasomal processing. Thus, we have identified a viral protein derived from host proteasomal processing, defined its notable role in liver fibrosis and highlighted the nature of an unanticipated host–HEV interaction that facilitates hepatitis E pathogenesis.

Hepatitis E virus (HEV) is one of the leading causes of viral-induced hepatitis in humans, with roughly 20 million infections every year and approximately 60,000 related deaths (1, 2). The major transmission pathway of genotype 1 (G1) HEV in developing countries is the fecal–oral route via virus-contaminated water, which occasionally triggers large HEV outbreaks. Notably, HEV infection causes severe symptoms in pregnant women including fulminant liver failure, hemorrhage, and stillbirth, with a mortality of up to 30% (3, 4). However, sporadic and cluster cases of hepatitis E disease in industrialized societies are caused mainly by spread of zoonotic G3 and G4 HEVs by food-borne transmission (5). Immunocompromised individuals are prone to chronic infection by G3 HEV (6–8), including patients with HIV (9) and those receiving chemotherapy or with rheumatic disorders (10). One-tenth of patients with chronic HEV infection undergo rapid progression of liver fibrosis and cirrhosis (11, 12), leading to deadly cirrhosis within 2 to 3 y (13). Nevertheless, it is not well understood how sustained HEV infection can trigger the occurrence of liver fibrosis.

HEV is a small, quasi-enveloped positive-strand RNA virus (14, 15), whose 7.2-kb genome contains three open reading frames (ORFs) that encode a roughly 190 kDa nonstructural protein (ORF1) responsible for viral replication, a capsid protein (ORF2), and a multifunctional protein (ORF3), respectively (16–21). ORF1 is a putative multidomain protein composed of a methyltransferase (Met), a Y-domain (Y), a putative papain-like cysteine protease (PCP), a hypervariable region (HVR), an X-domain (X) or macrodomain, a helicase (Hel), and an RNA-dependent RNA polymerase (RdRp) (18, 22). However, whether and how the ORF1 protein is processed for its functionality remains a long-standing question (18, 23).

Slow but significant progress has been made in understanding HEV biology, particularly with developing viral replicon cellular systems and animal infection models (2, 24). Based on our established RNA replicon system (25) and Mongolian gerbil infection model (26) of human HEV (G3) and HEV-infected patients (G4), in the current study, we observed a unique processing of viral ORF1 via a ubiquitin–proteasomal pathway, which leaves a stable C-terminally truncated ORF1 product localized in the nucleus. This soluble viral polypeptide, termed the HEV-derived SMAD activator (HDSA), profoundly activated fibrogenic TGF-β signaling by potentiating SMAD2/3, essential to mediate liver inflammation and fibrosis in viral-susceptible rodent models. Our findings collectively describe the unique nature of HEV–host interactions and uncover the critical molecular pathway leading to HEV pathogenicity.

Results

HEV ORF1 Is Processed to Produce HDSA.

It is long debated whether the ORF1 of HEV requires processing for its functionality (18, 23). Since we recently established a persistent RNA replicon of human G3 HEV that stably expresses both genomic and subgenomic RNAs and a sustained infection model of human G3 HEV in Mongolian gerbils (25, 26), it is now possible to precisely probe whether and how the viral ORF1 is processed during the viral lifecycle. Using an antibody targeting the X domain of ORF1, we observed an abundant, truncated isoform of ORF1, hereafter referred to as the HDSA, in HEV-infected gerbils liver specimens (Fig. 1A). This HDSA was similarly detected in HEV replicon cell lines, including both hepatocytes (S10-3) and baby hamster kidney (BHK) cells, with an approximate molecular weight of 120 kDa (Fig. 1B and SI Appendix, Fig. S1A). Notably, HDSA was evident by immunoblot in specimens of liver transplant patients with chronic HEV infection (G4), and the infection was confirmed by qPCR analyses (Fig. 1C and SI Appendix, Fig. S1B). HDSA displayed a significant protein level in hepatocytes, comparable to that of the intact ORF1 (Fig. 1 A–C), indicating it may play a biological role.

Fig. 1.

HEV ORF1 is processed to produce HDSA. (A), HDSA, with an approximate m.w. of 120 kDa, was visualized in gerbil liver specimens infected with G3 HEV by immunoblotting using a monoclonal antibody against the X domain. Gerbils were injected intraperitoneally (i.p.) with 1 mL of infectious virus stocks containing approximately 6.83 × 10⁷ genome equivalents (GE) of viral RNA, and at 10 d postinfection (dpi), liver lysates were immunoprecipitated by the antibody targeting the X domain of ORF1 and subjected to immunoblotting analyses. (B), HDSA was similarly detected by immunoblot after immunoprecipitation in human HEV replicon hepatocytes (S10-3-EZ). (C), HDSA was detected by immunoblot upon immunoprecipitation by anti-X domain monoclonal antibody in livers from three cirrhotic patients (G4 HEV) after liver transplantation. (D and E), The HSP90 inhibitor NVP-HSP990 (1 μM in BHK-EZ, and 5 mg/kg in gerbils) destabilized HEV ORF1 in HEV replicon BHK-EZ cells (D) or gerbil liver specimens (E); HDSA was detected by immunoblot using an anti-X domain monoclonal antibody. (F), HEK293 cells with ectopic expression of HA-ORF1-Myc (24 h posttransfection) were treated with NVP-HSP990 (1 μM; 3 h), and protein expression was visualized by immunoblot targeting the HA or the X domain, but not targeting Myc. A schematic diagram of ORF1 with the N-terminal HA and C-terminal Myc tags (HA-ORF1-Myc) is shown. Applied to Figs. 1–7: Unless otherwise specified, the mean ± s.e.m. is shown, n = 3 independent experiments, *P < 0.05, **P < 0.01, and ***P < 0.001, by ANOVA tests with Bonferroni correction. The statistics source data are provided in Dataset S1, and unprocessed images of the blots are shown in SI Appendix, Fig. S6).

In our recent work, we found that HEV ORF1 maintains its stability and avoids degradation by forming a complex with HSP90; upon treatment with an HSP90 inhibitor (HSP90i), we observed a rapid degradation of ORF1 (27). Based on these findings, we hypothesized that HSP90 may also protect ORF1 from processing and the subsequent production of HDSA. As expected, robust HDSA expression was observed in HepG2 and S10-3 with ORF1 expression (SI Appendix, Fig. S1 C and D), and in HEV replicon BHK cells (Fig. 1D), consistent with observations in liver specimens (Fig. 1E), following HSP90i treatment. This indicates that HSP90i is a suitable tool for detailed analyses of HDSA production. A dual-tagged (HA and Myc) ORF1 was then generated for this investigation. Using specific epitope targeting, we confirmed that HDSA is a truncation of ORF1 that includes the N-terminal and X domain (Fig. 1F). We also examined production of the HDSA in other HEV subtypes. Besides G3 ORF1, we found that ORF1 sourced from G1 and G4 HEV also shared this feature, suggesting this unique ORF1 processing is conserved across evolution (SI Appendix, Fig. S1E). Notably, the G3 ORF1 was sourced from the Kernow-C1 p6 strain of HEV, which contains an insertion of the human S17 ribosomal protein in the HVR (28). Although the insertion of S17 significantly enhances viral replication, it does not affect ORF1 processing, indicating that ORF1 processing is an intrinsic property of HEV. (SI Appendix, Fig. S1E).

ORF1 Undergoes Proteasomal Processing to Produce HDSA.

Many proteinases are responsible for protein processing in cells, including caspases, matrix metalloproteinases (MMPs), proteasomes, and other proteinases. We initially used various inhibitors to probe the proteinase involved in ORF1 processing. Surprisingly, this ORF1 processing was prevented entirely by MG132 and MG-101, two well-defined proteasomal inhibitors (Fig. 2A). Consistent with this observation, five distinct proteasomal inhibitors were found to block this ORF1 processing and HDSA production (Fig. 2B). In contrast, inhibitors of caspases, MMPs, and other proteinases failed to influence the HDSA production process (Fig. 2A). Proteasome-dependent protein processing, wherein a portion of a protein is degraded to release a biologically active fragment, has been documented for other proteins, including p105 (NF-κB) (29), Gli3 (Gli3-83) (30), and Def1 (pr-Def1) (31). As proteasome processing generally requires a K48-linked polyubiquitination signal, we introduced lysine-to-arginine (KR) mutations in the X, Hel, or RdRp domains of ORF1 to determine the effect on HDSA production. Notably, mutating all 18 K residues to R in the RdRp domain of ORF1 (ORF1-18KR), but not mutating the K residues in X or Hel domain, entirely abolished the proteasomal processing of ORF1 and the production of HDSA (Fig. 2C), while mutating 8 or 10 out of 18 K residues in the RdRp domain failed to prevent ORF1 processing (Fig. 2D). Remarkably, ORF1-18KR mutation blocked HDSA production but did not affect ORF1 degradation, whereas mutation of all 26 K residues found within the ORF1 C-terminal domain (CTD) prevented both ORF1 degradation and HDSA production (Fig. 2C). This suggests that ubiquitination modifications at various sites within the ORF1 CTD may regulate HDSA production or lead to its complete degradation.

Fig. 2.

ORF1 ubiquitination directs proteasomal processing at the X domain to produce HDSA. (A and B), HEV replicon cells (BHK-EZ) were treated with ten distinct proteinase inhibitors (10 μM; 4 h), including proteasome inhibitors (MG132, MG101), a calpain inhibitor (Calpeptin), a cysteine protease inhibitor (E-64), a serine protease inhibitor (Leupeptin Hemisulfate), an MMP inhibitor (Batimastat), and caspase inhibitors (Z-VAD-FMK, VX-765, Q-VD-Oph, and Z-DEVD-FMK) (A), or in the presence of six distinct inhibitors of the proteasome (10 μM; 4 h), including MG132, bortezomib, delanzomib, ixazomib, carfilzomib, and oprozomib (B). ORF1 and HDSA were clearly detected in cell lysates by immunoblot against the X domain in the absence or presence of NVP-HSP990 (1 μM; 4 h). (C), Mutating K residues to R in the X domain (ORF1-3KR), Hel domain (ORF1-5KR), RdRp domain (ORF1-18KR), or throughout the entire CTD of ORF1 (ORF1-26KR) affected the production of HDSA, as detected by anti-HA and anti-Myc immunoblotting in ORF1-KR mutants stably expressed in HEK293 cells. (D), Mutating 18 K residues (ORF1-18KR), but not 8 or 10 K residues, prevented ORF1 processing from producing HDSA. s.e. (short exposure). (E), HA and Myc dual-tagged ORF1, ORF1 del X, and ORF1 del Hel were ectopically expressed, and HDSA was visualized by immunoblotting using antibodies against HA or Myc, in the presence of NVP-HSP990 (1 μM; 3 h). (F), HA and Myc dual-tagged ORF1, ORF1 del RdRp, and ORF1 del Hel-RdRp were ectopically expressed, and HDSA was visualized by immunoblotting using antibodies against HA, X, or Myc. (G), HA and Myc dual-tagged ORF1, ORF1 point mutants PAA-3C, AAA-3C, and AWF-3C were ectopically expressed, and HDSA was visualized by immunoblotting using antibodies against HA or Myc in the presence of NVP-HSP990 (1 μM; 3 h). In ORF1 point mutants, the P⁹⁸⁹-A⁹⁹⁰-A⁹⁹¹-A⁹⁹²-W⁹⁹³-F⁹⁹⁴ residues at the C-terminus of the X domain were sequentially replaced by cysteines, as shown in the lower panel. (H), BHK-21 cells were transfected with capped genomic RNA transcripts of human G3 HEV containing a GFP tag, including WT and ORF1 mutants (PAA-3C, AAA-3C, and AWF-3C). GFP levels represent the replication state of HEV replicons that were measured at 7 d posttransfection.

HDSA Is a C-Terminal Truncation of ORF1, Lacking Hel and RdRp Domains.

To characterize the precise region of ORF1 proteasomal processing, we generated and analyzed a series of ORF1 truncations and point mutations, finding that deletion of the X domain but not the Hel domain blocked HDSA production entirely (Fig. 2E). This suggests that HDSA is likely to be generated by the removal of the Hel and RdRp domains from ORF1, which aligns with our experimental results (Fig. 2F). Based on the results of the ORF1 truncation experiments, we speculated that the cleavage site of ORF1 is located at the distal of the X domain (Fig. 2 E and F). Sequence analysis indicated a region at the distal of the X domain (986 to 995 a.a.) containing multiple hydrophobic amino acids, which is preferred by chymotrypsin-like proteasome processing (32, 33). Every attempt to disrupt ORF1 processing by introducing single-point mutations in this region failed, which suggests that ORF1 processing does not occur at a specific amino acid, as has also been found for other processed proteins (31, 34). Intriguingly, substitution of three hydrophobic amino acids in this region (P989-A990-A991-A992-W993-F994) into cysteines could substantially abolish the proteasomal processing of ORF1 (Fig. 2G). The conservation analysis of the 989-PAAAWF-994 sequence across HEV genotypes 1 to 8 indicates that G3, G4, and G6 exhibit relatively high conservation, while other genotypes display some variability (SI Appendix, Fig. S2A). Notably, at least four of the six amino acids in this sequence are identical among different genotypes. To further clarify the sequence of HDSA, we performed mass spectrometry analysis on expressed ORF1 and HSP90i-induced HDSA, identifying 125 peptides for ORF1 and 36 peptides for HDSA, which are highlighted in yellow (SI Appendix, Fig. S2 B and C). Alignment of these sequences with ORF1 revealed that the identified HDSA peptide sequences are closely located near the PAAAWF region, highlighted in red (SI Appendix, Fig. S2C). The mass spectrometry data provide valuable insights, consistent with our point mutation experiments, supporting the hypothesis that the cleavage site is near the PAAAWF region. When these mutations were introduced into the genomic RNA of the HEV replicon, they unfortunately curbed viral genome replication (Fig. 2H). These data collectively suggest that the cellular proteasome processes ORF1, eliminating its C-terminal Hel and RdRp domains to generate HDSA, with the cleavage site defined by a hydrophobic signal sequence at the C-terminal of the X domain.

HDSA from G3 HEV Is a Stable Nuclear Protein That Impacts Various Signaling Pathways.

To characterize HDSA, we constructed expression plasmids with a C-terminal Myc tag for expressing both HDSA and the full-length ORF1. Unlike the full-length ORF1 that was noticeably unstable, the HDSA displayed remarkable protein stability, even under inhibition of global protein synthesis or molecular chaperone (Fig. 3 A and B and Fig. 1D). Intriguingly, HDSA derived from G3 HEV localized in the nucleus of various cells, in sharp contrast to its parent ORF1 protein, which was mainly present in the cytoplasm (Fig. 3C). We also examined the localization of HDSA and ORF1 derived from G1 HEV in HepG2 cells. Unlike G3 HDSA, which predominantly localizes in the nucleus, G1 HDSA is primarily found in the cytoplasm (Fig. 3D). Next, we examined the effect of HDSA on a subset of signaling pathways related to liver diseases and virus–host interactions, including TGF-β/SMAD3, BMP/SMAD1, Hippo/YAP, RLR/IRF3, and Hedgehog (Hh)/Gli1. Surprisingly, reporter assays looking at the various corresponding promoters indicated a dramatic potentiation of HDSA on the TGF-β/SMAD pathway, in contrast to its suppressive effects on YAP, SMAD1, and Gli1 (Fig. 3E). In addition, transcriptome analyses indicated HDSA could significantly promote TGF-β signaling and upregulate the expression of receptor-activated SMAD (R-SMAD) target genes (Fig. 3F). The capability of HDSA to potentiate TGF-β/SMAD signaling is extraordinarily interesting, considering the central roles of TGF-β/SMAD signaling in liver fibrosis and hepatic stellate cell (HSC) activation, and the fact that chronic HEV infection frequently leads to liver fibrosis.

Fig. 3.

HDSA from G3 HEV is a stable nuclear protein that impacts various signaling pathways. (A and B), The lifespan and stability of whole ORF1 and HDSA proteins, tagged with C-terminal Myc, were revealed by immunoblotting during protein synthesis arrest by cycloheximide (CHX) (10 μM; 0/3/6/9 h) and revealed using antibodies against Myc tag in both ectopically expressed cells (A) and the stably expressed cells (B). l.e. (long exposure). (C), Immunofluorescence confocal imaging using an anti-Myc antibody revealed a prominent localization of tagged HDSA in the nucleus of human hepatocytes, markedly distinct from the cytosolic localization of ORF1; (Scale bars, 20 μm.) (D), Immunofluorescence confocal imaging using an anti-Myc antibody revealed the localization of G1 and G3 ORF1/HDSA in HepG2 cells; (Scale bars, 20 μm.) (E), HDSA was coexpressed with reporters and signaling effectors of various signaling pathways in HEK293 cells, including those of TGF-β, BMP, YAP, RLR, and Hedgehog (Hh). The potent potentiating effect of HDSA on SMAD3 transactivation was detected. (F), Transcriptome analyses were performed in HEK293 cells with or without TGF-β (5 ng/mL; 1 h) and HDSA expression. HDSA markedly enhanced the mRNA expression of a subset of TGF-β target genes such as ID1, SMAD7, SKIL, FURIN, and KLF10 and affected the expression of some TGF-β signaling regulatory genes.

HDSA Potently Facilitates TGF-β/SMAD Signaling.

Further experiments revealed that HDSA substantially facilitates three different SMAD3-responsive promoters in various cells (Fig. 4 A–C). Intact HDSA, but not each of its domains alone, markedly drove SMAD3 signaling (Fig. 4D), suggesting that the complete HDSA comprises a functional unit in this molecular event. On the other hand, HDSA did not directly activate, but rather enhanced SMAD3 signaling (Fig. 4 A and E). Intriguingly, HDSAs from the G3 and G4 HEVs that had a capacity for chronic HEV infection (2), but not that from G1 HEV, could potentiate TGF-β/SMAD signaling (Fig. 4F). Although S17 insertion does not affect the ability of ORF1 to produce HDSA (SI Appendix, Fig. S1E), it enhances HDSA’s ability to potentiate TGF-β/SMAD signaling (Fig. 4G). Profoundly, stable expression of HDSA in human HSC LX2 or epithelial cells (HaCaT) enhanced the mRNA expression of TGF-β target genes such as SMAD7, PAI1, and collagens (Fig. 4H and SI Appendix, Fig. S3A). Notably, ORF1 of G3 HEV, but not its processing-null mutants, enhanced the SMAD3-responsive promoter (Fig. 4I). This suggests that the ability of ORF1 to potentiate TGF-β/SMAD signaling is a gain-of-function resulting from its unique proteasomal processing by the host. Meanwhile, the other two proteins encoded by HEV, ORF2 and ORF3, do not enhanced the SMAD3-responsive promoter (SI Appendix, Fig. S3B). These intriguing observations unveil an unexpected relevance between the viral HDSA, its unique virus–host interactions, and a major fibrogenic signaling pathway.

Fig. 4.

HDSA potently facilitates TGF-β/SMAD signaling. (A), 4SBE-A _ Luc is a luciferase reporter responsive to SMAD3, indicating activation of the TGF-β signaling pathway. Coexpression of HDSA potentiated TGF-β/SMAD3 signaling in S10-3, HepG2, and HaCaT cells. (B), HDSA potentiated TGF-β/SMAD3 signaling in a dose-dependent manner. (C), HDSA markedly enhanced TGF-β/SMAD signaling in HEK293 cells, as indicated by reporter assays using three distinct SMAD3-responsive promoters (3TP_Luc, PAI1_Luc, and SMAD7_Luc). (D), SMAD3-responsive reporter assays indicated that intact HDSA, but not its domains, potentiated TGF-β/SMAD signaling. (E), HDSA failed to potentiate TGF-β signaling in the absence of SMAD3 activation. Mutant 2SA (S423A/S425A) is an inactive SMAD3. (F), Distinct effects of HDSAs derived from HEV genotypes 1, 3, and 4 on TGF-β/SMAD3 signaling were revealed by SMAD3-responsive reporter assays. Contrary to G3 HDSA, HDSA from G1 HEV failed to enhance TGF-β signaling. (G), The impact of the S17 insertion in G3 HDSA on TGF-β/SMAD3 signaling was assessed using SMAD3-responsive reporter assays. (H), HDSA, stably expressing in human hepatocytes (LX2 cells) via a lentiviral vector, substantially enhanced TGF-β-induced transcription of PAI1, SMAD7, COL1A1, and COL3A1, which are well-defined target genes of TGF-β. (I), Reporter assays indicated that ORF1 mutants incapable of HDSA production, including PAA-3C, AAA-3C, and AWF-3C, could not enhance TGF-β/SMAD signaling.

HDSA Tethers R-SMAD to Facilitate Its Promoter Binding and Transcription.

Considering the nuclear localization of HDSA (Fig. 3C) and its enhanced transactivation of constitutively active SMAD3 (caSMAD3) (Fig. 5A), we first investigated the association of HDSA with various SMAD proteins. Robust interaction between HDSA and the two R-SMADs, SMAD2 and SMAD3, but not SMAD1 or SMAD4, was observed in ectopically expressed proteins (Fig. 5B). Notably, G1 HDSA neither potentiates TGF-β/SMAD signaling (Fig. 4F) nor interacts with SMAD3 (SI Appendix, Fig. S4A). Additionally, we found that HDSA localizes predominantly in the nucleus, with a small fraction in the cytoplasm, in both wild-type (WT) and SMAD2/3 knockout HaCaT cells, indicating that HDSA has an intrinsic nuclear import ability (SI Appendix, Fig. S4B). Furthermore, HDSA showed a stronger binding affinity for caSMAD3 (Fig. 5C). The HDSA–SMAD3 complex was detected in cells stably expressing HDSA, and this interaction was significantly enhanced upon TGF-β pathway activation (Fig. 5D). Immunofluorescence analysis confirmed the nuclear colocalization of the HDSA–SMAD3 complex after TGF-β activation in HaCaT cells (Fig. 5E). Domain mapping assays by co-IP revealed that the N-terminal Met and Y domains of HDSA (Fig. 5F) and the MH2 domain of SMAD3 (Fig. 5G) were responsible for their mutual interaction. However, HDSA did not significantly elevate the levels of phosphorylated (activated) SMAD3 (SI Appendix, Fig. S4C), suggesting that HDSA might not participate in receptor-induced R-SMAD phosphorylation/dephosphorylation or the control of stability of R-SMAD proteins. HDSA also did not change the formation of SMAD3–SMAD4 complexes (SI Appendix, Fig. S4D) or the nuclear translocation of SMAD3 (SI Appendix, Fig. S4E). However, recruitment of the transcription coactivator p300, an essential step in SMAD3-driven transcription (35), was facilitated by HDSA (Fig. 5H). Notably, HDSA effectively facilitated the binding of SMAD3 to the SBE element (Fig. 5I), as well as endogenous promoters, including those of SMAD7 and PAI1 (Fig. 5J), as evidenced by DNA pull-down and ChIP assays, respectively. These consistent observations identified HDSA as a virus–host interaction–derived SMAD2/3 partner to improve their role in transcription.

Fig. 5.

HDSA tethers R-SMAD to facilitate its promoter binding and transcription. (A), Reporter assays using a SMAD3-responsive SBE promoter showed that HDSA enhanced TGF-β/SMAD3 signaling in HEK293 cells stimulated by either constitutively active TβRI (caTβRI) or SMAD3 (caSMAD3). (B), Coimmunoprecipitation (co-IP) assays revealed an interaction between HDSA with SMAD2/3 rather than SMAD1/4, performed in HEK293 cells at 24 h posttransfection. (C), Co-IP assays revealed a stronger interaction between HDSA and caSMAD3; but not with the inactive mutant SMAD3-2SA. (D), HDSA was stably expressed in HaCaT cells by a lentiviral system. Co-IP assays revealed a complex between stably expressed HDSA and endogenous SMAD3; with an enhanced interaction observed following TGF-β treatment (5 ng/mL; 0.5 h). (E), HDSA, stably expressed in HaCaT cells, colocalized with endogenous SMAD3 in the nucleus in response to TGF-β treatment (5 ng/mL; 0.5 h); (Scale bars, 20 μm.) (F), Co-IP assays between SMAD3 and HDSA, as well as the individual domain of HDSA, showed a robust interaction of SMAD3 and the intact HDSA, Met, and Y. (G), Co-IP assays between HDSA and SMAD3 truncations indicated that the C-terminal MH2 domain of SMAD3 was responsible for and sufficient for HDSA recruitment. A schematic diagram of the interaction domains between HDSA and SMAD3 is shown (Right panel). (H), Co-IP assays between p300 and caSMAD3 suggested that HDSA enhanced the formation of the SMAD3–p300 complex. A tripartite complex of SMAD3, HDSA, and p300 was also detected. (I), DNA pull-down assay showing increased binding of the SBE sequence by SMAD3 in the presence of HDSA. (J), ChIP assay using the anti-SMAD3 antibody in HEK293 cells revealed an enhanced SMAD3 binding on SMAD7 (Left) and PAI1 (Right) promoters in the presence of HDSA.

To better understand how HDSA interacts with and regulates Smad3 at the structural level, we utilized AlphaFold to predict the structure of HDSA (36). The protein sequence of HEV HDSA was input into the AlphaFold algorithm, and the best-ranked model was shown (SI Appendix, Fig. S4F). The figure presents three angular views of HDSA, with the upper portion predominantly occupied by the Met and Y domains, exhibiting relatively compact structures while the PCP spans across the top and bottom. In the lower portion, the HVR and X regions are situated, with the HVR displaying greater flexibility (SI Appendix, Fig. S4F). Our predicted structure of HDSA in complex with Smad3 revealed a more compact configuration of HDSA within the complex, with Smad3 adopting an inverted form chimerized with HDSA (SI Appendix, Fig. S4G). Notably, both the Met and Y domains of HDSA in the complex form interacting interfaces with Smad3 (SI Appendix, Fig. S4H), which is also consistent with the findings of our immunoprecipitation experiments (Fig. 5F). These observations suggest that HDSA can tightly associate with Smad3, thus exerting a regulatory function.

Persistent HEV Infection Induces Liver Fibrosis in Gerbils.

The frequent occurrence of liver fibrosis upon chronic HEV infection and the presence of endogenous HDSA in the livers of HEV-infected animals and patient specimens suggest a causal role for HDSA in HEV pathogenicity. To investigate the physiological functions of ORF1/HDSA in HEV-induced liver fibrosis, we initially examined whether persistent G3 HEV infection in HEV-susceptible gerbils could induce liver fibrosis. Based on our previous studies using the gerbil model, G3 HEV can sustain replication with a high viral load in the liver of gerbils for over 6 wk (26). Therefore, gerbils were infected with HEV and killed at the 6 wk postinfection and assessed via hematoxylin–eosin (H&E) staining, Sirius Red staining, and qPCR (Fig. 6A). In the liver of HEV-infected gerbils, the viral load remained at a high level (Fig. 6B), accompanied by noticeable immune cell infiltration and mild fibrotic phenotypes (Fig. 6 C and D). Consistent with these results, more severe liver disease scores (Ishak scoring) (37, 38) were seen in HEV-infected livers (Fig. 6E). Notably, elevated TGF-β target gene expression was detected in the livers of HEV-infected gerbils, including the mRNA expression of Smad7, Pai1, and collagens (Fig. 6F). These consistent observations suggest that sustained replication of G3 HEV in susceptible rodents can induce liver fibrosis.

Fig. 6.

ORF1 proteasomal processing is indispensable for HEV-induced liver fibrosis. (A), HEV-susceptible gerbils were infected intraperitoneally (i.p.) with 1 mL of the infectious genotype 3 HEV stock containing approximately 6.83 × 10⁷ GE of viral RNA on day 0, followed by regular feeding for 6 wk. (B), qRT-PCR measuring viral RNA levels in livers at 6 wk postinfection. (C–E) Livers from HEV-infected gerbils were tested by H&E staining (C), Sirius Red staining (D), and liver disease scoring (E), showing liver disease severity, immune cell infiltration, and liver fibrosis at 6 wk postinfection. (F), qRT-PCR was used to measure alterations of a few TGF-β target genes, including Pai1, Smad7, Col1a1, and Col3a1, at 6 wk postinfection in the liver of gerbils infected or not with G3 HEV (6.83 × 10⁷ GE of viral RNA). (G and H), Gerbils were inoculated intraperitoneally with 1 mL (approximately 5 × 10⁸ TCID₅₀) recombinant adenovirus type-5 (Ad5) stocks, including Ad5-GFP, Ad5-HDSA, Ad5-ORF1, and Ad5-ORF1 (PAA-3C), and were reinoculated every 2 wk (G). HA-tagged HDSA, WT ORF1, and mutant ORF1 (PAA-3C) proteins were detected in gerbil livers by anti-HA immunoblotting at 2 wk postinoculation. Levels of HDSA and ORF1 were at low abundance and comparable to those observed during live HEV infection in gerbil livers at 10 d postinfection (H). (I–K) Liver disease scoring (I), H&E staining (J), and Sirius Red staining (K) of gerbil livers with Ad5 inoculation showed liver disease severity, immune cell infiltration, and liver fibrosis at 8-wk postinoculation. Mutant ORF1 failing to produce HDSA (PAA-3C) showed marginal effects on liver disease. (L and M), mRNA levels of TGF-β target genes (Col1a1, Col3a1, Pai1, and Smad7) were measured by qRT-PCR assays in liver specimens from gerbils inoculated with Ad5, including Ad5-GFP, Ad5-HDSA, Ad5-ORF1, and Ad5-ORF1 (PAA-3C).

ORF1 Proteasomal Processing Is Indispensable for HEV-Induced Liver Fibrosis.

To investigate whether ORF1 proteasomal processing is physiologically necessary for HEV-induced liver fibrosis, we generated three ORF1 processing-null mutants. Unfortunately, it was impossible to recover live HEV carrying these mutations from replicon cell lines, thus making them unavailable for in vivo studies (Fig. 2H). The recombinant adenovirus type-5 (Ad5) system is a commonly used and reliable protein expression system for animal organs, capable of accommodating genes up to 8 kb in size, making it suitable for expressing ORF1 (>5 kb) (39). We next employed the Ad5 system in gerbil livers to express GFP, HDSA, WT ORF1, and the ORF1 processing-null mutant PAA-3C of G3 HEV. Twenty-four SPF gerbils were randomly divided into four groups and inoculated with Ad5 viruses and reinoculated every 2 wk (Fig. 6G). Hepatic expression of HDSA, ORF1, and the ORF1 processing-null mutant PAA-3C were found at low abundance, comparable to the levels observed during live HEV infection (Fig. 6H). Similar robust effects for HDSA on immune cell infiltration and liver fibrosis were seen (Fig. 6 J and K). Immunohistochemistry (IHC) for HA, as well as liver fibrosis markers such as α-SMA, Collagen I, and the epithelial–mesenchymal transition (EMT) marker N-Cadherin, were consistent with the observed immune cell infiltration and liver fibrosis (SI Appendix, Fig. S5A). WT ORF1, but not the processing-null mutant, triggered a mild-to-medium state of inflammation, fibrosis, and TGF-β/SMAD signaling in the livers of infected gerbils (Fig. 6 J–M). Finally, WT ORF1, but not the processing-null mutant, significantly affected liver disease score (Fig. 6I). These in vivo observations suggest that ORF1 proteasomal processing facilitates HEV pathogenicity.

G3 HDSA Facilitates Liver Fibrosis in HEV-Susceptible Gerbils.

Since G1 HEV induces acute hepatitis without causing fibrosis, whereas G3 HEV causes chronic hepatitis and liver fibrosis, we were able to use G1 HDSA as a parallel control. We generated gerbils bearing the hepatic expression of HDSA from G3 or G1 HEV by an adeno-associated virus 8 (AAV8) system and reinoculated them every 4 wk (Fig. 7A). Abundance of hepatic expression of HDSA was low, comparable to the levels observed during live HEV infection (Fig. 7B). Degrees of liver injury and fibrosis in gerbils were assessed at the 8 wk by visualizing fibroblast and immune cell accumulation via H&E staining, deposited extracellular matrix via Sirius Red staining, and ECM component expression via qPCR. Notably, increasing immune cell infiltration (Fig. 7C) and increased expression of chemokines (Fig. 7D) were seen in gerbil livers with the expression of HDSA from G3 HEV but not G1 HDSA or control protein. G3 HDSA significantly increased areas of extracellular matrix collagen deposition, with substantially enlarged Sirius Red positive tissues (Fig. 7E) and enhanced expression of collagens (Fig. 7F) in the gerbil livers, compared to the G1 HDSA or control protein expression. Consistent with these results, more severe liver disease scores (Ishak scoring) were seen in G3 HDSA-expressing livers (Fig. 7G). Meanwhile, IHC for HA, along with the liver fibrosis markers α-SMA, Collagen I, and the EMT marker N-Cadherin, also aligns with the results from H&E staining and qPCR (SI Appendix, Fig. S5B). As expected, elevated TGF-β target gene expression was detected in gerbil livers in the presence of G3 HDSA (Fig. 7H), validating the in vivo role of G3 HDSA in potentiating TGF-β/SMAD signaling. These consistent observations suggest that G3 HDSA, a viral protein derived from a unique virus–host interaction, can sufficiently drive liver fibrosis in HEV-susceptible rodents.

Fig. 7.

G3 HDSA is capable of and essential for driving fibrosis in HEV-susceptible gerbils. (A), HEV-susceptible gerbils were inoculated through the hepatic portal vein with 200 μL AAV8 stock (1 × 10¹² viral genomes (V.G.)/mL), including AAV8-GFP, AAV8-G3 HDSA, and AAV8-G1 HDSA, and reinoculated every 4 wk. (B), HA-tagged HDSA proteins were detected at 3 wk postinoculation by immunoblotting in liver specimens from gerbils administrated with AAV8-G3 HDSA and AAV8-G1 HDSA. Levels of HDSA were at low abundance and comparable to those observed during live HEV infection in gerbil livers at 10 d postinfection. (C and D), Liver histology by H&E staining of gerbil livers at 8 wk post-AAV inoculation showed immune cell infiltration (C) and the expression of chemokines was detected by qRT-PCR (D). (E–G), Liver histology by Sirius Red staining of gerbil livers at 8 wk post-AAV inoculation revealed enhanced liver fibrosis (E), upregulated mRNA expression of collagens (F), and exacerbated liver disease scoring (G) in gerbils with AAV-delivered G3 HDSA. Expression of G1 HDSA showed marginal effects on fibrosis of gerbil livers. (H), qRT-PCR assays indicated mRNA levels of TGF-β target genes (Smad7 and Pai1) in gerbil liver specimens following AAV inoculation. (I), Scheme showing pathways leading to HDSA formation and liver fibrosis. HEV viral replicase naturally undergoes a K48 polyubiquitination and unique proteasomal-mediated processing, which produces a stable and nuclear-localized viral polypeptide HDSA. HDSA profoundly potentiates TGF-β/SMAD3 fibrogenic signaling and substantially contributes to murine liver fibrosis, leading to the primary clinical symptom of chronic HEV infection.

Discussion

Hepatitis E is a significant threat to human health, especially in immunosuppressed individuals and pregnant women. Chronic HEV infection frequently occurs in immunosuppressed patients, resulting in life-threatening liver cirrhosis. This report reveals a very intriguing mechanism by which the viral replicase ORF1 is transformed into HDSA via ubiquitination-proteasomal processing. HDSA is relatively stable and translocates into the nucleus, where it interacts with SMAD2/3 and potentiates transcription activity. Remarkably, we found that liver-specific expression of either viral ORF1 or HDSA can trigger liver injury and fibrosis, thus suggesting a potential causative mechanism for HEV-induced liver fibrosis (Fig. 7I).

To date, details of how the ORF1 protein is processed and the precise functions of each putative domain are still poorly understood. Recently, one study utilizing a HEV replicon system with an HA tag inserted into the HVR detected the ORF1 protein but did not observe any ORF1 processing products (40). In contrast, another study employing a HEV replicon system with a V5 tag inserted into the HVR identified several processing products of ORF1, although the specific characteristics, proteases involved, processing mechanisms, and functions of these products remain unknown (23). In this study, we observed a rapid and proteasomal-dependent processing of HEV ORF1, leaving behind a visible C-terminal-truncated product, termed the HDSA, which comprised the putative Met, Y, PCP, HVR, and X domains. Inhibition of the proteasome activity, or mutagenesis of the residues in the hydrophobic amino acids–rich region (P989-A990-A991-A992-W993-F994), apparently attenuates HDSA production. Unlike previous reports and understandings, the production of HDSA does not rely on PCP or known proteases such as thrombin (41–43), but rather requires the involvement of host proteasomes. Notably, HDSA is a viral protein found to be produced by such a unique virus–host interaction, although similar processes have been reported in mammals, fruit flies, and yeast (30, 31, 44, 45). Nevertheless, it remains to be determined how the balance between viral ORF1 degradation and HDSA production is achieved and regulated. Acquiring host genome segments is a unique capability of HEV and critical in their virus–host interactions (46, 47), which significantly alters their infection and pathogenicity. G3 ORF1 was sourced from the Kernow-C1 p6 strain of HEV, which contains an insertion of the human S17 ribosomal protein. Although S17 insertion does not affect the ability of ORF1 to produce HDSA, it enhances HDSA’s ability to potentiate TGF-β/SMAD signaling. These findings suggest that the S17 insertion is dispensable but may contribute to the increased pathogenicity of HEV, and the specific molecular mechanisms still require further investigation.

Chronic infection by hepatitis viruses can result in severe disease symptoms, including inflammation, fibrosis, cirrhosis, and hepatocellular carcinoma (48). Significant liver fibrosis can be triggered by the sustained low-grade injury induced by persistent HBV, HCV, and HEV infections and their complex interactions with the immune system. Such fibrosis is characterized by excessive collagen deposition, extracellular matrix accumulation, functional hepatocyte death, and fibroblast overproliferation. HBV and HCV induce fibrosis by activating HSC, directly or indirectly, which triggers a wound-healing response (49). The release by liver and immune cells of various profibrinogenic factors, such as TGF-β, IL-6, TNF-α, IL-1, and PDGF, contributes to viral-induced fibrosis (50). Liver-resident macrophages and the recruitment of monocytes into the inflamed and damaged liver are also essential for initiation and progression of liver fibrosis (51). Here, we found that HDSA precisely and remarkably activates TGF-β/SMAD signaling, a master regulator of fibrotic diseases in various organs. Mechanistically, HDSA formed an endogenous complex with SMAD2/3 that facilitated the binding of R-SMADs to the promoter of TGF-β target genes and the recruitment of transcription coactivators such as p300. We further employed AAV and Ad5 delivery systems to validate the in vivo role of ORF1 processing and HDSA in HEV pathogenesis. We observed a marked increase in liver inflammation, injury, and fibrosis when ORF1 or HDSA was delivered into rodent livers, even without the requirements of other stimulations to trigger the liver fibrosis process. These dramatic phenotypes recapitulated the symptoms observed in HEV-susceptible gerbils, suggesting that HDSA, even expressed at marginal protein levels, has a profound role in driving liver fibrosis. Therefore, these intriguing data suggest that HDSA drives pathogenesis in patients with chronic HEV infections. Additionally, it remains to be determined whether HDSA is involved in regulating the HEV life cycle and if it is essential for HEV replication. In our previous small molecule screening, we found that the TGF-β inhibitor SB431542 can suppress HEV replication (27). Therefore, further investigation is needed to explore whether HDSA might promote viral replication by enhancing TGF-beta signaling.

In conclusion, our findings suggest a unique mechanism to explain HEV-induced pathogenesis, via production of a truncated viral protein, the HDSA, which potently activates TGF-β signaling and drives rodent livers into fibrosis. Identifying the HDSA reveals the intriguing biology behind HEV–host interactions, and sheds light on a mechanism that can help us better understand and possibly treat this devastating infectious disease.

Materials and Methods

Animal Experiments.

Mongolian gerbils were purchased from the Experimental Animal Center at the Zhejiang Academy of Medical Sciences and maintained in the specific-pathogen-free (SPF) environment. Before inoculation, a commercial ELISA kit confirmed the 6 to 8 wk male gerbils to be HEV seronegative (Wantai Biological Pharmacy). Gerbils were treated according to the experimental design and killed at the indicated times. Blood and liver samples were collected for subsequent detection and analysis. All animal experiments were performed strictly with the Experimental Animal Ethics Committee of Zhejiang University (IACUC approval no. ZJU20181049). Detailed grouping and treatment information is provided in SI Appendix.

Patients and Samples.

This study used archived tissue samples obtained from patients diagnosed with liver cirrhosis who provided informed consent as part of previous clinical treatments. The consent forms included information regarding the potential use of their samples for future research. All patient samples were anonymized to ensure confidentiality, and no personally identifiable information was included in the study. Patients 1 to 3 were not infected with HEV, while patients 4 to 6 were confirmed to have HEV infection. The liver samples underwent a grinding process, and a significant portion of the milled liver samples was lysed in a modified Myc lysis buffer (MLB). This lysate was then immunoprecipitated using an anti-X antibody targeting the HEV ORF1 and HDSA. After 3 to 4 washes with MLB, adsorbed proteins were resolved by SDS-PAGE (Bio-Rad) and subjected to immunoblotting with the indicated antibodies. Concurrently, a small portion of the milled liver samples was reserved for RNA extraction, followed by reverse transcription and qRT-PCR to detect HEV infection. The First Affiliated Hospital ethics committee approved the study, Zhejiang University School of Medicine (Ethics Approval: #IIT20230905A).

Detailed materials and methods are provided in SI Appendix.

Supplementary Material

Appendix 01 (PDF)

Dataset S01 (XLSX)

Dataset S02 (XLSX)

Dataset S03 (XLSX)

Dataset S04 (XLSX)

Data, Materials, and Software Availability

RNA-seq data were deposited in the GEO database under Accession No. GSE251924 (52). All other data are included in the article and/or supporting information.