The H5N1 avian influenza virus causes severe disease and high mortality, making it a major public health concern worldwide. The virus uses the host cellular machinery for several steps of its life cycle. In this report, we observed overexpression of the ubiquitin-like protein FAT10 following live H5N1 virus infection in BALB/c mice and in the human respiratory epithelial cell lines A549 and BEAS-2B. Further experiments demonstrated that FAT10 increased H5N1 virus replication and decreased the viability of infected cells. Total RNA extracted from H5N1 virus–infected cells, but not other H5N1 viral components, upregulated FAT10, and this process was mediated by the retinoic acid–induced protein I-NF-κB signaling pathway. FAT10 knockdown in A549 cells upregulated type I IFN mRNA expression and enhanced STAT1 phosphorylation during live H5N1 virus infection. Taken together, our data suggest that FAT10 was upregulated via retinoic acid–induced protein I and NF-κB during H5N1 avian influenza virus infection. And the upregulated FAT10 promoted H5N1 viral replication by inhibiting type I IFN.

The highly pathogenic H5N1 avian influenza virus had its first human outbreak in Hong Kong in 1997 (1). According to WHO reports, 826 confirmed cases occurred from 2003 to 2015, leading to 440 fatalities (2). In many cases, H5N1 virus infection resulted in an aggressive clinical course with rapid deterioration and high mortality (3). Intensive research on the H5N1 virus has been performed to elucidate the basis of its increased virulence. Viruses are capable of using the host cellular machinery to replicate, and considerable efforts to map the host cell machinery used by the virus have been made in the hope to uncover potential drug targets with a broader spectrum and a lower likelihood of drug resistance (4). Several host factors, such as IFITM genes, have been identified using functional genomics screening, although the interaction between the H5N1 virus and host cells is not fully understood (5).

Ubiquitin and ubiquitin-like proteins (ULPs) target cellular proteins involved in proteasomal degradation and play important roles in other biological processes. FAT10, also known as ubiquitin D (UBD), is a recently identified ULP member containing two ubiquitin-like domains (6). Similar to ubiquitination, protein FAT10ylation also appears to occur via a three-step enzymatic cascade, with UBA6 acting as a ubiquitin-activating E1 enzyme, USE1 acting as a ubiquitin-conjugating E2 enzyme and an unknown ligase functioning as E3 (7, 8). Because the FAT10 gene is in the MHC class I locus, and its expression is induced by IFN-γ and TNF-α, the gene has been proposed to play a role in immunity (9, 10). FAT10 overexpression in some cell lines results in caspase-dependent apoptosis (9, 11), and FAT10 is also involved in apoptosis during HIV-associated nephropathy (12). FAT10 is highly expressed in hepatocellular carcinomas and other types of cancers (13), and a recent report demonstrated dozens of potential substrates and interaction partners for FAT10 (14). However, the function of FAT10 in viral infection has not been elucidated.

In this study, we used a microarray assay to screen for host proteins involved in highly pathogenic H5N1 influenza virus infection and found that FAT10 was significantly upregulated during H5N1 virus infection. We provide evidence that FAT10 upregulation is mediated by the retinoic acid–induced protein I (RIG-I)-NF-κB pathway and plays a critical role in H5N1 virus replication by inhibiting type I IFN expression.

All of the procedures, including animal studies, were conducted following the National Guidelines for Care of Laboratory Animals (2006-398) and performed in accordance with institutional regulations after protocol review and approval by the Institutional Animal Care and Use Committee of the Institute of Military Veterinary, Academy of Military Medical Sciences (project number 2011-017).

293T, A549, Beas-2B, and Madin–Darby canine kidney (MDCK) cells were purchased from the American Type Culture Collection (Manassas, VA). CNE-2Z was obtained from the Cell Resource Center of Peking Union Medical College (Beijing, China). Human bronchial epithelial cells (HBEpiCs) were purchased from ScienCell Research Laboratories (Carlsbad, CA). The 293T cells, BEAS-2B, MDCK, and CNE-2Z were cultured in DMEM (Invitrogen) with 10% FBS (Invitrogen) and 100 U/ml penicillin/streptomycin (Invitrogen) at 37°C with 5% CO2. A549 cells were cultured in Ham’s F-12 (Invitrogen) medium supplemented with 10% FBS and 100 U/ml penicillin/streptomycin at 37°C with 5% CO2. HBEpiCs were cultured in bronchial epithelial cell medium (ScienCell) with 100 U/ml penicillin/streptomycin at 37°C with 5% CO2.

The primary Abs used in the analysis, anti-STAT1 and anti–phospho-STAT1, were purchased from Cell Signaling Technology (Boston, MA). Anti-FAT10 Ab was purchased from Enzo Life Sciences (Farmingdale, NY). Anti-nucleoprotein (NP) Ab was purchased from Millipore (Billerica, MA). Anti–β-actin Ab was purchased from Sigma-Aldrich (St. Louis, MO). HRP-conjugated secondary Abs and Western blotting luminal reagents were purchased from Santa Cruz Biotechnology (Dallas, TX).

Wild-type H5N1 virus A/Jilin/9/2004 (H5N1) and the seasonal H1N1 virus A/New Caledonia/20/1999 (H1N1) were grown in 10-d-old fertilized eggs. The working stocks were stored at −80°C as live virus or after inactivation by formaldehyde treatment. The viral titers were measured using the Reed and Muench method (15). Live virus experiments were performed in Biosafety Level 3 facilities at the Institute of Military Veterinary, Academy of Military Medical Sciences, under governmental and institutional guidelines.

For viral infection, cells were washed three times in DMEM to remove FBS and then incubated with influenza virus diluted in DMEM for 1 h at 37°C. After 1 h, the cells were washed and maintained in DMEM with 10% FBS for the indicated times.

Cell supernatants containing virus were 10-fold serially diluted with DMEM and applied in quadruplicate to 2 × 104 MDCK cells/well in a 96-well plate. The cells were supplied with DMEM containing TPCK-trypsin (Sigma-Aldrich) (1 μg/ml). On the fifth day postinfection (p.i.), the viral titer was determined by observing the cytopathogenic effect and was confirmed by hemagglutination. The TCID50 was determined based on the Reed–Muench method as described previously (15). The multiplicity of infection (MOI) was calculated with the following formula: MOI = 0.7*TCID50/cells.

BALB/c mice were purchased from the Institute of Laboratory Animal Science, Peking Union Medical College. The mice were fed normal mouse chow and water and caged in a pathogen-free facility with air filtration. All animal experiments were conducted in the animal facility at the Institute of Military Veterinary, Academy of Military Medical Sciences, in accordance with governmental and institutional guidelines.

After anesthesia induction with pentobarbital (100 mg/kg i.p.) and subsequent tracheotomy, 4-wk-old BALB/c mice were intratracheally inoculated with 106 TCID50 of live H5N1 virus or an identical volume of inactivated H5N1 virus, 106 TCID50 of live H1N1 virus, or allantoic fluid (AF) with a microsyringe. After 24 h, the mice were sacrificed. The mouse lung tissue was rapidly collected and preserved in liquid nitrogen prior to RNA extraction and purification. Transcriptional profiles were measured with a microarray assay using RNA isolated from lung tissue. For FAT10 mRNA measurements, lung tissue samples were collected 24, 48, and 72 h p.i., and the level of FAT10 was determined using real-time RT-PCR.

RNA quality was evaluated with a Bioanalyzer 2100 (Agilent Technologies, Santa Clara, CA), and RNA with minimal degradation and distinct 18S and 28S rRNA bands was used for analysis. Transcriptional profiling was performed using Agilent’s Whole Mouse Genome Microarray Kit 4644 K. The experiments were performed at Shanghai Biotechnology (Shanghai, China). Gene Spring GX software (Agilent Technologies) was used for data analysis (n = 3 mice/group). The results were deposited in the National Center for Biotechnology Information Gene Expression Omnibus (GEO; accession number GSE76719; http://www.ncbi.nlm.nih.gov/geo/).

The genes from the A/Jilin/9/2004(H5N1) virus as well as RIG-I, ΔRIG-I, IFN regulatory factor (IRF)3, and p65 were cloned into the pcDNA3.1 vector (Invitrogen) for efficient expression. PCR was performed for each gene with gene-specific primers. A C-terminal Flag tag was introduced into each construct using PCR with specific primers encoding the tag sequences.

The FAT10 promoter (−975/+209) (16) was cloned into pGL3-Promoter Vector (Promega, Madison, WI). The forward primer was 5′-GCTGGCTAGCTCAAGTTCCCATAAAATCATCT-3′, and the reverse primer was 5′-TCTGAAGCTTGCCAGAAACCAGAGACAGAA-3′. For the dual luciferase assay, 293T cells were transfected with the reporter vector (pGL3) and phRL expressing Renilla luciferase for normalization. After 24 h, the cells were transfected with RNA from H5N1-infected cells or a GFP control for another 24 h. The cells were collected, and the luciferase activity was measured using a Dual-Glo luciferase assay system (Promega), according to the manufacturer’s protocol. Mutation of the p65 binding site was achieved using the Easy Mutagenesis System (TransGen Biotech, Beijing, China), according to the manufacturer’s protocol.

Cells were seeded into 96-well plates at a density of 1 × 105/ml. Influenza virus or AF was added to the wells the next day. Each group was tested in triplicate wells. After incubation for the indicated times, 20 μl Cell Titer 96 AQueous One Solution Cell Proliferation Assay (Promega) was added to each well, and the cells were incubated for another 2 h. Then, the absorbance was recorded at 490 nm. In the siRNA knockdown group, the cells were first treated with siRNAs for 48 h.

Whole-cell lysates were prepared with lysis buffer (radioimmunoprecipitation assay lysis buffer with protease inhibitor mixtures) and boiled at 95°C for 10 min. The boiled lysates were analyzed by NaDodSO4 PAGE (SDS-PAGE) before transfer to nitrocellulose transfer membranes (Whatman Protran, Buckinghamshire, U.K.). The membranes were probed with primary Abs, and the proteins were visualized using HRP-conjugated Abs and a chemiluminescent substrate and then exposed to film. The band density was calculated using Quantity One software.

Total RNA was isolated from cells or mouse lungs using TRIzol reagent (Invitrogen), according to the manufacturer’s instructions. Reverse transcription of the purified RNA was performed using Moloney murine leukemia virus (Invitrogen) and random primers. PCR amplification assays were performed with LightCycler 480 SYBR Green I Master Max (Roche, Basel, Switzerland) in a LightCycler 480 Real-Time PCR System (Roche) using specific primers (see Supplemental Table I for sequences). Each value was calculated automatically by the LightCycler 480 software program using GAPDH as the reference.

Gene-specific small interfering RNA (siRNA) and nonspecific control siRNA were synthesized and supplied by RiboBio (Guangzhou, China). The siRNAs were transfected into cells with Lipofectamine RNAiMAX (Invitrogen), according to the manufacturer’s protocol. The cells were transfected with a pool of different gene-specific siRNAs or with a control siRNA for 48 h (see Supplemental Table I for sequences), and subsequent experiments were performed.

Full-length FAT10 was cloned into the lentiviral vector pWPXL (Addgene, Cambridge, MA), and cloning was confirmed by sequencing. This construct or an empty vector was cotransfected with the packaging vectors psPAX2 (Addgene) and pMD2.G (Addgene) into 293T cells to generate vesicular stomatitis virus-G–pseudotyped lentiviral particles. The lentiviral supernatants were harvested 48 h posttransfection, passed through a 0.45-μm filter, and stored at −80°C. The target cells were infected with lentiviral particles for 4 h in the presence of 8 μg/ml Polybrene (Sigma-Aldrich).

All of the data are shown as the mean ± SD. The statistical analyses were conducted using ANOVA analysis when comparing more than two groups and the Student t test for two groups. A p value <0.05 was considered significant.

To systematically investigate the interaction between the H5N1 avian influenza virus and its hosts, we performed a series of DNA microarray assays using mice. The mice were intratracheally administered the same dose of live or inactivated H5N1 virus, live H1N1 virus, or normal AF as a control. After 24 h, RNA extracted from mouse lungs was subjected to microarray analysis. The microarray data are available at GEO accession number GSE76719. Similar to previous studies, influenza virus infection dramatically changed the overall expression pattern in host cells (17, 18). The expression of hundreds of genes was highly upregulated or downregulated compared with AF. Among the affected host response pathways and gene ontologies, 15 members were detected of the 17 known ubiquitin and ubiquitin-like–encoding genes. Thirteen of these 15 genes only exhibited minimal changes in expression, but ISG15 and FAT10 were significantly upregulated in the H5N1 virus–infected group (Fig. 1A). The FAT10 fold change in the live H5N1 virus–infected group ranged up to 167.8-fold compared with the AF-treated group. And the live seasonal H1N1 infection increased FAT10 mRNA levels in mice by 56.012-fold compared with the AF control (Fig. 1A).

FIGURE 1.

FAT10 overexpression following H5N1 virus infection. (A) The relative gene expression levels of ubiquitin and ULPs in a DNA microarray analysis of mouse lungs infected with live (black) or inactivated (dark gray) H5N1 virus and live H1N1 virus (light gray). This experiment was performed once with three mice in each group. (B) In vivo kinetic expression of FAT10 in H5N1-infected mouse lungs. BALB/c mice were infected with 106 TCID50 H5N1 or an AF control. FAT10 mRNA in mouse lungs at 24, 48, and 72 h was quantified by real-time RT-PCR. The mRNA levels were normalized to GAPDH (n = 3 mice/group). (C) FAT10 mRNA expression levels increased in a time-dependent manner in A549 cells from 3 to 72 h p.i. A549 cells were infected with H5N1 at an MOI of 5. An equal volume of AF was used as the control. The FAT10 mRNA levels at 3, 6, 12, 24, 48, and 72 h were determined by real-time RT-PCR. The mRNA levels were normalized to GAPDH. (D) A549 cells were infected with H5N1 at an MOI of 1 or 5 for 24 h. FAT10 expression was analyzed by Western blotting analysis of cell lysates with anti-FAT10 Abs. Experiments (B–D) were independently performed three times. *p < 0.05, ** p < 0.01, *** p < 0.001.

FIGURE 1.

FAT10 overexpression following H5N1 virus infection. (A) The relative gene expression levels of ubiquitin and ULPs in a DNA microarray analysis of mouse lungs infected with live (black) or inactivated (dark gray) H5N1 virus and live H1N1 virus (light gray). This experiment was performed once with three mice in each group. (B) In vivo kinetic expression of FAT10 in H5N1-infected mouse lungs. BALB/c mice were infected with 106 TCID50 H5N1 or an AF control. FAT10 mRNA in mouse lungs at 24, 48, and 72 h was quantified by real-time RT-PCR. The mRNA levels were normalized to GAPDH (n = 3 mice/group). (C) FAT10 mRNA expression levels increased in a time-dependent manner in A549 cells from 3 to 72 h p.i. A549 cells were infected with H5N1 at an MOI of 5. An equal volume of AF was used as the control. The FAT10 mRNA levels at 3, 6, 12, 24, 48, and 72 h were determined by real-time RT-PCR. The mRNA levels were normalized to GAPDH. (D) A549 cells were infected with H5N1 at an MOI of 1 or 5 for 24 h. FAT10 expression was analyzed by Western blotting analysis of cell lysates with anti-FAT10 Abs. Experiments (B–D) were independently performed three times. *p < 0.05, ** p < 0.01, *** p < 0.001.

Close modal

We further analyzed FAT10 expression in a published gene-profiling dataset of human bronchial epithelial cells infected with a seasonal H1N1 virus vaccine strain BN/59, and identified a 1.58-fold increase in FAT10 mRNA expression in BN/59 virus–infected cells 36 h p.i. compared with control (GEO accession number GDS4855) (Supplemental Fig. 1A) (19). Analysis of another published microarray data on brain samples from vesicular stomatitis virus–infected mice showed that FAT10 was upregulated 2.9-fold compared with uninfected mice (GEO accession number GDS4842) (Supplemental Fig. 1B) (20).

To validate FAT10 upregulation after live H5N1 virus infection, we assayed FAT10 mRNA levels in the lung tissue of BALB/c mice using real-time PCR at different time points after intratracheal administration of live H5N1 virus or AF. The FAT10 mRNA levels in the live H5N1-infected group reached a peak 24 h p.i. and remained significantly high 48 h p.i., but the levels were comparable to the control group 72 h p.i. (Fig. 1B). We further assessed the FAT10 expression dynamics in A549 cells postinfection with the H5N1 virus. The FAT10 mRNA level slowly increased within 24 h after influenza virus infection and peaked 24 h p.i., then started to decrease from 48 h p.i. (Fig. 1C). FAT10 upregulation in A549 cells was confirmed by western blotting (Fig. 1D). The FAT10 protein level was significantly higher in H5N1-infected cells and was nearly undetectable in control cells. FAT10 overexpression was also observed in virus-infected BEAS-2B human bronchial epithelial cells (Supplemental Fig. 2A, 2B). These results were in accordance with the DNA microarray analysis and confirmed that H5N1 infection upregulated FAT10 expression.

These data showed that FAT10 expression was upregulated following live H5N1 virus infection, and suggested that FAT10 upregulation might be a general cellular response during virus infection.

Ubiquitin and ULPs not only target proteins for degradation but also participate in other biological processes, such as innate immunity and autophagy (21, 22). Therefore, we explored the functional role of FAT10 in virus/host interactions. One of the mechanisms underlying the host devastation caused by the avian influenza virus is epithelial cell death in the respiratory tract and lungs (23, 24). Therefore, we performed an MTS assay to evaluate the possible function of FAT10 in the viability of H5N1-infected A549 cells and primary HBEpiCs. First, we designed three siRNAs against the human FAT10 gene (siFAT10). A mixture of these siRNAs produced the best knockdown efficiency in transfection experiments based on the FAT10 mRNA level (Fig. 2A). Because the basal FAT10 protein level in A549 cells is rather low and beyond the detection limit of western blotting, we used IFN-γ and TNF-α to stimulate FAT10 expression 18 h prior to siFAT10 transfection (9, 10). The siFAT10 knockdown efficiency was confirmed by western blotting the IFN-γ– and TNF-α–stimulated groups, which ranged from clearly visible to not detectable (Fig. 2B). Then, A549 cells and primary HBEpiCs were transfected with siFAT10 or nonspecific control siRNA (siCtrl) and infected with live H5N1 48 h later. At 24 h p.i., the H5N1 virus induced significantly less cell death in the siFAT10-treated group compared with the siCtrl-treated group in both cell types (Fig. 2C, 2D). Virus-induced cell death was also reduced in siFAT10-treated BEAS-2B cells and CNE-2Z human nasopharyngeal carcinoma cells (Supplemental Fig. 2C, 2D), suggesting that the promotion of virus-induced cell death by FAT10 was a common mechanism.

FIGURE 2.

FAT10 promotes H5N1 virus replication in human lung epithelial cells. (A) The FAT10 mRNA level was significantly decreased by siFAT10. A549 cells were transfected with siFAT10 or siCtrl for 48 h. FAT10 mRNA was quantified by real-time RT-PCR. mRNA levels were normalized to GAPDH. (B) The FAT10 protein level was significantly knocked down by siFAT10. A549 cells were treated with 400 U/ml TNF-α and IFN-γ for 18 h and then transfected with siFAT10 or siCtrl for 48 h. FAT10 was detected by Western blotting. (C and D) siFAT10 significantly improved A549 (C) and HBEpiC (D) viability after H5N1 infection. The cells were transfected with siFAT10 or siCtrl for 48 h and then infected with H5N1 at an MOI of 5 for 24 h before viability assessment using the MTS assay. (E) M1 mRNA levels in H5N1 virus–infected cells increased in a time-dependent manner. A549 cells were infected with H5N1 at an MOI of 5. Viral M1 mRNA levels at different time points were quantified using real-time RT-PCR and were normalized to GAPDH. (F) M1 mRNA levels decreased in the siFAT10 group compared with the siCtrl group 12 and 24 h p.i. A549 cells were transfected with siFAT10 or siCtrl for 48 h and subsequently infected with H5N1 at an MOI of 5 for 12 or 24 h. (G) siFAT10 decreased NP protein levels in H5N1-infected cells. A549 cells were transfected with siFAT10 or siCtrl for 48 h and then infected with H5N1 virus for 24 h. The cell lysates were analyzed by Western blotting with Abs against the indicated proteins. Untreated A549 cells served as a blank control. (H) siFAT10 decreased viral titers 24 h p.i. The viral titers were determined at the indicated time points as the TCID50 per milliliter of supernatant for A549 cells pretreated with siFAT10 or siCtrl and then infected with H5N1 virus. (I) FAT10 was overexpressed using lentiviral vectors. A549 cells were transduced with lentiviral vectors encoding GFP (control) or FAT10. Then, the cells were infected with H5N1 for 12 h. The cell lysates were analyzed by Western blotting with Abs against the indicated proteins. (J and K) FAT10 overexpression using lentiviral vectors increased M1 mRNA levels (J) and viral titers (K) 24 h p.i. (L) H1N1 M1 mRNA levels were decreased in the siFAT10 group compared with the siCtrl group 24 h p.i. siUBA6 (M) and siUSE1 (N) reduced M1 mRNA levels 12 h p.i. A549 cells were transfected with UBA6- or USE1-specific siRNA or siCtrl and subsequently infected with H5N1 at an MOI of 5 for 12 h. Each experiment was independently performed three times. *p < 0.05, ** p < 0.01, *** p < 0.001.

FIGURE 2.

FAT10 promotes H5N1 virus replication in human lung epithelial cells. (A) The FAT10 mRNA level was significantly decreased by siFAT10. A549 cells were transfected with siFAT10 or siCtrl for 48 h. FAT10 mRNA was quantified by real-time RT-PCR. mRNA levels were normalized to GAPDH. (B) The FAT10 protein level was significantly knocked down by siFAT10. A549 cells were treated with 400 U/ml TNF-α and IFN-γ for 18 h and then transfected with siFAT10 or siCtrl for 48 h. FAT10 was detected by Western blotting. (C and D) siFAT10 significantly improved A549 (C) and HBEpiC (D) viability after H5N1 infection. The cells were transfected with siFAT10 or siCtrl for 48 h and then infected with H5N1 at an MOI of 5 for 24 h before viability assessment using the MTS assay. (E) M1 mRNA levels in H5N1 virus–infected cells increased in a time-dependent manner. A549 cells were infected with H5N1 at an MOI of 5. Viral M1 mRNA levels at different time points were quantified using real-time RT-PCR and were normalized to GAPDH. (F) M1 mRNA levels decreased in the siFAT10 group compared with the siCtrl group 12 and 24 h p.i. A549 cells were transfected with siFAT10 or siCtrl for 48 h and subsequently infected with H5N1 at an MOI of 5 for 12 or 24 h. (G) siFAT10 decreased NP protein levels in H5N1-infected cells. A549 cells were transfected with siFAT10 or siCtrl for 48 h and then infected with H5N1 virus for 24 h. The cell lysates were analyzed by Western blotting with Abs against the indicated proteins. Untreated A549 cells served as a blank control. (H) siFAT10 decreased viral titers 24 h p.i. The viral titers were determined at the indicated time points as the TCID50 per milliliter of supernatant for A549 cells pretreated with siFAT10 or siCtrl and then infected with H5N1 virus. (I) FAT10 was overexpressed using lentiviral vectors. A549 cells were transduced with lentiviral vectors encoding GFP (control) or FAT10. Then, the cells were infected with H5N1 for 12 h. The cell lysates were analyzed by Western blotting with Abs against the indicated proteins. (J and K) FAT10 overexpression using lentiviral vectors increased M1 mRNA levels (J) and viral titers (K) 24 h p.i. (L) H1N1 M1 mRNA levels were decreased in the siFAT10 group compared with the siCtrl group 24 h p.i. siUBA6 (M) and siUSE1 (N) reduced M1 mRNA levels 12 h p.i. A549 cells were transfected with UBA6- or USE1-specific siRNA or siCtrl and subsequently infected with H5N1 at an MOI of 5 for 12 h. Each experiment was independently performed three times. *p < 0.05, ** p < 0.01, *** p < 0.001.

Close modal

To determine whether the improved survival in the siFAT10-treated group was associated with viral replication, viral M1 gene mRNA levels (Fig. 2F), NP levels (Fig. 2G), and viral titers (Fig. 2H) were evaluated in H5N1 virus–infected A549 cells previously transfected with siFAT10. The quantity of M1 mRNA 3, 6, 12, and 24 h after H5N1 virus infection increased in a time-dependent manner (Fig. 2E). M1 mRNA levels were significantly reduced in the siFAT10 group compared with the control at 12 and 24 h p.i. (Fig. 2F). Specifically, an ∼75% reduction in M1 mRNA was observed 12 h p.i. in the siFAT10-treated A549 cells (Fig. 2F). In addition, FAT10 knockdown in BEAS-2B and CNE-2Z cells significantly reduced M1 mRNA levels (Supplemental Fig. 2E, 2F). Furthermore, the NP protein level was reduced in the siFAT10 group compared with the control by Western blotting (Fig. 2G, Supplemental Fig. 2G, 2H). The viral titer in siFAT10-treated cells was determined with a TCID50 assay 24 h p.i. and was also lower than the titer in siCtrl-treated cells (Fig. 2H). These results suggest that FAT10 knockdown inhibited the replication of live H5N1 virus. We determined the effect of FAT10 overexpression on viral replication using a lentivirus encoding FAT10 (Fig. 2I–K). The efficacy of FAT10 overexpression by the lentiviral vector was determined by immunoblotting (Fig. 2I). Both the M1 mRNA level and the viral titer as determined by TCID50 were higher in the FAT10-overexpressing groups compared with the control group (Fig. 2J, 2K). To test whether FAT10 increased the replication of other influenza viruses besides H5N1, we also used H1N1 infection in the FAT10 knockdown experiment (Fig. 2L). In H1N1 virus–infected A549 cells, FAT10 knockdown also inhibited H1N1 M1 gene expression (Fig. 2L). These data suggested that FAT10 was upregulated to promote viral replication during influenza A virus infection.

FAT10 targets FAT10ylated proteins for degradation via a proteasomal pathway that relies on specific enzymes, including the ubiquitin-activating E1 enzyme UBA6 (7) and the ubiquitin-conjugating E2 enzyme USE1 (8). To determine whether FAT10ylation was essential for FAT10 function during live H5N1 infection, we individually knocked down UBA6 and USE1 expression. M1 mRNA levels were notably reduced in the siUBA6 and siUSE1 groups 12 h p.i. (Fig. 2M, 2N). These data suggest that FAT10 promotes H5N1 virus replication and that the FAT10ylation system might be involved in this process.

Because FAT10 upregulation via H5N1 virus infection was important for viral replication, we investigated which component of the H5N1 virus was responsible for inducing FAT10 expression. We transfected 293T cells with plasmids encoding the H5N1 viral proteins HA, NA (N1), NP, PA, M1, M2, NS1, NS2, PB1-F2, PB1, and PB2 (Fig. 3A, Supplemental Fig. 3A) and a plasmid encoding GFP was used as a negative control. Although two of the viral proteins (M2 and HA) upregulated FAT10 mRNA levels slightly, the expression of each viral protein alone did not account for the FAT10 upregulation observed during influenza infection. Moreover, transfection with the viral nonstructural protein (NS1) significantly decreased FAT10 mRNA levels (Fig. 3A). Therefore, we further explored the possible role of H5N1 virus RNA in FAT10 induction. Total RNA extracted from H5N1 virus–infected cells was used as a model trigger, and total RNA from uninfected cells was used as a control (25). The FAT10 mRNA level in A549 cells 12 h posttransfection with RNA extracted from H5N1 virus–infected cells was significantly higher compared with the control (Fig. 3B). These data suggest that RNA from the H5N1 virus is the viral component inducing FAT10 upregulation.

FIGURE 3.

FAT10 expression is induced by RNA from H5N1 virus–infected cells via the RIG-I-NF-κB signaling pathway. (A) H5N1 viral proteins were screened to determine whether they had a role in FAT10 upregulation. Viral protein expression plasmids were transfected into 293T cells for 24 h. FAT10 mRNA was quantified by real-time RT-PCR. The mRNA levels were normalized to GAPDH. (B) Viral RNA transfection significantly increased FAT10 mRNA levels. A549 cells were transfected with 1 μg/ml viral RNA or control cellular RNA for 12 h. (C) siRIG-I and siP65 dramatically repressed FAT10 induction by viral RNA. A549 cells were transfected with the corresponding siRNAs and then incubated with 1 μg/ml viral RNA for 12 h. (DI) The effects of siRNAs on their corresponding targets in the knockdown assays shown in (C) were confirmed. (J) 293T cells were transfected with a wild-type (WT) or mutated (MT) FAT10 promoter–driven firefly luciferase reporter. Then, the cells were transfected with RNA from H5N1-infected cells for 12 h. (K) 293T cells were cotransfected with the corresponding protein-coding plasmid and a FAT10 promoter–driven firefly luciferase reporter. The luciferase activity is presented as the fold induction compared with vector-transfected cells. Each experiment was independently performed three times. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 3.

FAT10 expression is induced by RNA from H5N1 virus–infected cells via the RIG-I-NF-κB signaling pathway. (A) H5N1 viral proteins were screened to determine whether they had a role in FAT10 upregulation. Viral protein expression plasmids were transfected into 293T cells for 24 h. FAT10 mRNA was quantified by real-time RT-PCR. The mRNA levels were normalized to GAPDH. (B) Viral RNA transfection significantly increased FAT10 mRNA levels. A549 cells were transfected with 1 μg/ml viral RNA or control cellular RNA for 12 h. (C) siRIG-I and siP65 dramatically repressed FAT10 induction by viral RNA. A549 cells were transfected with the corresponding siRNAs and then incubated with 1 μg/ml viral RNA for 12 h. (DI) The effects of siRNAs on their corresponding targets in the knockdown assays shown in (C) were confirmed. (J) 293T cells were transfected with a wild-type (WT) or mutated (MT) FAT10 promoter–driven firefly luciferase reporter. Then, the cells were transfected with RNA from H5N1-infected cells for 12 h. (K) 293T cells were cotransfected with the corresponding protein-coding plasmid and a FAT10 promoter–driven firefly luciferase reporter. The luciferase activity is presented as the fold induction compared with vector-transfected cells. Each experiment was independently performed three times. *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

Viral RNA typically initiates intracellular antiviral responses, including type I IFN production, through an array of transmembrane and cytosolic receptors. TLR3 preferentially localizes to the endosome and recognizes viral dsRNA released by some viruses, such as retroviruses, inside endosomes after viral internalization (26). Two RNA helicases (RIG-I (27) and melanoma differentiation–associated gene 5 (MDA-5) (28)) recognize cytosolic viral dsRNA synthesized during viral replication. However, negative-strand RNA viruses, such as the influenza A virus, generate little dsRNA during replication. In this case, RIG-I is activated by uncapped viral genomic 5′-phosphate single-stranded RNA (ssRNA) (2931). To investigate the signaling pathways involved in FAT10 upregulation by H5N1 ssRNA, we first examined the roles of the three main cellular RNA sensors (RIG-I, MDA5, and TLR3) using their corresponding siRNAs (siRIG-I, siMDA5, and siTLR3) (Fig. 3C). Consistent with the evidence that no dsRNA signals were detected in cells infected with influenza virus (29, 30), the FAT10 upregulation induced by RNA extracted from H5N1-infected cells was dramatically repressed by treatment with siRIG-I but not siMDA5 or siTLR3 (Fig. 3C). Then, the transcription factors downstream of RIG-I (IRF-3, IRF-7, and NF-κB) were evaluated with knockdown assays using siRNAs (siIRF3, siIRF7, and sip65). p65 (Rel A) is a main subunit of the most widely studied and abundant form of the NF-κB heterodimer, p50 and p65. The upregulation of FAT10 by RNA extracted from H5N1 virus-infected cells was repressed by treatment with sip65 but not by siIRF3 or siIRF7 (Fig. 3C), suggesting that the ssRNA-RIG-I-NF-κB axis upregulated FAT10 expression. The effect of siRNAs on these genes was verified in additional experiments (Fig. 3D–I). The transcriptional effect of p65 on FAT10 was confirmed by mutating the p65 response element in the FAT10 promoter and assessing with the luciferase reporter system. Luciferase cDNA was placed under FAT10 promoter control. We mutated the putative p65-binding site between +142 and +151 bp downstream of the transcription start site in the FAT10 gene and tested luciferase activity using a dual luciferase reporter assay. Luciferase responsiveness to viral RNA was significantly reduced by this mutation, indicating that p65 directly regulated FAT10 gene transcription (Fig. 3J). We further confirmed the RIG-I-NF-κB-FAT10 signaling pathways by overexpressing RIG-I, ΔRIG-I, IRF3, or p65 together with the FAT10-Luc reporter plasmids in 293T cells (Fig. 3K, Supplemental Fig. 3B). Consistent with our previous data, p65 overexpression dramatically increased FAT10 promoter activity, whereas IRF3 overexpression did not (Fig. 3K). For RIG-I, full length RIG-I had no effect on FAT10 promoter activity (Fig. 3K). The RIG-I protein contains two N-terminal caspase recruitment domains (CARD), a helicase and a C-terminal regulatory domain. CARD transduces signals and leads to IRF-3 and NF-κB activation. The CARD domains have been proposed to be autoinhibited by other domains of the protein (32) (Supplemental Fig. 3C). Next, we constructed ΔRIG-I containing mainly the CARD domains, which constitutively activated downstream IRF3 and NF-κB signals according to previous studies (27) (Supplemental Fig. 3C). ΔRIG-I overexpression significantly increased FAT10 promoter-driven luciferase activity (Fig. 3K).

Altogether, these data suggest that FAT10 upregulation was mediated by RIG-I, the H5N1 genomic ssRNA sensor, and involved the activation of NF-κB (Fig. 4G). Thus, the ssRNA-RIG-I-NF-κB axis, which was originally a pathway to elicit antiviral responses during viral infection, was involved in a regulatory response following H5N1 virus infection to upregulate FAT10 and to promote viral replication.

FIGURE 4.

Knockdown of FAT10 promoted type I IFN expression and STAT1 phosphorylation. (AD) siFAT10 significantly increased the mRNA levels of IFN-α (A and C) and IFN-β (B and D) 24 h p.i. compared with the siCtrl. A549 cells were transfected with siFAT10 or siCtrl and subsequently infected with H5N1 (A and B) or H1N1 (C and D) at an MOI of 5 for 24 h. The IFN-α and IFN-β mRNA levels were quantified using real-time RT-PCR, and mRNA levels were normalized to GAPDH. (E and F) Western blotting analysis of phospho (p)-STAT1 in A549 cells infected with H5N1 at an MOI of 5 for 12 or 24 h (E) or transfected with siFAT10 or siCtrl and subsequently infected with H5N1 at an MOI of 5 for 12 h (F). The blots were analyzed with anti–p-STAT1, anti-STAT1 and anti–β-actin Abs. The bar graphs show the relative abundance of the p-STAT1 protein (normalized to STAT1) from three experiments. Untreated A549 cells served as a blank control. (G) A proposed model for the regulation of H5N1 infection by FAT10. H5N1 ssRNA induced FAT10 transcription via the RIG-I-NF-κB signaling pathway, and the NS1 protein from H5N1 negatively regulated FAT10 expression. FAT10 promoted viral replication and inhibited IFN-α/β expression and downstream STAT1 phosphorylation. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 4.

Knockdown of FAT10 promoted type I IFN expression and STAT1 phosphorylation. (AD) siFAT10 significantly increased the mRNA levels of IFN-α (A and C) and IFN-β (B and D) 24 h p.i. compared with the siCtrl. A549 cells were transfected with siFAT10 or siCtrl and subsequently infected with H5N1 (A and B) or H1N1 (C and D) at an MOI of 5 for 24 h. The IFN-α and IFN-β mRNA levels were quantified using real-time RT-PCR, and mRNA levels were normalized to GAPDH. (E and F) Western blotting analysis of phospho (p)-STAT1 in A549 cells infected with H5N1 at an MOI of 5 for 12 or 24 h (E) or transfected with siFAT10 or siCtrl and subsequently infected with H5N1 at an MOI of 5 for 12 h (F). The blots were analyzed with anti–p-STAT1, anti-STAT1 and anti–β-actin Abs. The bar graphs show the relative abundance of the p-STAT1 protein (normalized to STAT1) from three experiments. Untreated A549 cells served as a blank control. (G) A proposed model for the regulation of H5N1 infection by FAT10. H5N1 ssRNA induced FAT10 transcription via the RIG-I-NF-κB signaling pathway, and the NS1 protein from H5N1 negatively regulated FAT10 expression. FAT10 promoted viral replication and inhibited IFN-α/β expression and downstream STAT1 phosphorylation. *p < 0.05, **p < 0.01, ***p < 0.001.

Close modal

How does the FAT10 upregulation promote viral replication? IFNs are the most important proteins produced by host cells in response to pathogens. Type I IFNs, including IFN-α, IFN-β, and IFN-ω, are induced by viral infection. IFN-γ is a type II IFN that is induced by mitogenic or antigenic stimuli. Most virus-infected cells are capable of synthesizing IFNα/β as a host defense against viral infection and replication. When FAT10 expression was repressed by siFAT10 during H5N1 virus infection, the mRNA levels of type I IFNs rebounded significantly 24 h p.i. compared with the control group (Fig. 4A, 4B). FAT10 knockdown in H1N1 virus–infected A549 cells also promoted the expression of type I IFNs (Fig. 4C, 4D). Thus, FAT10 might promote viral replication by inhibiting the IFN response during influenza A viral infection.

IFN-mediated signaling and transcriptional activation occur primarily through the JAK-STAT signal transduction pathway. STAT1 plays a central role mediating the IFN-induced antiviral state. Many viruses encode proteins that impair the activity of this signaling pathway. STAT1 phosphorylation status is typically considered crucial to induce a cellular antiviral state, including the transcriptional activation of IFN-stimulated genes (33, 34). Therefore, we tested STAT1 phosphorylation levels in A549 cells. STAT1 phosphorylation was gradually upregulated during H5N1 virus infection (Fig. 4E). STAT1 phosphorylation was further increased in A549 cells infected by live H5N1 virus after FAT10 knockdown (Fig. 4F). Taken together, these data suggest that FAT10 inhibited IFN-α/β induction during viral infection and suppressed IFN signaling pathway activity by inhibiting STAT1 phosphorylation. Therefore, during H5N1 virus infection, FAT10 expression might be upregulated in a regulatory response to antagonize the IFN-induced antiviral response and promote viral replication.

The role of FAT10 and FAT10ylation in the interaction between host cells and influenza viruses such as the H5N1 virus was previously unknown. In this study, our experiments demonstrated that, during H5N1 virus infection, FAT10 expression was upregulated via the RIG-I-NF-κB signaling pathway, whereas upregulated FAT10 promoted H5N1 viral replication by inhibiting Type I IFN expression and STAT1 phosphorylation.

There are many similarities between our microarray data and data from other groups examining the influenza A virus infection model (17, 19, 35, 36). The robust upregulation of cytokine genes including CXCL10 observed in our microarray data were also detected in microarray experiments performed by Lee et al. (35), Cameron et al. (17), Chakrabarti et al. (36), and Gerlach et al. (19). Type I IFN–stimulated genes were also observed in these reports. Because cytokine response dysregulation and cell death is important for H5N1 virus pathogenesis, many studies have intensively analyzed the pathways and related genes. The importance of CXCL10 in the pathogenesis of influenza infection was recently reported by our laboratory (37). However, there are also some discrepancies between these published data because of differences in experimental designs and models.

Our microarray data detected dramatic FAT10 upregulation in the H5N1 and H1N1 virus–infected group compared with the AF control. When we further investigated the role of FAT10 upregulation, we found that it was critical in influenza A virus replication. FAT10 knockdown not only inhibited H5N1 and H1N1 viral gene expression but also restricted the H5N1 viral titer in the supernatants of infected cells. Correspondingly, FAT10 overexpression by lentiviral transduction increased the H5N1 viral titer. Moreover, data mining of the gene array data in the GEO database found that in the alphavirus M1 susceptible human hepatocellular carcinoma cell line Hep3B, FAT10 was significantly upregulated (332-fold) as compared with the alphavirus M1 resistant human fetal hepatic cell line L-02 (GEO accession number GSE54342) (Supplemental Fig. 1C) (38). In addition, another microarray data demonstrated a 2-fold FAT10 upregulation in ribavirin-resistant HCV-infected cell lines compared with the original ribavirin-sensitive HCV RNA-replicating cell line (GEO accession number GSE60948) (Supplemental Fig. 1D). Upregulated FAT10 seems to be responsible for the cell susceptibility to virus and the cell resistance to ribavirin. These results also shed light on our hypothesis that FAT10 upregulation promotes influenza A viral replication. Thus, FAT10 might serve as a common host response factor during influenza viral infection and possibly during infection by other viruses to promote viral replication, to make cells more easily infected by viruses or more resistant to antiviral drugs. Nevertheless, more viral strains must be investigated to validate this conclusion.

FAT10 upregulation during H5N1 virus infection was finally attributed to viral ssRNA, whereas NS1 alone controversially inhibited FAT10 expression in our study. The interactions between RIG-I and RNA viruses have been intensively studied (27, 32). RIG-I normally triggers the antiviral response upon activation by dsRNA or ssRNA and activates downstream transcription factors (i.e., IRF3 and NF-κB) to drive type I IFN production, thereby eliciting an innate immune response to counteract viral infection (Fig. 4G) (29, 32, 39). However, our results demonstrated that following H5N1 virus infection, a regulatory response via the host antiviral ssRNA-RIG-I-NF-κB signaling pathway upregulated FAT10 to enhance viral replication (Fig. 4G). On the other hand, a previous study found that the influenza A virus NS1 protein formed a complex with RIG-I and inhibited the RIG-I signaling cascade in response to viral infection (29, 40). Therefore, it is reasonable that NS1 was found to inhibit RIG-I-mediated FAT10 upregulation in our study (Fig. 4G).

We demonstrated that FAT10 knockdown ameliorated the cell death caused by H5N1 virus infection in different cell lines and in primary human cell HBEpiC. This result was notable because severe cell death is an important aspect of H5N1 virus pathogenesis, and these data provide an interesting area for exploration that was not addressed here. Future studies should investigate the precise mechanism underlying this finding. The observed reduction in cell death could be due to decreased viral replication or interference with the apoptosis pathway in the siFAT10-treated group. The identification of the mechanistic details will require further investigation.

IFN-α and IFN-β expression are important host cell responses to influenza infection. STAT1 is a key member of the downstream JAK/STAT pathway, which is an important host cell pathway that fights against pathogen invasion. Many viruses develop strategies to antagonize and block IFN-α/β signaling, including mimicking the cellular components involved in IFN signal transduction and inhibiting the expression of the STAT1 and JAK-2 proteins (41). In our study, we demonstrated the inhibitory effect of FAT10 on IFN-α and IFN-β expression and on STAT1 phosphorylation, because FAT10 knockdown promoted type I IFN expression and STAT1 phosphorylation during H5N1 infection. We also showed that STAT1 phosphorylation was elevated during H5N1 virus infection, demonstrating a role for STAT1 in immunity against H5N1 virus infection, which is consistent with recent data (42). Therefore, the inhibitory effect of FAT10 on type I IFN expression and STAT1 phosphorylation represents an important mechanism underlying the enhancement of H5N1 virus replication by FAT10.

In this study, we discovered that, following live H5N1 virus infection, FAT10 expression was upregulated in both mouse in vivo and human respiratory epithelial cell lines in vitro. FAT10 promoted influenza A replication and decreased the viability of infected cells. Viral RNA, not viral proteins, significantly increased FAT10 expression through the RIG-I-NF-κB signaling pathway. We examined the functional roles of FAT10 during live H5N1 virus infection in A549 cells and found that FAT10 inhibited type I IFN expression and downstream STAT1 phosphorylation. We hypothesized a negative response model for the antiviral immune response through the RIG-I-NF-κB-type I IFN signaling pathway involving regulation of FAT10 expression during live H5N1 infection (Fig. 4G). Our results provide new insights into the role of the ULP FAT10 in the interaction between host cells and influenza virus and suggest that the FAT10 protein is a potential new therapeutic target for the treatment of H5N1 virus infection.

We thank Annette Aichem for providing FAT10 expression plasmid and Ting Xie (Cedars Sinai Medical Center, Los Angeles, CA) for assistance with language editing.

This work was supported by Ministry of Science and Technology of China Grant 2015CB553400, Natural Science Foundation of China Grants 81230002 and 81570077, 111 Project Grant B08007, the Peking Union Medical College Youth Fund, and Fundamental Research Funds for the Central Universities Grant 3332013120. C.J. is a Hsien Wu Professor of Biochemistry.

The sequences presented in this article have been submitted to the National Center for Biotechnology Information's Gene Expression Omnibus (http://www.ncbi.nlm.nih.gov/geo/) under accession number GSE76719.

The online version of this article contains supplemental material.

Abbreviations used in this article:

AF

allantoic fluid

CARD

N-terminal caspase recruitment domain

GEO

Gene Expression Omnibus

HBEpiC

human bronchial epithelial cell

IRF

IFN regulatory factor

MDA-5

melanoma differentiation–associated gene 5

MDCK

Madin–Darby canine kidney

MOI

multiplicity of infection

MT

mutated

NP

nucleoprotein

NS1

nonstructural protein 1

p.i.

postinfection

RIG-I

retinoic acid–induced protein I

siRNA

small interfering RNA

UBD

ubiquitin D

ULP

ubiquitin-like protein

WT

wild-type.

1
Subbarao
K.
,
Klimov
A.
,
Katz
J.
,
Regnery
H.
,
Lim
W.
,
Hall
H.
,
Perdue
M.
,
Swayne
D.
,
Bender
C.
,
Huang
J.
, et al
.
1998
.
Characterization of an avian influenza A (H5N1) virus isolated from a child with a fatal respiratory illness.
Science
279
:
393
396
.
2
World Health Organization. 2015. Cumulative number of confirmed human cases of avian influenza A(H5N1) reported to WHO. Available at: http://www.who.int/influenza/human_animal_interface/EN_GIP_201503031cumulativeNumberH5N1cases.pdf?ua=1
.
3
Yuen
K. Y.
,
Chan
P. K.
,
Peiris
M.
,
Tsang
D. N.
,
Que
T. L.
,
Shortridge
K. F.
,
Cheung
P. T.
,
To
W. K.
,
Ho
E. T.
,
Sung
R.
,
Cheng
A. F.
.
1998
.
Clinical features and rapid viral diagnosis of human disease associated with avian influenza A H5N1 virus.
Lancet
351
:
467
471
.
4
Stertz
S.
,
Shaw
M. L.
.
2011
.
Uncovering the global host cell requirements for influenza virus replication via RNAi screening.
Microbes Infect.
13
:
516
525
.
5
Brass
A. L.
,
Huang
I. C.
,
Benita
Y.
,
John
S. P.
,
Krishnan
M. N.
,
Feeley
E. M.
,
Ryan
B. J.
,
Weyer
J. L.
,
van der Weyden
L.
,
Fikrig
E.
, et al
.
2009
.
The IFITM proteins mediate cellular resistance to influenza A H1N1 virus, West Nile virus, and dengue virus.
Cell
139
:
1243
1254
.
6
Raasi
S.
,
Schmidtke
G.
,
Groettrup
M.
.
2001
.
The ubiquitin-like protein FAT10 forms covalent conjugates and induces apoptosis.
J. Biol. Chem.
276
:
35334
35343
.
7
Chiu
Y. H.
,
Sun
Q.
,
Chen
Z. J.
.
2007
.
E1-L2 activates both ubiquitin and FAT10.
Mol. Cell
27
:
1014
1023
.
8
Aichem
A.
,
Pelzer
C.
,
Lukasiak
S.
,
Kalveram
B.
,
Sheppard
P. W.
,
Rani
N.
,
Schmidtke
G.
,
Groettrup
M.
.
2010
.
USE1 is a bispecific conjugating enzyme for ubiquitin and FAT10, which FAT10ylates itself in cis.
Nat. Commun.
1
:
13
.
9
Raasi
S.
,
Schmidtke
G.
,
de Giuli
R.
,
Groettrup
M.
.
1999
.
A ubiquitin-like protein which is synergistically inducible by interferon-γ and tumor necrosis factor-α.
Eur. J. Immunol.
29
:
4030
4036
.
10
Liu
Y. C.
,
Pan
J.
,
Zhang
C.
,
Fan
W.
,
Collinge
M.
,
Bender
J. R.
,
Weissman
S. M.
.
1999
.
A MHC-encoded ubiquitin-like protein (FAT10) binds noncovalently to the spindle assembly checkpoint protein MAD2.
Proc. Natl. Acad. Sci. USA
96
:
4313
4318
.
11
Ross
M. J.
,
Wosnitzer
M. S.
,
Ross
M. D.
,
Granelli
B.
,
Gusella
G. L.
,
Husain
M.
,
Kaufman
L.
,
Vasievich
M.
,
D’Agati
V. D.
,
Wilson
P. D.
, et al
.
2006
.
Role of ubiquitin-like protein FAT10 in epithelial apoptosis in renal disease.
J. Am. Soc. Nephrol.
17
:
996
1004
.
12
Snyder
A.
,
Alsauskas
Z.
,
Gong
P.
,
Rosenstiel
P. E.
,
Klotman
M. E.
,
Klotman
P. E.
,
Ross
M. J.
.
2009
.
FAT10: a novel mediator of Vpr-induced apoptosis in human immunodeficiency virus-associated nephropathy.
J. Virol.
83
:
11983
11988
.
13
Lee
C. G. L.
,
Ren
J.
,
Cheong
I. S. Y.
,
Ban
K. H. K.
,
Ooi
L. L. P. J.
,
Yong Tan
S.
,
Kan
A.
,
Nuchprayoon
I.
,
Jin
R.
,
Lee
K. H.
, et al
.
2003
.
Expression of the FAT10 gene is highly upregulated in hepatocellular carcinoma and other gastrointestinal and gynecological cancers.
Oncogene
22
:
2592
2603
.
14
Aichem
A.
,
Kalveram
B.
,
Spinnenhirn
V.
,
Kluge
K.
,
Catone
N.
,
Johansen
T.
,
Groettrup
M.
.
2012
.
The proteomic analysis of endogenous FAT10 substrates identifies p62/SQSTM1 as a substrate of FAT10ylation.
J. Cell Sci.
125
:
4576
4585
.
15
Reed
L. J.
,
Muench
H.
.
1938
.
A simple method of estimating fifty per cent endpoints.
Am. J. Hyg.
27
.
16
Zhang
D. W.
,
Jeang
K. T.
,
Lee
C. G.
.
2006
.
p53 negatively regulates the expression of FAT10, a gene upregulated in various cancers.
Oncogene
25
:
2318
2327
.
17
Cameron
C. M.
,
Cameron
M. J.
,
Bermejo-Martin
J. F.
,
Ran
L.
,
Xu
L.
,
Turner
P. V.
,
Ran
R.
,
Danesh
A.
,
Fang
Y.
,
Chan
P. K. M.
, et al
.
2008
.
Gene expression analysis of host innate immune responses during lethal H5N1 infection in ferrets.
J. Virol.
82
:
11308
11317
.
18
Cilloniz
C.
,
Pantin-Jackwood
M. J.
,
Ni
C.
,
Goodman
A. G.
,
Peng
X.
,
Proll
S. C.
,
Carter
V. S.
,
Rosenzweig
E. R.
,
Szretter
K. J.
,
Katz
J. M.
, et al
.
2010
.
Lethal dissemination of H5N1 influenza virus is associated with dysregulation of inflammation and lipoxin signaling in a mouse model of infection.
J. Virol.
84
:
7613
7624
.
19
Gerlach
R. L.
,
Camp
J. V.
,
Chu
Y. K.
,
Jonsson
C. B.
.
2013
.
Early host responses of seasonal and pandemic influenza A viruses in primary well-differentiated human lung epithelial cells.
PLoS One
8
:
e78912
.
20
Hou
Y. J.
,
Banerjee
R.
,
Thomas
B.
,
Nathan
C.
,
García-Sastre
A.
,
Ding
A.
,
Uccellini
M. B.
.
2013
.
SARM is required for neuronal injury and cytokine production in response to central nervous system viral infection.
J. Immunol.
191
:
875
883
.
21
Harty
R. N.
,
Pitha
P. M.
,
Okumura
A.
.
2009
.
Antiviral activity of innate immune protein ISG15.
J. Innate Immun.
1
:
397
404
.
22
Bedford
L.
,
Lowe
J.
,
Dick
L. R.
,
Mayer
R. J.
,
Brownell
J. E.
.
2011
.
Ubiquitin-like protein conjugation and the ubiquitin-proteasome system as drug targets.
Nat. Rev. Drug Discov.
10
:
29
46
.
23
Sun
Y.
,
Li
C.
,
Shu
Y.
,
Ju
X.
,
Zou
Z.
,
Wang
H.
,
Rao
S.
,
Guo
F.
,
Liu
H.
,
Nan
W.
, et al
.
2012
.
Inhibition of autophagy ameliorates acute lung injury caused by avian influenza A H5N1 infection.
Sci. Signal.
5
:
ra16
.
24
Daidoji
T.
,
Koma
T.
,
Du
A.
,
Yang
C. S.
,
Ueda
M.
,
Ikuta
K.
,
Nakaya
T.
.
2008
.
H5N1 avian influenza virus induces apoptotic cell death in mammalian airway epithelial cells.
J. Virol.
82
:
11294
11307
.
25
Kato
H.
,
Takeuchi
O.
,
Mikamo-Satoh
E.
,
Hirai
R.
,
Kawai
T.
,
Matsushita
K.
,
Hiiragi
A.
,
Dermody
T. S.
,
Fujita
T.
,
Akira
S.
.
2008
.
Length-dependent recognition of double-stranded ribonucleic acids by retinoic acid-inducible gene-I and melanoma differentiation-associated gene 5.
J. Exp. Med.
205
:
1601
1610
.
26
Kawai
T.
,
Akira
S.
.
2010
.
The role of pattern-recognition receptors in innate immunity: update on Toll-like receptors.
Nat. Immunol.
11
:
373
384
.
27
Yoneyama
M.
,
Kikuchi
M.
,
Natsukawa
T.
,
Shinobu
N.
,
Imaizumi
T.
,
Miyagishi
M.
,
Taira
K.
,
Akira
S.
,
Fujita
T.
.
2004
.
The RNA helicase RIG-I has an essential function in double-stranded RNA-induced innate antiviral responses.
Nat. Immunol.
5
:
730
737
.
28
Yoneyama
M.
,
Kikuchi
M.
,
Matsumoto
K.
,
Imaizumi
T.
,
Miyagishi
M.
,
Taira
K.
,
Foy
E.
,
Loo
Y. M.
,
Gale
M.
 Jr.
,
Akira
S.
, et al
.
2005
.
Shared and unique functions of the DExD/H-box helicases RIG-I, MDA5, and LGP2 in antiviral innate immunity.
J. Immunol.
175
:
2851
2858
.
29
Pichlmair
A.
,
Schulz
O.
,
Tan
C. P.
,
Näslund
T. I.
,
Liljeström
P.
,
Weber
F.
,
Reis e Sousa
C.
.
2006
.
RIG-I-mediated antiviral responses to single-stranded RNA bearing 5′-phosphates.
Science
314
:
997
1001
.
30
Weber
F.
,
Wagner
V.
,
Rasmussen
S. B.
,
Hartmann
R.
,
Paludan
S. R.
.
2006
.
Double-stranded RNA is produced by positive-strand RNA viruses and DNA viruses but not in detectable amounts by negative-strand RNA viruses.
J. Virol.
80
:
5059
5064
.
31
Hornung
V.
,
Ellegast
J.
,
Kim
S.
,
Brzózka
K.
,
Jung
A.
,
Kato
H.
,
Poeck
H.
,
Akira
S.
,
Conzelmann
K. K.
,
Schlee
M.
, et al
.
2006
.
5′-Triphosphate RNA is the ligand for RIG-I.
Science
314
:
994
997
.
32
Loo
Y. M.
,
Gale
M.
 Jr.
2011
.
Immune signaling by RIG-I-like receptors.
Immunity
34
:
680
692
.
33
Durbin
J. E.
,
Hackenmiller
R.
,
Simon
M. C.
,
Levy
D. E.
.
1996
.
Targeted disruption of the mouse Stat1 gene results in compromised innate immunity to viral disease.
Cell
84
:
443
450
.
34
Meraz
M. A.
,
White
J. M.
,
Sheehan
K. C.
,
Bach
E. A.
,
Rodig
S. J.
,
Dighe
A. S.
,
Kaplan
D. H.
,
Riley
J. K.
,
Greenlund
A. C.
,
Campbell
D.
, et al
.
1996
.
Targeted disruption of the Stat1 gene in mice reveals unexpected physiologic specificity in the JAK-STAT signaling pathway.
Cell
84
:
431
442
.
35
Lee
S. M.
,
Gardy
J. L.
,
Cheung
C. Y.
,
Cheung
T. K.
,
Hui
K. P.
,
Ip
N. Y.
,
Guan
Y.
,
Hancock
R. E.
,
Peiris
J. S.
.
2009
.
Systems-level comparison of host-responses elicited by avian H5N1 and seasonal H1N1 influenza viruses in primary human macrophages.
PLoS One
4
:
e8072
.
36
Chakrabarti
A. K.
,
Vipat
V. C.
,
Mukherjee
S.
,
Singh
R.
,
Pawar
S. D.
,
Mishra
A. C.
.
2010
.
Host gene expression profiling in influenza A virus-infected lung epithelial (A549) cells: a comparative analysis between highly pathogenic and modified H5N1 viruses.
Virol. J.
7
:
219
.
37
Wang
W.
,
Yang
P.
,
Zhong
Y.
,
Zhao
Z.
,
Xing
L.
,
Zhao
Y.
,
Zou
Z.
,
Zhang
Y.
,
Li
C.
,
Li
T.
, et al
.
2013
.
Monoclonal antibody against CXCL-10/IP-10 ameliorates influenza A (H1N1) virus induced acute lung injury.
Cell Res.
23
:
577
580
.
38
Lin
Y.
,
Zhang
H.
,
Liang
J.
,
Li
K.
,
Zhu
W.
,
Fu
L.
,
Wang
F.
,
Zheng
X.
,
Shi
H.
,
Wu
S.
, et al
.
2014
.
Identification and characterization of alphavirus M1 as a selective oncolytic virus targeting ZAP-defective human cancers.
Proc. Natl. Acad. Sci. USA
111
:
E4504
E4512
.
39
Panne
D.
2008
.
The enhanceosome.
Curr. Opin. Struct. Biol.
18
:
236
242
.
40
Marc
D.
2014
.
Influenza virus non-structural protein NS1: interferon antagonism and beyond.
J. Gen. Virol.
95
:
2594
2611
.
41
Samuel
C. E.
2001
.
Antiviral actions of interferons.
Clin. Microbiol. Rev.
14
:
778
809
.
42
Iwasaki
A.
,
Pillai
P. S.
.
2014
.
Innate immunity to influenza virus infection.
Nat. Rev. Immunol.
14
:
315
328
.

The authors have no financial conflicts of interest.

Supplementary data