PARP1 Enhances Influenza A Virus Propagation by Facilitating Degradation of Host Type I Interferon Receptor

Influenza A virus (IAV) infections cause seasonal and pandemic influenza outbreaks, which pose a devastating global health concern. Despite the availability of antivirals against influenza, new IAV strains continue to persist by overcoming the therapeutics. Therefore, much emphasis in the field is placed on identifying new therapeutic targets that can more effectively control influenza. IAV utilizes several tactics to evade host innate immunity, which include the evasion of antiviral type I interferon (IFN) responses. Degradation of type I IFN receptor (IFNAR) is one known method of subversion, but the molecular mechanism for IFNAR downregulation during IAV infection remains unclear. Here, we have found that a host protein, poly(ADP-ribose) polymerase 1 (PARP1), facilitates IFNAR degradation and accelerates IAV replication. The findings reveal a novel cellular target for the potential development of antivirals against influenza, as well as expand our base of knowledge regarding interactions between influenza and the host innate immunity.

ABSTRACT

Influenza A virus (IAV) utilizes multiple strategies to confront or evade host type I interferon (IFN)-mediated antiviral responses in order to enhance its own propagation within the host. One such strategy is to induce the degradation of type I IFN receptor 1 (IFNAR1) by utilizing viral hemagglutinin (HA). However, the molecular mechanism behind this process is poorly understood. Here, we report that a cellular protein, poly(ADP-ribose) polymerase 1 (PARP1), plays a critical role in mediating IAV HA-induced degradation of IFNAR1. We identified PARP1 as an interacting partner for IAV HA through mass spectrometry analysis. This interaction was confirmed by coimmunoprecipitation analyses. Furthermore, confocal fluorescence microscopy showed altered localization of endogenous PARP1 upon transient IAV HA expression or during IAV infection. Knockdown or inhibition of PARP1 rescued IFNAR1 levels upon IAV infection or HA expression, exemplifying the importance of PARP1 for IAV-induced reduction of IFNAR1. Notably, PARP1 was crucial for the robust replication of IAV, which was associated with regulation of the type I IFN receptor signaling pathway. These results indicate that PARP1 promotes IAV replication by controlling viral HA-induced degradation of host type I IFN receptor. Altogether, these findings provide novel insight into interactions between influenza virus and the host innate immune response and reveal a new function for PARP1 during influenza virus infection.

IMPORTANCE Influenza A virus (IAV) infections cause seasonal and pandemic influenza outbreaks, which pose a devastating global health concern. Despite the availability of antivirals against influenza, new IAV strains continue to persist by overcoming the therapeutics. Therefore, much emphasis in the field is placed on identifying new therapeutic targets that can more effectively control influenza. IAV utilizes several tactics to evade host innate immunity, which include the evasion of antiviral type I interferon (IFN) responses. Degradation of type I IFN receptor (IFNAR) is one known method of subversion, but the molecular mechanism for IFNAR downregulation during IAV infection remains unclear. Here, we have found that a host protein, poly(ADP-ribose) polymerase 1 (PARP1), facilitates IFNAR degradation and accelerates IAV replication. The findings reveal a novel cellular target for the potential development of antivirals against influenza, as well as expand our base of knowledge regarding interactions between influenza and the host innate immunity.

INTRODUCTION

Influenza A viruses (IAV) cause seasonal and pandemic influenza outbreaks leading to unpredictably severe morbidity and mortality worldwide (1, 2). The type I interferon (IFN) innate immune response acts as one of the first lines of host defense against IAV infection (3–6). Influenza viruses are known to utilize multiple tactics to evade host innate immune responses. The most well-characterized strategies aim to inhibit synthesis of type I IFNs (7–9). However, during IAV infection, IFN-α/β molecules are still produced to some extent. These molecules bind to the cognate IFN receptor (IFNAR), activating the JAK/STAT pathway, which generates antiviral responses. The continued presence of IFN-α/β suggests that IAV could overcome IFN receptor-mediated antiviral innate responses by utilizing a method that does not target type I IFN molecule synthesis.

Our previous findings have shown that upon infection, IAV promotes the degradation of subunit 1 of the type I and type II IFN receptors, IFNAR1 and IFNGR1, respectively, using viral hemagglutinin (HA) (10, 11). Viral HA induces the phosphorylation of IFNAR1 and IFNGR1, which is followed by polyubiquitination of these receptors. Ubiquitinated IFNAR1 and IFNGR1 are then degraded, resulting in decreased IFN receptor levels on the cell surface and decreased IFN sensitivity for that cell (10, 11). Receptor degradation results in a broad-spectrum downregulation of receptor-mediated interferon responses, creating more optimal conditions for IAV survival and replication. However, the molecular mechanism for HA’s triggering of IFN receptor degradation remains elusive.

Casein kinase 1α (CK1α) was previously shown to be critical for inducing the phosphorylation and ubiquitination of IFNAR1/IFNGR1, regulating receptor levels upon IAV infection (10). CK1α regulates IFNAR1 levels and in turn, controls IAV propagation in epithelial cells (10). However, HA-mediated IFNAR1 degradation does not follow known cellular mechanisms of IFNAR1 degradation, such as PERK-mediated endoplasmic reticulum (ER) stress or high levels of IFN molecules (10–16). This led us to hypothesize that there could be an unknown cellular factor(s) that mediates this receptor degradation process. Utilizing mass spectrometry (MS), we identified poly(ADP-ribose) polymerase 1 (PARP1) as a host protein that binds to IAV HA.

PARP1 is a member of the PARP family, which regulates various important cellular processes, such as cellular differentiation and cell proliferation (17–21). Of note, PARP1 is well-known as a regulator during tumor development (17), and a PARP1 inhibitor is approved as a therapeutic option for patients with cancers derived from BRCA germ line mutations, such as breast or ovarian cancer. PARP1 and PARP13.1 have been shown to regulate influenza A virus polymerase activity, which in turn affects IAV replication (22–24). Additionally, PARP1 depletion in human cancer cells induces the expression of interferon-stimulated genes (ISGs), suggesting the possible involvement of PARP1 in IFN signaling (25). However, the exact role of PARP1 in regulating the host type I IFN signaling pathway and its resultant effects on viral replication remain unknown.

In this study, we identified PARP1 as an IAV HA interacting partner during IAV infection. Furthermore, PARP1 was proven to be important for mediating IAV HA-induced degradation of IFNAR1, regulating host type I IFN responses and, consequently, promoting IAV propagation. These findings help us to gain novel insight into host-influenza interactions and highlight a promising therapeutic target to control influenza virus infection.

RESULTS

PARP1 is an IAV HA interacting protein.

We have previously shown that the IAV HA subunit HA1 induces IFNAR1 degradation (11). To investigate the molecular mechanisms behind this HA-induced degradation, we sought to identify the cellular proteins involved by employing a mass spectrometric screening approach. The lysates of cells expressing either a FLAG control vector or a FLAG-tagged HA1 domain of influenza A/CA/04/09 (H1N1) virus HA were subjected to coimmunoprecipitation (co-IP) experiments in order to identify cellular proteins that specifically bind to FLAG-HA1 but not to the FLAG control (Fig. 1A). MS analysis of the co-IP precipitates revealed the presence of viral HA and several host proteins, including poly(ADP-ribose) polymerase 1 (PARP1) protein, which specifically appeared in the FLAG-HA1-expressing cells but not in the FLAG control cells (Tables 1 and 2).

FIG 1.

PARP1 interacts with HA upon HA expression or IAV infection. (A) A schematic representation of the MS approach for identification of influenza A/New Caledonia/20/99 (H1N1) viral HA1 subunit-binding proteins in HEK293T cells. (B) HEK293T cells were transfected with plasmids encoding FLAG-tagged HA1 of A/New Caledonia/20/99 (H1N1) or an empty vector (FLAG-V) as a control. At 24 h posttransfection, the cells were harvested and subjected to a co-IP experiment. The levels of immunoprecipitated PARP1 and FLAG-tagged HA1 were detected by Western blotting using the indicated antibodies. The levels of PARP1, FLAG-HA1, and GAPDH in the cell lysates are also shown. (C) HEK293T cells were transfected with plasmids encoding FLAG-HA of A/New Caledonia/20/99 (H1N1) or FLAG-V. Cells were harvested and subjected to a co-IP experiment 24 h posttransfection. The levels of immunoprecipitated PARP1 were detected by Western blotting. (D) HEK293T cells were transfected with FLAG-tagged PARP1 and/or HA of influenza A/New Caledonia/20/99 (H1N1) virus as indicated. At 24 h posttransfection, a co-IP experiment was performed and the levels of HA in the immunoprecipitates were detected. (E) HEK293T cells were transfected with FLAG-PARP1. At 24 h posttransfection, the cells were mock infected or infected with IAV at an MOI of 1. Co-IP was performed 24 h postinfection, and the levels of immunoprecipitated HA of influenza virus were detected by Western blot analysis. (F and G) HEK293T cells (F) or A549 cells (G) were infected with IAV at an MOI of 1. At 24 h postinfection, co-IP experiments were performed. The levels of immunoprecipitated HA of IAV were detected by Western blotting. (H) HEK293 cells were mock infected or infected with IAV at an MOI of 1. At 18 or 36 h postinfection, the cells were subjected to co-IP experiments and the levels of immunoprecipitated HA were detected by Western blot analysis. MW, molecular weight (in thousands); IB, immunoblotting; IP, immunoprecipitation.

TABLE 1.

A short list of HA-interacting proteins identified by MS analysis

Protein identified	Mol wt (kDa)	No. of MS/MS spectra
		FLAG-vector	FLAG-HA1
Hemagglutinin	36	0	13
Tubulin alpha-1B chain	50	0	11
Poly(ADP-ribose) polymerase 1	113	0	9
Heat shock protein 90-beta	83	0	8
ADP/ATP translocase 2	33	0	7
Stress-induced phosphoprotein 1	68	0	4
BCL2-associated athanogene 2	24	0	4

TABLE 2.

PARP1 peptides identified by MS analysis

Peptide sequence identified	Best SEQUEST resulta
	XCorr score	ΔCn score
FYTLIPHDFGMK	3.1	0.138
KPPLLNNADSVQAK	3.99	0.181
MAIMVQSPMFDGK	2.6	0.282
SDAYYCTGDVTAWTK	3.92	0.0934
TTNFAGILSQGLR	3.36	0.317
VFSATLGLVDIVK	2.47	0.272
VVSEDFLQDVSASTK	4.32	0.496

^aThese data have been filtered for >95% confidence on peptide identification and >99% confidence on protein identification. The percentage of amino acid sequence coverage was 9.37. XCorr, cross-correlation value; ΔCn, delta correlation value.

Since PARP1 is known to regulate multiple cellular signaling events, we further investigated the interplay between PARP1 and viral HA/HA1. We confirmed this interaction by performing reciprocal co-IP experiments. Transiently expressed FLAG-tagged HA1 (Fig. 1B) or FLAG-tagged full-length HA (Fig. 1C) pulled down endogenous PARP1 (Fig. 1B and C). In the reverse experiment, immunoprecipitation of FLAG-tagged PARP1 also pulled down transiently expressed IAV HA (Fig. 1D). Importantly, this PARP1-HA interaction also occurred during infection. As shown by the results in Fig. 1E, upon IAV infection, viral HA was pulled down by overexpressed FLAG-tagged PARP1. Furthermore, endogenous PARP1 was shown to associate with HA during IAV infection in both HEK293 cells (Fig. 1F) and A549 cells (Fig. 1G), and the interaction appeared to increase over time during infection (Fig. 1H). These data indicate that PARP1 specifically interacts with IAV HA during influenza viral infection.

IAV HA changes PARP1 localization.

PARP1 is a well-known DNA damage response protein which localizes predominantly within the nucleus of human cells (26). However, IAV HA is located within the cytoplasm and plasma membrane of host cells. These contrasting localization patterns of PARP1 and HA led us to ask if viral HA expression affects the localization of endogenous PARP1. To determine if PARP1 localization changes upon IAV HA expression, IAV HA was transiently expressed in A549 cells and immunofluorescence and confocal microscopy were performed to detect PARP1 and IAV HA. Endogenous PARP1 localization was nuclear in control cells but was strikingly nuclear-cytoplasmic in cells expressing IAV HA (Fig. 2A). Quantification of the PARP1 signal in HA-expressing A549 cells indicates that over 20% of quantified PARP1 was cytoplasmic compared to the amount in the control (Fig. 2B). Colocalization between overexpressed HA and endogenous PARP1 was determined by the Manders colocalization coefficient (MC). The MC between overexpressed HA and endogenous PARP1 was 0.3 (Fig. 2C), indicating that 30% of HA also colocalized with PARP1. This positive correlation was significantly higher than the result for the untransfected control. To confirm that this localization can also occur during IAV infection, we next performed immunofluorescence analysis of infected cells. Immunofluorescence analysis of IAV HA and PARP1 during infection revealed a similar nuclear-cytoplasmic localization of endogenous PARP1 in A549 cells (Fig. 2D). Quantification of the PARP1 cytoplasmic signal, as described above, in IAV-infected A549 cells revealed a higher percentage of cytoplasmic PARP1 than in the uninfected control, similar to that in the cells overexpressing HA (Fig. 2E). MC measurements of infected cells also determined positive levels of colocalization between IAV HA and endogenous PARP1, which were similar to the levels in the HA-transfected samples (Fig. 2F). These data indicate that IAV HA induces PARP1 localization to the cytoplasm in addition to the nucleus.

FIG 2.

PARP1 localization changes upon IAV HA expression or IAV infection. (A) A549 cells were transfected with plasmids encoding myc-tagged IAV HA or a control vector. At 24 h posttransfection, cells were fixed and stained using Draq5 for nuclei, as well as anti-PARP1 and antimyc antibodies. Samples were visualized using confocal microscopy. (B and C) PARP1 fluorescence from confocal images was quantified using ImageJ, and PARP1 cytoplasmic fluorescence (B) and the Manders colocalization coefficient (MC) between HA and PARP1 (C) were determined. (D) A549 cells were infected with 1 MOI of IAV PR8 and were fixed at 24 hpi. Cells were stained using Draq5 for nuclei, as well as anti-PARP1 and anti-IAV HA antibodies. Samples were visualized using confocal microscopy. (E and F) PARP1 fluorescence from confocal images was quantified using ImageJ, and PARP1 cytoplasmic fluorescence (E) and MC between IAV HA and PARP1 (F) were determined. Scale bars represent 10 μm. Data are representative of at least two independent experiments. Eight to 10 images per condition per experiment were used. Statistical analysis was conducted using a two-tailed unpaired t test. The data represent mean values ± SD (*, P < 0.05; **, P < 0.01; ***, P < 0.001).

PARP1 mediates IAV HA-induced degradation of IFNAR1.

We next studied whether PARP1 mediates the IAV- or HA-induced degradation of IFNAR1. To test this, we utilized a knockdown approach using a small interfering RNA (siRNA) targeting PARP1 (si-PARP1). When endogenous PARP1 was subjected to siRNA knockdown, the HA-induced downregulation of IFNAR1 was strongly inhibited both in HEK293 cells (Fig. 3A) and in A549 cells (Fig. 3B). Knockdown of PARP1 also suppressed IFNAR1 degradation triggered by H5N1 IAV HA (Fig. 3C). Consistent with these results, knockdown of PARP1 impaired IAV infection-induced downregulation of IFNAR1 at both 8 and 24 h postinfection (hpi) in A549 cells (Fig. 3D). Virus infection triggers the production of type I IFNs, which in turn, decreases IFNAR1 levels through the ligand-dependent pathway when present in large amounts (27, 28). To eliminate the possibility that PARP1 affects IFN production and indirectly affects IFNAR1 degradation, we utilized Vero cells, which have type I IFN receptors but are unable to synthesize type I IFNs. IAV infection strongly induced IFNAR1 downregulation in Vero cells (Fig. 3E), and knockdown of PARP1 partially rescued IFNAR1 levels at both 8 and 24 hpi in Vero cells (Fig. 3E). This indicates that PARP1 regulates IFNAR1 levels independently of type I IFN production or signaling during IAV infection. To determine if PARP1 affects protein levels of IFNAR1, we next compared IFNAR1 mRNA levels between si-PARP1-transfected cells and control cells during IAV infection. Knockdown of PARP1 did not alter the mRNA expression of IFNAR1 at 8 hpi or 24 hpi (Fig. 3F), suggesting that PARP1 modulates IFNAR1 at the protein level.

FIG 3.

Knockdown of PARP1 results in impaired degradation of IFNAR1 during HA expression or IAV infection. (A and B) HEK293T cells (A) or A549 cells (B) were transfected with siRNA specific to PARP1 (si-PARP1) or nonspecific scrambled control siRNA (SCR). At 24 h posttransfection, the cells were transfected with a control vector (−) or plasmids encoding HA (+). Western blotting was performed 24 h after HA transfection, and the levels of IFNAR1, HA, and PARP1 were detected. The levels of GAPDH were used as an internal loading control. (C) HEK293 cells were transfected with SCR or si-PARP1. At 24 h posttransfection, the cells were transfected with a control vector (−) or plasmids encoding HA from influenza A/Thailand/1(KAN-1)/2004 (H5N1) (H5). Western blotting was performed to detect the levels of IFNAR1, HA, PARP1, and GAPDH. (D and E) A549 cells (D) and Vero cells (E) were transfected with SCR or si-PARP1. At 24 h posttransfection, cells were left uninfected (Mock) or infected with IAV at an MOI of 1 for the indicated time periods. The levels of IFNAR1, PARP1, NS1, HA, and GAPDH were analyzed by Western blotting. The relative intensity of each band of IFNAR1 was determined by densitometry based on the GAPDH levels and is depicted below each blot. The relative level of IFNAR1 from the mock-infected sample was set as 1.0. The relative intensities of each band of NS1 and HA in panel D were also determined. (F) HEK293T cells were transfected with SCR or si-PARP1. At 24 h, cells were left uninfected (Mock) or infected with IAV at an MOI of 1. The relative mRNA levels of IFNAR1 were analyzed by real-time qPCR at 8 and 24 hpi. Error bars represent mean values ± SD calculated from the results for three individual samples. NS, not significant.

In order to further investigate the regulatory effect of PARP1 on IFNAR1 degradation, cells were pretreated with a PARP1 inhibitor (ABT-888) (17, 29), followed by transient expression of HA. PARP1 inhibition resulted in almost no downregulation of IFNAR1 in the presence of HA, while the levels of HA were not affected by the inhibitor (Fig. 4A). To examine the specificity of inhibition, we utilized XAV-939, a compound known to inhibit PARP family member tankyrase 1/2 (PARP5) but not PARP1 (30). As shown by the results in Fig. 4B, upon HA expression, tankyrase 1/2 inhibition did not alter IFNAR1 levels, whereas PARP1 inhibition strongly suppressed IFNAR1 downregulation. Of note, both inhibitors did not affect p38 mitogen-activated protein (MAP) kinase activation, which is known to regulate IFNAR1 degradation during the host ER stress response (13). To support our conclusion that PARP1 activity is critical for IAV HA-induced IFNAR1 degradation, another PARP1 inhibitor (4-AN [4-amino-1,8-naphthalimide]) was utilized. Consistent with the result from experiments using ABT-888, 4-AN inhibited HA-induced IFNAR1 downregulation (Fig. 4C). Collectively, these data suggest that host factor PARP1 is crucial in mediating IAV HA-induced degradation of IFNAR1.

FIG 4.

Pharmacological inhibition of PARP1 leads to impaired downregulation of IFNAR1 upon viral HA expression. (A) HEK293T cells were transfected with a control vector (−) or plasmids encoding HA. The cells were then treated with solvent (dimethyl sulfoxide [DMSO]) or a PARP1 inhibitor (ABT-888) at a concentration of 20 μM. At 24 h posttransfection, the levels of IFNAR1, HA, and GAPDH were detected by Western blotting. (B) HEK293T cells were transfected with a control vector (−) or plasmids encoding HA. Cells were then treated with the indicated reagents for a time period of 24 h. The levels of IFNAR1, HA, p-p38, and GAPDH were detected by Western blotting. (C) HEK293T cells were transfected with a control vector (−) or plasmids encoding HA. Cells were then treated with DMSO, ABT-888 (20 μM), or 4-AN (4-amino-1,8-naphthalimide) (20 μM) for 24 h. Western blotting was performed to detect the levels of IFNAR1, HA, PARP1, and GAPDH.

PARP1 positively regulates IAV replication by impairing type I IFN receptor signaling.

Given that IFNAR1 expression is essential for the type I IFN-mediated antiviral response, we hypothesized that PARP1 promotes IAV replication in cells by controlling the levels of IFNAR1. To study the regulatory effects of PARP1 on IAV replication, A549 cells were pretreated with PARP1 inhibitor (ABT-888), followed by IAV infection. The effect of the inhibitor on IAV replication was evaluated by comparing the expression levels of viral proteins. IAV HA protein expression decreased at 12, 24, and 36 hpi with the PARP1 inhibitor treatment (Fig. 5A), and virus-induced downregulation of IFNAR1 was suppressed following PARP1 inhibitor treatment (Fig. 5A). In support of this, knockdown of endogenous PARP1 notably inhibited IAV protein expression following infection in comparison to the IAV protein expression in the control siRNA-transfected group (Fig. 5B). To further evaluate the effects of PARP1 on IAV replication, PARP1 was transiently overexpressed in cells, followed by IAV infection. PARP1 overexpression enhanced virus replication at the two time points measured (Fig. 5C), confirming the proviral activity of PARP1. Of note, an inverse correlation between PARP1 and IFNAR1 levels was observed when PARP1 was downregulated (Fig. 5B) or overexpressed (Fig. 5C), supporting the important role of PARP1 in IFNAR1 downregulation during infection. Furthermore, overexpressed PARP1 increased the production of infectious IAV (Fig. 5D), while downregulation of PARP1 using siRNA decreased IAV production (Fig. 5E). These results were consistent with the regulation of viral protein levels by PARP1.

FIG 5.

PARP1 positively regulates IAV replication. (A) A549 cells were infected with pandemic influenza A/CA/04/09 virus at an MOI of 1. Cells were then treated with DMSO or ABT-888 (20 μM). The levels of viral HA, IFNAR1, and GAPDH were detected at 12, 24, and 36 h postinfection. (B) A549 cells were transfected with SCR or si-PARP1. At 24 h posttransfection, cells were mock infected (−) or infected with IAV (+) at an MOI of 1. Cells were harvested at 24 hpi, and the levels of IFNAR1, PARP1, and viral HA and M1 proteins were detected by Western blotting. (C) A549 cells were transfected with FLAG-tagged PARP1 or a control FLAG vector as indicated. At 24 h posttransfection, cells were infected with IAV at an MOI of 1 for an additional 12 or 24 h. The levels of FLAG-PARP1, IFNAR1, viral NP, viral HA, and GAPDH were analyzed by Western blotting. (D and E) A549 cells were transfected with a control vector or FLAG-PARP1 (D) or SCR or si-PARP1 (E). At 24 h posttransfection, cells were infected with IAV at an MOI of 0.01. At 2 days postinfection, the titers of infectious virus in the supernatant of the culture were measured by plaque assay. The data represent the mean values ± SD from three independent samples (*, P < 0.05).

Considering that PARP1 has multiple functions in cells, we sought to determine whether the proinfluenza activity of PARP1 is linked to the receptor-mediated type I IFN signaling. To this end, cells were treated with PARP1 inhibitor and then infected with IAV. Cells treated with PARP1 inhibitor had more robust type I IFN responses upon infection, indicated by both the RNA levels of IFIT1 and ISG15 (Fig. 6A) and the protein expression levels of RIG-I and IFIT1 (Fig. 6B). To further clarify the role of type I IFN signaling in PARP1-mediated IAV replication, A549 cells were transfected with either scrambled control siRNA (SCR) or IFNAR1-specific siRNA (Fig. 6C). The siRNA-transfected cells were then infected with IAV in the presence or absence of PARP1 inhibitor. PARP1 inhibitor treatment reduced the production of infectious IAV from cells transfected with SCR at 1, 2, or 3 days postinfection (dpi) (Fig. 6C), which is consistent with the suppression of viral protein levels (Fig. 5). However, when endogenous IFNAR1 was knocked down, treatment with PARP1 inhibitor failed to significantly suppress virus propagation in comparison to that in solvent-treated samples (Fig. 6C). Downregulation of endogenous IFNAR1 was confirmed by Western blot analysis (Fig. 6D). Thus, PARP1 inhibition substantially restricts IAV replication, which requires intact IFNAR expression in A549 cells. Altogether, these data suggest the regulatory effect of PARP1 on IAV propagation is closely associated with the type I IFN signaling pathway.

FIG 6.

PARP1 inhibition suppresses IAV propagation by elevating type I IFN receptor signaling. (A and B) A549 cells were infected with IAV at 0.1 MOI. Cells were then treated with DMSO or ABT-888. At 24 hpi, relative RNA levels of IFIT1 and ISG15 were calculated by RT-qPCR (A) and the protein expression levels of RIG-I, IFIT1, and GAPDH were detected by Western blot analysis (B). (C) A549 cells were transfected with nonspecific scrambled control siRNA (SCR) (solid lines) or siRNA specific to IFNAR1 (dashed lines). At 24 h posttransfection, cells were treated with DMSO or ABT-888 (20 μM) and infected with IAV at an MOI of 0.001. The titers of infectious virus in the supernatants of the culture were assessed by plaque assays on MDCK cells at 1, 2, or 3 days postinfection (dpi). Each data point on the curve represents the mean value from three independently obtained samples. The data represent mean values ± SD. P values were obtained by comparing the indicated group values in a two-tailed, unpaired Student’s t test. p' indicates the P values of siIFNAR1 with either solvent or ABT-888 treatment. Furthermore, treatment groups were analyzed utilizing a two-way ANOVA with Tukey pairwise comparison. ANOVA results comparing treatment types, irrespective of time point, are indicated by lowercase letters to the right of the graph. Groups that do not share letters had significantly different results. Both analyses indicate that virus titers decrease following ABT-888 treatment, but this effect is not seen following transfection with si-IFNAR1. (D) A549 cells were left untransfected (control) or transfected with SCR or si-IFNAR1, and at 48 h posttransfection, the downregulation of the endogenous IFNAR1 was confirmed by Western blotting.

To further investigate the role of HA-PARP1 interaction in regulating the IFNAR signaling pathway, recombinant IFN-α (rIFN-α) was added to the cells exogenously in the presence of HA and/or PARP1 inhibitor (ABT-888). HA expression inhibited IFN-induced activation of STAT1 (phosphorylated STAT1 [pSTAT1]), presumably because of IFNAR1 degradation (Fig. 7A). However, PARP1 inhibition impaired HA-induced IFNAR1 downregulation and restored IFN-induced STAT1 activation (Fig. 7A). Similarly, treatment with the PARP1 inhibitor blocked HA-induced inhibition of the expression of IFIT1, which is an ISG, in response to rIFN-α (Fig. 7B). The inhibitor ABT-888 alone, however, did not affect STAT1 activation (Fig. 7C) or IFIT1 expression (Fig. 7D) when cells were stimulated by rIFN-α. Therefore, the data indicate that PARP1 regulates HA-induced IFNAR degradation to impact the IFN signaling pathway.

FIG 7.

Inhibition of PARP1 abolishes HA-induced suppression of type I IFN responses. (A and B) HEK293T cells were transfected with a control vector (control) or plasmids encoding HA. Cells were then cultured with or without ABT-888 (20 μM) as indicated. At 24 h posttransfection, cells were treated with recombinant human IFN-α2 (rIFN-α2) (1,000 U/ml) for an additional 1 h (A) or 20 h (B). The levels of pSTAT1, STAT1, IFNAR1, and GAPDH were detected by Western blotting (A), and the mRNA levels of IFIT1 were detected by RT-qPCR (B). (C and D) HEK293T cells were cultured with DMSO or ABT-888 (20 μM) for 24 h. Then, cells were treated with rIFN-α2 (1,000 U/ml) for 1 h (C) or 20 h (D). The levels of pSTAT1, STAT1, IFNAR1, and GAPDH were detected by Western blotting (C), and the mRNA levels of IFIT1 were detected by RT-qPCR (D). The data represent mean values ± SD (***, P < 0.001; NS, not significant).

DISCUSSION

Successful influenza virus replication relies heavily on the subversion and evasion of host type I IFN signaling (4–6). We have previously shown that IAV infection induces the degradation of type I IFN receptor IFNAR1 using viral HA protein. This results in the decreased sensitivity of infected cells to type I IFNs, which in turn, facilitates viral propagation. However, the mechanisms behind HA-induced IFNAR1 degradation remain obscure. Here, we demonstrate that PARP1 interacts with viral HA and plays a key role in IAV HA-induced degradation of IFNAR1. Importantly, PARP1 was shown to regulate IAV replication by manipulating receptor-mediated type I IFN antiviral responses.

The degradation of IFNAR1 is regulated by several cellular mechanisms, such as the ligand (IFN)-dependent pathway or the PERK-dependent ER stress response (also known as unfolded protein response [UPR]) (27, 28, 31). We have previously determined that influenza viral HA eliminates IFNAR1 using CK1α, but this degradation process is independent of both the ligand-dependent pathway and the ER stress response (11). In the UPR-induced IFNAR1 phosphorylation pathway, the primary phosphorylation of IFNAR1 by p38 is crucial to ensure subsequent IFNAR1 phosphorylation and ubiquitination (13). Given that p38 activation is dispensable for IFNAR1 downregulation upon HA expression (11), there could be an undiscovered mechanism utilized by viral HA to activate the signaling pathway for IFNAR1 degradation.

Performing mass spectrometry and utilizing viral HA1 as bait led to the identification of PARP1, which interacts with IAV HA (Fig. 1A and Tables 1 and 2). PARP1 is commonly known as a chromatin-associated protein and mainly exists in the nucleus under normal conditions (19). However, it has long been questioned whether it could also localize in other cellular compartments (32). Certain conditions, such as oxidative stress, have been reported to induce the nucleus-to-cytoplasm release of PARP1, which relies on the interaction between PARP1 and RNF146 (18). PARP1 has also been shown to interact with mitofilin, a mitochondrial protein, and it localizes in mitochondria, where it plays a role in the maintenance of mitochondrial DNA integrity (21). Nuclear-cytoplasmic expression of PARP1 has also been shown to occur in breast cancer (33) and pancreatic cancer (34) in human patients. Additionally, PARP1 localization is cytoplasmic during HIV-1 Vpr expression in cells (35), which could indicate that proteins of other viruses also have the potential to change PARP1 localization. These findings suggest that PARP1 could have several functions in the host cell cytoplasm during different viral infections, which requires further examination.

PARP1 and PARP13.1 have been previously shown to regulate the function of influenza viral polymerases (22–24). However, under our experimental settings, PARP1 bound to HA during IAV infection (Fig. 1E to H) and facilitated IFNAR1 degradation, regulating host antiviral IFN responses. Notably, PARP1 localization became nuclear-cytoplasmic during both IAV HA expression and IAV infection (Fig. 2). Our results and the findings of others indicate the possibility that PARP1 is utilized in multiple ways to enhance influenza virus infection. Our results show an approximately 2-fold difference in IAV production during the downregulation of IFNAR1 and treatment with PARP1 inhibitor (Fig. 6C). While these results are not statistically significant under our experimental conditions, they could correspond to the known effects of IAV polymerase interacting with PARP1 (23). It is likely that the presence of PARP1 in the nucleus and cytoplasm could also have different but necessary effects for robust influenza virus replication.

IAV HA was shown to partially colocalize with PARP1 in the cytoplasm (Fig. 2), suggesting that the interaction occurs transiently to elicit further signaling events in order to induce IFNAR1 degradation. It is currently unclear how IAV HA manipulates PARP1 to trigger IFNAR1 degradation. PARP1 can regulate various cellular processes (17–20) and modify other proteins through poly(ADP-ribosyl)ation, a posttranslational modification (19). IAV HA-PARP1 interaction could trigger the activation of PARP1 ADP-ribose polymerase, leading to the modification of an unknown protein involved in the IFNAR1 degradation process. The results of using inhibitors that block PARP1 activation indicate that PARP1 activity is crucial for HA-induced IFNAR1 degradation (Fig. 4). Poly(ADP-ribosyl)ation leads substrates to proteasomal degradation (36–38) under certain conditions. Since IAV HA-induced IFNAR1 degradation is dependent on both proteasomal and lysosomal pathways (11), it will be interesting to study how PARP1 is involved in the IFNAR1 degradation pathways. It is possible that poly(ADP-ribosyl)ation of IFNAR1 or its unknown regulatory protein is required prior to the degradation process during IAV infection. A recent study has shown that another PARP family member, PARP11, inhibits the type I interferon-mediated response by catalyzing mono(ADP-ribosyl)ation of E3 ligase β-TrCP and, in turn, promotes the ubiquitination and degradation of IFNAR1 (39). It would therefore be of interest to investigate whether PARP1 and PARP11 cooperate, leading to IFNAR1 degradation, or if they have distinct functions in the regulation of IFNAR1, enhancing viral infection. Another possibility is that PARP1 induces the activation of a kinase, such as CK1α, by poly(ADP-ribosyl)ation. The kinase could then phosphorylate IFNAR1, leading to degradation of the receptor. PARP1 may have an undiscovered function in regulation of the phosphorylation, ubiquitination, or degradation of IFNAR1. This area of research requires further investigation.

Cell surface levels of IFNAR1 are correlated with type I IFN-mediated antiviral responses (40). Lack of IFNAR1 on the cell surface desensitizes cells to type I IFNs, creating a more favorable environment for viral replication. Here, we identified host PARP1 as a pro-IAV factor that regulates viral propagation via controlling IFNAR1 degradation. Pharmacologic inhibition or PARP1 knockdown strongly suppressed IAV replication (Fig. 5 and 6). Thus, PARP1 may represent a new cellular target for controlling influenza virus propagation. Since known PARP1 inhibitors are currently used to treat cancer patients in the clinic, it would be worthwhile to test the effects of these inhibitors on influenza virus infections.

In summary, our work reveals the importance of host protein PARP1 in mediating IAV HA-induced degradation of IFNAR1, therefore regulating IAV propagation. This study expands our knowledge of the interactions between IAV and the host and unveils novel cellular targets that have the potential to be novel therapeutics to control influenza viral replication.

MATERIALS AND METHODS

Viruses and cells.

Influenza A/WSN/33 (H1N1) virus, pandemic influenza A/CA/04/09 (H1N1) virus, and influenza A/Puerto Rico/8/34 (H1N1) virus were used as previously reported (10, 11, 41, 42). Viruses were amplified on Madin-Darby canine kidney (MDCK) cells as previously described (10, 11, 41). Briefly, cells were incubated with virus at different multiplicities of infection (MOI), as indicated in the figures, for 1 h. The cells were then washed with phosphate-buffered saline (PBS) and incubated with fetal bovine serum (FBS)-free medium containing 0.3% bovine serum albumin (BSA) and TPCK (tosylsulfonyl phenylalanyl chloromethyl ketone)-trypsin (1 μg/ml) for certain time periods optimal for amplification. Titration of virus was performed using plaque assay. Briefly, supernatants containing viruses were harvested and diluted into serial dilutions. The diluted supernatants were then adsorbed onto 4 × 10⁵ MDCK cells/well in a 6-well plate for at least 1 h. Cells were then incubated with 2× Eagle’s minimum essential medium (EMEM) (Gibco) mixed with an equal portion of 1% agarose (Seakem). Human embryonic kidney 293 (HEK293) cells, human lung epithelial A549 cells, and African green monkey kidney epithelial cells (Vero cells) have been previously reported (10, 11, 43–45). The HEK293 cells, A549 cells, and Vero cells were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Gibco), while the MDCK cells were cultured in minimum essential medium Eagle (MEM; Mediatech), as previously described (10, 11, 43, 44). All the cells were cultured in a CO₂ incubator at 37°C, and all media were supplemented with 10% FBS (HyClone) and penicillin (100 U/ml)-streptomycin (100 μg/ml) (Invitrogen).

Constructs and transfection.

The plasmid encoding FLAG-tagged human PARP1 was purchased from GenScript; the plasmid encoding FLAG-tagged human IFNAR1 was provided by Serge Fuchs (University of Pennsylvania) (46); and the plasmids encoding influenza viral hemagglutinin (HA) from the A/New Caledonia/20/99 (H1N1) and A/Thailand/1(KAN-1)/2004 (H5N1) viruses have been previously described and were provided by the NIH Vaccine Research Center (10, 11, 47). To construct the FLAG-tagged HA and FLAG-tagged HA1, the coding sequences of full-length HA and HA1 subunit were amplified by PCR from the DNA of full-length HA of influenza A/New Caledonia/20/99 (H1N1) using the primers 5′-CGG AAT TCG ATG AAG GCC AAA CTG CTG -3′ and 5′-GGG GTA CCC GTC AGA TAC AGA TCC TGC ACT GCA-3′ for HA and 5′-CGG AAT TCG ATG AAG GCC AAA CTG CTG-3′ and 5′-GGG GTA CCC GTC ATC TGC TCT GGA TGC TAG GGA-3′ for HA1. For transfection of cultured cells, cells were seeded onto 6-well plates or 24-well plates at densities of 10⁶ cells/well or 2.5 × 10⁵ cells/well 24 h prior to transfection. Cells were then transfected with the plasmids indicated in the figures using Lipofectamine 2000 transfection reagent (Thermo Scientific) at 80% to 90% confluence following the protocols recommended by the manufacturer. A concentration of 500 ng/ml DNA was used for the transfection experiments unless specifically indicated in the figures. Empty vector plasmids were used as a control in all transfection experiments to ensure that each transfection sample received the same amount of total DNA.

Reagents and antibodies.

Anti-DYKDDDDK (FLAG) G1 antibody affinity resin (GenScript), IP lysis buffer (Thermo Scientific), protease inhibitor PMSF (phenylmethylsulfonyl fluoride; Gold Bio), cycloheximide (CHX; Sigma-Aldrich), recombinant human IFN-α2 (BioLegend), protein A/G agarose (Thermo Fisher), PARP1 inhibitor ABT-888 (Cayman) (17, 29), PARP1 inhibitor 4-AN (4-amino-1,8-naphthalimide; Sigma-Aldrich), and TNKS inhibitor XAV939 (Cayman) were purchased from the indicated manufacturers. Antibody against influenza viral HA (H1N1) was purchased from GeneTex and Santa Cruz; antibodies against human IFNAR1, phospho-IFNAR1 (S535/S539), and influenza virus M1 protein were purchased from Abcam, and antibodies against human PARP1, human GAPDH (glyceraldehyde-3-phosphate dehydrogenase), RIG-I, IFIT1, phospho-p38, FLAG tag, and Myc tag were purchased from Cell Signaling Technology. Fluorophore-labeled secondary antibodies against mouse and rabbit IgG were purchased from Invitrogen.

Confocal microscopy and image analysis.

A549 cells were seeded in 24-well plates on poly-l-lysine-coated coverslips (Neuvitro) and were either transfected with IAV HA-encoding plasmid or infected with influenza A/Puerto Rico/8/34 (H1N1) virus. Cells were fixed with 4% paraformaldehyde (Alfa Aesar) and then permeabilized, blocked with 10% FBS and 1% BSA for 1 h, and incubated with primary antibodies, i.e., anti-IAV HA (Santa Cruz) or anti-Myc tag (Cell Signaling Technology) antibody and anti-PARP1 antibody (Cell Signaling Technology) overnight at appropriate concentrations in 1% BSA. After washing, samples were incubated with the appropriate secondary antibodies, such as Alexa Fluor 488 goat anti-mouse antibody (Invitrogen) or Alexa Fluor 546 goat anti-rabbit antibody (Invitrogen), for 1 h and then washed. Cell nuclei were stained using Draq5 according to the manufacturer’s instructions (Thermo Fisher) and then mounted on glass slides with Prolong gold antifade mountant (Thermo Fisher). Confocal images of thousands of cells were acquired at the University of Missouri Molecular Cytology Core facility using a Leica SP8 TCP confocal microscope. For quantification purposes, imaging was performed using a 40× objective, capturing 8 to 10 random images per condition per experiment. Quantification was performed using Fiji (48) of ImageJ (49). Levels of cytoplasmic PARP1 were determined by using the ImageJ Intensity Ratio Nuclei Cytoplasm Tool (50, 51), where Draq5 was used as the nuclear stain. The threshold, select area, and ROI (region of interest) manager functions of ImageJ were used to restrict measurements of the Intensity Ratio Nuclei Cytoplasm Tool to cells that only expressed HA. The Manders colocalization coefficient (MC) between HA and PARP1 was determined using the ImageJ plugin JACoP (52).

Co-IP and Western blotting.

293T cells or A549 cells seeded in 6-well plates were transiently transfected or cotransfected with plasmids indicated in the figures (1 μg DNA in total). For detection of interaction between HA and transfected PARP1 during IAV infection, cells were infected with IAV at an MOI of 1 at 24 h posttransfection. For detection of interaction between HA and endogenous PARP1 during infection, cells were infected with IAV at an MOI of 1. Cells were then harvested and lysed in 1 ml immunoprecipitation (IP) lysis buffer at 48 h posttransfection or at the time points indicated in the figures. For co-IP experiments, cell lysates containing protease inhibitor (PMSF) (1 mM) were incubated with 20 μl of anti-DYKDDDDK (FLAG) G1 antibody affinity resin or 20 μl protein A/G agarose (premixed with anti-PARP1 antibody) overnight with rotation at 4°C. The beads were washed three times with 1 ml IP lysis buffer, and precipitates were analyzed by standard Western blot analysis. Western blotting was performed as previously described (10, 11, 43, 44). Briefly, the denatured polypeptides from cell lysates or co-IP were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and were transferred to nitrocellulose membranes (Bio-Rad). Membrane-bound antibodies were detected using IRDye secondary antibodies (Li-Cor). The signals were imaged using an Odyssey Fc imaging system (Li-Cor), and the resulting data were analyzed using Image Studio software version 5.2 (Li-Cor). Similar results were obtained from at least two independent experiments.

RNA interference.

SMARTpool On-Target plus siRNA targeting human PARP1 (L-006656-03-0005) was purchased from Dharmacon. siRNA targeting human IFNAR1 was purchased from RiboBio. The 27-mer universal scrambled negative-control siRNA duplex (SCR) was purchased from Origene. All siRNAs were used at a final concentration of 20 nM to transfect HEK293 cells or A549 cells using Lipofectamine RNAiMax transfection reagent (Thermo Scientific) according to the manufacturer’s instructions. Cells were harvested at 48 h posttransfection, and the knockdown of PARP1 was confirmed by Western blot analysis.

Real-time quantitative PCR.

Total cellular RNA was purified by using Tri Reagent (Sigma-Aldrich) according to the manufacturer’s instructions. Purified RNA was reverse transcribed with random primers (Invitrogen), and the resulting cDNA was analyzed by real-time quantitative PCR (RT-qPCR) using gene-specific primers. Primers for human IFNAR1 (5′-CAC TTC TTC ATG GTA TGA GGT TGA CT-3′ and 5′-ATT GCC TTA TCT TCA GCT TCT AAA TGT-3′), human IFIT1 (5′-AGA AGC AGG CAA TCA CAG AAA A -3′ and 5′-CTG AAA CCG ACC ATA GTG GAA AT-3′), human ISG15 (5′-CGC AGA TCA CCC AGA AGA TCG-3′ and 5′-TTC GTC GCA TTT GTC CAC CA-3′), and human GAPDH (5′-TCA CCA CCA TGG AGA AGG-3′ and 5′-GAT AAG CAG TTG GTG GTG CA-3′) were used. The qPCRs were performed with Power SYBR green PCR master mix (Applied Biosystems) using a Step One real-time PCR instrument. cDNA quantities were normalized to quantities of GAPDH RNA measured from the same samples.

MS analysis.

FLAG-HA1 or FLAG was transiently expressed in HEK293 cells using transfection. Twenty-four hours posttransfection, 100 million FLAG-HA1 or FLAG-expressing cells were harvested and lysed in 10 ml IP lysis buffer for 30 min on ice. Cell lysates were centrifuged at 10,000 × g for 15 min at 4°C to remove intact cells. The supernatant was then incubated with 50 μl anti-FLAG G1 antibody affinity resin overnight with rotation at 4°C. The beads were intensively washed four times to remove nonspecific binding. The precipitates were resuspended in an equal amount of urea-HEPES buffer and were analyzed using mass spectrometry (MS). The MS experiment and data processing were done at the University of Missouri Proteomics Core Facility. Briefly, the precipitated protein samples were digested with trypsin, and the peptides were purified with a C₁₈ tip (Pierce; Thermo Fisher). The peptides were then analyzed by liquid chromatography (LC)-MS on a ThermoFisher linear trap quadrupole (LTQ) Orbitrap XL instrument. Proteins that specifically bound to FLAG-HA1 but not to the FLAG control were determined and analyzed.

Statistical analysis.

Data were analyzed and compared using a bidirectional, unpaired Student’s t test or two-way analysis of variance (ANOVA) with Tukey’s post hoc test. All error bars in figures represent mean values ± standard deviations (SD). Data are representative of at least two independent experimental repetitions.