ABSTRACT
Highly pathogenic avian influenza A(H5N8) viruses first emerged in China in 2010 and in 2014 spread throughout Asia and to Europe and the United States via migrating birds. Influenza A(H5N8) viruses were first detected in the Netherlands in 2014 and caused five outbreaks in poultry farms but were infrequently detected in wild birds. In 2016, influenza A(H5N8) viruses were reintroduced into the Netherlands, resulting in eight poultry farm outbreaks. This outbreak resulted in numerous dead wild birds with severe pathology. Phylogenetic analysis showed that the polymerase genes of these viruses had undergone extensive reassortment between outbreaks. Here, we investigated the differences in virulence between the 2014-15 and the 2016-17 outbreaks by characterizing the polymerase complex of influenza A(H5N8) viruses from both outbreaks. We found that viruses from the 2014-15 outbreak had significantly higher polymerase complex activity in both human and avian cell lines than did those from the 2016-17 outbreak. No apparent differences in the balance between transcription and replication of the viral genome were observed. Interestingly, the 2014-15 polymerase complexes induced significantly higher levels of interferon beta (IFN-β) than the polymerase complexes of the 2016-17 outbreak viruses, mediated via retinoic acid-inducible gene I (RIG-I). Inoculation of primary duck cells with recombinant influenza A(H5N8) viruses, including viruses with reassorted polymerase complexes, showed that the polymerase complexes from the 2014-15 outbreak induced higher levels of IFN-β despite relatively minor differences in replication capacity. Together, these data suggest that despite the lower levels of polymerase activity, the higher 2016-17 influenza A(H5N8) virus virulence may be attributed to the lower level of activation of the innate immune system.
IMPORTANCE Compared to the 2014-15 outbreak, the 2016-17 outbreak of influenza A(H5N8) viruses in the Netherlands and Europe was more virulent; the number of dead or diseased wild birds found and the severity of pathological changes were higher during the 2016-17 outbreak. The polymerase complex plays an important role in influenza virus virulence, and the gene segments of influenza A(H5N8) viruses reassorted extensively between the outbreaks. In this study, the 2014-15 polymerase complexes were found to be more active, which is counterintuitive with the observed higher virulence of the 2016-17 outbreak viruses. Interestingly, the 2014-15 polymerase complexes also induced higher levels of IFN-β. These findings suggest that the higher virulence of influenza A(H5N8) viruses from the 2016-17 outbreak may be related to the lower induction of IFN-β. An attenuated interferon response could lead to increased dissemination, pathology, and mortality, as observed in (wild) birds infected during the 2016-2017 outbreak.
INTRODUCTION
Highly pathogenic avian influenza (HPAI) viruses of hemagglutinin (HA) subtypes H5 and H7 cause widespread epidemics in poultry and wild birds globally, with high morbidity and mortality. These outbreaks lead to substantial economic losses and are a continuous pandemic threat to humans (1–3). Since they were first isolated from a domestic goose in Guangdong Province in China in 1996, the influenza A/Goose/Guangdong/1/1996 (H5N1) lineage viruses have continued to evolve through genetic drift and reassortment events with other avian influenza A viruses, resulting in multiple genetic clades and H5Nx subtypes, like HPAI A(H5N1), A(H5N6), and A(H5N8) viruses (1, 3). Highly pathogenic avian influenza A(H5N8) viruses of clade 2.3.4.4 were first detected in mallards in 2009 and 2010 in China (4). In 2013 and 2014, two different influenza A(H5N8) lineages were detected in China, later designated clade 2.3.4.4 groups A and B (5). In 2014, viruses belonging to clade 2.3.4.4 group A disseminated throughout Asia, North America, and Europe via wild migrating birds and the migratory flyways connecting these geographic areas (5–8). In November 2014, HPAI A(H5N8) viruses were first detected in Europe. In the Netherlands, these viruses caused outbreaks in five poultry holdings (9), resulting in the death of over 300,000 domestic birds upon culling of infected birds or to prevent further spreading. During this outbreak, only a few wild birds tested positive for HPAI A(H5N8) viruses despite intensified active surveillance efforts (10, 11). Although highly virulent in chickens, varied disease severities, including subclinical infection, in wild birds and domestic waterfowl were reported (12). Continued serological and virological surveillance after the 2014-15 (i.e., 2014-2015) outbreak indicated that the 2014-15 HPAI A(H5N8) viruses did not persist in wild bird populations in the Netherlands (11). HPAI A(H5N8) viruses belonging to clade 2.3.4.4 group B continued to circulate in China from 2013 to 2014 onward, and in late 2016, these viruses were reintroduced to Europe via migrating birds (5, 7). In the Netherlands, there were eight influenza A(H5N8) virus outbreaks in commercial poultry holdings, and an unprecedented high mortality among wild birds was observed (13–15). Over 13,000 dead wild birds belonging to 71 species were reported during the 2016-17 (i.e., 2016-2017) outbreak in the Netherlands, of which the majority were tufted ducks (Aythya fuligula) and Eurasian wigeons (Anas penelope) (14). Across Europe, 80 times more poultry outbreaks were observed during the 2016-17 outbreak than during the 2014-15 outbreak. Similarly, throughout Europe, the reported number of dead wild birds and spectrum of species affected was far greater during the 2016-17 outbreak than during the 2014-15 outbreak (16) (https://www.oie.int/animal-health-in-the-world/update-on-avian-influenza). In addition, Pohlmann et al. observed increased pathological changes in dead wild birds found during the 2016-17 outbreak not observed during the 2014-15 outbreak (17).
The observed differences in virulence between the viruses of the 2014-15 and 2016-17 outbreaks were recently confirmed by Grund et al. (18). They experimentally infected domestic ducks, geese, and chickens with influenza A(H5N8) viruses from the 2014-15 and 2016-17 outbreaks in Germany. Compared to the 2014-15 virus, infection of ducks and geese with the 2016-17 influenza A(H5N8) virus led to more explicit clinical signs, with substantial mortality. Interestingly, although the replication kinetics of the 2014-15 virus were comparable to those of the 2016-17 virus in ducks (18), ducks infected with the 2016-17 virus revealed gross disseminated pathology compared to that of ducks infected with the 2014-15 virus, with pronounced neuro- and hepatotropism, illustrating the higher virulence of the 2016-17 influenza A(H5N8) viruses in a well-controlled experimental setting (18).
Taken together, these findings indicate a marked increase in virulence in avian hosts of the HPAI A(H5N8) viruses from the 2016-17 outbreak compared to the virulence of the 2014-15 HPAI A(H5N8) viruses.
We aimed to resolve the differences in observed virulence between the HPAI (A)H5N8 viruses of the 2014-15 and 2016-17 outbreaks by characterizing the polymerase complex of these viruses. We focused on the polymerase complex for several reasons. Genetic analysis of HPAI A(H5N8) viruses from the 2014-15 and 2016-17 outbreaks indicated that multiple reassortment events with low-pathogenic avian influenza (LPAI) viruses occurred along the migration flyways between China and Europe between the outbreak periods. Notably, the internal genes, including the polymerase complex gene segments, had undergone reassortment (5, 13), suggesting a possible role for the different polymerase complex compositions in the observed difference in virulence between outbreaks. Changes in polymerase complex activity may directly affect virus replication and thereby affect virulence (19). The polymerase complex can also affect virulence through the generation of (aberrant) viral RNAs, like miniviral RNA or defective interfering influenza virus RNA, which can trigger an innate immune response via retinoic acid-inducible gene I (RIG-I) and Toll-like receptors (20–22). Here, we describe the functional characterization of the polymerase complexes from several viruses from the 2014-15 and 2016-17 outbreaks in the Netherlands and several other European countries and their relation to virulence. Influenza A(H5N8) viruses of the 2014-15 outbreak had a higher polymerase complex activity in human and avian cell lines and induced higher levels of interferon beta (IFN-β) mediated via RIG-I than viruses from the 2016-17 outbreak. Likewise, inoculation of primary duck cells with a recombinant influenza A(H5N8) virus from the 2014-15 outbreak induced higher levels of IFN-β than with an influenza A(H5N8) virus from the 2016-17 outbreak, despite relatively minor differences in replication capacity. When the polymerase complex genes were exchanged between these recombinant viruses, the higher induction of IFN-β in primary duck cells could be directly related to the 2014-15 influenza A(H5N8) polymerase complex.
RESULTS
Origin and evolution of the influenza A(H5N8) viruses isolated from outbreaks in the Netherlands.The influenza A(H5N8) viruses isolated during the 2014-15 outbreaks in domestic poultry in the Netherlands clustered with other influenza A(H5N8) group A clade 2.3.4.4 viruses circulating in Europe, East Asia, and North America between 2013 and 2015 in the phylogeny of the hemagglutinin (HA) glycoprotein (Fig. 1), as well as the internal genes PB2, PB1, PA, and NP (Fig. 2), MP, and NS (data not shown) (8). This lineage encompasses the HPAI clade 2.3.4.4 A(H5N8) reassortant viruses that were first detected in China in 2010 before subsequent intercontinental spread within that year (8, 23). The 2014-15 outbreak influenza A(H5N8) viruses from the Netherlands have an internal gene cassette acquired through dynamic reassortment with influenza A/Goose/Guandong/1996-like descendant A(H5Nx) viruses circulating in East Asia (24). Clade 2.3.4.4 influenza A(H5N2) and A(H5N5) viruses circulating in China from 2008 to 2014 were located ancestrally to the Netherlands 2014-15 outbreak clade for the polymerase complex segments (Fig. 1 and 2). The influenza A(H5N8) viruses isolated from the 2016-17 outbreaks in the Netherlands formed distinct subclades from the 2014-15 outbreaks in the Netherlands in the phylogenies of all gene segments, reinforcing evidence that the 2016-17 outbreaks in Europe did not result from sustained local circulation but from a novel introduction of influenza A(H5N8) clade 2.3.4.4 viruses (11, 13, 16, 17, 25). The phylogenies of the internal gene segments have distinct evolutionary histories, suggesting that the 2016 viruses were generated by dynamic intersubtype reassortment with cocirculating Eurasian influenza A(H5Nx) and LPAI viruses, as previously reported (13). In the ancestral trace of the HA, NS, PB1, and MP phylogenies, the Netherlands 2016-17 viruses are most closely related to a subclade of group B clade 2.3.4.4 influenza A(H5N8) viruses from China, Mongolia, and Russia, as previously reported (13) (Fig. 1 and 2). The PB2, PA, and NP segments of the Netherlands 2016-17 viruses were acquired by reassortment from cocirculating Eurasian LPAI viruses into the Russia-Mongolia influenza A(H5N8) virus backbone (5, 13). Diversity between the 2014-15 and 2016-17 clades was quantified by the mean interclade distance (Table 1). The diversity between the 2014-15 and 2016-17 clades was far lower for HA than for the internal genes, excluding NS. In summary, this suggests that the complex reassortment of influenza A(H5N8) viruses between the 2014-15 and 2016-17 outbreaks resulted in a higher level of divergence for the internal genes than for the HA and NA glycoprotein genes.
The HA gene segments of the 2014-15 and 2016-17 outbreaks belong to clade 2.3.4.x but cluster differently. A phylogenetic tree of the hemagglutinin gene of the influenza A/Goose/Guangdong/1996 (H5N1)-like H5N8 viruses included in the study is shown. The leaf tips of H5N8 viruses are colored in green, and the HPAI A(H5N8) viruses of the 2014-15 and 2016-17 outbreaks in the Netherlands are highlighted in yellow and purple, respectively. All other H5Nx viruses are represented in black. Notable WHO/OIE/FAO clades (H5N1) are annotated.
The polymerase complex genes of the 2014-15 and 2016-17 outbreak HPAI A(H5N8) viruses cluster differentially. (A to D) Distinct evolutionary history of the internal genes PB2 (A), PB1 (B), PA (C), and NP (D) of the clade 2.3.4.4 influenza A(H5N8) viruses isolated during the 2014-15 and 2016-17 outbreaks in the Netherlands. Trees are rooted to the influenza A/Goose/Guangdong/1996 (H5N1) virus. The leaf tips of H5N8 viruses are colored in green, and the HPAI A(H5N8) viruses of the 2014-15 and 2016-17 outbreaks in the Netherlands are highlighted in yellow and purple, respectively. All other H5Nx viruses are indicated in black. Non-H5Nx viruses included by BLAST search to trace pansubtypic reassortment pathways (see Materials and Methods) are displayed in various colors, excluding green.
Intercluster patristic distance and interclade patristic distance for each segment of viruses from the 2014-15 and 2016-17 HPAI A(H5N8) outbreaks
HPAI influenza A(H5N8) viruses from the 2014-15 outbreak had higher polymerase complex activity.To investigate the role of the polymerase complex in the observed differences in virulence between the 2014-15 and 2016-17 outbreaks of influenza A(H5N8) viruses, we measured the polymerase complex activities for 15 viruses from these outbreaks in the Netherlands and several other European countries (Table 2) using a minigenome assay. Briefly, the PB2, PB1, PA, and NP segments were cloned in expression vectors and transfected into cells together with a reporter plasmid consisting of a firefly luciferase gene containing the noncoding regions of influenza A virus segment 8, serving as viral RNA (vRNA) for the influenza A polymerase complex. The measured luciferase signal quantifies the influenza A polymerase complex transcriptional activity. In order to obtain an understanding of the polymerase activity of influenza A(H5N8) viruses from these outbreaks, taking the genetic diversity of virus isolates into account, the polymerase complex activity of six influenza A(H5N8) viruses from the 2014-15 outbreak and nine viruses from the 2016-17 outbreak were measured in a polymerase activity reporter assay (Fig. 3A). Although there was a range of polymerase complex activities between the various viruses tested in both groups, the median polymerase complex activity of viruses from the 2014-15 outbreak is significantly higher (18-fold, P < 0.05) than that for viruses from the 2016-17 outbreak. The polymerase complex of viruses from the 2014-15 outbreak showed a wider spread in activity than viruses from the 2016-17 outbreak.
Influenza A(H5N8) viruses from the 2014-15 and 2016-17 influenza A(H5N8) outbreaks used to investigate polymerase activity
HPAI A(H5N8) viruses from the 2014-15 outbreak show higher polymerase complex activity in human and avian cells. (A to G) Polymerase complex activity of six influenza A(H5N8) viruses from the 2014-15 and nine from the 2016-17 outbreaks, as measured in minigenome assays in human HEK293T cells (A) and avian cell lines, LMH cells (B), DF-1 cells incubated at 37°C (C), DF-1 cells incubated at 39°C (D), QT-6 cells (E), CGBQ cells (F), and duck embryonic fibroblasts (G). Data are normalized to the negative control. The mean values from at least 3 independent experiments, with each done in quadruplicate, are shown. The median values for the group of viruses are shown as horizontal bars. Closed symbols indicated viruses from the 2014-15 outbreak, and open symbols indicate viruses from the 2016-17 outbreak. Unique symbols represent specific viruses, as indicated in Table 2. (H and I) Polymerase complex activity of influenza A(H5N8) viruses from 2014 (2014-15A, light gray) and 2016 (2016-17H, dark gray) and reassortants thereof in human HEK293T (H) and avian QT6 (I) cell lines. Data are normalized to the negative control (white). Bar graphs show means with standard deviation from 3 independent experiments, each done in quadruplicate. Asterisks indicate significant differences in median polymerase complex activity between the 2014-15 and 2016-17 viruses. *, P < 0.05; **, P < 0.005, Mann-Whitney test.
Since the viruses from the 2014-15 and 2016-17 outbreaks (Table 2) were derived from various bird species, we wanted to investigate the polymerase complex activity in different avian cell types. Therefore, we tested the polymerase complex activity of these viruses in a panel of available avian cell lines from chicken (LMH and DF-1), quail (QT6), goose (CGBQ), and duck (duck embryo) origin (Fig. 3B to G). Compared to HEK293T cells, the overall activity of all isolates was lower in all avian cell lines, which could be explained by lower transfection efficiency in avian cells. Similar to HEK293T cells, there was a considerable range of polymerase complex activities between viruses in avian cell lines. However, consistently across all avian cell lines tested, the viruses of the 2014-15 outbreak had significantly higher median polymerase complex activity than the viruses of the 2016-17 outbreak.
To investigate the effect of temperature on the activity of the polymerase complex, we tested the panel of influenza A(H5N8) viruses in DF-1 cells at 37°C and at 39°C, resembling the higher body temperature in birds (Fig. 3C and D). Overall, the polymerase complex activity was higher in DF-1 cells at 39°C than at 37°C, yet at both temperatures, the viruses from the 2014-15 outbreak had a significantly higher median polymerase complex activity than did those from the 2016-17 outbreak.
To identify which segment(s) of the polymerase complex contributed most to the higher polymerase complex activity of the 2014-15 viruses than of the 2016-17 viruses, we reassorted the polymerase complex genes from a representative virus of the 2014-15 outbreak (2014-15A) with a representative virus of the 2016-17 outbreak (2016-17H). We tested the reassorted polymerase complexes in minigenome assays in human HEK293T cells and avian QT6 cells (Fig. 3H and I). The low polymerase complex activity of the 2016-17H virus was markedly increased with either the PB2 or PA of the 2014-15A isolate in both HEK293T cells and QT6 cells, and no apparent improvements were observed with the PB1 or NP of the 2014-15A virus. When both the PB2 and PA from the 2014-15A virus are combined with the 2016-17H PB1 and NP, the activity of the reassorted polymerase complex was similar to that of the 2014-15A wild-type virus in both HEK293T and QT6 cells. The addition of the 2014-15A PB1 or NP did not increase polymerase activity further. Conversely, combining the PB2 and/or PA of the 2016-17H virus with the remaining 2014-15A polymerase complex segments resulted in a marked decrease in polymerase complex activity not seen when PB1 or NP was reassorted either alone or combined. Which domains or residues within the polymerase subunits are responsible for the observed differences cannot be concluded because of the large genetic differences of all 4 segments between viruses originating from the two outbreaks.
Quantitative PCR indicated no difference in the replication/transcription balance.During infection, the influenza virus polymerase complex transcribes mRNA for protein synthesis and a positive-sense cRNA intermediate in order to generate negative-sense viral RNA (vRNA) for virus replication. Although not fully understood, a balanced replication/transcription process is believed to be important for virus replication and fitness and, thereby, possibly virulence. We wanted to investigate whether there were differences in the balance between replication and transcription for the polymerase complexes of viruses of the 2014-15 and 2016-17 outbreaks which could affect the difference in virulence observed between these outbreaks. To test this, we transfected the polymerase complex genes together with the luciferase reporter plasmid in HEK293T cells, and total RNA was isolated 48 h after transfection. Using gene-specific primers for luciferase vRNA and cRNA and oligo(dT)20 for mRNA (26, 27), we synthesized cDNA which was used for quantitative PCR (qPCR). We selected five viruses from the 2014-15 outbreak (2014-15A, -B, -D, -E, and -F) and four from the 2016-17 (2016-17D, -F, -G, and -H) outbreak to characterize the transcription/replication balance using qPCR. Figures 4A to C show the expression levels of luciferase vRNA, cRNA, and mRNA, respectively. The viruses tested from the 2014-15 outbreak have significant higher median vRNA (P < 0.0005), cRNA (P < 0.005), and mRNA (P < 0.05) expression levels than those in the group of 2016-17 viruses tested. The expression of vRNA, cRNA, and mRNA corresponded to the activity of the polymerase complex activity as measured in minigenome assays, i.e., viruses with high or low activity in the minigenome assay demonstrate similar high or low levels of vRNA, cRNA, and mRNA expression. To investigate whether the ratio between luciferase vRNA and cRNA expression and the ratio between vRNA and mRNA expression were different between the groups of 2014-15 and 2016-17 viruses tested, we normalized the expression data of cRNA and mRNA to that of vRNA (Fig. 4D and E). When normalized, the median expression levels of cRNA and mRNA were no longer significantly different between the two groups, indicating that although there were differences in overall polymerase activities leading to different expression levels of vRNA, cRNA, and mRNA, the ratios between vRNA, cRNA, and mRNA were not significantly altered. Therefore, there was no apparent shift toward either transcription or replication in viruses from the 2014-15 and 2016-17 outbreaks.
Polymerase complexes from 2014-15 and 2016-17 HPAI A(H5N8) viruses do not show apparent differences in the balance between transcription and replication. Shown are quantitative PCR results for luciferase vRNA, cRNA, and mRNA in HEK293T cells transfected with the polymerase complex of influenza A(H5N8) viruses from the 2014-15 and 2016-17 outbreaks. (A to C) The median expression of the 2014-15 and 2016-17 viruses is shown for vRNA (A), cRNA (B), and mRNA (C). (D and E) Ratios between vRNA and cRNA (D) and vRNA and mRNA (E) were calculated by normalizing to vRNA levels. Closed symbols indicated viruses from the 2014-15 outbreak, and open symbols indicate viruses from the 2016-17 outbreak. Unique symbols represent specific viruses, as indicated in Table 2, with cross. *, P < 0.05; **, P < 0.005; ***, P < 0.0005; ns, not significant, Mann-Whitney test.
Higher IFN-β induction by the polymerase complexes of the 2014-15 outbreak HPAI A(H5N8) viruses.The polymerase complex of influenza viruses can induce type I interferons (IFN-α/β) when viral RNAs produced by the polymerase complex activate RIG-I or Toll-like receptors (TLRs) (20–22, 28). We used an IFN-β reporter assay to test whether the polymerase complex of influenza A(H5N8) viruses induced IFN-β and whether there were differences between the viruses of the 2014-15 and 2016-17 outbreaks. Plasmids encoding the polymerase complex proteins were transfected in HEK293T cells together with a plasmid encoding the vRNA of neuraminidase to form ribonucleoproteins (RNPs). Total RNA was extracted 48 h after transfection and was transfected into HEK293T cells together with an IFN-β reporter plasmid consisting of a firefly luciferase gene under the control of the human IFN-β promoter to test whether formed RNA was able to induce an IFN-β response. As a positive control, we transfected poly(I·C), which is a known activator of RIG-I and TLR3, inducing IFN-β signaling. We selected five representative viruses from the 2014-15 outbreak (2014-15A, -B, -D, -E, and -F) and four from the 2016-17 (2016-17D, -F, -G, and -H) outbreak to quantify the IFN-β response. Figure 5A shows the IFN-β induction by the polymerase complexes of representative influenza A(H5N8) viruses from the 2014-15 and 2016-17 outbreaks. Clearly, the majority of the 2014-15 viruses strongly induce IFN-β to high levels compared to that with the viruses tested from the 2016-17 outbreak. The median IFN-β activity of the 2014-15 group was significantly higher (48-fold, P < 0.05) than that of the 2016-17 group in HEK293T cells.
Polymerase complexes of 2014-15 HPAI A(H5N8) viruses induce higher IFN-β levels. (A and B) Shown is the IFN-β reporter activity in wild-type (A) and RIG-I-deficient (B) HEK293T cells normalized to the negative control (white), with poly(I·C) as a positive control (black). Viruses from 2014 to 2015 are shown in light gray, and viruses from 2016-17 are shown in dark gray. (C and D) The polymerase complex gene segments of an influenza A(H5N8) virus from 2014 (2014-15A, light gray) and 2016 (2016-17H, dark gray) were reassorted, and the induction of IFN-β activity was measured in wild-type (C) and RIG-I-deficient (D) HEK293T cell lines. All bar graphs show means and standard deviations of the results from at least 3 independent experiments, each done in quadruplicate. Asterisks indicate significant differences in mean IFN-β induction between the 2014-15 and 2016-17 viruses. *, P < 0.05; ns, not significant; Mann-Whitney test.
To identify which segment(s) of the polymerase complex contribute most to the higher median IFN-β induction by the polymerase complex of the 2014-15 isolates, we reassorted the polymerase complex genes of the 2014-15A virus with the 2016-17H virus and tested IFN-β induction in HEK293T cells in IFN-β reporter assays. As for the overall polymerase complex activity, the induction of IFN-β was dependent on both the 2014-15 PB2 and PA (Fig. 5B), although the effect of the individual genes was not as apparent as in the polymerase activity assays.
To test whether the IFN-β induction was dependent on RIG-I, we performed the same experiments in a RIG-I-deficient HEK293T cell line (Fig. 5C and D). The IFN-β response induced by the 2014-15 influenza A (H5N8) virus polymerase complexes were completely absent in RIG-I-deficient cells.
Minor differences in replication kinetics of recombinant A(H5N8) viruses from the 2014-15 and 2016-17 outbreaks.To test whether the higher polymerase complex activity of the 2014-15 outbreak viruses also resulted in higher replication of these viruses, we investigated the replication kinetics of recombinant viruses in primary duck cells. We selected two representative virus isolates, one from each outbreak, representing a virus containing a polymerase complex with a high polymerase activity (A/Chicken/Netherlands/emc-3/2014 [2014-15A]) and one with a low polymerase activity (A/Eurasian Wigeon/Netherlands/4/2016 [2016-17H]). Wild-type recombinant viruses were constructed, as well as reassortant viruses containing the PB2, PB1, PA, and NP gene segments derived from the virus originating from one outbreak with the other 4 segments derived from a virus originating from the other outbreak.
Duck embryo fibroblast (DEF) and duck lung homogenate (DLH) cells were isolated from embryonated duck eggs and inoculated with recombinant influenza A(H5N8) viruses from the 2014-15 outbreak (2014-15A) and 2016-17 outbreak (2016-17H), as well as two recombinant reassortant viruses in which the polymerase complexes were exchanged. This experiment revealed that the higher polymerase complex activity of the 2014-15A polymerase complex only modestly affected virus replication under these conditions (Fig. 6A and D). Viruses with the 2014-15 polymerase complex grew to slightly higher viral titers, although this was only clearly observed in DEF cells (Fig. 6B and E). Viruses with the 2016–2017 polymerase complex replicated slightly more slowly, with a peak at 48 h, but replicated to similar viral titers.
Recombinant HPAI A(H5N8) viruses from different outbreaks show similar replication kinetics in DEF and DLH cells, but viruses containing the 2014-15 polymerase complex induce higher levels of IFN-β. Primary DEF and DLH were inoculated with an MOI of 0.001 with one of the 4 following recombinant HPAI A(H5N8) viruses: one from the 2014-15 outbreak (2014-15A, ●), one from the 2016-17 outbreak (2016-17H, ■), a recombinant 2014-15A virus containing the 2016-17H polymerase complex (▲), and a recombinant 2016-17H virus containing the 2014-15A polymerase complex (▼). Viral titers were determined from samples taken up to 72 h after inoculation, and IFN-β expression was measured at 24 h after inoculation and normalized to the 2014-15A virus. (A) Virus replication on DEF cells. (B) Viral titers on DEF cells at 24 h postinoculation. (C) IFN-β expression at 24 h after inoculation on DEF cells. (D) Virus replication on DLH cells. (E) Viral titers on DLH cells at 24 h postinoculation. (F) IFN-β expression at 24 h after inoculation on DLH cells. *, P < 0.05; **, P < 0.005; ***, P < 0.0005; ns, not significant, unpaired t test.
Higher IFN-β induction by the 2014-15 polymerase complex in inoculated primary duck cells.To test whether the IFN-β induction by the polymerase complex of influenza A(H5N8) viruses as seen in IFN-β reporter assays also occurred in virus-infected cells, IFN-β induction was measured in primary duck cells upon inoculation with 2014-15A, 2016-17H, and reassortant viruses thereof with exchanged polymerase complexes. DEF and DLH cells were inoculated, and cells were harvested 24 h after inoculation, after which total RNA was isolated and IFN-β mRNA levels were measured. The mean IFN-β expression level was normalized to the expression level of the 2014-15A virus. In DEF cells (Fig. 6C), the 2016-17 virus induces significantly less IFN-β than does the 2014-15 virus, although the differences are not as substantial as in the reporter assays shown earlier (2-fold, P < 0.005). The 2014-15 virus with the 2016-17 polymerase complex also induced a low level of IFN-β comparable to that of the full 2016-17 virus and significantly lower than that of the full 2014-15 virus (0.6-fold, P < 0.005). The 2016-17H virus with the 2014-15A polymerase complex induced IFN-β expression levels higher than those in the full 2014-15A virus (1.5-fold, P < 0.05) and much higher than those in the full recombinant 2016-17H virus (2.5-fold P < 0.005) and the 2014-15A virus with the 2016-17H polymerase complex (2.4-fold, P < 0.005). The results in DEF cells were similar (Fig. 6F).
DISCUSSION
The striking differences in the numbers of affected birds by the 2014-15 and 2016-17 outbreaks of influenza A(H5N8) viruses in the Netherlands indicate differences in virus virulence. The aim of our study was to investigate these differences between the 2014-15 and 2016-17 outbreaks by characterizing their polymerase complexes. Although influenza virus HA is extremely important for virulence, as receptor specificity, avidity, and HA stability are key factors during virus entry and thereby affect viral replication and virulence, no genetic differences are present in the receptor binding sites between viruses from these two outbreaks. Therefore, differences in receptor binding, resulting in altered tissue tropism, for example, are unlikely.
Nearly all studies investigating the role of the polymerase complex in avian influenza viruses focus on virulence markers related to host adaptation of avian influenza viruses to mammalian hosts, either to humans or model systems like mice (29–31). So far, no studies have identified specific polymerase complex virulence markers in HPAI viruses like influenza A(H5N8) virus related to disease severity in birds. Our results showed that the polymerase complex activity of viruses from the 2014-15 outbreak is significantly higher than that in viruses from the 2016-17 outbreak in both human and avian cell lines. This observation is in contrast with the idea that increased polymerase activity would result in higher virus titers and therefore higher virulence. There are mismatches between species of hosts from which the viruses were obtained and the cell lines in which we tested the polymerase complex activity, as well as discrepancies between the cellular origins of cell lines and the tissue tropisms of these viruses. Nonetheless, consistently across all tested cell lines, the 2014-15 outbreak influenza A(H5N8) viruses had significantly higher polymerase complex activity than did the 2016-17 outbreak influenza A(H5N8) viruses, and no apparent host-specific or temperature effects on polymerase complex activity were observed. Viruses obtained from different geographic locations across Europe from both outbreaks had activities similar to those of viruses from the corresponding outbreaks in the Netherlands, suggesting a more general phenomenon.
For five viruses from the 2014-15 and four viruses from the 2016-17 outbreak, we quantified vRNA, cRNA, and mRNA expression levels. No apparent differences in the transcription and replication balance were observed that could play a role in the observed virulence differences between outbreaks. Although the exact role in virulence for the transcription and replication balance is unknown, it was previously shown that during host adaptation of HPAI H5N1 in the ferret model, PB2-E627K altered the transcription and replication balance, which was restored by a mutation in PB1 (H99Y) that was selected upon growth of the virus in ferrets (32). The findings from that study indicate that there might be an optimal balance for efficient replication and virulence. However, as viruses of 2014-15 and 2016-17 outbreaks show very similar transcription and replication balances, it is unlikely that this plays a role in the observed differences in virulence.
Interestingly, our IFN-β reporter assays demonstrated that the polymerase complexes of viruses from the 2014-15 HPAI A(H5N8) virus outbreak induce higher levels of IFN-β than the polymerase complexes of viruses from the 2016-17 outbreak, mediated through RIG-I.
To further investigate the differences in pathogenicity, we selected one virus from each outbreak, representing a polymerase complex with either high (2014-15A) or low (2016-17H) polymerase complex activity, to study replication kinetics and IFN-β induction in primary duck cells. Despite the significant differences in polymerase complex activity between these viruses, there were only in minor replicative differences in primary duck cells. This could be due to the fact that viral replication is governed by multiple factors other than polymerase complex function, including receptor binding, virus entry, and modularly functions by other internal proteins. However, experiments where we exchanged polymerase complexes between the 2014-15 and 2016-17 viruses suggest that the HA and NA (and NS and M segments) of the 2016-17 virus with the 2014-15 polymerase complex (PB2, PB1, PA, and NP segments) did not affect replication significantly. This was at least partially expected from the very similar receptor binding site sequences of HA. Another explanation for the small difference in virus replication is that the replication of these viruses in primary duck cells under laboratory conditions is extremely efficient, making it difficult to observe differences in replication. Other tests, such as plaque assays or, eventually, in vivo animal models, to obtain a better resolution of the differences in replication kinetics and virulence between these viruses may be required.
Despite the relatively small differences in replication in primary duck cells, we observed a small but significantly higher IFN-β induction in primary duck cells by viruses of the 2014-15 outbreak than in viruses from the 2016-17 outbreak. Since our virus inoculation experiments included viruses with exchanged polymerase complexes, the IFN-β induction can be attributed to the polymerase complex itself, irrespective of the effects of other regulatory viral proteins on IFN-β induction. An obvious shortcoming here is that we did not study the dynamics of IFN-β expression over the course of infection. Recently, it has been shown for influenza A(H5N1) viruses that the IFN-β response in Peking ducks is strongly regulated, and it peaks at 24 h after inoculation and declines shortly thereafter (33), which suggests that the 24-h IFN-β measurements are representative. However, earlier and later time points could be useful to determine whether the IFN-β response to influenza A(H5N8) viruses from the 2016-17 outbreak are attenuated or delayed compared to that in viruses from the 2014-15 outbreak.
Important factors determining the outcome of HPAI virus infection are the host species infected, with ducks being generally more resilient than chickens; age of the host, with young animals being more susceptible; and virus strain-specific properties (34). Ducks are important natural hosts of LPAI viruses and permissive to LPAI virus replication in their intestines, with minimal cytokine and type I IFN induction (34). In contrast, HPAI viruses replicate more abundantly in the respiratory tract, and infections spread systemically with an immediate and robust induction of cytokines and type I IFN expression of interferon-stimulated genes (34). Compared to ducks, chickens are extremely susceptible to HPAI virus infection, likely due to the lack of RIG-I in chickens (34, 35), emphasizing the importance of the innate immune response in controlling HPAI virus infections. This difference in disease severity between chicken and ducks was confirmed for the HPAI A(H5N8) viruses in two recent studies in which chickens showed more severe disease than did several water bird species, like mallards and geese, which only showed mild pathology and low mortality upon infection (12, 18). The RIG-I receptor is a type of intracellular pattern recognition receptor involved in the recognition of viruses as part of the innate immune system. RIG-I recognizes short double-stranded RNA with a di- or triphosphate motif at the 5′ end, as found at the noncoding regions of influenza virus genome segments (36, 37). The intrinsic low fidelity of the influenza polymerase complex leads to the generation of aberrant viral RNAs, like miniviral RNAs and defective interfering influenza virus RNA, which can induce IFN signaling via RIG-I (20, 37). RIG-I is crucial for early type I IFN-α/β responses to influenza A virus infections (22, 36, 38, 39). Low levels of RIG-I-mediated IFN-α/β signaling could lead to an inadequate innate immune response and severe infections. Indeed, rare somatic mutations in the RIG-I gene lead to severe influenza infection in humans (40), and RIG-I knockout studies in mice demonstrate impaired type I IFN-α/β induction as well as a diminished adaptive immune response (41). Given the fact that viruses from the 2016-17 influenza A(H5N8) outbreak induced lower levels of IFN-β expression, it is tempting to speculate that the polymerase complex of the 2016-17 outbreak allowed for immune evasion during early infection by reduced IFN-β induction. The absence of a strong (initial) immune response may lead to dysregulation of the IFN-β cascade and could result in prolonged infection and shedding and increased dissemination, pathology, and mortality of the 2016-17 influenza A(H5N8) viruses compared to those with the 2014-15 viruses, as seen in the experimentally infected ducks and geese with the 2016 influenza A(H5N8) virus (18).
Future animal experiments comparing viruses from the 2014-15 and 2016-17 outbreaks, including viruses where the polymerase complexes genes are reassorted, could further elucidate the role of the polymerase complex in the virulence of these viruses. Monitoring the levels of IFN, other cytokines, and IFN-stimulated genes during infection could provide insight into how the innate immune response develops over the course of infection and how the polymerase complex from virus strains determines dissemination and pathology as well as tissue tropism.
Taken together, our data suggest a role for the polymerase complex of HPAI A(H5N8) viruses in virulence related to the induction of a protective IFN response. In the context of HPAI A(H5N1) virus infection in humans, a strong induction of IFN can result in hypercytokinemia, which has been shown to be associated with a fatal outcome in human cases of HPAI (H5N1) virus infection (42). However, innate immunity has a strong protective role against virus infections in both humans and avian species. It was shown that the induction of type I IFNs is essential in defending the host against viral infections in avian species (43). Therefore, a more robust, but balanced, induction of the innate immune system is important in the control of influenza virus infections.
MATERIALS AND METHODS
Phylogenetic analysis influenza A(H5Nx) viruses.The sequences of all influenza A(H5Nx) subtype viruses collected between 1996 and 2018 were downloaded from GISAID (44). The sequence data set was curated to remove sequences with more than 1% ambiguous nucleotides, less than 80% sequence length, or sequences that had any of the eight segments missing. The sequence set was annotated with the WHO/OIE/FAO H5 nomenclature using LABELv0.5.2 with module H5v2015 (45). The influenza A(H5N1) A/goose/Guangdong/1/1996 (H5N1) virus-like lineage was systematically downsampled by a WHO/OIE/FAO-designated clade to obtain a representative influenza A/Goose/Guangdong/1996 (H5N1) virus-like background data set (n = 500 viruses), with the influenza A(H5N8) viruses from the Netherlands isolated in the 2014-15 and 2016-17 outbreaks automatically included in the final data set for each segment. To account for the possibility of reassortment with the internal genes of LPAI viruses, each of the influenza A(H5N8) viruses from the Netherlands was compared to the complete global database of avian influenza viruses for each segment with BLAST+ (46). The 50 most closely related LPAI sequences were included in the final data set for each segment (n = 597), with the exception of HA and NA. The final data sets were aligned with MAFFT v7.397 and trimmed to the start of the mature protein (47). Maximum likelihood phylogenetic trees were constructed for each segment with RAxML 8.2.12 under the GTR+GAMMA substitution model with 100 bootstrap replicates and rooted to the influenza A/Goose/Guangdong/1996 (H5N1) virus (48). Phylogenetic trees were annotated and visualized using ggtree (49). Pairwise mean interclade patristic distance distributions were calculated using the ape package in R (50). The files are available at https://github.com/AMC-LAEB/avflu_h5n8_polymerase.
Cell culture.Human HEK293T (ATCC CRL-3216) cells and HEK293T RIG-I-deficient cells (kindly provided Ben Berkhout, Amsterdam UMC) were cultured in Dulbecco’s modified Eagle’s medium (DMEM; Gibco) supplemented with 10% fetal calf serum (FCS; Sigma), 1% nonessential amino acids (NEAA; Gibco), 100 IU/ml penicillin, and 100 IU/ml streptomycin (Pen/Strep; Gibco) incubated at 37°C and 5% CO2. The HEK293T RIG-I knockout cells were produced using CRISPR-Cas gene knockout. HEK293T cells were transfected with three px458 plasmids expressing the CRISPR-enhanced green fluorescent protein (CRISPR-eGFP) protein and three independent guide RNA sequences targeting genomic RIG-I at a ratio of 1:1:1. After 48 h, cells were harvested, and single eGFP fluorescent cells were sorted into a 96-well plate. Expanding cell colonies were cultured for 6 weeks under the same conditions as for regular HEK293T cells. Cells were harvested and frozen in aliquots. Gene knockout was checked by gene-specific PCR that confirmed the absence of the genomic region containing the RIG-I gene, using a positive control for DNA extraction efficiency and PCR. Neither luciferase background levels nor absolute polymerase complex activity was significantly different between wild-type HEK293T and RIG-I knockout cells (data not shown). DF-1 cells from Gallus gallus (ATCC CRL-12203) were cultured in DMEM supplemented with 10% FCS, 1% NEAA, 100 IU/ml penicillin, and 100 IU/ml streptomycin and incubated at either 37°C and 5% CO2 or 39°C and 5% CO2. LMH cells from Gallus gallus (ATCC CRL-2218) were cultured in DMEM supplemented with 10% FCS, 1% NEAA, 100 IU/ml penicillin, and 100 IU/ml streptomycin and incubated at 37°C and 5% CO2. QT6 cells from Coturnix coturnix japonica (ATCC CRL-1708) were cultured in medium 199 with Earle salts and 2 mM l-glutamine (M199; Gibco) supplemented with 5% FCS, 1% chicken serum (Sigma), 1% of 100× GlutaMAX-I (Gibco), 5% tryptose phosphate broth (TPB; Gibco), 100 IU/ml penicillin, and 100 IU/ml streptomycin and incubated at 37°C and 5% CO2. CGBQ cells from Anser anser (ATCC CRL-169) were cultured in Ham’s F-12 nut mix with 2 mM l-glutamine (Ham’s F-12K; Gibco) supplemented with 15% FCS, 1% NEAA, 0.15% sodium bicarbonate (Gibco), 100 IU/ml penicillin, and 100 IU/ml streptomycin and incubated at 37°C and 5% CO2. Duck embryo cells from Anas platyrhynchos domesticus (ATCC CRL-2118) were cultured in minimum essential medium with Hanks salts and 2 mM l-glutamine (MEM; Gibco) supplemented with 10% FCS, 1 mM sodium pyruvate (100 mM; Gibco), 100 IU/ml penicillin, and 100 IU/ml streptomycin and incubated at 37°C and 5% CO2. Madin-Darby canine kidney (MDCK) cells from Canis familiaris (ATCC CRL-2935) were cultured in Eagle’s minimal essential medium (EMEM; Lonza) supplemented with 10% FCS, 100 IU/ml penicillin, 100 IU/ml streptomycin, 2 mM l-glutamine (Gibco), 0.15% sodium bicarbonate (Gibco), 10 mM HEPES (Lonza), and 1% NEAA (Gibco) and incubated at 37°C and 5% CO2.
Primary duck embryo fibroblast were isolated from 13-day-old embryonated eggs, cultured in medium 199 (M199; Lonza), 10% tryptose phosphate broth (TPB; MP Biomedicals), 10% FCS, 10% 100 IU/ml penicillin, and 100 IU/ml streptomycin, and incubated at 37°C and 5% CO2. Duck lung homogenate (DLH) cells were isolated from the lungs of 21-day-old duck embryonated eggs, grown in DMEM (Gibco) supplemented with 10% FCS (Sigma), 1% NEAA (Gibco), 100 IU/ml penicillin, and 100 IU/ml streptomycin (Pen/Strep; Gibco), and incubated at 37°C and 5% CO2.
Polymerase activity/minigenome assay.The open reading frames (coding region) of the PB2, PB1, PA, and NP genes were cloned into the pPPI4 expression vector (51) using Gibson Assembly (New England BioLabs). A plasmid coding for a model viral RNA (vRNA), consisting of the firefly luciferase (FF) open reading frame flanked by the noncoding regions of segment 8 of influenza A virus under the control of a human pPolI was used for minigenome assays (52). The production of mRNA of this FF vRNA is solely possible by transcription of the produced vRNA by the influenza virus polymerase complex. Therefore, the FF activity is a measure of the amount of mRNA produced and thereby of the polymerase complex activity. Transfection of pRL (Promega), a plasmid containing the Renilla luciferase gene under a cytomegalovirus promoter which is stably expressed in avian and mammalian cells independent from the influenza polymerase complex, served as an internal control to normalize variation in transfection efficiency and sample processing. Human (HEK293T) or avian (DF-1, LMH, QT6, CGBA, or duck embryo) cells were seeded 1 day prior to the experiment into 96-well plates. Twenty-five nanograms of the FF reporter plasmid, 50 ng of each of the plasmids encoding PB2, PB1, and PA, and 100 ng of NP, and 2 ng of the pRL expression plasmid in 50 μl Opti-MEM (Gibco, Thermo Fisher) were mixed with 50 μl Opti-MEM containing either Lipofectamine 2000 (Invitrogen, Thermo Fisher) for HEK293T cells or TransIT-X2 (Mirus) for avian cells in a 1:3 ratio and incubated for 20 to 30 min at room temperature. Twenty microliters of the transfection mixture was added to each well. Each transfection was performed in quadruplicate in at least three independent experiments. Twenty-four hours after transfection, or 48 h for duck embryo cells, luminescence was measured using the dual-luciferase reporter assay system (Promega) using a GloMax luminometer, according to the manufacturer’s instructions (Turner BioSystems).
Quantitative PCR luciferase vRNA, cRNA, and mRNA.Transfection of cells was performed as described above for the minigenome experiments. Briefly, 25 ng of the FF reporter plasmid, 50 ng of each of the plasmids encoding PB2, PB1, and PA, 100 ng of NP, and 2 ng of the pRL expression plasmid in 50 μl Opti-MEM (Gibco, Thermo Fisher) were mixed with 50 μl Opti-MEM containing Lipofectamine 2000 (Invitrogen, Thermo Fisher). HEK293T cells cultured in a 24-well format were transfected with 100 μl transfection mixture and incubated for 48 h; after this, medium was aspirated, and cells were lysed in 500 μl TRIzol (Thermo Fisher) and stored at –80°C until RNA isolation. Transfections were performed in duplicate, and for each transfection, separate cDNA synthesis and qPCR were performed twice. RNA isolation from TRIzol was performed according to the manufacturer’s protocol. To ensure complete removal of any residual DNA (genomic/plasmid), the isolated RNA was treated with 100 units of Turbo DNase (Thermo Fisher) for 2 h, and RNA was isolated a second time using TRIzol.
For cDNA synthesis, 1 μg of purified total RNA was used, to which 1 μl of 50 μM oligo(dT)20 (Thermo Fisher) or 1 μl of 2 μM gene-specific primer and 1 μl of 10 mM dinucleoside triphosphate (dNTP) mix (Thermo Fisher) were added in a total volume of 13 μl. Oligo(dT)20 was used to generate mRNA, gene-specific primers for luciferase vRNA (5′-AGTAGAAACAAGGGTGTTTTTTATCA-3′), and cRNA (5′-TATGGGCATTTCGCAGCCTACCGTGGTGTT-3′) (adapted from references 26 and 27). This mixture was heated to 65°C for 5 min and then incubated on ice for at least 1 min. Next, 4 μl of 5× first-strand buffer (FSB; Thermo Fisher), 1 μl of 0.1 M dithiothreitol (DTT; Thermo Fisher), 1 μl RNaseOUT (Thermo Fisher), and 1 μl of 200 U/μl SuperScript III reverse transcriptase (Thermo Fisher) were added to each reaction mixture, incubated at 55°C for 60 min and 75°C for 15 min, and cooled to 4°C. For the real-time quantitative PCR, 2 μl of cDNA was used, along with 10 μl of 2× SYBR FAST qPCR master mix for LightCycler480 (LC480) (catalog no. KK4610; Kapa Biosystems), 4 μl PCR-grade water; 2 μl of 2 μM forward primer and 2 μl of 2 μM reverse primer were used for each reaction. For the detection of luciferase vRNA, cRNA, and mRNA luciferase, qPCR primers were used (forward, 5′-TATGAACATTTCGCAGCCTACCGTAGTGTT-3′; reverse, 5′-CCGGAATGATTTGATTGCCA-3′) (26, 27). For quantification of the household gene glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA, qPCR primers were used (forward, 5′-AAAATCAAGTGGGGCGATGCT-3′; reverse, 5′-GGGCAGAGATGATGACCCTTT-3′) (53). Real-time quantitative PCR using Kapa SYBR FAST qPCR mix for LC480 was performed on a LC480 (Roche), according to the manufacturer’s protocol. Gene expression relative to the negative control was calculated using the ΔΔCT method (54), using GAPDH as a household gene.
IFN-β reporter assay.The influenza A virus ribonucleoprotein complex (RNP) was reconstituted in HEK293T cells using vRNA encoding NA of the A/Puerto Rico/8/34 (H1N1) virus instead of the FF reporter. Similar to the minigenome assays, 50 ng of each of the plasmids encoding PB2, PB1, and PA, 100 ng of NP, 25 ng of NA of A/Puerto Rico/8/34 (H1N1) virus, and 2 ng of the Renilla luciferase expression plasmid in 50 μl Opti-MEM were mixed with 50 μl Opti-MEM containing Lipofectamine 2000 in a 1:3 ratio and incubated for 20 to 30 min at room temperature. HEK293T cells grown in a 24-well format were transfected at 80% to 90% confluence and incubated for 48 h, after which cells were aspirated and resuspended in 500 μl TRIzol. Total RNA was extracted using the Direct-Zol RNA miniprep kit (Zymo Research), according to the manufacturer’s instructions. Four hundred nanograms of total RNA with 25 ng of IFN-β reporter plasmid (kindly provided A. J. W. te Velthuis [20, 28]) and 2 ng of the pRL expression plasmid in 50 μl Opti-MEM were mixed with 50 μl Opti-MEM containing Lipofectamine 2000 in a 1:3 ratio and incubated for 30 min. As a positive control, we used poly(I·C) (Thermo Fisher) in 10-fold dilutions at final concentrations of 1 μg to 10 ng in 20 μl of transfection mixture. HEK293T and HEK293T RIG-I knockout cells grown in a 96-well format were transfected at 80% to 90% confluence with 20 μl of the transfection mixture for each well. Each transfection was performed in quadruplicate in at least three independent experiments. At 48 h after transfection, luminescence was measured using the dual-luciferase reporter assay system (Promega) using a GloMax luminometer, according to the manufacturer’s instructions (Turner BioSystems).
Viruses.Rescue plasmids for an isolate from the 2014-15 outbreak [A/Chicken/Netherlands/emc-3/2014 (H5N8)] were described previously (29, 55). The eight gene segments of an isolate from the 2016-17 outbreak [A/Eurasian Wigeon/Netherlands/4/2016 (H5N8)] were amplified from RNA by one-step reverse transcription-PCR (RT-PCR; Thermo Fisher) using specific primers and were cloned in the bidirectional reverse-genetics plasmid pHW2000, as described before (56, 57). Recombinant viruses were rescued by reverse genetics upon transfection of HEK293T cells, as previously described (57). For the infection experiments, 4 recombinant viruses were generated, as follows: full recombinant A/Chicken/Netherlands/emc-3/2014 virus; full recombinant A/Eurasian Wigeon/Netherlands/4/2016 virus; recombinant A/Chicken/Netherlands/emc-3/2014 virus with the PB2, PB1, PA, and NP gene segments derived from A/Eurasian Wigeon/Netherlands/4/2016 virus; and a recombinant A/Eurasian Wigeon/Netherlands/4/2016 virus with PB2, PB1, PA, and NP gene segments derived from A/Chicken/Netherlands/emc-3/2014 virus. Virus stocks were propagated in and titrated on MDCK cells, as described below.
Titrations.MDCK cells were inoculated with 10-fold serial dilutions of virus stocks or supernatant of DLH- or DEF-inoculated primary cells. The cells were washed with phosphate-buffered saline (PBS) 1 h after inoculation, cultured in infection medium consisting of EMEM supplemented with 100 IU/ml penicillin, 100 IU/ml streptomycin, 2 mM l-glutamine (Gibco), 0.15% sodium bicarbonate (Gibco), 10 mM HEPES (Gibco), and 1% NEAA (Gibco), and incubated at 37°C and 5% CO2. Three days after inoculation, the supernatants of the cell cultures were tested for agglutinating activity using turkey red blood cells (tRBCs) as an indicator of virus replication. Infectious virus titers were calculated by the method of Reed and Muench (58).
Viral replication kinetics.DEF and DLH cells were inoculated with a multiplicity of infection (MOI) of 0.001. The supernatant was harvested at 12, 24, 36, 48, and 72 h postinfection. Viral loads were determined by titration in MDCK cells, as described above.
IFN-β qPCR on HPAI A(H5N8) virus-inoculated primary duck cells.DEF and DLH primary cells were plated in 6-well plates at 1 to 2 days prior to inoculation. Cells were washed with PBS and inoculated in duplicate for DEF cells and triplicate for DLH cells with an MOI of 0.001 for each of the 4 recombinant viruses in infection medium consisting of M199 and DMEM, respectively, containing 100 IU/ml penicillin and 100 IU/ml streptomycin. Uninfected DEF and DLH cells were used as a negative control. Cells were aspirated at 24 h postinfection and were lysed in 500 μl lysis buffer using the Total RNA isolation kit (Roche). RNA was isolated according to the manufacturer’s protocol. cDNA synthesis was performed as described above, except that 250 ng total RNA was used for cDNA synthesis using 1 μl of 50 μM oligo(dT)20 (Thermo Fisher). Quantitative PCR was performed as described above, using duck IFN-β qPCR primers (forward, 5′-ACCTCCTCAACCAGCTCAA-3′; reverse, 5′-GAAGTGTTGGATGCTCCTGA-3′) and duck GAPDH qPCR primers (forward, 5′-ATGTTCGTGATGGGTGTGAA-3′; reverse, 5′-CTGTCTTCGTGTGTGGCTGT-3′) (53). IFN-β gene expression relative to the negative control was calculated with the ΔΔCT method (54) using GAPDH as a household gene.
Statistical analysis.The polymerase complex activity of viruses from the 2014-15 and 2016-17 outbreaks, the polymerase complex activity of reassortants thereof, the IFN-β reporter activity, and the expression levels of vRNA, cRNA, and mRNA were compared by using a Mann-Whitney t test (Prism 8.0.2; GraphPad). IFN-β expression levels in inoculated primary duck cells and viral titers were compared by using a two-tailed unpaired t test (Prism 8.0.2; GraphPad). A P value of <0.05 was considered statistically significant. *, P < 0.05; **, P < 0.005; ***, P < 0.0005, ns not significant.
ACKNOWLEDGMENTS
We thank Theo Bestebroer and Stefan van der Vliet (Erasmus MC, Rotterdam, the Netherlands) for technical assistance. We thank Ronald Dijkman from the University of Bern, Institute of Virology and Immunology, for kindly sharing virus sequences and cDNA for cloning purposes. We gratefully acknowledge the originating and submitting laboratories of the sequences from GISAID’s EpiFlu Database on which part of this research is based. All submitters of data may be contacted directly via the GISAID website (https://www.gisaid.org/).
This work was funded by a European Union H2020 Marie Curie International Incoming Fellowship to D.E., R.A.M.F. and N.S.L. are supported by NIAID/NIH contract HHSN272201400008C (CEIRS), and C.A.R. is supported by an ERC consolidator grant. E.P. is supported by the Gates Cambridge Trust (Bill and Melinda Gates Foundation; OPP1144).
FOOTNOTES
- Received 4 March 2020.
- Accepted 4 March 2020.
- Accepted manuscript posted online 1 April 2020.
- Copyright © 2020 American Society for Microbiology.