Hemagglutinin Stability Regulates H1N1 Influenza Virus Replication and Pathogenicity in Mice by Modulating Type I Interferon Responses in Dendritic Cells

Diverse influenza A viruses circulate in wild aquatic birds, occasionally infecting farm animals. Rarely, an avian- or swine-origin influenza virus adapts to humans and starts a pandemic. Seasonal and many universal influenza vaccines target the HA surface protein, which is a key component of pandemic influenza viruses. Understanding the HA properties needed for replication and pathogenicity in mammals may guide response efforts to control influenza. Some antiviral drugs and broadly reactive influenza vaccines that target the HA protein have suffered resistance due to destabilizing HA mutations that do not compromise replicative fitness in cell culture. Here, we show that despite not compromising fitness in standard cell cultures, a destabilizing H1N1 HA stalk mutation greatly diminishes viral replication and pathogenicity in vivo by modulating type I IFN responses. This encourages targeting the HA stalk with antiviral drugs and vaccines as well as reevaluating previous candidates that were susceptible to destabilizing resistance mutations.

ABSTRACT

Hemagglutinin (HA) stability, or the pH at which HA is activated to cause membrane fusion, has been associated with the replication, pathogenicity, transmissibility, and interspecies adaptation of influenza A viruses. Here, we investigated the mechanisms by which a destabilizing HA mutation, Y17H (activation pH, 6.0), attenuates virus replication and pathogenicity in DBA/2 mice compared to wild-type (WT) virus (activation pH, 5.5). The extracellular lung pH was measured to be near neutral (pH 6.9 to 7.5). WT and Y17H viruses had similar environmental stability at pH 7.0; thus, extracellular inactivation was unlikely to attenuate the Y17H virus. The Y17H virus had accelerated replication kinetics in MDCK, A549, and RAW 264.7 cells when inoculated at a multiplicity of infection (MOI) of 3 PFU/cell. The destabilizing mutation also increased early infectivity and type I interferon (IFN) responses in mouse bone marrow-derived dendritic cells (DCs). In contrast, the HA-Y17H mutation reduced virus replication in murine airway murine nasal epithelial cell and murine tracheal epithelial cell cultures and attenuated virus replication, virus spread, the severity of infection, and cellular infiltration in the lungs of mice. Normalizing virus infection and weight loss in mice by inoculating them with Y17H virus at a dose 500-fold higher than that of WT virus revealed that the destabilized mutant virus triggered the upregulation of more host genes and increased type I IFN responses and cytokine expression in DBA/2 mouse lungs. Overall, HA destabilization decreased virulence in mice by boosting early infection in DCs, resulting in the greater activation of antiviral responses, including the type I IFN response. These studies reveal that HA stability may regulate pathogenicity by modulating IFN responses.

IMPORTANCE Diverse influenza A viruses circulate in wild aquatic birds, occasionally infecting farm animals. Rarely, an avian- or swine-origin influenza virus adapts to humans and starts a pandemic. Seasonal and many universal influenza vaccines target the HA surface protein, which is a key component of pandemic influenza viruses. Understanding the HA properties needed for replication and pathogenicity in mammals may guide response efforts to control influenza. Some antiviral drugs and broadly reactive influenza vaccines that target the HA protein have suffered resistance due to destabilizing HA mutations that do not compromise replicative fitness in cell culture. Here, we show that despite not compromising fitness in standard cell cultures, a destabilizing H1N1 HA stalk mutation greatly diminishes viral replication and pathogenicity in vivo by modulating type I IFN responses. This encourages targeting the HA stalk with antiviral drugs and vaccines as well as reevaluating previous candidates that were susceptible to destabilizing resistance mutations.

INTRODUCTION

Influenza A virus (IAV) isolates vary in fitness and pathogenicity because of differences in viral genetics and in the ability of the viruses to stimulate or evade host responses. In the H1 to H16 subtypes, the influenza virus hemagglutinin (HA) surface glycoprotein binds sialic acid-terminating moieties on the host cell plasma membrane (1, 2), is triggered by endosomal low pH to undergo irreversible conformational changes that cause membrane fusion (3), and contributes to virus assembly and morphology (4, 5). Genetic differences between strains or due to mutations that alter the fundamental properties of the HA protein can alter virus replication, tropism, host range, and pathogenicity (reviewed in references 6 and 7). For example, isolates containing an HA protein with a single Arg residue in its cleavage site are restricted to replication in the lungs, where extracellular proteases are available, while isolates encoding the furin protease consensus sequence Arg-X-(Lys/Arg)-Arg^v (where "v" is the site of cleavage) undergo cleavage maturation intracellularly in the secretory pathway, expanding tropism and increasing pathogenicity in chickens and mammals (8, 9). Mutations in the HA receptor-binding pocket that switch preferential binding from α2,3- to α2,6-linked sialic acid are necessary for avian influenza viruses to adapt to humans (10 – 12). Two recent studies showed that a mutation that increases HA stability (by decreasing the pH threshold at which the protein is activated) is necessary for the gain-of-function airborne transmissibility of H5-subtype viruses in ferrets (13, 14). A stable HA protein was also found to be necessary for the airborne transmission of influenza A/H1N1/2009 virus in ferrets and for pandemic potential in humans (15). Numerous mutations that alter HA stability are located in the conserved stalk region (16), which is a target for antiviral drugs and broadly immunogenic vaccines (17). Susceptibility to stalk-based antivirals and vaccines can be altered by changes in HA stability (18, 19). In view of these facts, an understanding of how HA stability regulates IAV replication and pathogenesis bears upon the emergence of pandemic influenza and the development of HA-based control efforts.

During IAV entry into cells, receptor binding at the plasma membrane induces virion internalization (20) via clathrin-dependent or -independent endocytosis (21, 22). Within 5 min, internalized virions are exposed to a pH of 6.0 to 6.5 in early endosomes, and over a period of 30 to 40 min or more they are exposed to pHs of 5.0 to 5.5 in late endosomes and 4.6 to 5.0 in lysosomes (23, 24). At a threshold pH, HA trimers are triggered to undergo irreversible conformational changes that cause membrane fusion (reviewed in reference 3). The threshold pH for HA activation (and subsequent inactivation) ranges from approximately 4.8 to 6.2 (16, 25, 26). In general, avian IAVs have less stable HA proteins with higher activation pH values and human-adapted IAVs have more stable HA proteins. Less stable avian-origin HA proteins can enable virus replication in macrophages (27) and allow virus entry into early endosomes, thereby avoiding virion interactions with the IFITM2 and IFITM3 proteins that stimulate interferon (IFN)-induced antiviral activity (28).

Dozens of mutations throughout the HA trimer have been identified to alter the HA activation pH, and many of these mutations are associated with adaptations (reviewed in references 16, 29, and 30). Destabilizing mutations enhance virus replication in the presence of ammonium chloride and high concentrations of amantadine, which raise the endosomal pH (31 – 34), and have been associated with adaptation of egg-grown X-31 virus in Madin-Darby canine kidney (MDCK) and Madin-Darby bovine kidney (MDBK) cells (35). The replication of A/Puerto Rico/8/34 (H1N1) virus, which has an HA activation pH of 5.0 to 5.1 (26), is enhanced in Vero cells by a destabilizing mutation that increases the activation pH by 0.2 pH unit (36). NS1-deleted vaccine candidates with HA activation pH values of 5.5 to 5.8 have infectivities and immunogenicities higher than those of related variants triggered at pH 6.0 to 6.3 (37, 38). In H5 viruses with relatively unstable HA proteins, stabilizing mutations have been shown to increase upper respiratory tract replication in mice and ferrets (39 – 41) and to enable gain-of-function transmissibility in ferrets (13, 14); however, these mutations decrease the replication, virulence, and transmissibility in avian species (42 – 44).

Adaptations of IAVs to mice suggest that there is an optimal HA activation pH range of approximately 5.4 to 5.8 for replication in the lungs. In the context of 20th century seasonal H1N1 and H3N2 viruses that are activated at pH 5.2 to 5.3, adaptation to mouse lungs yielded destabilization mutants that elevated the HA activation pH to 5.6 to 5.8 (45 – 47). However, the adapted viruses also exhibited changes in receptor binding. For H5N1 and A/2009/H1N1 viruses containing point mutations that did not alter receptor-binding specificity or avidity, HA proteins that were activated at pH 5.4 to 5.5 boosted replication in mouse lungs and increased the pathogenicity compared to the pathogenicity of those that were activated at pH 5.9 to 6.3 (15, 40).

The mechanisms by which HA stability regulates influenza virus replication, pathogenesis, transmissibility, and host range are unclear. As viruses containing unstable HA proteins (with an activation pH of >5.8) have less environmental stability (42), we hypothesized that a relatively stable HA protein is required for maximum in vivo infectivity and replication to avoid extracellular inactivation in the respiratory tract (16). To test this hypothesis, we studied infection with A/Tennessee/1-560/2009 (H1N1), a 2009 pandemic virus, and two viruses with HA stability-altering mutations (Y17H and R106K). The wild-type (WT) HA protein is activated at pH 5.5, whereas a Y17H mutation in the HA1 fusion peptide pocket increases the activation pH to 6.0 and an R106K mutation in the HA2 coiled-coil core decreases the activation pH to 5.3 (15). Neither mutation altered HA protein expression, cleavage, maturation, receptor-binding avidity, or receptor-binding specificity. Both mutant viruses exhibited replication kinetics similar to those of the WT virus in MDCK cells when inoculated at a multiplicity of infection (MOI) of 0.01 PFU/cell; however, the Y17H virus had reduced replication and was less lethal than the WT virus in mice (15). The objective of this study was to use a mouse model to determine the mechanism by which HA stability regulates A/H1N1/2009 replication and pathogenicity.

(This article was submitted to an online preprint archive [48].)

RESULTS

Y17H virus is attenuated for infectivity, replication, and virulence in mice.

The HA protein of the A/Tennessee/1-560/2009 (H1N1) WT virus was previously shown to be activated for membrane fusion, or in the absence of target cells inactivated by low-pH buffer, at a midpoint pH of 5.5. HA stalk mutations HA1-Y17H and HA2-R106K altered the HA stability to pH 6.0 and 5.3, respectively, yet these mutated proteins retained similar expression levels, cleavage, and receptor-binding specificities (15, 49). To investigate the mechanisms by which HA stability alters infectivity and pathogenicity, we inoculated groups of DBA/2 mice intranasally with various doses of WT, Y17H, and R106K viruses generated by reverse genetics (r.g.) approaches. The mouse 50% infectious dose (MID₅₀) of these viruses decreased with decreasing HA activation pH (Table 1); thus, increased HA stability was associated with increased infectivity. WT virus had a mouse 50% lethal dose (MLD₅₀) value of 11,000 PFU, the R106K mutation increased the MLD₅₀ to 20,100 PFU, and 80% of Y17H virus-infected mice survived infection with 375,000 PFU, the highest dose tested (Table 1). At equivalent doses, the WT and R106K viruses induced similar weight loss, whereas the Y17H virus caused substantially less weight loss (Fig. 1A to C). For example, at a dose of 750 PFU, mice in the WT and R106K virus-infected groups exhibited approximately 10% weight loss, whereas mice in the Y17H virus-infected group lost only approximately 2% of their weight. We euthanized additional groups of mice at 2 and 5 days postinfection (dpi) and measured the viral loads. The WT and R106K viruses yielded similar titers (P > 0.05) at all inoculated doses except for 750 PFU in the trachea at 2 dpi (Fig. 1D to I). As infection with the R106K virus yielded viral loads and weight loss similar to those observed with the WT virus, the R106K virus was excluded from subsequent mechanistic studies. The titers of Y17H virus were only 10% of those of WT virus for many of the inoculation doses used (Fig. 1D to I). The maximal nasal titer of WT virus at 2 dpi was approximately 10⁵ 50% tissue culture infective doses (TCID₅₀)/ml, whereas that of Y17H virus was lower by a factor of approximately 100 (Fig. 1D). At 2 dpi with a 750-PFU dose, WT virus reached its maximal lung titer of approximately 10⁶ TCID₅₀/ml, whereas the corresponding lung titer of Y17H virus was lower by a factor of approximately 200. Overall, these studies showed that the HA-stabilizing mutation R106K had little effect on pathogenicity, whereas the HA-destabilizing mutation Y17H was highly attenuating.

TABLE 1.

Infectious and lethal dose values in mice compared to HA activation pH values

Virus	Activation pHa	MID50 (PFU)b	MLD50 (PFU)c
Y17H	6.0	48	>375,000
WT	5.5	4.7	11,000
R106K	5.3	0.68	20,100

^aHA activation pH values were measured by a syncytium formation assay and pH of inactivation of virus infectivity assay, both of which were in numerical agreement.

^bMID₅₀, mouse 50% infectious dose value, calculated by the method of Reed and Muench (79).

^cMLD₅₀, mouse 50% lethal dose value, calculated by the method of Reed and Muench (79). For the Y17H virus-infected group, 80% of mice survived infection after inoculation with a dose of 375,000 PFU, the highest dose tested.

FIG 1.

Weight loss and tissue viral titers in mice. (A to C) Changes in starting weight of DBA/2 mice after inoculation with 750 (A), 7,500 (B), or 75,000 (C) PFU of virus. Values are the combined mean (±SD) from two independent experiments with 10 mice total. (D to I) Nasal, tracheal, and lung tissue titers at 2 dpi (D to F) or 5 dpi (G to I). The inoculated doses were 75, 750, 7,500, 75,000, or 375,000 PFU. Y17H virus was not inoculated at a dose of 75 PFU, and WT and R106K viruses were not inoculated at a dose of 375,000 PFU. Values are the mean (±SD) for three mice. The key in panel A shows the symbols used in all panels. For panels D to I, the statistical significance was determined by Student's t test, comparing each set of results to those for WT virus at the equivalent dose. *, P < 0.05.

Reference and normalized inoculation doses of WT and Y17H viruses.

For subsequent in vivo studies, we selected 750 PFU as the reference dose. This dose yielded a 10-fold difference in weight loss and 10- to 100-fold differences in lung viral titers between WT and mutant viruses (Fig. 1), and it had previously resulted in differences in histopathology, cellular infiltration, and cytokine/chemokine expression (15). To normalize the viral titers and weight loss for subsequent studies of host responses, we considered two additional groups. A 75-PFU inoculum of WT virus resulted in an average lung viral titer that was lower by a factor of approximately 10 at 2 dpi but 10-fold higher at 5 and 7 dpi (Fig. 2A) than the corresponding titer obtained with a 10-fold higher 750-PFU inoculum of Y17H virus at those time points. With respect to weight loss, a 75-PFU dose of WT virus yielded a maximum weight loss of approximately 9%, whereas a 750-PFU dose of Y17H virus caused significantly less weight loss (P < 0.05 on days 7 to 10) (Fig. 2C). Thus, WT virus at a reduced dose of 75 PFU caused substantially more weight loss than Y17H virus at a dose of 750 PFU. We next inoculated mice with a 500-fold higher dose (375,000 PFU) of Y17H virus and compared the results to those obtained in mice inoculated with 750 PFU of WT virus. These two groups of mice exhibited similar weight loss (P > 0.05) (Fig. 2D). Compared to the lung titers achieved with a 750-PFU dose of WT virus, a 375,000-PFU dose of Y17H virus resulted in similar lung titers at 2, 7, and 9 dpi (P > 0.05) and an approximately 1-log higher lung titer at 5 dpi (P = 0.0114) (Fig. 2B). Overall, the data show that increasing the dose of Y17H virus to 375,000 PFU results in weight loss and lung viral titers (except at 5 dpi) similar to those seen with a dose of 750 PFU of WT virus. Thus, in addition to the groups of mice infected with 750 PFU of WT or Y17H virus, we included a group infected with 375,000 PFU of Y17H virus in our subsequent experiments.

FIG 2.

Results for two pairs of groups inoculated with different virus doses with the aim of normalizing the lung viral loads and weight changes. (A and C) Mice were inoculated with 75 PFU of WT virus or 750 PFU of Y17H virus. (B and D) Mice were inoculated with 375,000 PFU of Y17H virus or 750 PFU of WT virus. (A, B) Lung tissue titers at 2, 5, 7, and 9 days postinfection (dpi) for groups of DBA/2 mice. Values are the mean (±SD) for three to five replicates. (C, D) Changes in starting weight. Values are the combined mean (±SD) from two experiments with a total of 10 mice. For panels A and B, P values were determined at a given time point by comparing the results for WT and Y17H virus-infected mice by Student's t test. *, P < 0.05; **, P < 0.01.

Y17H virus retains WT virus-like infectivity at the extracellular pH of murine lungs.

One mechanism by which the HA-destabilizing Y17H mutation could attenuate virus growth in vivo is by extracellular inactivation if the respiratory tract is mildly acidic (16). To measure the extracellular pH in murine lungs, groups of mice were inoculated intranasally with WT virus (750 PFU), Y17H virus (750 or 375,000 PFU), or phosphate-buffered saline (PBS). In PBS-treated control mice, the average lung pH was 7.04 (Fig. 3A). The average lung pH in infected mice at 2 and 5 dpi was increased to approximately 7.3 to 7.4 (P < 0.02 or less compared to the results for the PBS-treated controls) for all groups except mice infected for 5 days with 750 PFU of Y17H virus, in which the lung pH recovered to 7.04. By 35 dpi, the lung pH in all groups had recovered to approximately 7.0 to 7.1. Overall, this experiment showed that the lungs of DBA/2 mice remain at a nearly neutral pH whether they are uninfected, are infected with a pH1N1 virus, or have recovered from an infection.

FIG 3.

Murine extracellular lung pH and virion environmental stability. (A) Lung pH values in DBA/2 mice. Mice were inoculated intranasally with PBS, 750 PFU of WT virus, 750 PFU of Y17H virus, or 375,000 PFU of Y17H virus. After 2, 5, or 35 days of infection, extracellular pH values were measured in the lungs. Reported values are means (n = 6) with standard deviations. (B, C) Environmental stabilities of WT and Y17H viruses. Virus stocks were thawed and diluted to approximately 10⁶ TCID₅₀/ml in PBS with the pH adjusted to 7.0 (B) or 6.4 (C) and then incubated for the specified times at 37°C before the residual infectivity was measured by the TCID₅₀ assay. Groups included those infected with WT virus or Y17H virus. For panels B and C, the values are reported as the means (n = 3) with standard deviations. Statistical significance tests compared the results to those for PBS-treated (A) or WT virus-infected (B, C) mice. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001.

To measure the persistence of virion infectivity (environmental stability) at neutral pH, we incubated aliquots of WT and Y17H viruses at 37°C in buffer of pH 7.0 or 6.4 and measured the residual infectivity as a function of time by the TCID₅₀ assay (Fig. 3B and C). At pH 7.0, the titers of both viruses dropped from approximately 5.7 to 1.7 log₁₀ TCID₅₀/ml over a period of 72 h. When virus aliquots were incubated at pH 6.4, the titer of Y17H virus decayed at an accelerated rate compared to that of WT virus. The 90% reduction time (Rt value) is the time required for a 90% (1 log₁₀) decrease in the viral titer (50). At pH 7.0, the Rt values for the WT and Y17H viruses were similar (21.7 ± 1.5 h and 20.4 ± 2.1 h, respectively); however, at pH 6.4, the Rt values for the WT and Y17H viruses were 26.6 ± 1.8 h and 7.4 ± 0.6 h, respectively. In summary, Y17H virions were inactivated at a rate 3.6 times faster than that for WT virions when exposed to pH 6.4 but were inactivated at a rate similar to that for WT virions when exposed to pH 7.0. As the measured pH of the murine lungs did not drop below pH 7.0, a value at which the environmental stability of Y17H virus showed little to no accelerated decay, we concluded that the attenuation of Y17H virus in murine lungs was not due to extracellular inactivation.

Replication in cultured cells.

We found previously that Y17H virus exhibited replication kinetics like those of WT virus in MDCK, A549, and differentiated normal human bronchial epithelial (NHBE) cells at 37°C when inoculated at a low MOI (0.01 PFU/cell) (15). Here, we performed additional growth-curve experiments at MOI values of 0.01 and 3 PFU/cell. In MDCK cells inoculated at an MOI of 0.01 PFU/cell and incubated at 33 or 39.5°C, Y17H virus grew in a manner similar to that for WT virus (Fig. 4A and B). Y17H and WT viruses also had similar replication kinetics (at low and high MOI) in LA-4 murine lung adenoma epithelial cells (Fig. 4C and F). In differentiated murine nasal epithelial cells (mNECs) and murine tracheal epithelial cells (mTECs) grown at an air-liquid interface, the replication of Y17H virus was delayed by approximately 12 to 24 h (Fig. 4D and E). Additionally, the maximal growth of Y17H virus in mTECs was less than 10% of that of WT virus. In RAW 264.7 mouse macrophage cells, WT virus did not produce infectious virions, whereas Y17H virus replicated (Fig. 4I). For completeness, we performed high-MOI growth-curve experiments using MDCK and A549 cells. Surprisingly, the data showed that the growth of Y17H virus was enhanced relative to that of WT virus (Fig. 4G and H). In summary, Y17H virus replicated faster than WT virus when inoculated at an MOI of 3 PFU/cell in MDCK, A549, and RAW 264.7 cells; Y17H virus exhibited growth similar to that of WT virus in MDCK, A549, and LA-4 cells inoculated at an MOI of 0.01 PFU/cell; but the growth of Y17H virus was delayed in comparison to that of WT virus in mNECs and mTECs.

FIG 4.

Virus growth in cell cultures. (A, B) Virus replication (MOI, 0.01 PFU/cell) in MDCK cells in culture at 33°C (A) or 39.5°C (B). (C) Virus replication (MOI, 0.01 PFU/cell) in LA-4 mouse lung adenoma cells in culture at 37°C. (D, E) Virus replication (MOI, 0.01 PFU/cell) in immortalized murine epithelial cells in culture at an air-liquid interface at 37°C. The differentiated nasal mNECs (D) and tracheal mTECs (E) were grown on transwell plates at an air-liquid interface. (F) Virus replication (MOI, 3 PFU/cell) in LA-4 mouse lung adenoma cells in culture at 37°C. (G) Virus replication (MOI, 3 PFU/cell) in MDCK cells in culture at 37°C. (H) Virus replication (MOI, 3 PFU/cell) in A549 cells in culture at 37°C. (I) Virus replication (MOI, 0.01 PFU/cell) in RAW 264.7 murine macrophage cells in culture at 37°C. Virus titers are given as the mean (±SD) for three or four replicates of infection with WT virus and Y17H virus. Statistical significance tests compared the results to those for WT virus. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Pathogenicity at equivalent and normalized doses.

To investigate how the HA-destabilizing Y17H mutation affected in vivo spread and pathogenesis, we inoculated groups of five mice intranasally with PBS, WT virus (750 PFU), or Y17H virus (750 or 375,000 PFU) and recovered the lungs at 2 and 5 dpi. Compared to the characteristics of infection with WT virus, infection with Y17H virus at the same dose (750 PFU) resulted in significantly less spread of infection, growth of virus, severity of infection, cellular infiltration, and expression of interleukin-6 (IL-6), gamma interferon (IFN-γ)-induced protein 10, granulocyte colony-stimulating factor (G-CSF), IL-1α, and IFN-γ expression in the lungs (Fig. 5 and 6). Thus, Y17H virus was highly attenuated in vivo, recapitulating in vitro replication in mNECs and mTECs, the only differentiated cells used for the growth-curve experiments. We also wished to compare the results of a 750-PFU infection with WT virus and a 375,000-PFU infection with Y17H virus, because in a previous experiment, these two virus exposures induced similar weight loss and the viruses grew to similar lung titers (Fig. 2). By 5 dpi, infection in the 375,000-PFU Y17H virus-infected group covered 14% less area, the virus grew to titers approximately 1 log higher, and the infection induced approximately 19% more cellular infiltration and was scored as approximately 17% more severe than the infection in the 750-PFU WT virus-infected group. However, the 750-PFU WT virus-infected and 375,000-PFU Y17H virus-infected groups did not differ significantly (P > 0.05) at 2 dpi or 5 dpi with respect to virus growth, spread of infection, the severity of infection, cellular infiltration, and expression of the proinflammatory cytokines IL-6, CXCL10, G-CSF, IL-1α, and IFN-γ in the lungs (Fig. 5 and 6). Expression of a total of 25 cytokines and chemokines was measured by the Milliplex assay. The remaining 20 cytokines for which the results are not shown in Fig. 5 (GM-CSF, IL-1β, IL-2, IL-4, IL-5, IL-7, IL-9, IL-10, IL-12P40, IL-12P70, IL-13, IL-15, IL-17, keratinocyte-derived chemokine, monocyte chemotactic protein 1, macrophage inflammatory protein 1α [MIP-1α], MIP-1β, MIP-2, RANTES, and tumor necrosis factor alpha [TNF-α]) had levels similar to the background levels, consistent with previous reports that pH1N1/2009 induces relatively weak inflammatory cytokine responses compared to those induced by other strains of influenza A viruses (51). Overall, inoculation of mice with Y17H virus at a dose 500-fold higher than that used for WT virus induced an infection that was sufficiently robust to overcome the attenuation of Y17H virus. This suggested that normalizing Y17H virus infection with a dose 500-fold higher than the WT virus dose could be used to investigate the differential triggering and/or counteracting of antiviral host responses by the two viruses.

FIG 5.

Infection, histopathology, infiltration, and cytokine induction in lungs. Groups of mice were infected with 750 PFU of Y17H virus, 750 PFU of WT virus, or 375,000 PFU of Y17H virus. After 2 and 5 days of infection, the lungs were recovered to assess the spread of infection, histopathology, cellular infiltration in BALF, and cytokine expression in BALF. (A) Lung homogenates were plaque titrated by TCID₅₀ assay on MDCK cells. (B, C) Lungs were fixed, sectioned, and analyzed microscopically to determine the percentage of cells infected (B) and the severity of the infection (C). (D) Cellular infiltration in BALF was measured by flow cytometry. (E to I) Cytokine and chemokine expression in BALF was measured by the Milliplex assay. Displayed values are the mean (±SD) for five mice per group. Multiple comparisons to the 750-PFU WT virus-infected group at each time point were analyzed by two-way ANOVA (time and group), followed by Tukey’s post hoc test. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001; ns, not significant (P > 0.05).

FIG 6.

Microscopic analyses of lung sections. Mice were inoculated intranasally with PBS, WT virus (750 PFU), or Y17H virus (750 or 375,000 PFU). At 5 dpi, the lungs were collected and sections were stained and photographed. (A) The extent of active infection (red) and cleared infection (yellow) was visually traced by a histopathologist blind to the treatment group identity. (B) Hematoxylin and eosin (H&E) staining of lung sections. Magnifications, ×20. (C) Immunohistochemical staining (IHC) of infected lungs, using a polyclonal antibody against the NP protein.

Impact of HA stability on antiviral responses.

We examined next if the Y17H virus induces host gene expression different from that induced by WT virus. We inoculated groups of three mice each with PBS, WT virus (750 PFU), or Y17H virus (750 or 375,000 PFU). At 2 and 5 dpi, we collected the lungs and processed the samples for Affymetrix microarray analysis. At an equivalent dose of 750 PFU, infection with Y17H virus resulted in 21% fewer genes (127 versus 161) being upregulated than infection with WT virus after 2 days and 54% fewer genes (142 versus 308) being upregulated after 5 days (Fig. 7A). The upregulation of fewer genes by infection with Y17H virus was consistent with the results of prior experiments showing less virus growth and cytokine expression with the attenuated virus (Fig. 5). Despite mice in the 375,000-PFU Y17H virus-infected group showing virus spread, infiltration, and cytokine expression in the lungs that were similar to those in the 750-PFU WT virus-infected group (Fig. 5), inoculation with Y17H virus at the normalized 375,000-PFU dose induced the upregulation of 80% (289 versus 161) and 104% (629 versus 308) more genes at 2 dpi and 5 dpi, respectively (Fig. 7B and C). We also analyzed the number of shared and unique genes upregulated after infection with WT and Y17H viruses. At 5 dpi, 90% (308/344) of the genes upregulated in the 750-PFU WT virus-infected group were also upregulated in the 375,000-PFU Y17H virus-infected group (Fig. 7C). In contrast, only 44% (275/629) of the genes upregulated in the 375,000-PFU Y17H virus-infected group were also upregulated in the 750-PFU WT virus-infected group. A total of 386 genes were uniquely upregulated in the Y17H virus-infected groups at 5 dpi, whereas only 29 were uniquely upregulated in the WT virus-infected group. Gene enrichment analysis showed that the genes most differentially upregulated were involved in the IFN pathway, the inflammatory response, and/or the antiviral state (Fig. 7D). The transcription profiles of the genes involved in the IFN pathway were largely similar for the 750-PFU WT virus-infected and 375,000-PFU Y17H virus-infected groups (Fig. 7E).

FIG 7.

Differential gene expression in DBA/2 mouse lungs after infection. Mice were inoculated intranasally with PBS, WT virus (750 PFU), or Y17H virus (750 or 375,000 PFU). At 2 and 5 dpi, lungs were collected for analysis of RNA expression by Affymetrix microarray analysis (n = 3). (A) Numbers of upregulated genes (log₂ ratio of virus-infected group/PBS-treated group > 0.5). (B, C) Venn diagrams of upregulated genes. (D) Gene Ontology (GO) enrichment analysis in the biological process category for differentially expressed genes common to all three groups (P < 0.05). (E) Heat map of genes involved in the interferon pathways. The log₂ ratio of the mean for the virus-infected group to the mean for the PBS-treated group is shown. pos., positive; reg., regulation; neg., negative; LPS, lipopolysaccharide.

To investigate further the effect of the HA-Y17H mutation on type I IFN signaling, we performed Western blot analyses by using antibodies to STAT1, phosphorylated STAT1 (pSTAT1), influenza virus M1 (as a control for infectivity), and β-actin (Fig. 8A). Comparing the results for the 375,000-PFU Y17H virus-infected group and 750-PFU WT virus-infected group at 5 dpi, we found that pSTAT1 expression in the Y17H virus-infected group was 2-fold higher than that in the WT virus-infected group (Fig. 8D). At 2 dpi, the 750-PFU WT virus-infected group had higher viral titers and higher levels of M1 expression in the lungs than the 375,000-PFU Y17H virus-infected group (Fig. 2B and 8D). Lung viral titers and M1 expression for the 750-PFU WT virus-infected group decreased from days 2 to 5, but those for Y17H virus-infected group increased from days 2 to 5, such that the lung viral titers and the levels of M1 expression in the 375,000-PFU Y17H virus-infected group exceeded those in the 750-PFU WT virus-infected group on day 5. In summary, Y17H virus induced the upregulation of more host genes and a higher level of pSTAT1 response in murine lungs than did WT virus when mice were inoculated with Y17H at a dose 500-fold higher than that used for WT virus. The higher dose was necessary to boost the growth of Y17H virus to a level comparable to that of WT virus.

FIG 8.

Protein expression by Western blotting. (A) Mice were inoculated intranasally with PBS, WT virus (75 or 750 PFU), or Y17H virus (750 or 375,000 PFU). At 2 and 5 dpi, lungs were collected for Western blot analysis of protein expression (n = 3). (B, C) Bone marrow-derived macrophages (B) or dendritic cells (C) were infected at an MOI of 3 PFU/cell. After 6, 12, or 24 h, protein was harvested for Western blot analysis. SDS-PAGE gels were labeled with antibodies specific for pSTAT1, STAT1, A/H1N1 influenza virus M1, or β-actin. (D to F) Quantitation of relative protein expression of pSTAT1 (left) and M1 (right) by ImageJ software. In panel D, the results for the 750-PFU WT virus-infected and 375,000-PFU Y17H virus-infected groups from DBA/2 mouse lungs were normalized to the level of WT virus expression at 5 dpi (D5). In panels E and F, the results for WT and Y17H virus infection (at equivalent MOIs in macrophages and dendritic cells) were normalized to the level of WT expression at 12 h postinfection. Values for WT virus and Y17H virus represent the mean (±SD) for a single gel in panel A and the average for two gels in panels B and C. Significant differences compared to the results for the WT were determined by multiple Student’s t tests. *, P < 0.5; **, P < 0.01.

Type I IFN responses are enhanced in Y17H virus-infected DCs.

Dendritic cells (DCs) and macrophages recognize IAV and other pathogens via a subset of intracellular receptors, such as Toll-like receptors 7/8 (TLR7/8) and retinoic acid-inducible gene I (RIG-I) helicase, and they release high levels of antiviral mediators, such as type I IFN and other inflammatory cytokines/chemokines (52). This process leads to more immune cells being recruited to the site of the infection and to the initiation of an adaptive immune response. As the previous analyses of type I IFN responses in murine lungs required a 500-fold higher dose of Y17H virus to normalize the level of infection, we wished to investigate these responses to infection at an equal MOI in the immune cells most likely to display potential differences.

We derived DCs and macrophages from the bone marrow of DBA/2J mice by using standard protocols (53). After 6 to 7 days of differentiation, we infected cells with WT or Y17H virus at an MOI of 3 PFU/cell. In macrophages, M1 expression was more than 3-fold higher in Y17H virus-infected cells than in WT virus-infected cells at 12 to 24 h postinfection (Fig. 8E), consistent with macrophages supporting productive infection by Y17H virus but not WT virus (Fig. 4I). However, pSTAT1 expression in infected macrophages infected with Y17H and WT viruses was similar at between 6 and 24 h postinfection (Fig. 8E). In DCs, pSTAT1 responses were the highest at 12 h postinfection, at which time pSTAT1 expression in Y17H virus-infected cells was 3.3-fold higher than that in WT virus-infected cells (Fig. 8F). Thus, pSTAT1 expression in DCs, but not in macrophages, was elevated by infection with Y17H virus compared to that by infection with WT virus.

We also measured the induction of inflammatory mediators in bone marrow-derived DCs (BMDCs) and macrophages infected with WT and Y17H viruses at an MOI of 3 PFU/cell. Infection with either virus resulted in the increased expression of IFN-β, CXCL10, IL-6, TNF-α, and IL-1β at 24 h postinfection (Fig. 9A to E). Compared to the levels of induction by WT virus, Y17H virus induced 2.2-fold higher levels of expression of IFN-β (P < 0.0001), 2.3-fold higher levels of expression of CXCL10 (P < 0.001), and 1.5-fold higher levels of expression of IL-1β (P < 0.05) in DCs but not in macrophages. In DCs, NP was expressed at a level 1.13-fold higher during Y17H virus infection than during WT virus infection (P = 0.0313) at 4 h postinfection (Fig. 9I). At 6 and 12 h postinfection, DCs infected with Y17H virus also had higher levels of viral RNA (vRNA), cRNA, and mRNA transcripts than did DCs infected with WT virus (Fig. 9F to H). Overall, Y17H virus infected DCs, transcribed viral genes, and translated viral proteins at higher levels than WT virus early in infection, resulting in the increased induction of protective responses, including elevated expression of pSTAT1 and the inflammatory mediators IFN-β and CXCL10.

FIG 9.

Host responses in bone marrow-derived dendritic cells (DC) and macrophages (M) from DBA/2J mice. Cells were infected with WT and Y17H viruses (at an MOI of 3 PFU/cell) or left untreated (Mock). Three experiments were performed independently, with each having two or three replicates. (A to E) Cytokine levels in the supernatants were measured by ELISA at 24 h postinfection. (F to H) vRNA, cRNA, and mRNA levels in dendritic cells were measured by real-time RT-PCR. (I) Dendritic cells were stained with an anti-NP antibody at 4 h after infection and imaged in a plate reader. Statistical analysis was performed using one-way ANOVA, followed by Tukey’s post hoc tests (A to E), or Student’s t tests (F to I) separately for each time point. *, P < 0.05; **, P < 0.01; ***, P < 0.001; ****, P < 0.0001. h.p.i., hours postinfection; OD, optical density.

To investigate if the WT and Y17H viruses differentially induced RIG-I-like receptors (RLR) and Toll-like receptor (TLR) signaling, we infected bone marrow-derived DCs from WT mice, mice lacking the mitochondrial antiviral-signaling protein (MAVS), and mice lacking the two major adaptor proteins MyD88 and TRIF. After 24 h of infection, we collected supernatants and measured the induction of the cytokines IFN-β, IL-6, and TNF-α (Fig. 10). Consistent with previous results, IFN-β expression was 32% higher (P = 0.0069) in Y17H virus-infected bone marrow-derived dendritic cells (BMDCs) from WT mice than in BMDCs from WT mice infected with WT virus (Fig. 10A). MAVS is the adaptor protein required for retinoic acid-inducible gene I (RIG-I)-mediated sensing of IAV RNA (54, 55). Bone marrow-derived DCs from mice lacking MAVS were unable to produce IFN-β after infection with either virus, but these cells did induce the expression of IL-6 and TNF-α (Fig. 10). The effects were similar for infection with both WT virus and Y17H virus. These data confirm that the RIG-I-mediated sensing of IAV is required for interferon signaling in DCs and that both viruses stimulate the RIG-I/MAVS pathway to trigger the type I IFN response in DCs. In bone marrow-derived DCs from mice lacking MyD88 and TRIF, IFN-β expression was reduced compared to IFN-β expression in DCs from WT mice. In these cells, expression of IL-6 and TNF-α was eliminated, showing that the MyD88/TRIF signaling pathway is required to induce IL-6 and TNF-α inflammatory cytokines. Overall, the results suggest that WT and Y17H viruses are sensed by the same pathway in DCs and that the increased type I IFN response observed during Y17H virus infection is due to increased replication by the mutant virus.

FIG 10.

Cytokine production in BMDCs from WT and knockout C57BL/6 mice. BMDCs from WT, MAVS^−/−, and Myd88^−/− TRIF^−/− mice were infected with WT and Y17H viruses at an MOI of 3 PFU/cell or with PBS. Supernatants were collected at 24 h after infection so that the levels of expression of IFN-β (A), IL-6 (B), and TNF-α (C) could be measured by ELISA. The displayed values are the mean (±SD) for 3 mice. Statistical analysis was performed using one-way ANOVA, followed by Tukey’s post hoc tests. **, P = 0.0069; ns, not significant (P > 0.4).

DISCUSSION

We have investigated several potential mechanisms by which a destabilizing mutation in the HA protein might attenuate the replication of A/H1N1/2009 virus, reduce its infectivity (as demonstrated by an increased 50% infectious dose), and diminish its pathogenesis in mice. The extracellular pH in the lungs of DBA/2J mice ranged from 6.9 to 7.5, a nearly neutral pH range in which changes in HA stability did not affect the persistence of A/H1N1/2009 infectivity. The HA stability altered the virus tropism in cell culture. A destabilizing HA mutation boosted virus replication in MDCK, A549, and RAW 264.7 cells inoculated at an MOI of 3 PFU/cell but reduced multistep replication in murine airway mNEC and mTEC cultures. In DBA/2J mice, a destabilizing HA mutation substantially reduced virus replication, virus spread in the lungs, the severity of infection, and cellular infiltration into bronchoalveolar lavage fluid (BALF). Increasing the inoculation dose of the destabilized HA-Y17H mutant virus 500-fold compared to the dose of WT virus boosted replication, pulmonary spread, the severity of infection, and cellular infiltration to levels comparable to those seen with WT virus. Normalized infection with the destabilized mutant triggered the upregulation of more host genes and increased type I IFN responses and cytokine expression in DBA/2 mouse lungs and bone marrow-derived DCs. Collectively, the data show that HA destabilization attenuates virulence in mice not by decreasing the extracellular virion stability but by enhancing early viral transcription and translation in DCs, which results in stronger activation of antiviral responses, including the type I IFN response.

Extracellular environments with an acidic pH (pH < 6.6), warmer temperatures, and/or a higher salinity decrease the environmental persistence of IAVs (50, 56). Environmental stability can also be decreased by destabilizing HA mutations (42), i.e., those that increase the pH at which the HA protein becomes activated to undergo irreversible structural changes (16, 30). In the present study, the A/H1N1/2009 virus containing a destabilizing HA1-Y17H mutation (activation pH, 6.0) had reduced environmental stability over 72 h at pH 6.4 but not at pH 7.0. From this, we hypothesize that such a destabilizing HA mutation will attenuate influenza virus replication in acidic environments but not at a nearly neutral pH. The respiratory pH in the lungs of DBA/2J mice ranged from 6.9 to 7.5. In previous work, the nasal pH and tracheal pH in mice were reported to range from 6.5 to 7.0 and from 6.7 to 7.5, respectively (57 – 59). Therefore, we conclude that extracellular inactivation is unlikely to be the cause of A/H1N1/2009 HA-Y17H attenuation in mice. The HA-Y17H mutation may decrease the environmental persistence of A/H1N1/2009 in the human nasal cavity, the pH of which is reported to range from 5.3 to 7.0 in healthy adults and from 5.5 to 6.7 in healthy children (60 – 63).

The HA-Y17H mutation studied here increases the HA activation pH of A/H1N1/2009 to pH 6.0, most likely enabling quicker virion entry into early endosomes. The wild-type HA protein is activated at pH 5.5, most likely delaying virion entry until the progression into late endosomes. The earlier entry by the Y17H mutant virus is consistent with the accelerated replication kinetics of the virus in MDCK, A549, RAW 264.7, and bone marrow-derived dendritic cells when inoculated at an MOI of 3 PFU/cell. Entry into early endosomes via a relatively unstable HA protein, such as HA-Y17H, may also enable the virus to avoid functional interactions with the IFITM2 and IFITM3 proteins in late endosomes. In general, the optimal growth of H1N1 in mice may be favored by the presence of a moderately stable HA protein (activation pH, approximately 5.5) that avoids both the early entry and more robust interferon responses of destabilized HA (pH 6.0) and interferon-induced transmembrane proteins in late endosomes by stabilized HA (pH 5.3), as has been demonstrated recently (64). For viruses containing the six internal genes of A/H1N1/PR8 and two surface genes generated by reverse genetics approaches, less stable HA proteins from avian influenza viruses were shown to confer increased infectivity in MDCK cells, and the viruses were less sensitive to antiviral IFN responses (28). If these effects contribute to the increased pathogenicity of H7 and H9 viruses in humans, as has been suggested (28), the effect may be lost for seasonal IAVs. On the background of A/H1N1/2009, any early replication advantage by the less stable HA-Y17H protein after high-MOI (3-PFU/cell) inoculation was not observed at later time points in MDCK cells, mNECs, and mTECs inoculated at a low MOI (0.01 PFU/cell). In previous studies, destabilizing HA mutations have also resulted in reduced virus growth in DBA/2J mice, ferrets, and human airway epithelium (15, 64). Although the impact of HA destabilization on virulence in humans is unknown, the HA-Y17H mutation was previously associated with a loss of pandemic potential in A/H1N1/2009 (15).

Wild-type A/Tennessee/1-560/2009 (H1N1) virus with an activation pH of 5.5 was replication incompetent in macrophages, whereas the destabilized Y17H mutant with an activation pH of 6.0 produced infectious virions. Similarly, the closely related wild-type virus A/California/04/2009 (H1N1) (activation pH, 5.4) was replication incompetent in RAW 264.7 cells, whereas a 7 + 1 reassortant encoding a less stable H5 HA protein (pH 5.9) produced infectious virions (27). In the present study, gain-of-function replication in macrophages did not correlate with altered antiviral IFN responses or with increased virus replication and pathogenicity in mice. Thus, expanded tropism does not necessarily result in gain-of-function pathogenicity.

Dendritic cells often elicit immunity by capturing and phagocytosing influenza virus-infected airway cells (65, 66). Additionally, IAVs have been shown to directly infect primary human myeloid DCs, human blood monocyte-derived DCs, mouse splenic DCs, and mouse bone marrow-derived DCs (51, 67 – 75), although these infections do not necessarily produce infectious virions (75). A/H1N1/09 wild-type virus was previously shown to induce weak cytokine responses in DCs but to be highly sensitive to the antiviral activities of IFNs (51). In the present study, the destabilizing HA-Y17H mutation increased early infection and type I IFN responses in mouse bone marrow-derived DCs, but not in macrophages, and amplified IFN responses in mice were associated with attenuated virus replication and pathogenesis. Thus, the present work shows that the destabilization of the A/H1N1/2009 HA protein to an activation pH of 6.0, a relatively high value typically associated with avian influenza viruses, helps elicit stronger type I IFN responses by accelerating infection in DCs.

We have discovered that HA stability influences IAV replication and pathogenesis in mice by altering the virus infection kinetics and type I IFN responses in DCs. This provides a link between a fundamental property of the viral surface glycoprotein, namely, acid-induced activation of the HA protein, and the ability of the virus to evade antiviral host responses. Further studies are needed to assess the impact of HA stability on the infection kinetics in DCs of other IAV strains and the resulting antiviral responses, especially for highly pathogenic avian influenza viruses that have HA proteins with relatively high activation pH values that are similar to the activation pH for the Y17H virus studied here. Future investigations should also investigate the impact of HA stability on IFN responses in pigs, ferrets, and humans. Overall, we hypothesize that narrow ranges of HA activation pH support robust replication by a particular IAV in a specific host.

MATERIALS AND METHODS

Cells.

The following cell lines were obtained from the American Type Culture Collection (ATCC): Madin-Darby canine kidney (MDCK; ATCC CCL-34), murine lung adenoma LA-4 (ATCC CCL-196), human lung epithelial carcinoma A549 (ATCC CCL-185), and murine macrophage Abelson murine leukemia virus-transformed RAW 264.7 (ATCC TIB-71). MDCK cells were propagated in culture in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 5% fetal bovine serum (FBS) and 1% penicillin-streptomycin (Pen-Strep) at 37°C in 5% CO₂. Both LA-4 and A549 cells were propagated in culture in Kaign’s modification of Ham’s F-12 medium with l-glutamine (F-12K) supplemented with 10% FBS and 1% Pen-Strep at 37°C in 5% CO₂. RAW 264.7 cells were propagated in culture in RPMI 1640 medium supplemented with 10% FBS and 1% Pen-Strep at 37°C in 5% CO₂. To prepare and propagate murine nasal epithelial cells (mNECs) and murine tracheal epithelial cells (mTECs), the nasal turbinate and trachea were collected and cells were isolated using collagenase (200 U/ml) and dispase (2.5 U/ml) in Ham’s F-12 medium containing Pen-Strep or 0.15% filter-sterilized pronase diluted in Ham’s F-12 medium containing Pen-Strep, respectively. After appropriate incubation, the cells were pelleted by centrifugation, washed, and resuspended in ammonium chloride potassium (ACK) lysis buffer to remove the red blood cells. Following resuspension in medium, the cells were pelleted by centrifugation and plated at 3.3 × 10⁴ cells per 0.32-cm² transwell insert precoated with rat tail collagen and grown in DMEM–Ham’s F-12 medium supplemented with 200 mM GlutaMAX, 7.5% sodium bicarbonate, insulin (10 μg/ml), transferrin (5 μg/ml), cholera toxin (100 ng/ml), murine epidermal growth factor (25 ng/ml), bovine pituitary extract (30 μg/ml), 5% fetal bovine serum, retinoic acid (5 × 10⁻¹⁰ M), and Pen-Strep until confluence. Once the cells were confluent, the apical medium was removed and the basal medium was replaced with ALI medium (DMEM–Ham’s F-12 medium containing 200 mM GlutaMAX, 7.5% sodium bicarbonate, Pen-Strep, 2% NuSerum, retinoic acid). Cells were used after full differentiation (transepithelial/transendothelial electrical resistance > 1,000), which was typically after 1 week for mNECs and 1 to 3 weeks for mTECs. Bone marrow-derived macrophages (BMDMs) and bone marrow-derived dendritic cells (BMDCs) were prepared as described previously (53). Briefly, primary BMDMs were grown for 6 days in IMDM (catalog number 12440-053; Gibco) supplemented with 1% nonessential amino acids (Gibco), 10% FBS (catalog number S1620; lot number 221C16; BioWest), 30% medium conditioned by L929 mouse fibroblasts, and 1% penicillin-streptomycin (Gibco). Primary BMDCs were grown in RPMI medium (catalog number 10-040-CV; Cellgro; Corning) supplemented with 10% FBS (catalog number S1620; lot number 221C16; BioWest), 1% nonessential amino acids (Gibco), 1% sodium pyruvate (Gibco), 1% penicillin-streptomycin (Gibco), 0.1% β-mercaptoethanol, and 20 ng·ml⁻¹ granulocyte-macrophage colony-stimulating factor for 7 days. BMDMs and BMDCs were seeded in antibiotic-free medium at concentrations of 1 × 10⁶ and 3 × 10⁶ cells, respectively, onto 12-well plates.

Viruses.

Influenza A/Tennessee/1-560/2009 (H1N1) WT, Y17H, and R106K viruses were previously generated by reverse genetics, sequenced, and characterized (15, 49). The Y17H mutation is located in HA1 at residue 17 in the H3 numbering, which is at position 7 from the N-terminal cleavage site (76), and at residue 24 in the full-length H1 sequence. The R106K mutation is at HA2 position 106 in both the H1 and H3 numbering.

Mice.

Six-week-old female DBA/2J mice were obtained from The Jackson Laboratory. MAVS^−/− and Myd88^−/− TRIF^−/− mice were described previously (77, 78). All mice were backcrossed to the mice of the C57BL/6 mouse background and bred in-house.

Animal ethics statement.

Animal experiments were conducted in an animal biosafety level 2+ facility, using procedures compliant with NIH requirements and with the Animal Welfare Act. The St. Jude Animal Care and Use Committee reviewed and approved the animal experiments (protocol numbers 459 and 464).

Animal experiments.

Seven-week-old DBA/2J mice were anesthetized with isoflurane and inoculated intranasally with virus in 30 μl of PBS. Clinical symptoms (including weight loss and mortality) were monitored daily. To enable the collection of respiratory tissues, groups of mice were euthanized with CO₂ after 2, 5, 7, and 9 days of infection. Bronchoalveolar lavage fluid (BALF) was collected by washing the lungs three times with 0.5 ml of PBS containing 2 mM EDTA (for a total volume of 1.5 ml) via a catheter. The BALF was centrifuged, and then the total number of infiltrating cells was counted by flow cytometry and the supernatants were stored at −80°C. Lung tissues were collected for use in the various assays described below.

Virus titrations and environmental persistence assays.

At 2, 5, 7, and 9 days postinfection (dpi), mice were euthanized and nasal turbinates, tracheae, and lungs were collected. After the tissues had been homogenized using a Qiagen Tissue Lyser II homogenizer (30/s frequency for 30 s, operated twice), the homogenates were clarified by centrifugation at 8,000 rpm for 15 min at 4°C in an Eppendorf 5417R centrifuge. The infectivity of the supernatants was titrated by 50% tissue culture infective dose (TCID₅₀) assays. One day before the virus infection, 3 × 10⁴ to 5 × 10⁴ MDCK cells in culture medium were dispensed into each well of a 96-well tissue culture plate and incubated at 37°C in 5% CO₂. On the day of infection, the tissue supernatants were serially diluted 10-fold in virus infection medium supplemented with 1 μg/ml l-1-tosylamide-2-phenylethyl chloromethyl ketone (TPCK)–trypsin (Worthington Biochemical Corporation, Lakewood, NJ). The plates were rinsed twice with PBS, and 200 μl of diluted tissue supernatant sample was added to each well containing MDCK cells. The plates were incubated at 37°C in 5% CO₂ for 3 days, then 50 μl of the culture from each well was transferred to a new 96-well round-bottom plate, and then 50 μl of 0.5% turkey red blood cells in PBS was added to each well. The plates were incubated for 30 min at room temperature. The highest dilution that tested positive for virus was recorded. TCID₅₀ values were calculated using the method of Reed and Muench (79).

For plaque assays, viruses were harvested from the culture or collected from homogenized tissue supernatant. At 1 day before infection, 6-well tissue culture plates were seeded with 0.3 × 10⁶ to 0.5 × 10⁶ MDCK cells per well in 3 ml culture medium. On the day of virus infection, virus was serially diluted 10-fold in PBS or DMEM. MDCK cells were rinsed twice with PBS buffer, and 0.5 ml containing diluted viruses was loaded into assigned wells containing MDCK cells. The plates were incubated at 37°C in a 5% CO₂ incubator for 1 h and gently rocked every 15 min during the incubation. Meanwhile, 0.7% agarose gel in water was melted completely in a microwave oven and incubated in a 45°C water bath. Virus infection medium (VIM; 2×) with TPCK-treated trypsin was prewarmed in a 37°C water bath. After infection, the virus-containing medium was aspirated and the plate was washed three times with PBS. Equal volumes of 0.7% agarose gel and 2× virus infection medium were mixed well and immediately loaded onto plates (3 ml/well). The plates were kept in a biological safety cabinet at room temperature for 15 min and then moved to an incubator. After 3 days of incubation at 37°C in a 5% CO₂ incubator, the plates were fixed, stained with 1% crystal violet in 10% formalin buffer for 30 min, and rinsed with tap water, and plaques were counted to calculate the virus titer in terms of the number of PFU.

To measure virus persistence, thawed virus stocks were diluted to approximately 10⁶ TCID₅₀/ml in pH-adjusted PBS (pH 6.4 or 7.0). The viruses were incubated at 37°C and collected at the time points specified above to measure the residual infectivity by the TCID₅₀ assay. The time required for a 10-fold decrease in virus titer (reduction time [Rt]) was calculated by least-squares linear regression of values above the limit of detection, using GraphPad Prism (version 7.03) software.

Lung pH measurements.

A pH 1 micro-fiber-optic pH transmitter and a pH microsensor (a needle-type fiber-optic microsensor) were purchased from PreSens Precision Sensing GmbH (Regensburg, Germany). The microsensor was calibrated manually according to the instructions in the manufacturer’s manual before experimental measurements. In brief, the calibration details listed on the final inspection protocol of the microsensors, which were provided together with each sensor, were typed into the pH 1-View, Manual Calibration under Calibration. DBA/2J mice were inoculated intranasally with 30 μl of PBS or the virus inoculum (750 or 375,000 PFU). At 2 and 5 dpi, the mice were euthanized, their chests were immediately opened, and the pH microsensor was inserted into their lungs. In each case, the pH was recorded 30 s after probe insertion.

Virus replication curves.

For low-MOI virus growth-curve experiments, MDCK cells, LA-4 cells, mNECs, and mTECs were infected with virus at an MOI of 0.01 PFU/cell and incubated at a temperature of 33°C, 37°C, or 39.5°C in 5% CO₂ for 1 h, after which they were rinsed three times with PBS. Infection medium was then added, and the cells were incubated at a temperature of 33°C, 37°C, or 39.5°C in 5% CO₂. Supernatants were collected at the time points indicated above, stored at −80°C, and titrated in MDCK cells by the TCID₅₀ assay. For high-MOI growth-curve experiments, MDCK, A549, and RAW 264.7 cells were infected with virus at an MOI of 3 PFU/cell for 1 h at 37°C, after which the infected cells were rinsed once with saline buffer (pH 2.2) to inactivate extracellular virus that did not enter cells and then twice with PBS to reneutralize. The cells were incubated at 37°C in 5% CO₂ until harvested at the time points specified above. Samples were stored at −80°C until they could be titrated in MDCK cells by the TCID₅₀ assay.

Histopathology.

The lungs from control and virus-infected mice were fixed via intratracheal infusion of and then immersion in 10% buffered formalin solution. Tissues were embedded in paraffin and sectioned, and immunohistochemical labeling of viral antigen was completed by using a primary goat polyclonal antibody (United States Biological, Swampscott, MA) against influenza A virus USSR (H1N1) at 1:1,000 and a secondary biotinylated donkey anti-goat immunoglobulin antibody (catalog number sc-2042; Santa Cruz Biotechnology, Santa Cruz, CA) at 1:200 on tissue sections subjected to antigen retrieval for 30 min at 98°C. The extent of virus spread was quantified by first capturing digital images of whole-lung sections stained for viral antigen by using an Aperio ScanScope XT slide scanner (Aperio Technologies, Vista, CA) and then manually outlining the fields with alveolar areas containing virus antigen-positive pneumocytes highlighted in red (defined as active infection). Those lesioned areas containing minimal or no antigen-positive cells were highlighted in yellow (defined as inactive infection). The percentage of each lung field with infection/lesions was calculated using Aperio ImageScope software. In addition, a pathologist blind to the treatment group identity evaluated pulmonary lesions in hematoxylin-eosin (H&E)-stained histologic sections and assigned scores based on their severity and extent as follows: 0, no lesions; 1, minimal, focal to multifocal, and barely detectable lesions; 15, mild, multifocal, and small but conspicuous lesions; 40, moderate, multifocal, and prominent lesions; 80, marked, multifocal coalescing, and lobar lesions; and 100, severe, diffuse lesions with extensive disruption of normal architecture and function.

Chemokine and cytokine assays.

IFN-β was measured by use of an enzyme-linked immunosorbent assay (ELISA) kit (BioLegend). Other chemokines and cytokines were measured by the Milliplex mouse magnetic bead assay (Millipore) according to the manufacturer’s instructions.

Microarray analyses.

Pieces of lung tissue were preserved in RNAlater (Ambion), and lung tissues were homogenized using a Qiagen Tissue Lyser II apparatus. RNA was extracted from lung tissue samples with an RNeasy kit (Qiagen), and microarray analyses were performed by the Hartwell Center for Bioinformatics and Biotechnology at St. Jude Children’s Research Hospital. Briefly, total RNA (100 ng) was converted to biotin-labeled cRNA with an Ambion wild-type expression kit (Life Technologies) and was hybridized to a Mouse Gene (version 2.0) ST GeneChip microarray (Affymetrix). After the microarray was stained and washed, the array signals were normalized and transformed into log₂ transcript expression values with the multiarray average algorithm (Partek Genomics Suite, version 6.6). Differential expression was defined by applying a difference in expression of 0.5-fold (log₂ signal) between conditions. Lists of differentially expressed transcripts were analyzed for functional enrichment with the Gene Ontology Resource at geneontology.org (80, 81). Visualization of the gene expression profile (heat map creation) was done using TIBCO Spotfire software (version 10.1.0).

Real-time quantitative RT-PCR.

RNAlater-preserved tissues were homogenized, and total RNA was extracted using an RNeasy minikit (Qiagen). The levels of virus-specific vRNA, cRNA, and mRNA, as well as GAPDH (glyceraldehyde-3-phosphate dehydrogenase) mRNA, were analyzed by semiquantitative real-time PCR analysis on an Applied Biosystems 7500 Fast real-time PCR system (Applied Biosystems, Waltham, MA) as previously described (82). Briefly, total RNA was reverse transcribed using virus-specific reverse transcription (RT) primers or oligo(dT) primers and a SuperScript III first-strand synthesis system (Invitrogen, Carlsbad, CA). Real-time quantitative PCR was performed on an Applied Biosystems 7500 system with QuantiTect SYBR green PCR master mix (Qiagen) and the appropriate primers. After normalization of the results to the GAPDH expression levels, the fold change ratio of expression in virus-infected samples to that in control samples was calculated for each gene by using the ΔΔC_T threshold cycle (C_T) method and was expressed as 2^−ΔΔCT.

Immunoblot analysis.

Lung tissues or bone marrow-derived DCs were lysed in radioimmunoprecipitation assay lysis buffer and sample loading buffer. Proteins were separated by electrophoresis on 8% to 12% polyacrylamide gels. After the proteins were electrotransferred onto polyvinylidene difluoride membranes, nonspecific binding was blocked by incubating the membranes with 5% skim milk or bovine serum albumin. The membranes were then incubated with primary antibody (anti-pSTAT1 Y701 [1:100 dilution; clone D4A7; Cell Signaling Technologies], anti-STAT1 p84/p91 [1:1,000 dilution; clone E23; Santa Cruz Biotechnology], anti-M1 [1:1,000 dilution; catalog number GTX127356; GeneTex], or anti-actin [1:10,000 dilution; catalog number SC-47778; Santa Cruz Biotechnology]) and then with secondary antibody (horseradish peroxidase-conjugated anti-rabbit IgG [catalog number 7074S; Cell Signaling Technologies] or anti-mouse IgG [catalog number 7076S; Cell Signaling Technologies], both at a 1:5,000 dilution).

Immunofluorescence microscopy.

Bone marrow-derived DCs were infected as described above and seeded in a 96-well plate. At 4 h postinfection, cells were fixed in PBS–4% paraformaldehyde for 15 min. They were then permeabilized with PBS–0.5% Triton X-100 for 10 min, blocked in PBS–10% FBS for 1 h, and stained with an antibody to influenza A virus NP (antibody MAB8257; Millipore) overnight at 4°C. After the plate had been washed, it was incubated with anti-mouse IgG-Alexa Fluor 488 secondary antibody (Invitrogen) in 10% FBS in PBS for 1 h. The plate was then imaged in a fluorescence multiplate reader (Synergy 2 multimode microplate reader; BioTek).

Statistical analyses.

GraphPad Prism (version 7.03) software was used for data analysis. Statistical significance for two groups was determined by Student’s t test (two-tailed). Statistical significance for three or more groups was determined by one-way analysis of variance (ANOVA), followed by Tukey’s multiple-comparison test. We considered P values of less than 0.05 to indicate statistical significance.

Data availability.

The data from this study will be made fully available and without restriction.