Highly pathogenic avian influenza A(H5N1) viruses have circulated continuously in Bangladesh since 2007, and active surveillance has detected viral evolution driven by mutation and reassortment. Recently, three genetically distinct A(H5N1) reassortant viruses were detected in live poultry markets in Bangladesh. Currently, we cannot assign pandemic risk by only sequencing viruses; it must be conducted empirically. We found that the H5Nx highly pathogenic avian influenza viruses exhibited high virulence in mice and chickens, and one virus had limited capacity to transmit between ferrets, a property considered consistent with a higher zoonotic risk.
KEYWORDS: Bangladesh, influenza viruses, live poultry markets, influenza
ABSTRACT
The genesis of novel influenza viruses through reassortment poses a continuing risk to public health. This is of particular concern in Bangladesh, where highly pathogenic avian influenza viruses of the A(H5N1) subtype are endemic and cocirculate with other influenza viruses. Active surveillance of avian influenza viruses in Bangladeshi live poultry markets detected three A(H5) genotypes, designated H5N1-R1, H5N1-R2, and H5N2-R3, that arose from reassortment of A(H5N1) clade 2.3.2.1a viruses. The H5N1-R1 and H5N1-R2 viruses contained HA, NA, and M genes from the A(H5N1) clade 2.3.2.1a viruses and PB2, PB1, PA, NP, and NS genes from other Eurasian influenza viruses. H5N2-R3 viruses contained the HA gene from circulating A(H5N1) clade 2.3.2.1a viruses, NA and M genes from concurrently circulating A(H9N2) influenza viruses, and PB2, PB1, PA, NP, and NS genes from other Eurasian influenza viruses. Representative viruses of all three genotypes and a parental clade 2.3.2.1a strain (H5N1-R0) infected and replicated in mice without prior adaptation; the H5N2-R3 virus replicated to the highest titers in the lung. All viruses efficiently infected and killed chickens. All viruses replicated in inoculated ferrets, but no airborne transmission was detected, and only H5N2-R3 showed limited direct-contact transmission. Our findings demonstrate that although the A(H5N1) viruses circulating in Bangladesh have the capacity to infect and replicate in mammals, they show very limited capacity for transmission. However, reassortment does generate viruses of distinct phenotypes.
IMPORTANCE Highly pathogenic avian influenza A(H5N1) viruses have circulated continuously in Bangladesh since 2007, and active surveillance has detected viral evolution driven by mutation and reassortment. Recently, three genetically distinct A(H5N1) reassortant viruses were detected in live poultry markets in Bangladesh. Currently, we cannot assign pandemic risk by only sequencing viruses; it must be conducted empirically. We found that the H5Nx highly pathogenic avian influenza viruses exhibited high virulence in mice and chickens, and one virus had limited capacity to transmit between ferrets, a property considered consistent with a higher zoonotic risk.
INTRODUCTION
Influenza A viruses continue to present a challenge to human and animal health. Wild aquatic birds are the primary natural reservoir for influenza A viruses, and they play a major role in their global distribution and the emergence of novel viruses (1). Avian influenza viruses (AIVs) can be classified into low-pathogenicity avian influenza viruses (LPAIVs) or highly pathogenic avian influenza viruses (HPAIVs) according to their virulence in chickens. Both LPAIVs and HPAIVs sporadically cross the species barrier from birds to mammals, although they have not yet acquired the capacity for sustained transmission between humans without reassortment with human-adapted viruses (2–5). Reassortment is an important mechanism in the generation of novel influenza viruses with unique phenotypes and has been a hallmark of the evolution of HPAIV A/goose/Guangdong/1/96 (H5N1)-lineage (Gs/GD lineage) viruses, from their emergence in China through their spread to Europe, Africa, and the North American continent via migratory birds (6–8).
HPAIVs of the Gs/GD lineage first emerged in China in 1996 and subsequently evolved into multiple distinct hemagglutinin (HA) clades over the ensuing years, with dissemination of clade 2.3.2.1a viruses to Bangladesh and to Bhutan and India (9–11). The first human case of A(H5N1) was detected in Bangladesh in January 2008, with total cases now numbering eight, one of which was fatal (12). A(H5N1) viruses are geographically widespread in poultry across Bangladesh, and coinfections with A(H9N2) viruses of the G1 genetic lineage and other LPAIVs are common (13, 14); subsequent reassortment between these viruses has been consistently detected. In 2012, reassortant clade 2.3.2.1a Gs/GD lineage viruses possessing the M gene or PB1 gene from an A(H9N2) virus were detected (11, 15). In 2015, another novel genotype of A(H5N1) virus (designated H5N1-R1) emerged; this virus had HA, M, and NA genes of circulating Bangladeshi A(H5N1) viruses and five genes from Eurasian-lineage LPAIVs (16). Continued surveillance identified another new reassortant, H5N1-R2, in 2017 that was derived from H5N1-R1 with a PA gene from a Eurasian-lineage LPAIV (17). In 2018, we detected a transient reassortant distinct from H5N1-R2 viruses, in that it possessed NA and M genes from the Bangladeshi A(H9N2) viruses (17). While reassortment has increased viral diversity in Bangladesh, its influence on the relative zoonotic and pandemic risks of HPAIVs is unknown.
To better understand the evolution and emergence of these novel reassortants, we characterized the genetic properties of four H5Nx reassortment AIVs, determined their pathogenicity in a mouse model, and evaluated the pathogenicity and transmissibility of the H5Nx viruses in chickens and ferrets.
RESULTS
Genetic analysis of multiple subgroups of H5 HPAIVs.
Active surveillance for AIVs in live poultry markets (LPMs) in Bangladesh detected three reassortant influenza A(H5N1) clade 2.3.2.1a virus genotypes. Two of these genotypes, designated H5N1-R1 and H5N1-R2, contained genes (HA, NA, and M gene) from circulating A(H5N1) clade 2.3.2.1a viruses; all other viral genes clustered with those from Eurasian viruses. The third reassortant group, H5N2-R3, contained an HA gene from circulating A(H5N1) clade 2.3.2.1a viruses, NA and M genes from concurrently circulating A(H9N2) influenza viruses, and all other genes from Eurasian viruses (Fig. 1A).
FIG 1.
(A) Genome constellations representative of the multiple subgroups of clade 2.3.2.1a highly pathogenic (H5Nx) avian influenza A virus characterized in this study. The eight gene segments (from top to bottom) in each virus are polymerase basic 2, polymerase basic 1, polymerase acidic, hemagglutinin, nucleoprotein, neuraminidase, matrix, and nonstructural. (B) Phylogenetic trees of the HA gene. Phylogenetic analysis was done using the neighbor-joining algorithm with the Kimura two-parameter model. The reliability of phylogenetic inference at each branch node was estimated by the bootstrap method with 1,000 replications; evolutionary analyses were conducted in MEGA 7.
To better understand the genetic relationships of the H5 viruses, we compared the complete genomes of H5 AIVs that represented four genotypes: A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), and A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) (Fig. 1B). Based on the phylogenetic analysis and antigenic characteristics, the HA gene of the four genotypes belonged to clade 2.3.2.1a and clustered with other H5N1-subtype viruses isolated in Bangladesh (see Fig. S1 in the supplemental material).
A series of basic amino acids at the cleavage site of the HA protein (PQRERRRKR↓GLF) was present and similar in viruses of the four genotypes, which is consistent with observations of other HPAIVs. Q222 and G224 (H5 numbering) were present at the receptor-binding pockets of all viruses, suggesting that they preferentially bind to avian-like receptors. No virus had oseltamivir resistance markers E119, H275, R293, and/or N295 in NA (N1 numbering).
None of the viruses selected contained the human adaptation marker PB2 627K or 701N. However, a PB2 K702R mutation, which increases polymerase activity and enhances pathogenicity in mice, was present in A/duck/Bangladesh/26042/2015 H5N1 (subgroup R0). G309D and T339K mutations in the PB2 gene were detected in all viruses except A/duck/Bangladesh/26042/2015 H5N1 (subgroup R0). H5N1-R1, H5N1-R2, and H5N2-R3 possessed additional amino acid changes not present in the parental H5N1-R0 virus, including the PB2 K526R substitution, which enhances the effects of E627K on influenza virus replication (18).
The H5N1-R0, H5N1-R1, and H5N2-R3 viruses expressed a PB1-F2 protein consisting of 90 amino acids, and that of the H5N1-R2 virus consisted of 76 amino acids. Three genotypes (H5N1-R0, -R1, and -R2) were sensitive to M2 ion-channel blockers, and we found the mutation S31N in the H5N2-R3 virus. Previously, A(H5N1) viruses isolated from Bangladesh were characterized by a 5-amino-acid deletion at positions 80 to 84 of the NS1 protein. This deletion occurred in all viruses (Fig. 1B). Amino acid substitutions P42S and V149A in the NS1 protein were detected in all viruses, and the C-terminal motif ESEV was present in all viruses except that from the H5N2-R3 genotype, which contained EPEV.
Replication and pathogenesis of multiple subgroups of H5 HPAIVs in mice.
To assess the impact of reassortment on the biological characteristics of H5 HPAIVs, we first compared their virulence in mice. Viral pathogenicity in mice was determined by generating 50% mouse lethal dose (MLD50) values. The MLD50 values of the mice inoculated with A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), or A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) viruses were >106, 105.4, 104.4, and 103.5 LD50, respectively (Fig. 2).
FIG 2.
Pathogenicity and replication of the multiple subgroups of clade 2.3.2.1a highly pathogenic (H5Nx) avian influenza A viruses in mice. Groups of 5- to 6-week-old BALB/c mice (n = 5) were inoculated intranasally with the indicated doses (102, 103, 104, 105, or 106 EID50) of A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), or A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3). (A and B) Body weight loss (A) and survival (B) were evaluated daily for 14 days. Groups of mice (n = 3) that were infected with 106 EID50 were euthanized at day 3 or 5 postinfection. (C) Lung, nasal turbinate, heart, brain, and intestine were harvested, homogenized, and used to quantify viral titers by EID50 assay. Viral titers expressed as the log10 EID50/ml were plotted as the mean. Statistical analysis was performed using two-way ANOVA (*, P < 0.05; **, P < 0.01; ***, P < 0.001). A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0) is shown in red, A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1) is shown in blue, A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2) is shown in green, and A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) is shown in pink.
At a dose of 106, the H5N1-R0 virus caused 20% mortality, the H5N1-R1 virus caused 80% mortality, and the H5N1-R2 and H5N2-R3 viruses caused 100% mortality (Fig. 2B).
To assess the viruses’ replication kinetics in organs, we inoculated mice (n = 3) with 106 50% egg infectious doses (EID50) of virus and determined the viral replication in lungs, nasal turbinate, heart, brain, and intestines at 3 and 5 days postinfection (dpi). All of the viruses tested replicated in mouse lung (Fig. 2C). Virus titers in the lungs of mice infected with the H5N1-R2 or H5N2-R3 virus were 5.0 ± 0.4 log10 and 5.1 ± 0.4 log10, respectively, at 5 dpi, which were significantly higher than those in mice infected with the H5N1-R0 or H5N1-R1 virus. We also detected variable extrapulmonary dissemination of the viruses in other organs (Fig. 2C).
Histologic changes in mouse lung tissues.
We next studied the pathology in the respiratory tract of inoculated mice (Fig. 3). There were no notable lesions and only a few positive epithelial foci in the upper respiratory tract (URT) of mice infected with A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0) at 3 or 5 dpi. At 3 dpi, there were no notable lesions in the lower respiratory tract (LRT), and infection was limited to multifocal clusters of bronchiolar epithelium. Pulmonary lesions at 5 dpi consisted of small, clearly demarcated foci surrounding a few terminal bronchioles. Clusters of virus-positive bronchiolar epithelial cells were scattered widely in airways, but virus-positive cells and debris in alveoli were restricted to small peribronchiolar lesions.
FIG 3.
Pulmonary lesions and viral spread in the lungs of H5Nx-infected mice. Mice (n = 3) were infected intranasally with 106 EID50 of A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), or A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3). Mice were euthanized at 3 dpi (A) or 5 dpi (B). Lungs were harvested and fixed in 10% neutral buffered formalin and stained with hematoxylin and eosin (HE), subjected to IHC staining with anti-NP antiserum, and analyzed by histomorphometry. Magnifications: 60× (HE), 20× (IHC), and 2× (histomorphometry). For histomorphometry, total lung areas are outlined in green, and areas with antigen-positive cells are shown in red.
Scattered virus-positive epithelial cells were detected in the URT of mice infected with A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1) at 3 dpi but not at 5 dpi (Fig. 3). At 3 dpi, there were extensive areas of alveolar inflammation and damage. Virus distribution in the LRT was characterized by extensive infection of type II pneumocytes but only very rare infection of bronchiolar epithelium. By 5 dpi, prominent perivascular/peribronchiolar lymphoid aggregates were associated with extensive alveolar inflammation and damage. Small numbers of virus-positive type II pneumocytes were present at the margins of parenchymal lesions.
There were no notable lesions or virus-positive epithelial cells in the URT of any mice infected with A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2) at 3 dpi (Fig. 3). However, at 5 dpi, one of three mice had abundant virus-positive neurons in the olfactory neuroepithelium and within a mediastinal ganglion. At 3 dpi, there were no notable lesions in the LRT and only a few clusters of virus-positive bronchiolar epithelial cells and alveolar pneumocytes. The extent and severity of virus infection was highly variable in the mice at 5 dpi. One mouse had a single clearly demarcated alveolar focus and no infection of bronchiolar epithelium. In contrast, the other two had extensive infection of bronchiolar epithelium and alveolar pneumocytes; one mouse had an extensive infection of neurons in the olfactory neuroepithelium, olfactory bulb, and thoracic ganglion.
In the URT of mice infected with A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) virus, there were clusters of virus-infected respiratory epithelial cells at 3 and 5 dpi (Fig. 3). In the LRT at 3 dpi, virus distribution was characterized by extensive infection of bronchiolar epithelium extending into the peribronchiolar alveoli. By 5 dpi, the extent and severity of damage to bronchiolar epithelium was markedly increased, and there were extensive areas with denuded bronchiolar epithelium. Peribronchiolar/perivascular inflammation was widespread, and extensive coalescing areas of alveolar inflammation and damage contained abundant virus-positive macrophages and type II pneumocytes.
Pathogenicity and transmission of multiple genotypes of H5 HPAIVs in chickens.
We assessed the pathogenicity and transmission of the A(H5Nx) viruses in chickens. Groups of chickens were intranasally inoculated with 30 chicken LD50/0.5 ml of each virus and then were cohoused 1 day later with naive contact birds. Within 1 dpi, 2/3 of H5N1-R0-inoculated chickens and 3/3 of H5N1-R1-inoculated chickens died, compared to 1/3 and 1/3 for H5N1-R2- and -R3-infected birds, respectively. One bird from each of the H5N1-R2 and H5N2-R3 groups survived until the end of the experiment (Table 1).
TABLE 1.
Pathogenicity and transmission of H5Nx HPAI viruses in chickens
| Strain | Manifestations in no. D/S/totala chickens | Mean ± SD viral titer (log10 EID50/ml) on dayb
: |
|||||
|---|---|---|---|---|---|---|---|
| 3 |
5 |
7 |
|||||
| Oropharyngeal swabs | Cloacal swabs | Oropharyngeal swabs | Cloacal swabs | Oropharyngeal swabs | Cloacal swabs | ||
| A/duck/Bangladesh/26042/2015 (H5N1-R0) | |||||||
| Donor | 3/3/3 | 4 (1/1) | 3.5 (1/1) | 7.5 (1/1) | ND (1/1) | ||
| Contact | 3/3/3 | 1.87 ± 0.53 (2/3) | ND (0/3) | 6.5 (1/1) | 3.5 (1/1) | ||
| A/duck/Bangladesh/25683/2015 (H5N1-R1) | |||||||
| Donor | 3/3/3 | ||||||
| Contact | 0/0/3 | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) |
| A/duck/Bangladesh/33841/2017 (H5N1-R2) | |||||||
| Donor | 2/2/3 | 4.87 ± 2.2 (2/3) | 3 (1/3) | 7.25 (1/2) | ND (2/2) | ND (1/1) | ND (1/1) |
| Contact | 0/0/3 | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) | ND (3/3) |
| A/chicken/Bangladesh/34722/2018 (H5N2-R3) | |||||||
| Donor | 2/2/3 | 7.37 ± 0.17 (2/3) | 5.75 ± 0.7 (2/3) | ND (1/1) | ND (1/1) | ND (1/1) | ND (1/1) |
| Contact | 3/3/3 | 1.4 ± 0.14 (3/3) | ND (3/3) | 6.33 ± 0.52 (3/3) | 4.42 ± 0.76 (3/3) | ||
No. S/D/total shows the number of dead (D), number of sick (S), and total number of chickens from the observation period.
Numbers in parentheses are number of birds positive/total number of birds surviving. ND, not detected.
Oropharyngeal and cloacal swabs were collected on 1, 3, 5, and 7 dpi and titrated in eggs. As shown in Table 1, the H5N1-R0 virus was detected in cloacal swabs from one surviving chicken at 3 dpi (103.5/ml EID50) and from oropharyngeal swabs at 3 and 5 dpi (104 to 107.5/ml EID50). The H5N1-R2 virus was detected in cloacal swabs from inoculated chickens at 3 dpi (103/ml EID50) and from oropharyngeal swabs at 3 and 5 dpi (104.87 to 107.25/ml EID50). The H5N2-R3 virus was detected in cloacal (105.75/ml EID50) and oropharyngeal (107.37/ml EID50) swabs from inoculated chickens at 3 dpi.
To determine whether H5Nx viruses efficiently transmitted between chickens, we isolated viruses from oropharyngeal and cloacal swabs from contact birds (Table 1). H5N1-R0 and H5N2-R3 viruses transmitted to contact birds; H5N1-R1 and H5N1-R2 viruses did not. At 3 dpi, we detected the H5N1-R0 virus from oropharyngeal swabs only (101.87/ml EID50), while at 5 dpi, we detected it from cloacal swabs and oropharyngeal swabs (103.5/ml and 106.5/ml EID50, respectively). At 3 dpi, we detected the H5N2-R3 virus from oropharyngeal swabs only (101.4/ml EID50), while at 5 dpi, we detected the virus from cloacal swabs and oropharyngeal swabs (104.42/ml and 106.33/ml EID50, respectively). All H5N1-R0 and H5N2-R3 contact birds succumbed to infection.
Overall, viral titers were higher in oropharyngeal swabs than in cloacal swabs in all chickens infected with H5Nx viruses by inoculation or after contact with infected birds, but no virus was detected in either group after 5 dpi.
Pathogenicity and transmission of multiple genotypes of H5 HPAIVs in ferrets.
To determine the pathogenicity and transmissibility of the H5Nx viruses in ferrets, we used nine influenza-seronegative ferrets for each virus. Three ferrets were intranasally inoculated with 106/ml EID50 of virus. Twenty-four hours later, three inoculated ferrets were individually paired and cohoused with direct-contact ferrets to examine contact transmission; to examine airborne transmission, an airborne-contact ferret was housed in a wireframe cage adjacent to that of the infected ferret. To monitor virus shedding, we collected nasal washes from all inoculated, direct-contact, and airborne-transmission groups and titrated for virus in 9-day-old embryonated eggs. Inoculated ferrets shed virus in nasal washes that peaked at levels ranging from 102.75 to 106.7 EID50/ml (Fig. 4). In the H5N1-R0 and H5N1-R2 groups, virus was shed in nasal washes from inoculated ferrets at 2, 4, and 6 dpi, while in the H5N1-R1 group, the virus was detected at 2 and 4 dpi. The highest titers were detected in H5N2-R3-infected animals, and one animal still had detectable virus in nasal wash at 8 dpi (Fig. 4). Another H5N2-R3-infected ferret required euthanasia at day 8 dpi due to the development of neurologic symptoms (Fig. 5). To determine whether H5Nx viruses were efficiently transmitted between ferrets, we collected nasal washes from direct-contact and airborne-transmission groups (Fig. 4). None of the contact ferrets from the H5N1-R0, H5N1-R1, or H5N1-R2 groups had detectable virus in nasal wash specimens. One contact ferret from the H5N2-R3 group shed virus in nasal washes at 4, 6, 8, and 10 days postinfection (102.5 to 104.7/ml EID50) (Fig. 4).
FIG 4.
Pathogenicity and transmission of the multiple subgroups of clade 2.3.2.1a highly pathogenic (H5Nx) avian influenza A viruses in ferrets. Groups of 3 ferrets each were intranasally inoculated with 106 EID50 of each virus shown. Each ferret was paired with an individual naive ferret at 24 h postinoculation. Viral titers in nasal washes from individual inoculated ferrets (D; red), direct-contact ferrets (DC; blue), and aerosol-contact ferrets (AC; green) on the days postinoculation or postcontact were determined and presented as the log10 EID50/ml.
FIG 5.
Body weight loss (left) and changes in body temperatures (right) of individually inoculated (D), direct-contact (DC), or aerosol-contact (AC) ferrets up to 12 days postinoculation or postcontact. Ferrets were inoculated intranasally with 106 EID50 of A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), or A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3), and the body weight and temperature of the animals were monitored.
To evaluate the tissue tropism of H5N1-R2 and H5N2-R3 in ferrets, we collected lung, trachea, and nasal turbinates at 4 dpi and measured the level of infectious virus present in each tissue. We found that the two viruses were detected in ferret nasal turbinate and trachea, but only H5N2-R3 was detected in the lung (Fig. 6).
FIG 6.
Viral replication of multiple subgroups of clade 2.3.2.1a highly pathogenic (H5Nx) avian influenza A viruses in ferret tissues. Three ferrets were inoculated with 106 EID50 of A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2) or A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) virus and then humanely euthanized on day 4 postinoculation to collect tissues for viral titration in eggs by EID50 assay. Each bar represents a ferret, and the detection limit is shown by a dotted line.
Our findings show that the four reassortment viruses have not acquired the ability to transmit efficiently among ferrets; however, they do show a difference in replicative ability and tissue tropism.
DISCUSSION
For almost a decade, A(H5N1) HPAIVs have been endemic to Bangladesh, where multiple introductions of highly pathogenic A(H5N1) have been reported. The continued circulation of these viruses with other AIVs in poultry has provided an opportunity for reassortment and subsequent generation of novel genotypes. In this study, we characterized the genetic properties of four A(H5Nx) reassortant AIVs, determined the pathogenicity of these viruses in a mouse model, and evaluated their pathogenicity and transmissibility in chickens and ferrets.
The HA of the ancestral clade 2.3.2 virus evolved into clade 2.3.2.1 and further diversified into 2.3.2.1a, b, and c. Clade 2.3.2.1a viruses have become dominant in Bangladesh (16, 17, 19), and our recent studies have detected the emergence of HPAI A(H5N1) reassortant viruses in LPMs. The internal genes of these reassortant viruses were most closely related to those of Eurasian-lineage LPAIVs, which have been isolated from free-range ducks and LPMs in Bangladesh. Antigenic analyses showed a lack of significant antigenic diversity among the four reassortant H5Nx genotypes (17). Correspondingly, sequence analyses showed that the amino acids across the HA antigenic sites were conserved and that the four viruses all carried multiple basic amino acid residues at the cleavage site of the HA gene, which is a characteristic of HPAIVs. All four viruses were highly pathogenic in chickens. Molecular analysis revealed significant differences between these viruses, including an 80- to 84-residue deletion in the NS gene, which was present in all genotypes except H5N2-R3. This deletion is associated with increased pathogenicity in chickens (20). Globally, the major amantadine-resistant cluster of IAVs had S31N mutation (21, 22). We identified H5N2-R3 viruses with the S31N mutation, which was reported previously in clade 2.3.2.1a viruses.
We used BALB/c mice to evaluate the disease potential of H5Nx viruses in mammalian models. All AIVs used in the current study efficiently infected and replicated in mice. However, H5N1-R2 and H5N2-R3 viruses were more pathogenic than H5N1-R0 and H5N1-R1 viruses. The major difference between H5N1-R1 and H5N1-R2 viruses was the origin of the PA gene. The PA protein has been implicated in the pathogenesis of influenza virus (23, 24), and we found nine mutations in the PA gene of H5N1-R2 that were not found in the PA gene of H5N1-R1 (data not shown). Further studies are needed to explore the potential roles of any of these mutations. It should also be noted that although the other gene segments of H5N1-R1 and H5N1-R2 were of the same phylogenetic lineage, variations were also seen in these. The number of substitutions in the proteins expressed from these more conserved segments ranged from 0 (M2) to 11 (PB2). Thus, while the genotype-defining segments had the most differences, these more subtle changes may also have been critical for our observed phenotypic properties.
Several factors that affect AIV adaptation and virulence in mice have been identified. Avian influenza viruses become more virulent in mice when passaged in these animals. In our study, we noted a dramatic difference in the pathological consequences of H5N1-R2 infection in mouse lungs at day 3 compared to day 5. H5N1-R2-infected mice had no damage to the epithelial cells in the URT and only a few clusters of virus-positive bronchiolar epithelial cells and alveolar pneumocytes in the LRT at 3 dpi. At 5 dpi, mice exhibited extensive infection of both the bronchiolar epithelium and alveolar pneumocytes. Sequence data showed that the virus in the lung at 5 dpi acquired the E627K substitution within the PB2 gene and other mutations in the HA gene (data not shown), which were associated with the pathological changes of H5N1-R2 during replication in mice.
In our ferret study, all viruses caused relatively mild disease. These results are consistent with previous findings (25). All infected ferrets had detectable virus in nasal washes. Transmission to direct-contact or respiratory droplet-contact animals was not detected for three viruses (H5N1-R0, H5N1-R1, and H5N1-R2), like most A(H5N1) HPAIVs. We detected limited transmission via direct contact with the H5N2-R3 virus; the contact-infected ferret had nasal wash titers on days 4, 6, 8, and 10 postcontact. In addition to the distinct gene segments, the H5N2-R3 virus contained the HA substitution T156E. Previous studies have shown that the substitution T156A increases virus binding to both α2,6-SA and α2,3-SA, which may have contributed to the transmission event (26).
Previous studies have reported the diversity of AIVs in poultry in LPMs, free-range ducks, and wild birds in Bangladesh. We previously detected several subtypes of LPAIVs, and some gene segments from the LPAIVs isolated from LPMs were closely related to those of LPAIVs isolated from ducks on free-range farms and wild birds (27). In addition, there are genetic similarities between internal gene segments of free-range duck viruses and some genes of the HPAI A(H5N8) clade 2.3.4.4 origin (8), suggesting their potential role in the genesis of the HPAIV A(H5N8). LPAIVs and HPAIVs were isolated from a wide range of hosts in Bangladesh (13, 27–30), and, recently, the HPAIV A(H5N6) of clade 2.3.4.4 was detected in LPMs in Bangladesh (31). This diverse range facilitates the spread of the viruses and can contribute to high levels of genetic diversity.
The results described here highlight that the cocirculation of different viruses in the same susceptible host can give rise to novel reassortants that have variable phenotypes. As an example, reassortment between A(H5N1) and A(H9N2) viruses led to a virus with increased pathogenicity in mice and some, albeit limited and not significantly different, transmission by direct contact in ferrets. Thus, this study emphasizes the importance of active and continuous surveillance of AIVs in both wild birds and poultry to assess viral evolution and detect the generation of reassortant viruses that have the potential for virulence in mammalian hosts. It also highlights the continued need for empirical determination of biologic characteristics for continued risk assessment efforts.
MATERIALS AND METHODS
Ethics statement.
All mouse and ferret experiments were approved and performed according to the guidelines set by the Animal Care and Use Committee of St. Jude Children’s Research Hospital in an enhanced animal biosafety level 3 containment facility.
Viruses.
The H5Nx HPAIVs used in this study were isolated in Bangladesh during routine surveillance. Virus stocks of HPAI A/duck/Bangladesh/26042/2015 H5N1 (subgroup H5N1-R0), A/duck/Bangladesh/25683/2015 H5N1 (subgroup H5N1-R1), A/duck/Bangladesh/33841/2017 H5N1 (subgroup H5N1-R2), and A/chicken/Bangladesh/34722/2018 H5N2 (subgroup H5N2-R3) viruses were propagated and titrated for EID50 in the allantoic cavities of 10-day-old embryonated chicken eggs at 35°C for 48 h. Allantoic fluid was pooled from multiple eggs, clarified by centrifugation, and frozen in aliquots at −80°C. Virus titers were determined by injecting 100 μl of serial 10-fold dilutions of virus into the allantoic cavities of 10-day-old embryonated chicken eggs and then calculating the EID50 according to the method of Reed and Muench (32).
Deep amplicon sequencing and genetic analysis.
Viral RNA was extracted using an RNeasy kit (Qiagen); conventional two-step reverse transcription-PCR then was performed using a SuperScript IV first-strand synthesis kit (Invitrogen) with the Uni12 influenza primer. Multiplex PCR of all eight gene segments was conducted by using Phusion high-fidelity DNA polymerase (New England Biolabs) with the Uni12/13 primers. PCR products were purified using illustra GFX PCR DNA and gel band purification kit (GE Healthcare). DNA libraries were prepared by the staff of the Hartwell Center at St. Jude Children’s Research Hospital by using NEXTera XT DNA-Seq library preparation kits (Illumina) according to the manufacturer’s instructions. Pooled libraries were sequenced with an Illumina MiSeq personal genome sequencer by 150-bp paired-end reads. CLC Genomics Workbench, version 7.0.3 (CLC Bio, Qiagen), was used to analyze and process the sequencing reads. DNA Lasergene 15 and BioEdit7.0 (33) were used for multiple sequence alignment and genomic signature analysis using the Clustal W algorithm (34). MEGA 7 was used for the phylogenetic tree reconstruction by applying the neighbor-joining method with Kimura’s two-parameter distance model and 1,000 bootstrap replicates (35). Trees were visualized using FigTree version 1.4.2 (http://tree.bio.ed.ac.uk/software/figtree/).
Pathogenicity in mice.
The MLD50 was determined by inoculating groups of five mice with serial 10-fold dilutions of virus, ranging from 10 to 106 EID50. Mice were anesthetized with isoflurane and intranasally inoculated with 30 μl of each virus. Another group of mice was anesthetized and intranasally inoculated with 30 μl of phosphate-buffered saline (PBS) at pH 7.2 as uninfected controls. Upon virus challenge, mice were monitored for 14 dpi for disease symptoms, weight loss, and mortality. Mortality was recorded as actual death or loss of ≥25% body weight, which is the threshold at which animals are required to be euthanized according to our animal protocol. Additional groups of six mice were inoculated intranasally with 106 EID50 of each virus and euthanized via CO2 asphyxiation at 3 or 5 dpi. Lung, nasal turbinate, heart, brain, and intestine were collected to determine viral titers. All organs were homogenized in sterile PBS with a Qiagen Tissue Lyser II (Qiagen). Organ homogenates were centrifuged at 2,000 × g for 10 min, and the supernatants were transferred to clean tubes. Virus titers were determined by EID50 assay.
Histology and immunohistochemistry.
The lungs of mice (n = 3) were collected at 3 and 5 dpi and fixed via intratracheal infusion and immersion in a 10% neutral buffered formalin solution. Tissues were embedded in paraffin, sectioned, and stained with hematoxylin and eosin. Immunohistochemical staining was performed on serial histologic sections to determine the distribution of AIV antigens. A goat primary polyclonal antibody (US Biological) against influenza A, USSR (H1N1), was used (1:1,000) on tissue sections that were previously subjected to antigen retrieval for 30 min at 98°C (target retrieval solution, pH 9; Dako Corp.). Stained sections were scanned on an Aperio ScanScope XT slide scanner (Leica Biosystems) to acquire digital images of whole-lung sections. Both uninfected and virus-positive regions were then manually outlined.
Pathogenicity and transmission in chickens.
Five-week-old specific-pathogen-free White Leghorn chickens (3 chickens/group; 6 groups/virus) were used to determine the 50% lethal dose (LD50) by the method of Reed and Muench (32). To compare the pathogenicity of the 4 viruses in vivo in chickens, three donor SPF White Leghorn chickens (CRL Avian Products and Services) were infected with 0.5 ml of 30 chicken LD50 dilutions of virus stock in PBS via the intranasal, intraocular, and intratracheal routes. Two chickens inoculated with 0.5 ml sterile PBS were used as negative controls. To examine virus transmission, we added three naive chickens to each group after 24 h postinfection, and they shared food and drinking water. Chickens were observed for clinical signs over 14 days. Oropharyngeal and cloacal swabs were collected from all birds at days 3, 5, and 7 postinfection to determine virus shedding. Viral titers from oropharyngeal and cloacal swabs were determined by the EID50 assay.
Pathogenicity and transmission in ferrets.
Three-month-old ferrets were purchased (Triple F Farms) for the pathogenicity and transmission experiments. Each animal was serologically negative for currently circulating influenza viruses, as determined by standard hemagglutination inhibition assay. Three ferrets were infected intranasally with 1 ml of 106 EID50/ml of each virus under isoflurane sedation. To assess direct-contact transmission, we placed a naive ferret in each cage with an infected ferret. To assess airborne-contact transmission, we placed three naive ferrets in cages adjacent to those of the infected ferrets. The body weight and temperature of the animals were monitored, and nasal washes were collected every 2 days for 2 weeks. Three additional ferrets were inoculated and then euthanized 4 dpi to assess virus replication. Virus titrations of nasal washes and various tissues and organs were determined by EID50 assays.
Statistical analyses.
All statistical analyses were performed by two-way analysis of variance (ANOVA) in combination with Bonferroni multiple-comparison tests by using GraphPad Prism 5.0 (GraphPad Software). P values of <0.05 were considered statistically significant.
Supplementary Material
ACKNOWLEDGMENTS
We thank the staff and students in the Department of Zoology at Jahangirnagar University for sample collection in the LPMs of Bangladesh, as well as the poultry workers for their cooperation during this study. We thank James Knowles for administrative assistance and Angela McArthur (St Jude Children's Research Hospital, Memphis, TN) for scientific editing.
This work was funded by the National Institute of Allergy and Infectious Diseases, National Institutes of Health (CEIRS contract no. HHSN266200700006C). The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or the American Lebanese Syrian Associated Charities (ALSAC).
Footnotes
Supplemental material is available online only.
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