VIRUS CEDAR UN HENIPAVIRUS EN MURCIÉLAGOS AUSTRALIA

Cedar Virus: A Novel Henipavirus Isolated from Australian Bats Glenn A. Marsh1#, Carol de Jong2#, Jennifer A. Barr1, Mary Tachedjian1, Craig Smith2, Deborah Middleton1, Meng Yu1, Shawn Todd1, Adam J. Foord1, Volker Haring1, Jean Payne1, Rachel Robinson1, Ivano Broz1, Gary Crameri1, Hume E. Field2*, Lin-Fa Wang1* 1 CSIRO Livestock Industries, Australian Animal Health Laboratory, Geelong, Australia, 2 Queensland Centre for Emerging Infectious Diseases, Biosecurity Queensland, Coopers Plain, Australia Abstract Top The genus Henipavirus in the family Paramyxoviridae contains two viruses, Hendra virus (HeV) and Nipah virus (NiV) for which pteropid bats act as the main natural reservoir. Each virus also causes serious and commonly lethal infection of people as well as various species of domestic animals, however little is known about the associated mechanisms of pathogenesis. Here, we report the isolation and characterization of a new paramyxovirus from pteropid bats, Cedar virus (CedPV), which shares significant features with the known henipaviruses. The genome size (18,162 nt) and organization of CedPV is very similar to that of HeV and NiV; its nucleocapsid protein displays antigenic cross-reactivity with henipaviruses; and it uses the same receptor molecule (ephrin- B2) for entry during infection. Preliminary challenge studies with CedPV in ferrets and guinea pigs, both susceptible to infection and disease with known henipaviruses, confirmed virus replication and production of neutralizing antibodies although clinical disease was not observed. In this context, it is interesting to note that the major genetic difference between CedPV and HeV or NiV lies within the coding strategy of the P gene, which is known to play an important role in evading the host innate immune system. Unlike HeV, NiV, and almost all known paramyxoviruses, the CedPV P gene lacks both RNA editing and also the coding capacity for the highly conserved V protein. Preliminary study indicated that CedPV infection of human cells induces a more robust IFN-β response than HeV. Author Summary Top Hendra and Nipah viruses are 2 highly pathogenic paramyxoviruses that have emerged from bats within the last two decades. Both are capable of causing fatal disease in both humans and many mammal species. Serological and molecular evidence for henipa-like viruses have been reported from numerous locations including Asia and Africa, however, until now no successful isolation of these viruses have been reported. This paper reports the isolation of a novel paramyxovirus, named Cedar virus, from fruit bats in Australia. Full genome sequencing of this virus suggests a close relationship with the henipaviruses. Antibodies to Cedar virus were shown to cross react with, but not cross neutralize Hendra or Nipah virus. Despite this close relationship, when Cedar virus was tested in experimental challenge models in ferrets and guinea pigs, we identified virus replication and generation of neutralizing antibodies, but no clinical disease was observed. As such, this virus provides a useful reference for future reverse genetics experiments to determine the molecular basis of the pathogenicity of the henipaviruses. Citation: Marsh GA, de Jong C, Barr JA, Tachedjian M, Smith C, et al. (2012) Cedar Virus: A Novel Henipavirus Isolated from Australian Bats. PLoS Pathogens 8(8): e1002836. doi:10.1371/journal.ppat.1002836 Editor: Christopher F. Basler, Mount Sinai School of Medicine, United States of America Received: December 15, 2011; Accepted: June 19, 2012; Published: August 2, 2012 Copyright: © 2012 Marsh et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited. Funding: This study is supported in part by an OCE Postdoctoral Fellowship (GAM) and a CEO Science Leader Award (LFW) from the CSIRO Office of the Chief Executive, and a research grant from the Australian Government Wildlife and Exotic Diseases Preparedness Program (HEF). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript. Competing interests: The authors have declared that no competing interests exist. * E-mail: hume.field@qld.gov.au (HEF); linfa.wang@csiro.au (LFW) # These authors contributed equally to this work. Introduction Top Henipaviruses were first discovered in the 1990s following investigation of serious disease outbreaks in horses, pigs and humans in Australia and Malaysia [1], [2] and comprise the only known Biosafety Level 4 (BSL4) agents in the family Paramyxoviridae [3]. Depending upon the geographic locations of outbreaks, and the virus and animal species involved, case mortality is between 40% to 100% in both humans and animals [4], [5], making them one of the most deadly group of viruses known to infect humans. The genus Henipavirus in the subfamily Paramyxovirinae currently contains two members, Hendra virus (HeV) and Nipah virus (NiV) [6]. Fruit bats in the genus Pteropus, commonly known as flying foxes, have been identified as the main natural reservoir of both viruses although serological evidence suggests that henipaviruses also circulate in non-pteropid bats [7], [8], [9], [10]. The discovery of henipaviruses had a significant impact on our understanding of genetic diversity, virus evolution and host range of paramyxoviruses. Paramyxoviruses, such as measles virus and canine distemper virus, were traditionally considered to have a narrow host range and to be genetically stable with a close to uniform genome size shared by all members of Paramyxovirinae [3]. Henipaviruses shifted this paradigm on both counts having a much wider host range and a significantly larger genome [6]. Identification of bats as the natural reservoir of henipaviruses also played an important role in significantly increasing international scientific attention on bats as an important reservoir of zoonotic viruses, including Ebola, Marburg, SARS and Melaka viruses [11], [12], [13], [14]. Since the discovery of the first henipavirus in 1994, much progress has been made in henipavirus research, from identification of functional cellular receptors to the development of novel diagnostics, vaccine and therapeutics [15], [16], [17], [18], [19], [20], [21], [22], [23], [24], [25]. By contrast, there is little understanding of the pathogenesis of these highly lethal viruses. This is due in part to the requirement of a high security BSL4 facility for any live infection studies and in part to the limited range of research tools and reagents for the current small animal models. Research into the mechanisms of henipavirus pathogenesis is also hampered by the lack of related, but non-pathogenic or less pathogenic viruses, thus preventing targeted comparative pathogenetic studies. Early serological investigations in Australia and more recent studies in other regions (e.g., China) indicated the presence of cross-reactive, but not cross-neutralizing, antibodies to henipaviruses in bats of different species [8]. These findings were further supported by the detection of henipavirus-like genomic sequences in African bats [26]. Discovery and isolation of these related viruses will be highly important to our further understanding of henipavirus evolution, mechanism of cross-species transmission, and pathogenesis in different animal species. Here we report the isolation and characterization of a new bat henipavirus which, based on preliminary infection studies, is non-pathogenic in two of the small animal infection models currently used in henipavirus research. We believe that this new virus will provide a powerful tool to facilitate our future study into different aspects of henipaviruses, especially in the less advanced area of pathogenesis. Results Top Virus isolation from pooled bat urine samples As part of our on-going field studies on HeV genetic diversity and infection dynamics in the Queensland flying fox populations, urine samples were collected on a regular basis for PCR and virus isolation. Since the establishment of the Pteropus alecto primary cell lines in our group [27], we have intensified our effort to isolate live virus from these urine samples by routinely inoculating separate primary cell lines derived from kidney, spleen, brain, and placenta, as well as Vero cells. Syncytial CPE was observed in kidney cell (PaKi) monolayers 5 days post inoculation (dpi) with two different urine samples (Fig. S1) collected in September 2009 from a flying fox colony in Cedar Grove, South East Queensland (see Fig. S2 for map location). No CPE was observed in any of the four other cell lines. Supernatant harvested 6 dpi was used to inoculate fresh PaKi cell monolayers. After two passages in PaKi cells, the virus was able to infect and cause CPE in Vero cells. However, the CPE morphology of CedPV infection in Vero cells was different from that of HeV infection. Further analysis using HeV-specific PCR primers indicated that the new bat virus was not an isolate of HeV. Genome analysis of the newly isolated virus Considering the formation of syncytial CPE by this new virus and the previous success in isolating paramyxoviruses from bat urine [28], [29], [30], paramyxovirus family-specific and genus-specific primers were used to determine whether this new virus was a member of the family Paramyxoviridae. Positive PCR fragments of the expected sizes were obtained from the Paramyxovirinae and Respirovirus/Morbillivirus/Henipavirus primer sets developed by Tong et al [31]. Sequencing of the PCR products indicated that it was a new paramyxovirus most closely related to HeV and NiV. Based on these preliminary data, the virus was named Cedar virus (CedPV) after the location of the bat colony sampled. Full length genome sequence was determined by a combination of three different approaches, random deep sequencing using 454 technology, sequencing of PCR products obtained using degenerate primers designed based on known henipaviruses, and RACE to determine the precise genome terminal sequences. As shown in Fig. 1, the genome of CedPV is 18,162 nt in length most similar to that of HeV in the family. The full genome sequence has been deposited to GenBank (Accession No. JQ001776). The genome size is a multiple of 6, hence abiding by the Rule-of-Six observed for all known members of the subfamily Paramyxovirinae [3]. It has a 3-nt intergenic sequence of CTT absolutely conserved at all seven positions and highly conserved gene start and stop signals similar to those present in HeV and NiV (Fig. S3). Also similar to the HeV genome is the presence of relatively large non-coding regions in the CedPV genome (Fig. 1 and Table 1). The overall protein-coding capacity of the CedPV genome is 87.41% which is significantly lower than the average of 92.00% for other family members but higher than HeV at 82.12%. As the genome size of CedPV and HeV is very similar, the increased coding capacity of CedPV is attributed to an increase in protein sizes for five of the six major proteins, with the L protein being 257-aa larger (Table 1). At 2,501 aa, the CedPV L protein is the largest, not only in the family Paramyxoviridae but also for all known viruses in the order Mononegavirale. thumbnail Figure 1. Comparison of genome size and organization of CedPV to those of the prototype viruses of the five existing genera in the subfamily Paramyxovirinae. Each of the coding and non-coding regions is drawn to scale. The six major genes present in all paramyxovirus genomes are indicated as follows: light shaded = RNA polymerase and nucleocapsid genes (N, P and L); slanted = envelope membrane protein genes (F and attachment protein); dotted = matrix protein (M). The dark shaded box represents the gene (SH) not commonly shared among members of the subfamily. doi:10.1371/journal.ppat.1002836.g001 Download: PowerPoint slide | larger image (112KB PNG) | original image (367KB TIFF) thumbnail Table 1. Comparison of common genes among CedPV, HeV and NiV. doi:10.1371/journal.ppat.1002836.t001 Download: PowerPoint slide | larger image (63KB PNG) | original image (383KB TIFF) Phylogenetic analysis based on the full length genome sequence and the deduced amino acid sequences of each structural protein confirmed the initial observation that CedPV is most closely related to henipaviruses in the family. A phylogenetic tree based on the deduced sequences of the nucleocapsid protein (N) is presented in Fig. 2. Phylogenetic tree based on whole genome sequences gave similar results (Fig. S4). CedPV is more closely related to HeV and NiV than henipavirus-like sequences detected in African bats [26], [32] as shown in a phylogenetic tree based on the only sequences available of a 550-nt L gene fragment (Fig. S5). thumbnail Figure 2. Phylogenetic tree based on the N protein sequences of selected paramyxoviruses. Virus name (abbreviation) and GenBank accession numbers are as follows: Avian paramyxovirus 6 (APMV6) AY029299; Atlantic salmon paramyxovirus (AsaPV) EU156171; Beilong virus (BeiPV) DQ100461; Bovine parainfluenza virus 3 (bPIV3) AF178654; Canine distemper virus (CDV) AF014953; Cedar virus (CedPV) JQ001776; Fer-de-lance virus (FdlPV) AY141760; Hendra virus (HeV) AF017149; Human parainfluenza virus 2 (hPIV2) AF533010; Human parainfluenza virus 3 (hPIV3) Z11575; Human parainfluenza virus 4a (hPIV4a) AB543336; Human parainfluenza virus 4b (hPIV4b) EU627591; J virus (JPV) AY900001; Menangle virus (MenPV) AF326114; Measles virus (MeV) AB016162; Mossman virus (MosPV) AY286409; Mapeura virus (MprPV) EF095490; Mumps virus (MuV) AB000388; Newcastle disease virus (NDV) AF077761; Nipah virus, Bangladesh strain (NiV-B) AY988601; Nipah virus, Malaysian strain (NiV-M) AJ627196; Parainfluenza virus 5 (PIV5) AF052755; Peste-des-petits-ruminants (PPRV) X74443; Porcine rubulavirus (PorPV) BK005918; Rinderpest virus (RPV) Z30697; Salem virus (SalPV) AF237881; Sendai virus (SeV) M19661; Simian virus 41 (SV41) X64275; Tioman virus (TioPV) AF298895; Tupaia paramyxovirus (TupPV) AF079780. doi:10.1371/journal.ppat.1002836.g002 Download: PowerPoint slide | larger image (257KB PNG) | original image (473KB TIFF) A phosphoprotein (P) gene lacking RNA editing and coding capacity for the V protein First discovered for the parainfluenza virus 5 (PIV5, previously known as simian virus 5), almost all members of Paramyxovirinae have a P gene which produces multiple proteins through an RNA editing mechanism by addition of non-templated G residues leading to production of N-terminal co-linear proteins from different reading frames downstream from the editing site [3], [33]. These multiple gene products are known to play a key role in antagonizing the innate response of susceptible hosts [3]. A search of CedPV for open reading frames (ORF) in the P gene revealed a 737-aa P protein and a 177-aa C protein, but failed to find the highly conserved, cysteine-rich V ORF present in most other paramyxoviruses. The RNA editing site with the sequence of AAAAGGG, which is absolutely conserved in all known HeV and NiV isolates discovered to date, is also missing from the CedPV P gene sequence. To further verify that there are no multiple mRNAs produced from the CedPV P gene, direct sequencing of P gene transcripts was conducted from CedPV-infected Vero cells using multiple sets of primers generating overlapping fragments covering the entire coding region of the P gene. Each produced uniform trace files indicating a lack of RNA editing activities, which is very different from the mixed peaks generated by HeV and NiV immediately after the editing site (Fig. S7). To our knowledge, CedPV is the first member of Paramyxovirinae that lacks both RNA editing and any V-related coding sequence in its P gene. Further investigation is required to exclude the possibility that the P-gene editing in CedPV is cell- or tissue-specific and not present or present at an extremely low level in the current virus-cell system. Antigenic relatedness with henipaviruses The striking similarity in genome size and organization and the presence of highly conserved protein domains among the N, M and L proteins between CedPV and henipaviruses prompted us to investigate the antigenic relatedness of these viruses. Staining of CedPV- infected Vero cells using rabbit anti-henipavirus antibodies indicated the presence of cross-reactivity. This cross-reactivity was further confirmed in reverse by staining of HeV-infected Vero cells using a rabbit serum raised against a recombinant CedPV N protein (Fig. 3). However, analysis by virus neutralization test using either polyclonal or monoclonal antibodies found that henipavirus-neutralizing antibodies were unable to neutralize CedPV. Similarly, CedPV-neutralizing antibodies obtained in our infection studies (see below) also failed to neutralize either HeV or NiV. It can therefore be concluded that CedPV and henipaviruses share cross-reactive antigenic regions, but not cross-neutralizing epitopes. thumbnail Figure 3. Antigenic cross reactivity between CedPV and HeV. Vero cells infected with CedPV and HeV, respectively, were stained with rabbit sera raised against recombinant N proteins of each virus. doi:10.1371/journal.ppat.1002836.g003 Download: PowerPoint slide | larger image (1.64MB PNG) | original image (1.92MB TIFF) Use of ephrin-B2 as a functional receptor for membrane fusion and entry by CedPV To further investigate the relationship between CedPV and recognized henipaviruses, we investigated the use of the henipavirus receptors, the ephrin-B2 and -B3 host cell proteins, as potential receptors for CedPV infection. Our previous studies have demonstrated that the ephrin-B2 and -B3 expression negative HeLa-USU cell line could support henipavirus infection and formation of syncytial CPE only when either the ephrin-B2 or -B3 gene was transiently expressed in the cells [22], [34]. For CedPV, similar observations were made with respect to the ephrin-B2 receptor. As shown in Fig. 4, CedPV failed to infect HeLa-USU, but was able to infect and cause syncytial CPE when the human ephrin-B2 gene was expressed. In contrast, when ephrin-B3 molecule was introduced, there was no evidence of infection. thumbnail Figure 4. Functional testing of ephrin-B2 and -B3 as an entry receptor for CedPV. Infection of HeLa-USU cells by CedPV in the presence and absence of ephrin gene products. The susceptibility of infection, as an indirect measurement of receptor function, is demonstrated by the formation of syncytial CPE. doi:10.1371/journal.ppat.1002836.g004 Download: PowerPoint slide | larger image (1.18MB PNG) | original image (1.44MB TIFF) Pathogenicity for laboratory mammals Ferrets, guinea pigs, and mice exhibit differing responses to the previously described henipaviruses HeV and NiV, with ferrets and guinea pigs, but not mice developing severe disease characterized by systemic vasculitis [20], [35], [36], [37], [38]. In contrast, ferrets and guinea pigs exposed to CedPV by, respectively, oronasal and intraperitoneal routes remained clinically well although neutralizing antibody was detected in serum between 10 to 21 days pi (Table 2). Balb-C mice exposed to CedPV by the oronasal route remained clinically well and did not develop neutralizing antibody in serum by day 21 pi. In ferrets electively euthanized at earlier time-points, there was reactive hyperplasia of tonsillar lymphoid tissue, retropharyngeal and bronchial lymph nodes, accompanied by edema and erythrophagocytosis. CedPV antigen was detected in bronchial lymph node of one animal euthanized on day 6 pi, consistent with viral replication in that tissue; cross-reactive immunostaining against anti-NiV N protein antibodies was also noted (Fig. 5). No other significant histological lesions were identified. Viral RNA was detected in selected lymphoid tissues of 3 (of 4) ferrets sampled day 6 to 8 pi, including pharynx, spleen, and retropharyngeal and bronchial lymph nodes, as well as the submandibular lymph node of the ferret euthanized on day 20 pi. This pattern of lymphoid involvement suggests that there may be transient replication in the upper and lower respiratory tracts although CedPV genome was not recovered from nasal washes, oral swabs, pharynx or lung tissue of affected animals. Virus isolation was unsuccessful for all PCR positive tissues. thumbnail Figure 5. Immunohistochemical analysis of bronchial lymph node of CedPV infected ferrets. Bronchial lymph node of ferret #2, euthanized on day 6 pi, was stained with rabbit antiserum against recombinant N protein of CedPV (B) and NiV (D), respectively. Bronchial lymph node of an unrelated ferret (infected with influenza H5N1 from another experiment) was used as negative control and stained with the same anti-CedPV (A) and anti-NiV (C) antisera under identical conditions. doi:10.1371/journal.ppat.1002836.g005 Download: PowerPoint slide | larger image (8.3MB PNG) | original image (6.73MB TIFF) thumbnail Table 2. Antibody responses in CedPV-infected ferrets and guinea pigs. doi:10.1371/journal.ppat.1002836.t002 Download: PowerPoint slide | larger image (37KB PNG) | original image (142KB TIFF) Induction of IFN responses upon infection As a first step towards the understanding of the pathogenicity difference between CedPV and HeV, we examined the IFN responses in human HeLa cells upon virus infection. As shown in Fig. 6, while the induction of IFN-α was similar in cells infected with HeV or CedPV, there was a significant difference of IFN-β production upon infection by HeV or CedPV, with CedPV-infected cell producing a much higher level of IFN-β. thumbnail Figure 6. Induction of IFN responses upon henipavirus infection. HeLa cells were infected at an MOI 0.5 for 24 hours. Total RNA was isolated, and quantitative real-time PCR for IFN-α and IFN-β was performed. n = 2, with error bars indicating SEM. doi:10.1371/journal.ppat.1002836.g006 Download: PowerPoint slide | larger image (14KB PNG) | original image (86KB TIFF) Prevalence of neutralizing antibodies in Australian fruit bats To investigate the CedPV exposure status of pteropid bats in Queensland and potential co-infection (either concurrent or consecutive) of CedPV with HeV, we tested 100 flying fox sera collected previously for other studies for antibody against the two viruses. Due to the cross-reactivity observed above, virus neutralization tests were conducted to obtain more accurate infection data for each virus. Overall, 23% of the sera were CedPV-positive and 37% HeV-positive (Table S1). Co-infection was reflected in 8% of the sera tested. Discussion Top The emergence of bat-borne zoonotic viruses (including HeV, NiV, Ebola, Marburg, and SARS) has had a significant impact on public health and the global economy during the past few decades. With the rapidly expanding knowledge of virus diversity in bat populations around the world, it is predicted that more bat-borne zoonotic viruses are likely to emerge in the future. The discovery of a novel ebolavirus-like filovirus in Spanish microbats demonstrates that the potential for such spill over events is not limited to Africa or Asia [39]. It is therefore important to enhance our preparedness to counter future outbreaks by conducting active pre-emergence research into surveillance, triggers for cross-species transmission, and the science of identification of potential pathogens. Henipaviruses represent one of the most important bat-borne pathogens to be discovered in recent history. Although CedPV displays some differences from existing members of the genus Henipavirus, we propose that CedPV be classified as a new henipavirus based on the following shared features with known henipaviruses: 1) it is antigenically related to current henipaviruses; 2) its genome size and organization is almost identical to those of HeV and NiV; 3) it has a similar prevalence in flying foxes; and 4) it uses ephrin-B2 as the cell entry receptor. The lack of cross-neutralization between CedPV and HeV or NiV was not unexpected from the comparative sequence analysis of all the deduced proteins, especially the G protein (see Table 1). It is clear that the genetic relatedness of CedPV with HeV or NiV is much lower than between HeV and NiV. However, the percentage sequence identities of the major viral proteins between CedPV and HeV/NiV are on average at least 10% higher than that between HeV/NiV and any other known paramyxoviruses. Also, there was no antigenic cross-reactivity observed between CedPV and representative viruses of the other paramyxovirus genera in the subfamily Paramyxovirinae (Fig. S6). Like other paramyxoviruses, the P gene of henipaviruses produces multiple proteins which play a key role in viral evasion of host innate immune responses [4], [40], [41]. One of these is the Cys-rich V protein: all members of the subfamily Paramyxovirinae produce the V protein with the exception of the human parainfluenza virus 1 (hPIV1). Although a putative RNA editing sequence (AAGAGGG) is present at the expected editing site of the P gene, the hPIV1 RNA polymerase does not produce an edited mRNA of the P gene [42]. There are remnants of the V ORF easily detectable in the hPIV1 P gene although the predicted 68-aa ORF region is interrupted by multiple in-frame stop codons. Of the 7 Cys residues conserved between bovine parainfluenza virus 3 and Sendai virus, four are still present in the non-functional V ORF of hPIV1[42]. In contrast, an extensive ORF and sequence homology search of the CedPV P gene only identified one aa coding region with minimal sequence identity to the V ORFs of HeV and NiV (see Fig. S8). In this region, out of the 9 Cys residues conserved between HeV and NiV V proteins, only 2 are present in the CedPV P gene. Furthermore, the sequence (AGATGAG) upstream from this putative ORF V coding region does not match the consensus RNA editing site. It can therefore be concluded that CedPV is the only member of Paramyxovirinae which lacks both the functional V mRNA/protein and the coding capacity for the RNA editing site and ORF V. The evolutionary significance of this finding needs further investigation. Our in vitro study indicated that ephrin B2, but not ephrin B3, was able to restore CedPV infection in the ephrin B2-deficient HeLa cells. While this is highly suggestive that ephrin B2 is the functional entry receptor for CedPV, it should be emphasized that this was not a direct proof that ephrin B2 is the receptor. Further investigation is required to confirm this. In our preliminary studies, it was shown that CedPV was able to replicate in guinea pigs and ferrets, but failed to cause significant clinical diseases, unlike that of the closely related HeV and NiV. These first infection experiments were conducted with a high dose if virus to establish whether the CedPV could replicate in these animals and determine the degree of any clinical disease. A second experiment was then carried out in ferrets to determine the site of replication and tissue tropism in sequentially sacrificed animals. A lower dose was used to gain better comparison with similar infection experiments using HeV and NiV [18], [35]. Although these initial experimental infection studies indicate that CedPV is less or non-pathogenic in these species, it is possible that CedPV may be pathogenic in other hosts, such as horses. We hypothesize that the lack of a V protein may impact on the pathogenicity. In this regard, it was encouraging to observe that infection of human cells by CedPV induced a much more robust IFN-β response than HeV. Further study is required to dissect the exact molecular mechanism of this observed difference. Due to the close relationship between CedPV and HeV, it was important to investigate the possibility of co-infection by these two viruses in the Australian bat population. Based on the detection of neutralizing antibodies at 23% for CedPV, 37% for HeV and 8% for both, it can be concluded that the co-infection rate is very close to the theoretical rate of 8.5% (the product of the two independent infection rates). Based on this limited preliminary analysis, it appears that infection of bats by one henipavirus neither prevents nor enhances the likelihood of infection by the other. In summary, the discovery of another henipavirus in Australian flying foxes highlights the importance of bats as a significant reservoir of potential zoonotic agents and the need to expand our understanding of virus-bat relationships in general. Our future research will be directed at determining whether spill-over of CedPV into other hosts has occurred in the past in Australia, whether CedPV is pathogenic in certain mammalian hosts, and whether CedPV exists in bat populations in geographically diverse regions. Materials and Methods Top All animal studies were approved by the CSIRO Australian Animal Health Laboratory's Animal Ethics Committee and conducted following the Australian National Health and Medical Research Council Code of Practice for the Care and Use of Animals for Scientific Purposes guidelines for housing and care of laboratory animals. Cell culture Cell lines used this study were Vero (ATCC), HeLa-USU [22], and the P. alecto primary cell lines derived from kidney (PaKi), brain (PaBr), (spleen) PaSp and placenta (PaPl) recently established in our group [27]. Cells were grown in Dulbecco's Modified Eagle's Medium Nutrient Mixture F-12 Ham supplemented with double strength antibiotic-antimycotic (Invitrogen), 10 µg/ml ciprofloxacin (MP Biomedicals) and 10% fetal calf serum at 37°C in the presence of 5% CO2. Urine collection and virus isolation Urine (approximately 0.5–1 ml) was collected off plastic sheets placed underneath a colony of flying foxes (predominantly Pteropus alecto with some P. Poliocephalus in the mixed population) in Cedar Grove, South East Queensland, Australia and pooled into 2-ml tubes containing 0.5 ml of viral transport medium (SPGA: a mix of sucrose, phosphate, glutamate and albumin plus penicillin, streptomycin and fungizone). The tubes were temporarily stored on ice after collection and transported to a laboratory in Queensland, frozen at −80°C, and then shipped on dry ice to the CSIRO Australian Animal Health Laboratory (AAHL) in Geelong, Victoria for virus isolation. The samples were thawed at 4°C and centrifuged at 16,000×g for 1 min to pellet debris. Urine in the supernatant (approximately 0.5–1 ml) was diluted 1:10 in cell culture media. The diluted urine was then centrifuged at 1,200×g for 5 min and split evenly over Vero, PaKi, PaBr, PaSp and PaPl cell monolayers in 75-cm2 tissue culture flasks. The flasks were rocked for 2 h at 37°C, 14 ml of fresh cell culture media was added and then incubated for 7 d at 37°C. The flasks were observed daily for toxicity, contamination, or viral cytopathic effect (CPE). Molecular characterization Cells showing syncytial CPE were screened using published broadly reactive primers [31] for all known paramyxoviruses and a subset of paramyxoviruses. PCR products were gel extracted and cloned into pGEM T-Easy (Promega) to facilitate sequencing using M13 primers. Sequences were obtained and aligned with known paramyxovirus sequences allowing for initial classification. Whole genome sequence was determined using a combination of 454 sequencing [43] and conventional Sanger sequencing. Virions from tissue culture supernatant were collected by centrifugation at 30,000×g for 60 min and resuspended in 140 µl of PBS and mixed with 560 µl of freshly made AVL for RNA extraction using QIAamp Viral RNA mini kit (Qiagen). Synthesis of cDNA and random amplification was conducted using a modification of a published procedure [44]. Briefly, cDNA synthesis was performed using a random octomer-linked to a 17-mer defined primer sequence: (5′-GTTTCCCAGTAGGTCTCNNN NNNNN-3′) and SuperScript III Reverse Transcriptase (Invitrogen). 8 µl of ds-cDNA was amplified in 200 µl PCR reactions with hot-start Taq polymerase enzyme (Promega) and 5′-A*G*C*A*C TGTAGGTTTCCCAGTAGGTCTC-3′ (where * denotes thiol modifications) as amplification primers for 40 cycles of 95°C/1 min, 48°C/1 min, 72°C/1 min after an initial denaturation step of 5 min at 95°C and followed by purification with the QIAquick PCR purification kit (Qiagen). Sample preparation for Roche 454 sequencing (454 Life Sciences Branford, CT, USA) was according to their Titanium series manuals, Rapid Library Preparation and emPCR Lib-L SV. To obtain an accurate CedPV genome sequence, 454 generated data (after removing low quality, ambiguous and adapter sequences) was analysed by both de novo assembly and read mapping of raw reads onto the CedPV draft genome sequence derived from Sanger sequencing. For 454 read mapping, SNPs and DIPs generated with the CLC software were manually assessed for accuracy by visualising the mapped raw reads (random PCR errors are obvious compared to real SNPs and DIPs especially when read coverage is deep). Consensus sequences for both 454 de novo and read mapping assembly methods were then compared to the Sanger sequence with the latter used to resolve conflicts within the low coverage regions as well as to resolve 454 homopolymer errors. Sequences of genome termini were determined by 3′- and 5′-RACE using a protocol previously published by our group [45]. Briefly, approximately 100 ng of RNA was ligated with adaptor DT88 (see reference for sequence information) using T4 RNA ligase (Promega) followed by cDNA synthesis using the SuperScript III RT kit (Invitrogen) and an adaptor-specific primer, DT89. PCR amplification was then carried out using DT89 and one or more genome-specific primers. PCR products were sequenced directly using either DT89 or genome specific primers by an in-house service group on the ABI Sequencer 3100. Sequence analysis The CLC Genomics Workbench v4.5.1 (CLC Inc, Aarhus, Denmark) was used to trim 454 adapter and cDNA/PCR primer sequences, to remove low quality, ambiguous and small reads <15 bp and to perform de novo and read mapping assemblies all with default parameters. Clone Manager Professional ver 9.11 (Scientific and Educational Software, Cary, NC, USA) was used to join overlapping contigs generated by de novo assembly. Phylogenetic trees were constructed by using the neighbor-joining algorithm with bootstrap values determined by 1,000 replicates in the MEGA4 software package [46]. Real time PCR Quantitative PCR assays (qPCR) were established based on CedPV-specific sequences obtained from the high throughput sequencing. A TaqMan assay on the P gene was developed and used for all subsequent studies. The sequences of the primer/probe are as follows: forward primer, 5′-TGCAT TGAGC GAACC CATAT AC; reverse primer, 5′-GCACG CTTCT TGACA GAGTT GT; probe, 5′-TCCCG AGAAA CCCTC TGTGT TTGA-MGB. Production of recombinant antigen and rabbit sera The coding region for the CedPV N protein was amplified by PCR with a pair of primers flanked by AscI (5′ end) and NotI (3′ end) sites for cloning into our previously described GST-fusion expression vector [47]. The expression and purification by gel elution was conducted as previously described [48]. For antibody production, purified protein was injected subcutaneously into 4 different sites of 2 adult (at a dose of 100 µg per animal) New Zealand white female rabbits at days 0 and 27. The CSIRO's triple adjuvant [49] was used for the immunization. Animals were checked for specific antibodies after days 5 and 42 and euthanized at day 69 for the final blood collection. Antibody tests For immunofluorescence antibody test, Vero cell monolayers were prepared in 8-well chamber slides by seeding at a concentration of 30,000 cells/well in 300 µl of cell media and incubating over night at 37°C. The cell monolayers were infected with an MOI of 0.01 of CedPV, HeV or NiV and fixed with 100% ice-cold methanol at 24 h post-infection. The chamber slides were blocked with 100 µl/well of 1%BSA in PBS for 30 min at 37°C before adding 50 µl/well of rabbit sera against CedPV N or NiV N diluted 1:1000. After incubation at 37°C for 30 min, the slides were washed three times in PBS-T and incubated with 50 µl/well of anti-rabbit 488 Alexafluore conjugate (Invitrogen) diluted 1:1000 at 37°C for 30 min. The slides were then washed three times in PBS-T and mounted in 50% glycerol/PBS for observation under a fluorescence microscope. For virus neutralization test, serial two-fold dilutions of sera were prepared in duplicate in a 96-well tissue culture plate in 50 µl cell media (Minimal Essential Medium containing Earle's salts and supplemented with 2 mM glutamine, antibiotic-antimycotic and 10% fetal calf serum). An equal volume containing 200 TCID50 of target virus was added and the virus-sera mix incubated for 30 min at 37°C in a humidified 5% CO2 incubator. 100 µl of Vero cell suspension containing 2×105 cells/ml was added and the plate incubated at 37°C in a humidified 5% CO2 incubator. After 4 days, the plate was examined for viral CPE. The highest serum dilution generating complete inhibition of CPE is defined as the final neutralizing titer. Testing of receptor specificity Human ephrin B2 and B3 genes were cloned into pQCXIH (Clontech) and the resulting plasmids packaged into retrovirus particles in the GP2–293 packaging cell line (Clontech) and pseudotyped with vesicular stomatitis virus G glycoprotein (VSV-G) following the manufacturer's instructions. HeLa-USU cell line [22] was infected with the VSV-G pseudotyped retrovirus particles in the presence of 1 µg/ml polybrene (Sigma). 8 h post infection, the medium was changed and the cells were allowed to recover for 24 h, allowing time for the retroviral insert to be incorporated into the cell genome and for expression of the hygromycin resistance gene. 24 h post infection, cells transformed by the retrovirus were selected for by the addition of 200 µg/ml hygromycin in the media. Stocks of cells that were resistant to hygromycin were prepared and frozen. HeLa-USU and ephrin-expressing HeLa-USU cells were seeded in 6-well tissue culture plates at a density of 250,000 cells/well overnight. The viruses (HeV and CedPV) were diluted to give an MOI of 0.01 and inoculated into the wells. The cell monolayers were examined daily for syncytial CPE. Animal infection studies Animal studies were carried out in the BSL4 animal facility at AAHL. Ferrets, guinea pigs and mice were used on the basis of their known and varying responses to exposure to other henipaviruses. Firstly, 2×106 TCID50/ml CedPV passaged twice in bat PaKi cells was administered to 2 male ferrets (1 ml oronasally); 4 female guinea pigs (1 ml intraperitoneally); and 5 female Balb-C mice (50 µl oronasally). Guinea pigs and mice were implanted with temperature sensing microchips (LifeChip Bio-thermo, Destron Fearing) and weighed daily. Ferret rectal temperature and weight was recorded at sampling times. Animals were observed daily for clinical signs of illness and were euthanized at 21 d post-inoculation. Sera were collected on days 10, 15 and 21 to test for neutralizing antibody against CedPV. Secondly, on the basis of asymptomatic seroconversion to CedPV noted in ferrets in the first study, 7 further female ferrets were exposed by the oronasal route to a lower dose of 3×103 TCID50. Two animals were euthanized on each of days 6, 8 and 10 post-inoculation and one on day 20. Nasal washes, oral swabs, and rectal swabs were collected on days 2, 4, 6, 8 and 10 and urine was sampled on the day of euthanazia; each specimen was assessed for CedPV genome. A wide range of tissue samples were collected at post mortem examination and assessed by routine histology, immunohistochemistry (using rabbit antibodies raised against recombinant CedPV and NiV N proteins, respectively), qPCR (see above) and virus isolation using reagents and procedures previously established in our group [16]. Determination of IFN responses HeLa cells were infected with Hendra and Cedar viruses at an MOI 0.5 for 24 hours, at which time total cellular RNA was extracted and IFN-α and IFN-β mRNA levels were quantified by real-time PCR using Power SYBR Green RNA-to-CT 1-Step Kit (Applied Biosystems). Primers were as previously described [50]. Serological survey Sera from 100 flying foxes collected during 2003–2005 from Queensland, Australia were screened for neutralizing antibodies to CedPV. Virus neutralization test was conducted as described above (antibody tests). All serum samples were tested at a dilution of 1:20. Supporting Information Top Figure S1. Cytopathic effect (CPE) observed in Paki cells. This is the original syncytial CPE seen in Paki cells 5 days post inoculation. (TIF) Figure S2. Map location of the sampling site, Cedar Grove, in southeast Queensland. The location of the index Hendra virus outbreak in 1994 is shown by a green dot while the sampling site of the current study is marked by a red star. (TIF) Figure S3. Comparison of genomic features among different henipaviruses. (A) Alignment of leader and trailer sequences (antigenome sequences shown). (B) Sequences of intergenic regions (IGR) and transcriptional start and stop sties of CedPV in comparison with those of HeV and NiV. (DOCX) Figure S4. Phylogenetic trees of viruses in the subfamily Paramyxovirinae based on whole genome sequence. (TIF) Figure S5. Phylogenetic trees of viruses in the subfamily Paramyxovirinae based on a 550-nt region of the L-gene. (TIF) Figure S6. Sequencing trace files for the editing site of P genes for HeV and NiV in comparison to a putative editing site of the CedPV P gene. Trace files showing editing of the HeV and NiV P gene (indicated by the * sign) and lack of editing in CedPV P gene mRNA in infected cells. Sequencing of PCR products covering all potential editing sites in the P gene of CedPV did not reveal any RNA editing activity. A representative potential editing site (see Fig. S8) of the CedPV P gene is shown. (TIF) Figure S7. Determination of antigenic cross reactivity with other paramyxoviruses. Shown here are IFAT conducted with anti-CedPV serum on Vero cells infected with J paramyxovirus (JPV), Rinderpest virus (RPV), Sendai virus (SeV), Menangle virus (MenPV) and CedPV, respectively. Mock infected cell monolayer was included as a negative control. (TIF) Figure S8. Sequence alignments of putative V ORF (A) and mRNA editing site (B) among HeV, NiV and CedPV. (DOCX) Table S1. Prevalence of neutralizing antibodies to CedPV and HeV in Australian flying foxes. (DOCX) Acknowledgments Top We thank Kaylene Selleck, Jessica Haining, Lauren Dagley, Susanne Wilson, Honglei Chen and Tony Pye for technical assistance. We thank Drs. 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Emergencia de influenza aviar fatal en focas de New England Harbor

Emergence of Fatal Avian Influenza in New England Harbor Seals Emergence of Fatal Avian Influenza in New England Harbor Seals S. J. Anthony,a,b J. A. St. Leger,c K. Pugliares,d H. S. Ip,e J. M. Chan,f Z. W. Carpenter,f I. Navarrete-Macias,a M. Sanchez-Leon,a J. T. Saliki,g J. Pedersen,h W. Karesh,b P. Daszak,b R. Rabadan,f T. Rowles,i and W. I. Lipkina Center for Infection and Immunity, Mailman School of Public Health, Columbia University, New York, New York, USAa; EcoHealth Alliance, New York, New York, USAb; SeaWorld, San Diego, California, USAc; New England Aquarium, Boston, Massachusetts, USAd; National Wildlife Health Center, United States Geological Survey, Madison, Wisconsin, USAe; Center for Computational Biology and Bioinformatics, Department of Biomedical Informatics, Columbia University, New York, New York, USAf; Athens Veterinary Diagnostic Laboratory, University of Georgia, Athens, Georgia, USAg; National Veterinary Services Laboratory, United States Department of Agriculture, Washington, DC, USAh; and National Oceanic and Atmospheric Administration, Washington, DC, USAi ABSTRACT From September to December 2011, 162 New England harbor seals died in an outbreak of pneumonia. Sequence analysis of postmortem samples revealed the presence of an avian H3N8 influenza A virus, similar to a virus circulating in North American waterfowl since at least 2002 but with mutations that indicate recent adaption to mammalian hosts. These include a D701N mutation in the viral PB2 protein, previously reported in highly pathogenic H5N1 avian influenza viruses infecting people. Lectin staining and agglutination assays indicated the presence of the avian-preferred SA_-2,3 and mammalian SA_-2,6 receptors in seal respiratory tract, and the ability of the virus to agglutinate erythrocytes bearing either the SA_-2,3 or the SA_-2,6 receptor. The emergence of this A/harbor seal/Massachusetts/1/2011 virus may herald the appearance of an H3N8 influenza clade with potential for persistence and cross-species transmission. IMPORTANCE The emergence of new strains of influenza virus is always of great public concern, especially when the infection of a new mammalian host has the potential to result in a widespread outbreak of disease. Here we report the emergence of an avian influenza virus (H3N8) in New England harbor seals which caused an outbreak of pneumonia and contributed to a U.S. federally recognized unusual mortality event (UME). This outbreak is particularly significant, not only because of the disease it caused in seals but also because the virus has naturally acquired mutations that are known to increase transmissibility and virulence in mammals. Monitoring the spillover and adaptation of avian viruses in mammalian species is critically important if we are to understand the factors that lead to both epizootic and zoonotic emergence. Received 31 May 2012 Accepted 29 June 2012 Published 31 July 2012 Citation Anthony SJ, et al. 2012. Emergence of fatal avian influenza in New England harbor seals. mBio 3(4):e00166-12. doi:10.1128/mBio.00166-12. Editor Anne Moscona, Weill Medical College, Cornell University Copyright © 2012 Anthony et al. This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-Share Alike 3.0 Unported License, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited. Address correspondence to S. J. Anthony, sja2127@columbia.edu, or W. I. Lipkin, wil2001@columbia.edu; for questions on computational modeling, contact R. Rabadan, rabadan@dbmi.columbia.edu. S.J.A. and J.A.S.L. contributed equally to the study. T.R. and W.I.L. are joint senior authors. Fatal pulmonary epizootics of influenza have been observed previously in seal populations, including outbreaks of H7N7 in 1979 to 1980 (1, 2), H4N5 in 1983 (3) and H4N5 and H3N3 in 1991 to 1992 (4). Such outbreaks are significant not just because of the detriment they pose to animal health but because influenza in mammals can act as a source for human pandemics (5). In a _4-month period beginning in September 2011, 162 harbor seals (Phoca vitulina) were found dead or moribund along the New England coast. This number is approximately four times the expected mortality for this period. Most of the affected individuals were less than 6 months old, and common causes of death (including malnourishment) were ruled out. Five of the affected animals were investigated to identify a causative agent, and here we demonstrate that avian influenza virus subtype H3N8 was responsible for the observed clinical and pathological signs in these animals. Unlike any previous outbreak in seals, this H3N8 virus has naturally acquired mutations that reflect adaptation to mammalian hosts and that are known to increase virulence and transmissibility in avian H5N1 viruses infecting mammals. The virus has further acquired the ability to use the SA_-2,6 receptor commonly found in the respiratory tracts of mammals, including humans. The existence of a transmissible and pathogenic influenza is of obvious public concern. RESULTS AND DISCUSSION Five animals were submitted for anatomical and microbiological analysis. All were collected from the peak of the outbreak (late September to October) and had pneumonia and ulcerations of the skin and oral mucosa (Fig. 1). Nucleic acids extracted from lung, trachea, liver, kidney, thoracic lymph node, mesenteric lymph node, spleen, skin lesion, and oral mucosa were tested by PCR for the presence of a wide range of pathogens, including herpesviruses, poxviruses, adenoviruses, polyomaviruses, caliciviruses, paramyxoviruses, astroviruses, enteroviruses, flaviviruses, rhabdoviruses, orbiviruses, and influenza viruses. Influenza A virus was detected in several tissues from all five animals, and PCR cloning and sequencing of genes for hemagglutinin (HA) and neuraminidase (NA) revealed the subtype to be H3N8, a subtype RESEARCH ARTICLE July/August 2012 Volume 3 Issue 4 e00166-12 ® mbio.asm.org 1 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org typically associated with infection of avian, equine, and canine hosts (6-8). Influenza virus was isolated from the allantoic fluid of specific-pathogen-free (SPF) eggs inoculated with homogenates of PCR-positive tissues, including lung, lymph nodes, tonsil, and kidney, and all isolates were reconfirmed to be H3N8. In accord with conventional nomenclature, the virus is provisionally named A/harbor seal/Massachusetts/1/2011. In situ hybridization (ISH) using oligonucleotide probes for influenza virus H3N8 segments 4 (HA) and 7 (matrix) and immunohistochemistry using polyclonal antibodies against H3N8 HA antigen confirmed the presence of influenza virus in lung, where signal was concentrated in the bronchiole epithelium and mucosa of the pulmonary parenchyma (Fig. 2 and see Fig. S1 in the supplemental material). The average load of HA and NA RNAs in lung was 300 copies/100 ng of extracted RNA. ISH staining in nonrespiratory tissues was limited to sporadic infection of single cells in intestine, kidney, and lymph node, and the averageHA/NA RNA load was five copies/100 ng. These results are consistent with the histological observation that the main site of viral replication is the respiratory tract. Cellular morphology and terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) consistent with apoptosis was observed in virusinfected pulmonary epithelial cells (Fig. 3). TUNEL staining also revealed the widespread presence of apoptotic cells in areas where no virus was observed but not in negative-control tissue, suggesting an additional host-mediated response to the infection. Full-genome sequencing was completed following PCR cloning of all eight influenza genome segments directly from infected tissue. Sequences were submitted to GenBank and assigned accession numbers JQ433879 to JQ433882. Phylogenetic analyses of these nucleotide sequences with avian, canine, and equine H3N8 influenza virus genomes demonstrated the closest relationship to a virus identified in North American waterfowl in all 8 segments (Fig. 4). These data are consistent with the recent transmission of the H3N8 virus from wild birds to seals. The closest avian relative, A/blue-winged teal/Ohio/926/2002, had an overall 96.07% nucleotide sequence identity across the genome (Hamming distance), with no individual segment having less than 94% identity. This level of similarity across all segments with an isolate separated by a span of 10 years suggests that this virus has been circulating in the aquatic bird populations since at least 2002. A total of 37 amino acid substitutions separate the seal H3N8 and avian H3N8 viruses, which are summarized in Table 1. The corresponding amino acids found in other seal influenza viruses, in the canine and equine H3N8 viruses, and in selected human influenza viruses are included for comparison. Of these, mutations PB2-701N and HA-260M are shared exclusively by the seal and mammalian (canine/equine) H3N8 viruses, while mutations NA-399R, PB2-382V, and PA-184N are shared by the seal H3N8 and human H3N2 viruses, all of which suggests adaptation to mammalian hosts. Mutations PB2-60N, PB2-376R, PB1-174I, PB1-309G, PB1-359G, PB1-376V, PB1-377G, PB1-464E, NP- 63V, NP-128G, and NP-296H are all exclusively found in the seal H3N8 virus (Table 1). Future studies will be required to assess the functional significance of many of these mutations, especially in sealH3N8PB1, where a significant number of exclusive mutations were observed. Given the importance of PB1 in viral replication, it is probable that these mutations represent adaptive selection to accommodate host-specific differences in intracellular replication. The seal H3N8 genome was interrogated for any genetic features that might contribute to enhanced transmissibility and virulence in seals. Expression of a second peptide (PB1-F2) from segment 2 has been associated with an increase in pathogenicity by inducing apoptosis and increasing both inflammation and secondary bacterial pneumonia (9, 10). The seal H3N8 virus contains an intact open reading frame for the pathogenic version of this accessory protein, which includes a serine at amino acid position 66 (9). All five seals had evidence of apoptosis and secondary bacterial pneumonia. Glycosylation can also affect pathogenicity in influenza viruses (11-14). Six potential glycosylation sites were detected in the seal H3N8 HA, based on the sequence X_2X_1NX(S/T) X_1, at amino acid positions 24, 38, 54, 181, 301, and 499. None had features FIG 1 Hematoxylin and eosin staining of the lung at a_10 magnification. There is diffuse acute interstitial pneumonia and a mix of acute hemorrhagic alveolitis with necrotizing bronchitis. Multifocally, alveoli are either filled with hemorrhage and scant inflammatory cells or expanded with emphysema. There is irregularity to the bronchial mucosa due to necrosis, a mild to moderate edema, and mucous partially filling the bronchial lumen. There is mild to moderate expansion of the interlobular septa, with edema and hemorrhage. Anthony et al. 2 ® mbio.asm.org July/August 2012 Volume 3 Issue 4 e00166-12 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org suggestive of inactivity or reduced efficiency, as was previously demonstrated for H5N2 (12). Many H5N1 viruses have an additional glycosylation site at positions 158 to 160, and previous work demonstrated that the deletion of this site is critical for H5N1 viruses to bind to human-SA_2,6-like receptors and to transmit between mammals (15). This glycosylation site is missing in the seal H3N8 HA. In order to investigate the specificity of sialic acid binding, the seal virus was compared to avian H5 and swine H9, both of which bind to the sialyloligosaccharide SA_2,3 ligand in a structuralhomology model (Fig. 5). All structures confirmed the presence of a highly conserved serine at position 152 (corresponding to S136 by H3 numbering [16, 17]), which lies in a binding pocket, where its hydroxyl group contacts the axial carboxylate of sialic acid (17). While the seal virus contains the same conserved S152, it also harbors a neighboring G151E mutation (Table 1), which introduces a large residue capable of both donating and receiving hydrogen bonds with residues in close proximity to the ligandbinding pocket. Rotamer hydrogen bond analysis of the modeled seal structure indicates that HA’s altered conformation results in reduced hydrogen bonding between the conserved serine and SA_2,3 compared to that of H5 and H9 influenza viruses. Such changes in sialic acid binding play important roles in novel host adaptation (18). Mutations at positions 226 and 228 (H3 numbering) in the H3 HA can also affect receptor-binding preferences and can either completely abrogate (Q226L) or reduce (G228S) affinity for the avian-preferred SA_-2,3 interaction (18, 19). Seal H3N8 virus maintains the avian phenotype at positions 226 (Q) and 228 (G), which correlates with a continued ability to use SA_-2,3. Together, these findings suggest that the seal virus may still be able to use SA_2,3, but perhaps with less efficiency than in its original FIG 2 (A and B) Fluorescent in situ hybridization (ISH) of H3N8 virus-infected seal cells with DAPI counterstaining. A probe targeting the viral hemagglutinin demonstrates diffuse infection of the bronchial mucosal epithelium. ISH was also performed using probes for the matrix gene. Staining was identical to that shown here for HA. (C and D) Lectin staining to demonstrate the distribution of SA_-2,3 and SA_-2,6 in seal pulmonary parenchyma. The SA_-2,6 (green) was detected using fluorescein-labeled Sambucus nigra agglutinin (SNA) lectin, while SA_-2,3 (red) was detected using Maackia amurensis II (MAL II) lectin. Both infected and uninfected control tissues were stained, and the results were consistent for both. High levels of SA_-2,6 are observed on bronchiole and alveolar epithelial cells and on endothelial cells. The images in panels C and D were selected because they show staining for both sialic acids; however, the expression of SA_-2,3 was rarely observed (arrows) and limited to bronchiole luminal (C) and occasional alveolar (D) epithelia. (E and F) Costaining of SA_-2,6 and H3N8 HA. SA_-2,6 (green) is expressed on the respiratory epithelium of an intrapulmonary bronchus (E). H3N8 virus-infected cells (red) are present. A serial section was also stained for SA_-2,3, and none was detected. A high-magnification image of infected mucosa clearly shows H3N8 virus infection of cells expressing SA_-2,6 (arrows). All composite images are presented separately (single stains) in Fig. S1 to S3. Fatal Avian Influenza in New England Harbor Seals July/August 2012 Volume 3 Issue 4 e00166-12 ® mbio.asm.org 3 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org avian host. Given this, we investigated whether seal H3N8 may have adapted and acquired an additional or increased affinity for the SA_-2,6 receptors that are more prevalent in mammalian respiratory tissue (20-23). The pulmonary distribution of SA_-2,3 and SA_-2,6 influenza receptors was investigated using the receptor-specific lectins Sambucus nigra agglutinin (SNA) for SA_-2,6 and Maackia amurensis lectin II (MAL II) for SA_-2,3. SA_-2,6 was widely expressed in both infected and noninfected pulmonary parenchyma, with the highest concentration seen on endothelial cells, followed by alveolar/ bronchiole epithelia (Fig. 2 and see Fig. S2 in the supplemental material). SA_-2,3 was also observed, but less frequently, and was generally limited to the luminal surfaces of epithelial cells of the bronchioles. This broadly agrees with the expression of these SA saccharides in humans and pigs (20-23) and demonstrates that seals do express receptors that would allow avian viruses to initiate infection. This observation is supported by the H3N3 seal virus from the 1991 epizootic (4), which was shown to preferentially bind SA_-2,3 in vitro (19). However, the limited prevalence of SA_-2,3 in the lower lung suggests that the process of infection is inefficient and may help to explain why epizootics of avian influenza occur but are infrequent in harbor seals. Importantly, the rare expression of SA_-2,3 is insufficient to explain the diffuse infection seen throughout the pulmonary parenchyma (Fig. 2). In contrast, the wide distribution of SA_-2,6 is far more consistent with the level of infection observed. Costaining of infected lung with viral HA and SA_-2,3 or SA_-2,6 demonstrated clear infection of SA_-2,6-positive cells, in which no SA_-2,3 was seen (Fig. 2 and S3). Hemagglutination assays were also performed to confirm sialic acid binding preferences. Seal H3N8 isolates were first sequenced to confirm that passage in eggs had not altered the HA phenotype detected in the infected tissues, and the viruses were then tested for their ability to agglutinate erythrocytes that preferentially express SA_-2,3 (horse) or SA_-2,6 (guinea pig, pig) (24, 25), relative to several avian H3N8 viruses (Fig. 6). Average agglutination titers for seal H3N8 virus with horse erythrocytes (1:48) show that the virus can still bind to SA_-2,3. However, titers were appreciably higher with guinea pig (1:192) and pig (1:144) erythrocytes, demonstrating a preference for SA_-2,6. These findings show that seal H3N8 can use both avian and mammalian receptors and add to previous studies that have demonstrated changes in receptor preferences following a host switch event (26). The patterns of SA_- 2,3 and SA_-2,6 binding to seal H3N8 virus also agree with the patterns observed for H3 avian viruses adapting to humans (18). A further mutation was observed in HA, this time at position 110 (Table 1). In avian H3 viruses, phenylalanine (Phe/F) is consistently seen, while seal H3N8 uses Ser (F110S). The significance of this (if any) is currently unknown; however, previous work has suggested that this amino acid (position 110) is a critical component of the influenza fusion peptide (27), and given the essential role of fusion in viral replication and the host-specific differences that presumably exist in this process, the F110S substitution may well represent further adaption of this virus to mammalian replication. The ability of avian influenza viruses to adapt to SA_-2,6- mediated cell entry and replication is regarded as a significant driving force in the emergence of global pandemics (19, 28-30), especially for viruses with phenotypes that confer increased virulence. Such phenotypes are often, though not exclusively, dictated by mutations in segment 1 (PB2), which is an important determinant of host range for influenza viruses. Previous studies have experimentally demonstrated the effect of various PB2 substitutions on virulence and transmissibility in mammalian hosts (15, 31-38), including the modification of the aspartic acid (D) avian phenotype to an asparagine (N) mammalian phenotype at amino FIG 3 (A) Immunohistochemistry (IHC) of seal bronchus. Polyclonal antibodies were raised against H3N8 virus-specific HA antigen. Brown staining (DAB reporter system) indicates the presence of viral antigen. There is irregularity of the mucosal surface, with sloughed epithelium. IHC demonstrates the presence of viral antigen, pyknosis, and apoptosis (arrows). No viral antigen or apoptosis was seen on negative-control tissue. (Bi) ISH staining of H3N8 virus in lung epithelium (HA probe). (Bii) TUNEL staining (green) in the same region (serial section) of the lung, showing the presence of apoptotic cells. Comparison of virus and TUNEL staining shows localization of apoptosis to virally infected cells. (Biii) DAPI (with dihydrochloride) staining for cell nuclei. Anthony et al. 4 ® mbio.asm.org July/August 2012 Volume 3 Issue 4 e00166-12 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org acid 701 (15, 31, 32, 36, 39, 40). This D701N mutation has been experimentally introduced into an adapted version of the H7N7 seal influenza virus isolated in 1982 (1, 2) and was shown to increase the pathogenicity of the virus to mice (32). The seal H3N8 virus from the 2011 outbreak has naturally acquired this D701N substitution (Table 1), which was confirmed by clonal sequencing directly from infected tissue (50 clones/animal) to be the only phenotype present in all five animals. None of the previous outbreaks of influenza in seals showed this 701N phenotype, but it is consistently found in H3N8 viruses from horses and dogs, demonstrating further adaptation to replication in mammalian hosts. These observations raise significant concern about the virulence and transmission of this virus between mammals. Interestingly, analysis of HA sequences over the course of the outbreak show the introduction and maintenance of two nucleotide polymorphisms (Table 2), and while this is insufficient to convincingly demonstrate seal-to-seal transmission, it leads us to postulate that mammalian spread might already have occurred. Together, the adaptations observed in A/harbor seal/Massachusetts/ 1/11 suggest that it may be able to persist within the seal FIG 4 Phylogenetic trees of representative influenza H3N8 genome segments. Nucleotide sequence alignments for all genome segments were created using ClustalW, and trees were produced using neighbor-joining, maximum-likelihood, and Bayesian algorithms. Models of evolution were selected usingModelTest, and a tree was selected based on a consensus of the results of the three algorithms. Only the trees for HA and NA are shown; however, all eight segments showed strong association with sequences of avian origin. Trees are constructed with H3N8 viruses only, and published sequences were selected to represent variation in the year, host, and location of isolation. *, A/blue-winged teal/Ohio/926/2002; NY, New York; LA, Louisiana. Fatal Avian Influenza in New England Harbor Seals July/August 2012 Volume 3 Issue 4 e00166-12 ® mbio.asm.org 5 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org population and evolve into a new clade within the H3N8 group, as happened with the canine and equine viruses. An additional concern is the potential zoonotic threat that this virus poses, as it has already acquired mutations in both PB2 and HA that are often, though perhaps not exclusively, regarded as prerequisites for pandemic spread (19-23, 28, 30, 37) and it is uncertain how any persistence of the virus in mammals may continue to alter its phenotype. A comparison of A/harbor seal/Massachusetts/1/11 with human H3N2 viruses revealed three substitutions that are already common to both seal H3N8 and human H3N2 viruses. These are NA-W399R, PB2-I382V, and PA-S184N (Table 1). In all cases, these substitutions are shared by the seal H3N8 and human H3N2 viruses but are not found in influenza viruses isolated previously from seals, in avian, equine, or canine H3N8 viruses, or in either seasonal or pandemic H1N1 viruses. Further studies will be required to establish the functional significance of these substitutions; however, the natural epizootic emergence at this time of a pathogenic virus that can transmit between mammals, found in a species that can become infected with multiple influenza virus subtypes, must be considered a significant threat to both wildlife and public health. MATERIALS AND METHODS Extractions, PCR, and sequencing. RNA was extracted from all tissues using Trizol reagent, and cDNA was synthesized using Superscript III (Invitrogen) according to the manufacturer’s instructions. PCR for the detection of influenza A virus was performed using primers FLUAV-MU44 (GTCTTCTAACCGAGGTCGAAACG) and FLUAV-M-L287 (GCA TTTTGGACAAAGCGTCTACG), to produce a 243-bp product of seg ment 7 (coding for matrix protein). For full-genome sequencing, fulllength cDNAs were amplified for all eight influenza segments. Primers were designed to target terminal sequences for each segment, based on alignments of avian, canine, and equine H3N8 sequences from the Influ enza Research Database (http://www.fludb.org). All PCRs were performed using fast-cycling chemistry (Qiagen), according to the manufac- TABLE 1 List of amino acid substitutions between seal H3N8 and avian H3N8 virusesa Segment (protein) nt position aa position Amino acid substitution Seal H3N8 (2011) Avian H3N8 Seal H7N7 (1980) Seal H4N5 (1982) Seal H3N3 (1992) Equine H3N8 Canine H3N8 Human H3N2 Human H1N1 (seasonal) Human H1N1 (pandemic) 1 (PB2) 178 60 N D D D D D D D D D 441 147 M I I I I V V I I T 1127 376 R K K K K K K K K K 1144 382 V I I I I I I V I I 2101 701 N D D D D N N D D D 2 (PB1) 522 174 I M M M M M M M M M 925 309 G W W W W W W W W W 1075 359 G S S S S S S S S S 1126 376 V I I I I I I I I I 1130 377 G D D D D D D D D D 1392 464 E D D D D D D D D D 3 (PA) 253 85 A T T T T T T T T I 551 184 N S S S S S S N S S 794 265 L P P T P P P P P P 4 (HA) 242 81 (65) K T G D T T T T S N 323 108 (92) S N E T N S N K N S 329 110 (94) S F S V F F F Y E D 452 151 (135) E G A K G R R T V V 527 176 (160) V A A A A S S K L S 713 238 (222) L W Q W W W L R K K 778 260 (244) M V T V V M M L I T 802 268 (252) V I I I I V V I I V 859 287 (271) N D D A D D D D N D 1114 372 K Q Q Q Q Q Q Q Q Q 1247 416 L S T E S S S S N N 5 (NP) 187 63 V I I I I I I I I I 383 128 G D D D D D D D D D 886 296 H Y Y Y Y Y Y Y Y Y 1336 446 G R R R R R R R K R 6 (NA) 440 147 E V I None I V I V V I 849 283 D E N None D E E Y T S 937 313 R G Q None G G G S G G 958 320 S P L None H P P V F F 1186 396 D N N None N N D R I I 1195 399 R W W None W W W R W W 1295 432 A E A None N E E E R K 8 (NS1/NS2) 263 88 H R R R R R R R R R a A total of 40 amino acid substitutions were observed in a comparison of seal H3N8 virus with avian H3N8 virus. Sequences of other seal influenza viruses, canine and equine H3N8 viruses, and selected human influenza viruses were included for comparison. Amino acid positions presented in parentheses represent corresponding H3 numbering. None, no sequence available for comparison; nt, nucleotide; aa, amino acid. Anthony et al. 6 ® mbio.asm.org July/August 2012 Volume 3 Issue 4 e00166-12 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org turer’s instructions. Amplified products were cloned into the pGEM T-easy vector (Promega) and sent for commercial sequencing. Virus isolation and intravenous pathogenicity index test. Homogenates from PCR-positive tissues were inoculated into SPF embryonated chicken eggs, and virus growth was determined by PCR. Tissue homogenates were also used to infect the Vero andMDCKcell lines in the presence of trypsin. Virus isolates were sent to the National Veterinary Services Laboratory (Ames, IA), where the chicken intravenous pathogenicity index test was performed according to the OIE manual (41). Molecular pathology. Fluorescent in situ hybridization (FISH) was performed using the Quantigene ViewRNA ISH tissue assay (Affymetrix), according to the manufacturer’s instructions. FISH conditions were optimized to include a 10-min boiling and 20-min protease treatment. Oligonucleotide probes were designed commercially by Affymetrix using sequences of HA and M (accession numbers JQ433879 and JQ433882, respectively). Immunohistochemistry (IHC) was performed by pretreating deparaffinized tissue sections with a 1:10 dilution of antigen retrieval solution (DAKO) for 20 min in a steamer. Samples were then washed three times in distilled water (dH2O), incubated in 3% hydrogen peroxide (in phosphate-buffered saline [PBS]) for 10 min, washed again twice in dH2O and once in PBS, and then blocked (10% normal goat serum, 0.1% bovine serum albumin [BSA]) for 20 min. Sections were treated with HA polyclonal H3N8 antibody (Novus Biologicals; catalogue number NBP1- 46796) at a 1:250 dilution for 2 h at room temperature. Following three washes in PBS, sections were incubated in Signal Stain Boost IHC reagent (Cell Signaling; catalogue number 8112) for 30 min at room temperature. Sections were again washed three times in PBS, stained with 3,3- diaminobenzidine (DAB; Dako), and counterstained with hematoxylin. TUNEL staining was performed using the in situ cell death detection kit and fluorescein (Roche) with deparaffinization and protease treatment as described for the FISH protocol. Simultaneous detection of SA_-2,3 and SA_-2,6 glycans. Deparaffinized tissue sections (5 _M) were blocked with 1_Carbo-Free solution (Vector Laboratories; catalogue number SP-5040) for 1 h at room tem- FIG 5 Structural-homology model showing the interaction of influenza HA with the SA_2,3 ligand. Seal H3 (gray), avian H5 (orange) (Protein Data Bank [PDB] accession number 1JSO), and swine H9 (pink) (PDB accession number 1JSH) were compared. The mutation G151E causes a conformational shift and interrupts H bonding between seal H3 S152 and SA_2,3, which suggests a reduction in SA_2,3 binding efficiency. A lost H bond in seal H3 is depicted in green. Fatal Avian Influenza in New England Harbor Seals July/August 2012 Volume 3 Issue 4 e00166-12 ® mbio.asm.org 7 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org perature. Sections were then stained for SA_-2,6 using fluorescein SNA (Vector Laboratories; catalogue number FL-1301) at 10_g/ml for 30 min at room temperature, and rinsed twice for 3 min each time in PBS. Sections were then reblocked in 1_ Carbo-Free solution for 30 min, before being stained for SA_-2,3 with 10 _g/ml of biotinylated MAL II (Vector Laboratories; catalogue number B-1265) for 30 min at room temperature. The MAL II was poured off, and Texas Red streptavidin (Vector Laboratories; catalogue number SA-5006) was laid over the sections at 10 _g/ml for a further 30 min. Sections were rinsed twice for 3 min each time in PBS and mounted with Vectashield hard-set mounting solution with DAPI (4=,6-diamidine-2-phenylindole) counterstaining. Hemagglutination assays. Hemagglutination assays were performed according to the WHO diagnostic manual (42). Briefly, red blood cells (RBCs) from rooster chicken, guinea pig, horse, and pig were obtained from Lampire Biological Laboratories (Ottsville, PA). The RBCs were washed in PBS and resuspended to 0.5% (chicken) or 0.75% (guinea pig and pig). Equine RBCs were resuspended to 1% in PBS with 0.5% BSA (43). Viruses were diluted to 64 HA units using chicken red blood cells. Serial dilutions were then made, added to equal volumes of washed RBCs of each species, and incubated in U-bottom plates, with the exception of chicken RBCs, which were incubated in V-bottom plates. Reaction mixtures were incubated at room temperature for 1 h, with the exception of those with chicken RBCs, which were incubated for 30 min. The HA titer endpoint is the reciprocal of the highest dilution which causes complete hemagglutination. The seal H3N8 virus was compared with several avian H3N8 isolates, including A/common eider/Massachusetts/20507-001/ 2007 (H3N8), A/northern pintail/Oregon/44249-547/2006 (H3N8), A/mallard/Washington/44338-052/2007 (H3N8), A/blue-winged teal/ Kansas/44440-003/2008 (H3N8), A/American black duck/Maine/44411- 174/2008 (H3N8), and A/American black duck/Maine/44411-532/2008 (H3N8). An H5N2 virus was also included: A/turkey/Minnesota/3689- 1551/1981 (H5N2) virus. Sequence analysis. Nucleotide sequences were aligned using ClustalW. Phylogenetic trees were constructed using neighbor-joining, maximum-likelihood, and Bayesian algorithms. Models of evolution were selected using ModelTest, and a representative tree was selected based on FIG 6 Hemagglutination assays were performed on an isolate of the H3N8 seal virus (A/harbor seal/New Hampshire/179629/2011) to confirm sialic acid binding preferences. Viruses were tested for their ability to agglutinate erythrocytes that preferentially express SA_-2,3 (horse) or SA_-2,6 (guinea pig, pig). Average agglutination titers for seal H3N8 with horse erythrocytes (1:48) show that the virus can still bind to SA_-2,3, though weakly. Titers were appreciably higher with guinea pig (1:192) and pig (1:144) erythrocytes, demonstrating a preference for SA_-2,6. Given that horse erythrocytes express SA_-2,3, it is interesting that the avian H3N8 viruses did not agglutinate with horse RBCs (red blood cells) efficiently, even following repeated attempts. Ito et al. (25) showed that avian H3N8 viruses from Asia in the early 1980s could bind to horse RBCs, while Wiriyarat et al. (43) gave examples of avian viruses (albeit not H3 viruses) that did not bind to horse RBCs. It is not known whether the avian viruses included here simply have a preference for the N-acetyl (NeuAc) sialic acid species, which is not found on horse RBCs, while the seal virus uses N-glycolyl (NeuGc) SA_-2,3. Ck, chicken; GP, guinea pig; Eq, equine; Sw, swine; 179629 Seal H3N8, A/harbor seal/New Hampshire/179629/2011; 53968 COEI H3N8, A/common eider/Massachusetts/20507-001/2007 (H3N8) virus; 16232 NOPI OR 06 H3N8, A/northern pintail/Oregon/44249-547/2006 (H3N8) virus; 52290 MALLWA07 H3N8, A/mallard/Washington/44338-052/2007 (H3N8) virus; 93866BWTEKS 08 H3N8, A/blue-winged teal/Kansas/44440-003/2008 (H3N8) virus; 96016ABDUMA07 H3N8, A/American black duck/Maine/44411-174/2008 (H3N8) virus; 96647 ABDUMEH3N8, A/American black duck/Maine/44411-532/2008 (H3N8) virus; TurkeyMnH5N2, A/turkey/Minnesota/3689-1551/1981 (H5N2) virus. TABLE 2 Sequence analysis of the HA genes isolated from various tissuesa Animal Sample Date Nucleotide at position: 1347 1499 278-Pv Kidney 28 Sept 2011 C C 286-Pv Trachea 29 Sept 2011 C C Mes LN C C Kidney C C 295-Pv Mes LN 3 Oct 2011 C C Lung C C Tonsil T C Kidney T A Tonsil 3 Oct 2011 T A Trachea T A Cerv LN T A a Two polymorphisms were observed in HA at positions 1347 and 1499, relative to avian H3N8 sequence CY041887. Isolates from animals earlier in the outbreak showed C at position 1347 and C at position 1499. The variations C1347T and C1499A were observed in animal 295-Pv, in addition to the wild-type sequence. Animal 294-Pv showed only the variant genotype. Anthony et al. 8 ® mbio.asm.org July/August 2012 Volume 3 Issue 4 e00166-12 Downloaded from mbio.asm.org on August 6, 2012 - Published by mbio.asm.org a consensus of the results of the three algorithms. Published H3N8 sequences included in the analyses were selected to represent the diversity of year, host, and location of isolation. Structural modeling. To create a homology model of the seal 2012 outbreak HA sequence, 10 template models were selected based on their super-secondary structures, with use of the LOMETS meta-threading approach (44, 45). Continuous fragments excised from these templates were then reassembled into full-length models by replica exchange Monte Carlo simulations (44). Ab initio modeling of threaded unaligned regions was then used to complete the structure. Low free-energy states were subsequently identified through clustering of simulation decoys by the SPICKER near-native model selection algorithm (46). Chimera was utilized for structural analysis and visualization (47). Hydrogen bonding analysis was based on geometric criteria established through survey of small-molecule crystal systems and Dunbrack rotamer libraries (48, 49). Nucleotide sequence accession numbers. The sequences of all eight influenza genome segments were submitted to GenBank and assigned accession numbers JQ433879 to JQ433882. SUPPLEMENTAL MATERIAL Supplemental material for this article may be found at http://mbio.asm.org /lookup/suppl/doi:10.1128/mBio.00166-12/-/DCSupplemental. Figure S1, JPG file, 0.1 MB. Figure S2, JPG file, 0.2 MB. Figure S3, JPG file, 0.2 MB. ACKNOWLEDGMENTS We acknowledge funding from the NIH: AI57158 (NBC-Lipkin), LM010140, and CA121852; NIH/NSF TW005769; USAID PREDICT; and DTRA. We thank Nicole Arrigo for editing and Wendy Barclay and Jennifer Howard for comments on the manuscript. We thank Johnice Miller for technical assistance. REFERENCES 1. Geraci JR, et al. 1982. Mass mortality of harbor seals: pneumonia associated with influenza A virus. Science 215:1129 –1131. 2. Webster RG, et al. 1981. Characterization of an influenza A virus from seals. Virology 113:712–724. 3. Hinshaw VS, et al. 1984. 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VIRUS INFLUENZA, TEMIDO Y CAMBIANTE 1986

El virus influenza, temido y cambiante ( Publicado en Revista Creces, Agosto 1986 ) Son los receptores de las células respiratorias los que interactúan con la superficie externa del virus y le abren las puertas para que este llegue hasta apoderarse de la maquinaria metabólica de la célula infectada. Lo impredecible es la forma como cambia la superficie externa del virus, lo que ha dificultado hacer una vacuna que ofrezca protección eficiente y duradera. Los españoles solían llamarle "garrotazo", y lo cierto es que la influenza, y más concretamente el virus que la provoca, representa una seria amenaza para la humanidad. Durante el transcurso del siglo pasado el virus influenza causó varias epidemias en la población mundial, siendo la más devastadora la que dejó un saldo de 20 millones de muertos en 1918. A pesar de las frecuentes vacunas que se aprueban y salen al mercado en todos los países, hoy resulta imposible evitar que pueda volver a ocurrir algo como lo de 1918. El problema tiene también consecuencias económicas porque las ausencias laborales debido a cuadros de influenza dejan saldo de millones de dólares en pérdidas cada año, perjudicando la economía de cualquier país. Y la foca estornudó El virus causa epidemias, no tan sólo en el Hombre sino también en diversas especies de vertebrados entre las cuales se encuentran aves, focas, cerdos, caballos y primates. Se han reportado casos que parecen indicar la factibilidad de contagio de una especie a otra. Es decir virus de cualquiera de estas especies podría intercambiarse en la naturaleza y generar enfermedad. Un ejemplo que merece mención lo constituye un suceso acontecido en Islandia hace algunos años. Mientras un científico y su asistente examinaban una foca infectada por el virus influenza, una mala maniobra indujo el estornudo de la foca. Todo fue a dar en la cara del tecnólogo, quien sabia que el virus había causado la muerte de cientos de focas en ese lugar. Este contagio natural podría representar una seria amenaza para su salud. Afortunadamente, 48 horas después se produjo un cuadro de conjuntivitis, se aisló el virus a partir del ojo del asistente y no hubo mayor trascendencia con respecto a la enfermedad. Transmisiones de virus similares a éstas pueden ocurrir, representando un peligro potencial para la humanidad. El virus influenza infecta principalmente células del aparato respiratorio porque éstas presentan en su membrana estructuras específicas denominadas receptores, los cuales interactúan con la superficie externa viral (figura 1A). Luego de establecido el contacto virus-célula, el virus influenza penetra al citoplasma celular y se apodera de la maquinaria metabólica de la célula infectada. Como consecuencia se producen innumerables partículas virales y la lenta destrucción celular. Nuevos virus infectan células vecinas y se va comprometiendo el tejido pulmonar. Esta necrosis en desarrollo puede producir desde síntomas de resfrío común hasta el cuadro clínico fatal que todos conocemos como influenza. Defensa El sistema inmune de defensa humano actúa mediante el reconocimiento de las proteínas superficiales del virus. La estructura de la capa externa del virus influenza puede visualizarse como una membrana dinámica y polimórfica de la cual emergen 2 tipos de glicoproteínas: la hemaglutinina y la neuraminidasa (figura 2). La hemaglutinina es la más abundante y la encargada de establecer contacto con los receptores de las células a infectar. Al igual que toda proteína, ambas son codificadas por secuencias específicas de RNA (ácido ribonucleico). El genoma (conjunto de genes de los cromosomas), en el virus influenza está compuesto por ocho segmentos discretos de RNA, cada uno de los cuales originará al menos una proteína viral. Una característica singular del proceso de replicación de todos aquellos virus cuyo genoma es RNA, es su alta frecuencia de mutación. (Replicación es el proceso según el cual una molécula origina otra idéntica a la preexistencia). No existe un mecanismo de corrección para la incorporación de nucleótidos (los productos de la ruptura del ácido nucleico), efectuada por polimerasas que copian RNA. Este fenómeno mutacional genera innumerables alteraciones en el genoma viral y, por ende, proteínas diferentes a las del virus del que derivan. Tales mutaciones que afecten los genes involucrados en la producción de hemaglutinina y neuraminidasa pueden inducir cambios en estas glicoproteínas por lo que la superficie externa de virus ya no será la misma. Alteraciones peligrosas Dos tipos de cambios es posible advertir a este respecto: Mutaciones menores y mutaciones mayores. Estas últimas llamadas también reasociaciones genéticas, (Figura 2). Las mutaciones menores se deben al cambio de un nucleótido en el gen viral, lo que genera un cambio específico de un aminoácido en la glicoproteína superficial. Este fenómeno origina una superficie viral muy similar a la del virus nativo sin mutación. La relevación de la presencia de este nuevo agente infeccioso en la población no es trascendental. Individuos infectados por el virus nativo, ya sea mediante vacunación o por contagio natural, no serán afectados seriamente por este nuevo agente causal. La acumulación de mutaciones menores puede derivar en un cambio mayor. En este caso, las diversas alteraciones experimentadas por la hemaglutinina viral la transforman en un antígeno considerablemente distinto al original. El virus portador de esta glicoproteína mutante, nueva en la población, puede causar estragos a nivel mundial. Otro tipo de mutación mayor (la reasociación genética), se produce mediante intercambio de genes entre dos virus diferentes. Algo así como si en el ejemplo del tecnólogo y la foca, el tecnólogo también hubiera estado con influenza. Podría producirse infección de una sola célula, en cualquiera de los dos, por ambos tipos de virus. Como resultado se originaría progenie viral con características que son una mezcla entre aquellas de los virus participantes. En este caso el virus mutante podría diferir considerablemente con respecto a la partícula original. Un virus influenza humano con una membrana externa de un virus de foca podría representar un serio peligro de infección a nivel mundial. Este virus mutante podría replicarse en humanos, y la hemaglutinina, derivada de la foca, representaría un antígeno nuevo contra el cual no habría inmunidad protectora en el Hombre. En resumen, la superficie externa del virus influenza está cambiando frecuentemente. Los cambios, pueden ser sutiles o drásticos dependiendo del tipo de mutación. Cualquiera sea el tipo de cambio, éstos tienden a acumularse en ciertos sitios específicos de la hemaglutinina viral (figura 3). Por último, esta alta frecuencia mutacional dificulta el desarrollo de una efectiva vacuna antiinfluenza, considerado aún como el mejor método preventivo de una enfermedad viral. Vacunas virales Una vacuna viral es un virus que ha sido atenuado o inactivado en su virulencia. El individuo vacunado será expuesto a una dosis pequeña de antígeno inmunizante, la cual inducirá protección en lugar de causar enfermedad. Las vacunas contra la influenza en uso en la actualidad son hechas de virus inactivados. Su producción comienza con el aislamiento de la última cepa infectante, a partir de secreciones nasales de individuos con influenza. El virus se hace crecer en grandes cantidades en huevos embrionados de gallina para finalmente someterlo a tratamiento de inactivación con agentes químicos como cloformo o ácidos. Esta vacuna inducirá protección de corta duración y sólo contra virus muy semejantes en su superficie externa. Otro tipo de vacuna antiinfluenza es la por virus atenuados. Consiste en hacer crecer al virus artificialmente, a temperaturas más bajas que lo habitual. De este modo se selecciona un virus mutante que es resistente al frío, incapaz de replicarse eficazmente a la temperatura fisiológica normal. Se le combina en el laboratorio con el último virus influenza aislado de la población y el resultado de esta asociación será un virus que presenta las glicoproteínas superficiales del agente que está, produciendo enfermedad, pero que es incapaz de replicarse con eficacia a 37°C, la temperatura corporal. El virus se replicará en el organismo sólo lo suficiente para inducir protección. Una aproximación más en boga al respecto son los denominados péptidos sintéticos, pequeñas cadenas de aminoácidos que representan segmentos de una proteína completa. Las proteínas muestran estructuras conformacionales complejas, producto de su secuencia de aminoácidos. Se ha postulado que aquellos segmentos de una proteína que están expuestos al medio ambiente representan los péptidos que el organismo reconocerá como antígenos. Todos estos péptidos tienen la capacidad potencial de estimular una respuesta inmune, muchas veces efectiva contra la proteína completa. En el caso del virus influenza, se conoce en detalle la secuencia y la estructura de la hemaglutinina, proteína crucial en la interacción del virus con la célula blanco. El problema reside en la identificación de aquellos péptidos que inducirán la neutralización de la partícula viral completa. En el virus influenza, debido a los cambios mencionados, esto no es fácil. Sin embargo, estudios al respecto parecen indicar la existencia de un péptido neutralizante, común para varios virus influenza de distinto origen. Si esto fuera cierto, el péptido podría producirse a muy bajo costo mediante el uso de un sintetizador automático. La vacunación con este péptido sintético induciría protección eficiente y duradera. Finalmente, los últimos intentos consideran la producción de virus con mutaciones en aquellos genes que codifican la polimerasa viral, enzima encargada de la replicación del genoma. Algo similar a lo que se está haciendo para combatir el SIDA.