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1 Department of Biology, Memorial University of Newfoundland, St John's, NL A1B 3X9, Canada
2 Institute of Arctic Biology, PO Box 757000, University of Alaska Fairbanks, Fairbanks, AK 99775, USA
Correspondence
Jonathan A. Runstadler
j.runstadler{at}uaf.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The GenBank/EMBL/DDBJ accession numbers for the sequences reported in this paper are EU086918–EU087180.
A supplementary figure showing sampling locations for this study and supplementary tables identifying viruses included in the H3 and H11 phylogenetic analyses are available with the online version of this paper.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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, 2006; Ferguson et al., 2005; Liu et al., 2005; Olsen et al., 2006; Savill et al., 2006). Alaska is an intersection of migratory bird routes originating throughout the world, including North and South America, Asia, Australia and Africa. Millions of migratory birds arrive in Alaska to breed each spring, and many of these breeding migrants are waterfowl. There has been previous work to isolate and characterize influenza viruses from waterfowl in Alaska (Ito et al., 1995; Runstadler et al., 2007; Spackman et al., 2005), and these efforts continue. The information gained about influenza viruses in these birds is important for understanding the biology of influenza viruses and their spread amongst birds from different locations. This, in turn, is important for understanding the public health risks associated with these viruses and their potential transmission to agricultural species. Unfortunately, the role of the physical and biogeochemical environment as an integral part of this transmission is poorly understood.
We recently described samples collected from an important waterfowl-breeding wetland area, the Minto Flats State Game Refuge in the interior region of Alaska (Runstadler et al., 2007). Cloacal swabs were taken from 880 ducks in the summer of 2005 and screened for the presence of influenza viruses. Over 25 % of the samples were found to be positive for influenza virus (Runstadler et al., 2007). Culturing and subsequent subtyping of viruses from a subset of these samples revealed five different haemagglutinin (HA)/neuraminidase (NA) subtypes (H3N6, H3N8, H4N6, H8N4 and H12N5). This work demonstrated an overall high rate of virus infection in birds at this location and showed that a diverse array of subtypes was present. This particular location was also the focus of a previous study between the years 1991 and 1994 (Ito et al., 1995), where six virus subtypes were isolated from 391 waterfowl faecal samples from Minto Lake.
Here, we report efforts to characterize the occurrence and diversity of influenza viruses in the sediments of ponds used by a wide variety of migratory waterfowl. Located within the city of Fairbanks, Alaska, is the Creamer's Field Migratory Waterfowl Refuge, a location that is used heavily by migratory waterfowl during both the spring and fall migration periods. The waterfowl, regularly numbering in the thousands each day, are largely concentrated at three small ponds; samples were collected from these three ponds, beginning in the fall migration period of 2005, through the winter and into the spring migration period of 2006. We used RT-PCR to target the matrix (M) and HA genes of influenza viruses in RNA extractions from the sediment samples. The diversity of sequences in these samples was extremely high and included four different HA subtypes. Most, but not all, of the HA sequences were most similar to viruses isolated from ducks in Alaska in 2005. This study demonstrates that environmental sampling is a valuable technique to assess the diversity of influenza viruses in specific geographical or environmental locations to complement other approaches, but without the need for more difficult and time-dependent bird sampling and screening of cloacal swabs by real-time PCR or culturing. Viruses were readily detectable in samples collected in the middle of winter and early spring before the arrival of migrants. Therefore, as has been suggested for lake water (Webster et al., 1992), viruses could be persisting in sediments and be a source of infection for new birds in subsequent years.
A recent summary of influenza infections by subtype in different wild bird species (Olsen et al., 2006) shows a predilection for some viral subtypes in specific groups of mammals and birds. If, particularly in the case of migratory birds, animals are dispersed in time from a potential ‘hot zone’ of virus infection, such as a migratory stopover, then the environmental persistence of viruses may play a strong ecological role in transmission. Animals utilizing an area where persistence in environmental reservoirs is possible may experience increased viral exposure and therefore greater potential for infection and also reassortment. The behaviour of the virus outside the host could therefore play a major role in interspecies infection and viral ecology and evolution (Kuiken et al., 2006).
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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RNA was extracted from 2 g sediment sample by using an RNA PowerSoil Total RNA isolation kit (MoBio) according to the manufacturer's recommendations. The resulting nucleic acid eluate was then treated with 1 unit RNase-free DNase I (New England Biolabs) according to the manufacturer's instructions, followed by extraction with buffered phenol/chloroform (1 : 1) and ethanol precipitation (Sambrook & Russell, 2001). The precipitated RNA was dissolved in 20 µl DEPC-treated water (Ambion) and stored at –80 °C.
Amplification of influenza virus sequences.
RNA isolated from sediments was screened for the presence of influenza virus sequences by RT-PCR targeting part of the M gene. The primers M52C (5'-CTTCTAACCGAGGTCGAAACG-3') and M253R (5'-AGGGCATTTTGGACAAAKCGTCTA-3') (Fouchier et al., 2000) were used to amplify a 244 bp product from the M1 portion of the M segment. For the HA gene, the primers HA-1134F (5'-GGAATGATHGAYGGNTGGTATGG-3') (Phipps et al., 2004) and Bm-NS-890R (5'-ATATCGTCTCGTATTAGTAGAAACAAGGGTGTTTT-3') (Hoffmann et al., 2001) were used to amplify an approximately 640 bp region of the HA-2 portion of the HA segment. The above primers were used in RT-PCR with the SuperScript III One-Step RT-PCR system with Platinum Taq DNA polymerase (Invitrogen). The reactions contained 1 µl RNA, each primer at 0.4 µM and 1 µl enzyme mix in 1x buffer in a total volume of 25 µl. For M sequence amplification, the thermal cycler conditions were 42 °C for 30 min, then 94 °C for 4 min, followed by 35 cycles of 94 °C for 1 min, 45 °C for 1 min and 72 °C for 1 min, and a final 7 min incubation at 72 °C. For HA sequence amplification, the thermal cycler conditions were 42 °C for 30 min, then 94 °C for 4 min, followed by 35 cycles of 94 °C for 1 min, 45 °C for 1 min and 72 °C for 2 min, and a final 7 min incubation at 72 °C.
For the HA sequence amplification reactions, a second round of PCR was used to increase the amount of material available for analyses. These reactions were performed with either the same primers as were used in the RT-PCR (HA-1134F and Bm-NS-890R) or with primer HARKs, a shortened version of the HA-specific reverse primer HAR K (Bragstad et al., 2005) with non-influenza sequences removed (5'-AGTAGAAACAAGGCTGTTTT-3'), in place of Bm-NS-890R. Both primer sets were used in parallel and the reactions that gave better amplification were used for the subsequent cloning step. The entire initial RT-PCR product was run on a 1 % agarose gel and DNA was visualized with ethidium bromide staining. Plugs were removed from bands at the correct location on the gel (approx. 640 bp) with a sterile Pasteur pipette and collected in a sterile microfuge tube containing 100 µl sterile distilled water. The tubes were then heated at 80 °C for 20 min and 50 µl liquid was collected. This eluate was used for a second round of PCR with the following conditions: 1 µl eluate, each primer at 0.4 µM, 0.5 units Platinum Taq DNA polymerase, 0.4 mM each dNTP, 2 mM MgCl2 and 1x Platinum Taq polymerase buffer in a final volume of 25 µl. The thermal cycler conditions were 94 °C for 5 min, followed by 35 cycles of 94 °C for 1 min, 45 °C for 1 min and 72 °C for 1 min, and a final 7 min incubation at 72 °C.
During the initial screening of the RNA extracts for the M gene, a control reaction was also performed with every sample in parallel to test for potential inhibition of the PCR by factors in the purified RNA. The control reactions contained primers targeting small subunit rRNA, uni-for (5'-TGCCAGCAGCCGCGGTA-3') and uni-rev (5'-GACGGGCGGTGTGTACAA-3') (Vaisvila et al., 2001). It was assumed that every RNA extraction from the sediment samples would contain at least bacterial rRNA; therefore, a negative result in this reaction was taken as evidence of failure of the RT-PCR process itself and not necessarily a genuine influenza-negative sample.
The methods used here to amplify viral sequences with RT-PCR products do introduce errors into the final cloned sequences (Bracho et al., 1998), and these cannot be distinguished specifically from mutations introduced by the viral RNA polymerase, which has an estimated error rate of 2x10–3 per position per generation (Webster et al., 1992).
DNA cloning and sequencing.
The RT-PCR and PCR products were cleaned with a MinElute PCR purification kit (Qiagen) and cloned with the TOPO-TA system (Invitrogen) according to the manufacturers' recommendations. The resulting colonies were screened by colony PCR according to the manufacturer's directions (Invitrogen). Products of interest from the colony PCRs were sequenced with a BigDye Terminator v. 3.1 cycle sequencing kit (Applied Biosystems) according to the manufacturer's recommendations. The sequencing reactions were analysed on an ABI 3100 sequencer (Applied Biosystems) at the University of Alaska Fairbanks DNA Core Facility.
Sequence analyses, alignments and phylogenetics.
The nucleotide sequences of all M and HA clones analysed in this study have been deposited in GenBank under accession numbers EU086918–EU087180. Sequences were compared with those in GenBank by using the MEGABLAST BLASTN algorithm (Altschul et al., 1990). Nucleotide sequence alignments were done with CLUSTAL_X v. 1.81 (Chenna et al., 2003; Thompson et al., 1997) and these alignments were used for subsequent phylogenetic analyses. Bayesian maximum-likelihood trees were constructed with MrBayes v. 3.1.2 (Huelsenbeck & Ronquist, 2001) and 4 000 000 generations. Neighbour-joining trees were constructed with PAUP v. 4.0 (Swofford, 2000) and bootstrap values were calculated based on percentages of 10 000 replicates.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Ten of these samples (11 %) did not produce a product with the rRNA control primers and so we assume that these were inhibited for RT-PCR (these samples are still included in Table 1). Of the 81 remaining samples, 45 (55.6 %) were positive for the influenza virus M gene by RT-PCR screening. We cloned the M RT-PCR products from 12 of these positive samples and sequenced at least nine different clones from each cloned sample, with a total of 145 clones sequenced overall. In the 145 clones, we found 52 different sequences. Some sequences differ from each other at only a single base position, and it is possible that some of the observed differences in sequences resulted from errors introduced by the polymerase enzymes (Bracho et al., 1998).
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). In addition, multiple samples were sometimes collected on the same day from the same pond and some of these samples tested positive by the RT-PCR assay, whilst others were negative for the virus.
HA sequence prevalence and diversity
We amplified and cloned the HA gene RT-PCR products from 11 different samples. At least five clones were sequenced from each cloned sample, with a total of 116 clones sequenced. Within these 116 clones, there were 76 different nucleic acid sequences (65.5 %) and 44 different predicted protein sequences (37.9 %). Some sequences differ from each other at only a single base position, and it is possible that some of the observed differences in sequences resulted from errors introduced by the polymerase enzymes (Bracho et al., 1998).
Four HA subtypes were identified by molecular methods in these sequences: H3, H8, H11 and H12 (Tables 1 and 2; Fig. 1). The abundance of sequences recovered in the clones was H3>H12>H11>H8, with H3 and H12 together comprising approximately 84 % of the cloned sequences. Only one clone containing an H8 sequence was recovered. The H3 subtype was found in samples from all days that were analysed, and H12 was found on all days analysed except one. No distinct seasonal patterns were identified from the HA subtypes (Tables 1 and 2). In fact, when we looked at samples collected from the same pond on the same day (pond 3; 26 September 2005), we identified different subsets of viral subtypes; only the H11 subtype was found in the clones from one sample, whilst the other contained subtypes H3, H11 and H12. Thus, it seems that there is a persistent and diverse collection of influenza viruses in the ponds, including over the winter months when the ponds are frozen and covered in snow.
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Relationships of HA sequences to those of previously characterized viruses
We used our HA sequences to search against sequences in GenBank to identify the subtype group to which each belonged. We then used the sequences from the appropriate subtype group for phylogenetic analyses to examine the relationships of our sequences to those from previously characterized viruses (Figs 2 and 3). Incomplete sequences (relative to the HA-2 RT-PCR product) were excluded. In cases where more than one of our sequences had exactly the same nucleotide sequence, only one of these was included in the analyses for simplicity.
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) and another four are from Alberta, Canada. The single sequence from Asia falls within the North American sequences and so our analyses do not suggest that there is any geographical pattern to the relationships of these H8 sequences (Fig. 2). The sequence that we amplified branches most closely with a virus isolated from a mallard in Alaska in 2005, with strong support. The continued, yet rare, occurrence of this subtype will make reports of it valuable to understanding its role in influenza viral ecology.
H3.
There are many H3 sequences in GenBank. Of these, we used the top 100 sequences that were returned from a BLAST search with one of our sequences for the initial analyses. The sequences formed four strongly supported, independent clades (Fig. 3a), based on the Bayesian and neighbour-joining analyses. All of our RT-PCR-derived sediment sequences fall in the clade labelled ‘1’. All of the sequences in this clade except one are from North America (Fig. 4; Supplementary Table S1, available in JGV Online). The other clades show some geographical patterns, with all of the sequences in the clade labelled ‘2’ originating in North America and including two viruses isolated from swine and one virus isolated from a seal (Supplementary Table S1). The viruses in the clade labelled ‘3’ are from all over the world and those in clade ‘4’ are from Eurasia (Supplementary Table S1).
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). The sequences amplified from the sediments form three distinct clades within this larger clade, and all of these sequences fall into a larger, well-supported clade (by the Bayesian analysis) that is made up of viruses from a variety of locations across North America. The majority of the new sequences cluster in a strongly supported clade with six viruses isolated from mallards and pintails in Alaska in 2005. Another group of sediment sequences cluster strongly with another virus isolated from a pintail in Alaska in 2005. The third set of sediment sequences forms a distinct clade that does not cluster directly with any specific virus sequence in GenBank. These sequences are all from a single sediment sample (82405), but note that not all of the sequences from this particular sample are in this clade.
H12.
There were 19 H12 virus sequences in GenBank. Phylogenetic analyses show that the H12 sequences fall into two distinct, well-supported clades, with one clade being from North America and one from outside North America (Fig. 3b). Within the North American clade, the sediment sequences form a strongly supported clade with two viruses isolated from pintails in Alaska in 2005 (Fig. 5).
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; Supplementary Table S2). The H11 sediment sequences form a distinct cluster within a larger clade of sequences from a variety of species and locations in North America (Fig. 6).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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; Sivanandan et al., 1991; Zhang et al., 2006), but we have revealed the greatest sequence diversity to date in a single location and with a limited amount of total sequencing. The power of this technique as a way to study the diversity of influenza viruses present in an environmental location, and therefore present in birds that use the location, is well illustrated by one of the samples collected from pond 2 on 1 March 2006. In the RNA extracted from 2 g sediment, we were able to detect four different HA subtypes after sequencing only 10 clones.
We found a remarkable amount of M gene diversity with limited sequencing. The M primers that we used yield 200 bp internal sequence in a well-conserved region of the virus genome, and this region is a frequent target of PCR-based influenza assays (Fouchier et al., 2000; Runstadler et al., 2007). Therefore, with this RT-PCR technique, we are detecting populations of viruses with a high degree of diversity in the environmental samples. The results of the M gene screening of the samples also suggest that the distribution of viruses in the ponds is very patchy. There are instances where multiple samples were collected on the same day and both positive and negative results were found in different samples. This could result from patchy deposition of virus-containing faeces and subsequent lack of mixing, uneven degradation of virus particles, or other biogeochemical factors that could inhibit detection in some of the samples. Given this pattern, future studies should aim to collect a higher density of individual samples from a given location (e.g. five samples from each pond) each time that samples are taken.
In the RT-PCR assays that we used, the M gene target provided much better reliability, in terms of sensitivity and specificity, than the HA gene. This is probably due to several factors, such as the smaller size of the target (200 versus 600 bp) and better conservation of the primer target sequences. This makes the M gene assay ideal for initial screening. However, the sequence amplified in this case is much less informative about the viruses present. The HA RT-PCR was technically more problematic. Multiple rounds of amplification were usually required to obtain enough material for efficient cloning of the products. Multiple bands at sizes other than that expected, i.e. approximately 640 bp, were amplified frequently, and non-influenza products that were the same size as the expected influenza sequences were occasionally amplified, cloned and sequenced. These non-influenza sequences were fungal and bacterial in origin, based on BLAST searches of GenBank. The use of more specific primer pairs that target specific subsets of HA subtypes may alleviate some of these problems.
For the HA-2 results, we found a high diversity in the sequences recovered in the clone libraries, with 76 of 116 sequences (65.5 %) being unique. There were four different subtypes found, of which H3 and H12 were the most abundant. The H3 sequences showed the greatest overall abundance and diversity in our study, and this is also true of virus sequences present in GenBank (Figs 3a, 4) for these four subtypes. A more temporally intensive sampling scheme, combined with improving data on bird populations and movements, may be able to determine whether viral diversity detected in our sequences has any relation to the number and type of birds present at a given time or is characteristic of the viral populations in circulation at the time.
Unlike the other subtypes, there have been no H11 sequences reported previously from Alaska. Therefore, this culture-independent RT-PCR method of looking for avian influenza in sediments was able to recover a subtype that was not found in a large-scale sampling of waterfowl species (Runstadler et al., 2007). Creamer's Field is a migratory stopover site that attracts a great diversity of waterfowl, and also a great diversity of shorebirds and passerine species in close proximity. Perhaps these sequences originated from viruses deposited in the Creamer's Field sediments by species that have not been sampled (heavily enough or at all) in other studies, or the culturing techniques used previously were less sensitive to this particular virus subtype. This second explanation seems less likely, because many H11 viruses have been cultured successfully in the past. Alternatively, we could be detecting viruses that were deposited in the sediments in previous years and this subtype may not have been prevalent in birds in 2005. In addition, direct sampling of birds is limited to times at which birds can be captured efficiently and effectively; the viruses identified from such samples are thus temporally limited. Therefore, the H11 sequences that we obtained may have been from a virus that was prevalent in populations earlier in the season, but which had declined by the time of heavy waterfowl sampling. Moreover, they could represent viruses shed from birds that used locations other than the Minto Flats area.
The role of abiotic reservoirs in perpetuation and transmission of influenza viruses is unclear (Webster et al., 1992), but viruses have previously been isolated from and detected in water samples. Viruses were cultured successfully from 12 of 102 water samples collected between 1992 and 1994 from lakes in Alaska with breeding waterfowl (Ito et al., 1995). Three different subtypes were identified in these water samples. This shows that abiotic sources such as water (and sediment) could be acting as reservoirs of active viruses that can infect further birds. It has also been reported recently that influenza viruses have been detected in lake ice and water in Siberia (Zhang et al., 2006). This study only reported viruses with the H1 subtype, and perhaps the more general primers used here would allow the detection of other subtypes, if present. There was a high degree of diversity in the H1 sequences found, with 83 unique sequences recovered. These H1 sequences showed a monophyletic pattern, similar to results with our H12 and H11 sequences (Figs 5 and 6), but very distinct from the polyphyletic pattern that we found for the H3 sequences (Fig. 4). There are four H3 and three H11 virus sequences in GenBank included in our analyses that were isolated from environmental samples. The H11 environmental isolates all cluster strongly with two viruses isolated from shorebirds (Fig. 6), but the H3 viruses cluster most closely with viruses isolated from ducks and the sediment sequences reported here (Fig. 4). A strong set of environmental data could contribute a temporal and spatial scale for virus distributions, so that the identification of a pathogenic virus in a bird could be used to map likely contact with other species in other locations.
A long-term study of the different HA subtypes that are found in these ponds would be ideal for understanding the influx of novel virus subtypes and possible passage of others out of the system. It may be possible to use these sediments as an archive of historical influenza virus diversity in birds using these ponds by doing linear depth profiles with cores. Viable viruses can be found in old sediments (up to 40 cm depth) in marine systems (Lawrence et al., 2002), and they may be equally well preserved here. Even if not viable, characterizing the diversity present over past time would be extremely valuable for understanding the ecology of influenza viruses in Alaskan waterfowl. Some sediments can harbour extremely high amounts of free DNA (Dell'Anno & Danovaro, 2005) but, to our knowledge, similarly abundant free RNA has not been found in natural systems. We would not expect extracellular RNA to be stable in these sediments, where there is an abundant microbial community that would rapidly degrade any free RNA. Therefore, we interpret the amplification of influenza virus sequences to result from RNA extracted from intact virus particles, although we have not demonstrated the presence of infectious particles.
A limitation of this approach arises from the segmented nature of the influenza virus genome. RT-PCR can only provide information about a single gene sequence at a time, and therefore it is not possible to know what combinations of gene segments are in the actual viruses without culturing. It is clear that the combinations of gene segments in viruses are both important and dynamic (Brown et al., 1998; Ghedin et al., 2005; Hatchette et al., 2004; Holmes et al., 2005; Spackman et al., 2006; Webster et al., 1992). It is also unclear whether there might be a bias with the HA-2 primers for which subtypes are amplified well, but they have been used to amplify sequences from all of the subtypes H1–H15 (Phipps et al., 2004). In cases where there are mixtures of viruses, it is possible that some sequences would amplify better than others. This may explain the lack of any H4 subtype sequences in our clone collections, despite the fact that H4 viruses were prevalent in the species of waterfowl using Creamer's Field in 2005 (Runstadler et al., 2007). Alternatively, it could be that H4 subtypes are less stable in the sediments and do not persist as well, or were shed less prolifically than the other subtypes that we did detect.
Application of this culture-independent RT-PCR approach to more samples and locations will greatly increase our understanding of the prevalence and diversity of the influenza viruses circulating in host populations and the environments that they use. It could readily be expanded to cover analysis of more viral gene segments. In conjunction with the culturing of viruses, both from birds and from environmental samples, this approach will offer significant contributions to understanding influenza virus ecology and evolution.
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ACKNOWLEDGEMENTS |
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Received 10 August 2007;
accepted 14 September 2007.
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