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Short Communication |
Department of Primary Industries and Fisheries, Ritchie Building 64A, Level 3, C Wing, Research Road, St Lucia, QLD 4072, Australia
Correspondence
Paul Francis Horwood
paul.horwood{at}dpi.qld.gov.au
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ABSTRACT |
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The GenBank/EMBL/DDBJ accession numbers for the sequences reported in this paper are EF108221–EF108225, EU266069 and EU277658.
Supplementary figures are available with the online version of this paper.
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MAIN TEXT |
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TOPABSTRACTMAIN TEXTREFERENCES |
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). Other viruses in this group include human parainfluenza virus types 1 and 3 (HPIV-1 and HPIV-3) and Sendai virus. The clinical presentation of bovine infections with BPIV-3 can vary considerably, ranging from asymptomatic infections to severe respiratory illness. In the majority of cases where BPIV-3 is implicated in disease, mild clinical signs characterized by coughing, fever and nasal discharge are observed (Dinter & Morein, 1990). In some instances where animals are also subjected to high stress, infection with BPIV-3 can contribute to tissue damage and immunosuppression, resulting in severe bronchopneumonia from secondary bacterial infections (Haanes et al., 1997). The resulting disease is known as bovine respiratory disease complex (BRDC) and is considered the most significant illness associated with feedlot cattle in the USA (Snowder et al., 2006) and possibly worldwide. Other respiratory viruses such as bovine herpesvirus 1, bovine viral diarrhea virus (BVDV) and bovine respiratory syncytial virus (BRSV) have also been associated with BRDC development in feedlot cattle.
The complete nucleotide sequences for three isolates of BPIV-3 have previously been determined (Sakai et al., 1987; Bailly et al., 2000). The variation between these isolates is limited, with greater than 92 % nucleotide identity across the entire genome, and reflects similar levels of conservation displayed between HPIV-3 isolates. In this study, sequence analyses of the matrix (M) protein coding region for seven Australian isolates of BPIV-3 and the complete genome of a representative isolate, indicated that this viral species can be classified into two distinct genotypes, BPIV-3 genotype A (BPIV-3a) and BPIV-3 genotype B (BPIV-3b). Strains of BPIV-3 and HPIV-3 have previously been described with sequence polymorphisms (Coelingh & Winter, 1990; Swierkosz et al., 1995) and different characteristics in cell culture (Breker-Klassen et al., 1996; Shibuta et al., 1983). However, the low degree of nucleotide variation reported did not identify distinct groups of BPIV-3.
Seven BPIV-3 isolates (Table 1) from clinical samples and historical collections were grown in CRIB-1 cells (Flores & Donis, 1995) using standard methods. Viral nucleic acids were extracted from 200 µl of cell culture supernatant using the High Pure Viral Nucleic Acid Extraction kit (Roche) and stored at –80 °C until required. The failure of in-house BPIV-3 RT-PCR methods to yield expected products for four of the isolates (Q5592, BP4158, BP4169 and BP6128) prompted further investigations into the characteristics of these viruses.
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Primers specific to the M-protein gene of BPIV-3 were utilized to generate sequence data for phylogenetic analysis. The primers RspV1 (5'-GATCAGGAACTCTTAAAGGC-3') (Halpin, 2000) and MR2 (5'-TTTTCCCGACCCCTTCTAT-3') amplified a 739 bp product using the following RT-PCR conditions. Viral RNA was reverse transcribed in a 20 µl reaction mix, containing 5 µl nucleic acid sample, 1x StrataScript buffer (Stratagene), 20 U StrataScript RT (Stratagene), 5 U RNasin (Promega), 0.5 mM dNTPs (Promega), 1 µM each primer and sterile deionized water, at 42 °C for 60 min followed by 90 °C for 5 min. PCR amplification was carried out in a 50 µl reaction volume, containing 3 µl cDNA, 1 µM each primer, 25 µl HotStar Taq Mastermix (Qiagen), and sterile deionized water. The PCR cycling parameters were: 95 °C denaturing for 15 min, followed by 30 cycles of 95 °C for 1 min, 50 °C for 1 min and 72 °C for 1 min, and a final extension cycle at 72 °C for 10 min. The amplification products were resolved by 1 % (w/v) agarose gel electrophoresis.
RT-PCR with the RspV1 and MR2 primers produced an amplicon, consistent with the expected size of 739 bp, from all of the BPIV-3 isolates tested. Gel-purified amplification products were ligated into the pGEM-T Vector System (Promega) and subsequently cloned according to the manufacturer's instructions. The nucleotide sequences of plasmid inserts were determined by automated DNA sequencing using BigDye Terminator chemistry (ABI BigDye Terminator version 3.1). Contiguous nucleotide sequences were assembled using the computer software Sequencher (GeneCodes) and all nucleotide positions were confirmed by three or more independent sequencing reactions in both directions. Putative amino acid sequences were generated following BLASTX search routines (Altschul et al., 1990). BPIV-3 sequences were retrieved from GenBank (Table 1
) for phylogenetic analyses. Nucleotide and amino acid alignments were generated using the computer program GeneDoc (Nicholas et al., 1997).
Sequence analyses of the region amplified by the RspV1 and MR2 primer pair revealed that three of the isolates, BPI3JCU, BP6121 and BP7393 displayed greater than 93 % nucleotide identity when compared to the BPIV-3 type strain Kanas/15626/84 (Ka). In contrast, the nucleotide identity between the remaining four isolates sequenced (Q5592, BP4158, BP4169 and BP6128) and the Ka type strain varied from 83.9 to 84.5 %. This group of four isolates displayed greater than 79 unique base changes from previously reported BPIV-3 isolates and the BPI3JCU, BP6121 and BP7393 isolates that were also sequenced as part of this study. The level of conservation within this group of four isolates was high, with greater than 98 % nucleotide identity between isolates. Alignment of the putative amino acid sequences demonstrated a much higher conservation than nucleotide sequences between all BPIV-3 isolates and also the HPIV-3 isolates that were included in the alignment. The identity between the putative amino acid sequences of the BPIV-3 isolates was greater than 95 %, and greater than 93 % when BPIV-3 isolates were compared to the HPIV-3 isolates.
To ascertain if the observed nucleotide sequence variation was universal throughout the BPIV-3 genome, and not limited to the M-gene, the complete genome sequence was determined for the isolate Q5592. Twenty-two primer sets were designed to amplify overlapping regions of the complete BPIV-3 genome. Primers were designed manually using the complete genome alignments of BPIV-3 (AF178654
[GenBank]
, AF178655
[GenBank]
and D84095
[GenBank]
) and HPIV-3 (Z11575
[GenBank]
and NC_001796
[GenBank]
) sequences from GenBank. Genomic leader and trailer sequences were derived using the 5'–3' RACE kit (Roche) according to the manufacturer's instructions. Contiguous nucleotide sequences were assembled using the computer software Sequencher and all nucleotide positions were confirmed by three or more independent sequencing reactions for both cDNA strands. The complete Q5592 genome sequence was compiled from overlapping sequences of the Q5592 amplicons. Comparative analysis with the BPIV-3 strain Ka genome sequence from GenBank was used to identify coding regions, and putative amino acid sequences were generated following BLASTX search routines (Altschul et al., 1990). Nucleotide and putative amino acid sequence alignments were generated manually using the computer program GeneDoc with BPIV-3 and HPIV-3 sequences retrieved from GenBank (Table 1).
The Q5592 isolate varied considerably from the previously characterized BPIV-3 strains in all of the coding and non-coding regions of the genome (Table 2). The range of nucleotide identities for the BPIV-3 coding regions between Q5592 and other BPIV-3 strains was 82.0–85.8 %, whereas the range of nucleotide identities between the type strain Ka and other BPIV-3 strains was 91.7–98.8 %. The nucleotide identities for the complete genome were 82.6–83.1 % between Q5592 and other BPIV-3 strains, and 92.3–98.2 % between Ka and other BPIV-3 strains.
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). The longer genome length of the Q5592 isolate was due to two unique inserts, totalling 30 nt, in the untranslated region (UTR) between the fusion (F) and haemagglutinin/neuraminidase (HN) glycoproteins, and another unique insert of 12 nt that was present in the UTR between the M and F proteins.
Nucleotide sequence alignments of the M-protein coding regions and complete genomes were used for phylogenetic analyses using the computer program MEGA 3.1 (Kumar et al., 2004). Phylogenetic trees were determined by bootstrap analysis (500 replicates) using the neighbour-joining program with the Kimura two-parameter method for nucleotide data analysis. Phylogenetic reconstructions based on the alignment of the M-gene nucleotide sequences demonstrated that four of the isolates, Q5592, BP4158, BP4169 and BP6128, clustered into a very distinct group. This group was readily identified from the other BPIV-3 isolates and also from HPIV-3. The clustering is clearly demonstrated in the radiation-style tree illustrated in Fig. 1, where BPIV-3b is distinct from the previously characterized members of BPIV-3 represented by genotype A. Phylogenetic reconstructions based on the nucleotide sequences of BPIV-3 and HPIV-3 complete genomes demonstrated that the Q5592 isolate, representing BPIV-3b, formed a distinct lineage separate from the previously identified BPIV-3 and HPIV-3 groups (Fig. 1). The phylogenetic tree produced from the complete genome nucleotide sequence was almost identical to the phylogenetic tree produced from the M-gene nucleotide data (Fig. 1). This finding supports the use of the M-gene of PIV-3 as an appropriate target for the generation of informative phylogenetic reconstructions.
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; Bailly et al., 2000). To date, this study is the most comprehensive sequencing survey of clinical BPIV-3 isolates. Therefore, it is difficult to accurately assess whether the level of genetic variation identified here is typical of the BPIV-3 species as a whole, or if genotype B is an isolated variant lineage. However, it is interesting that genotype B has only been identified in Australia, which is geographically removed from other cattle breeding countries, and enforces strict quarantine measures. The recognition of a previously unidentified genotype of BPIV-3 is reliant upon sequence analysis of a greater number of BPIV-3 isolates from a wide geographical distribution. The validity of the two proposed genotypes for BPIV-3 may also require further immunological validation to determine their significance for vaccine development purposes, as has been recognized for the BRSV subgroups (Schrijver et al., 1997). However, even in the absence of significant immunological differences, variant genotypes from other viruses have been implicated in playing significant roles in disease. An example of this is BVDV, where there are seven recognized genotypes based on nucleotide sequence analyses (Mahony et al., 2005). Despite the large number of genotypes, there appears to be a correlation between particular genotypes and the development of BRDC in feedlot cattle (Fulton et al., 2005). The recognition of sequence variability, and hence two distinct BPIV-3 genotypes, is also significant because current RT-PCR methods have specifically been designed to detect BPIV-3 on the basis of existing sequence data and provides the basis for generating false-negative data.
The BPIV-3b genotype could hypothetically be a lineage from a strain that recently crossed from another host species into cattle. PIV-3 infections have been serologically demonstrated in a wide variety of mammals including cattle, humans, sheep (Lyon et al., 1997), goats (Yener et al., 2005), bison (Zarnke & Erickson, 1990), guinea pigs (Ohsawa et al., 1998), black and white rhinoceros (Fischer-Tenhagen et al., 2000), moose (Thorsen & Henderson, 1971), bighorn sheep (Parks et al., 1972) and camels (Eisa et al., 1979). Cross-species infections have been reported in numerous instances including HPIV-3 in guinea pigs (Ohsawa et al., 1998), BPIV-3 in a human (Ben-Ishai et al., 1980), BPIV-3 in sheep and ovine PIV-3 in cattle (Stevenson & Hore, 1970). Further sequence analysis of BPIV-3 isolates from a larger number of host species will be essential to fully elucidate the relationship between the various host strains of PIV-3. The relative divergence of the BPIV-3b genotype when compared with BPIV-3a and HPIV-3 suggests that viruses from this genotype may play an important role in further studies to elucidate host-specificity and viral gene function in the genus Respirovirus.
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Bailly, J. E., McAuliffe, J. M., Skiadopoulos, M. H., Collins, P. L. & Murphy, B. R. (2000). Sequence determination and molecular analysis of two strains of bovine parainfluenza type 3 that are attenuated for primates. Virus Genes 20, 173–182.[CrossRef][Medline]
Ben-Ishai, Z., Naftali, V., Avram, A. & Yatziv, S. (1980). Human infection by a bovine strain of parainfluenza virus type 3. J Med Virol 6, 165–168.[Medline]
Breker-Klassen, M. M., Yoo, D. & Babiuk, L. A. (1996). Comparisons of the F and HN gene sequences of different strains of bovine parainfluenza virus type 3: relationship to phenotype and pathogenicity. Can J Vet Res 60, 228–236.[Medline]
Coelingh, K. V. W. & Winter, C. C. (1990). Naturally occurring human parainfluenza type 3 viruses exhibit divergence in amino acid sequence of their fusion protein neutralization epitopes and cleavage sites. J Virol 64, 1329–1334.
Dinter, Z. & Morein, D. (editors) (1990). Virus Infections in Ruminants. New York: Elsevier Science Publishers BV.
Durbin, A. P., Siew, J. W., Murphy, B. R. & Collins, P. L. (1997). Minimum protein requirements for transcription and RNA replication of a minigenome of human parainfluenza virus type 3 and evaluation of the rule of six. Virology 234, 74–83.[CrossRef][Medline]
Eisa, M., Karrar, A. E. & Abdel Rahim, A. H. (1979). The occurrence of antibodies to parainfluenza 3 virus in sera of some domestic animals of the Sudan. Br Vet J 135, 192–197.[Medline]
Fischer-Tenhagen, C., Hamblin, C., Quandt, S. & Frolich, K. (2000). Serosurvey for selected infectious disease agents in free-ranging black and white rhinoceros in Africa. J Wildl Dis 36, 316–323.[Abstract]
Flores, E. F. & Donis, R. O. (1995). Isolation of a mutant MDBK cell line resistant to bovine viral diarrhea virus infection due to a block in viral entry. Virology 208, 565–575.[CrossRef][Medline]
Fulton, R. W., Ridpath, J. F., Ore, S., Confer, A. W., Saliki, J. T., Burge, L. J. & Payton, M. E. (2005). Bovine viral diarrohea virus (BVDV) subgenotypes in diagnostic laboratory accessions: distribution of BVDV1a, 1b and 2a subgenotypes. Vet Microbiol 111, 35–40.[CrossRef][Medline]
Haanes, E. J., Guimond, P. & Wardley, R. (1997). The bovine parainfluenza virus type-3 (BPIV-3) hemagglutinin/neuraminidase glycoprotein expressed in baculovirus protects calves against experimental BPIV-3 challenge. Vaccine 15, 730–738.[CrossRef][Medline]
Halpin, K. (2000). Genetic studies of Hendra virus and other novel paramyxoviruses. PhD thesis, University of Queensland, Australia.
Kumar, S., Tamura, K. & Nei, M. (2004). MEGA3: integrated software for molecular evolutionary genetics analysis and sequence alignment. Brief Bioinform 5, 150–162.
Lyon, M., Leroux, C., Greenland, T., Chastang, J., Patet, J. & Mornex, J. F. (1997). Presence of a unique parainfluenza virus 3 strain identified by RT-PCR in visna-maedi virus infected sheep. Vet Microbiol 57, 95–104.[CrossRef][Medline]
Mahony, T. J., McCarthy, F. M., Gravel, J. L., Corney, B., Young, P. L. & Vilcek, S. (2005). Genetic analysis of bovine diarrhoea viruses from Australia. Vet Microbiol 106, 1–6.[CrossRef][Medline]
Murphy, F. A., Fauquet, C. M., Bishop, D. H. L., Ghabrial, S. A., Jarvis, A. W., Martelli, A. W., Mayo, M. A. & Summers, M. D. (editors) (1995). Virus Taxonomy: Sixth Report of the International Committee on Taxonomy of Viruses. Wien, New York: Springer-Verlag.
Nicholas, K. B., Nicholas, H. B., Jr & Deerfield, D. W., II (1997). GeneDoc: analysis and visualization of genetic variation. EMBnet News 4, 1–4. http://www.embnet.org/files/shared/EMBnetNews/embnet_news_4_2.pdf
Ohsawa, K., Yamada, A., Takeuchi, K., Watanabe, Y., Miyata, H. & Sato, H. (1998). Genetic characterisation of parainfluenza virus 3 derived from guinea pigs. J Vet Med Sci 60, 919–922.[CrossRef][Medline]
Parks, J. B., Post, G., Thorne, T. & Nash, P. (1972). Parainfluenza-3 virus infection in Rocky Mountain bighorn sheep. J Am Vet Med Assoc 161, 669–672.[Medline]
Sakai, Y., Suzu, S., Shioda, T. & Shibuta, H. (1987). Nucleotide sequence of the bovine parainfluenza genome: its 3' end and the genes of NP, P, C and M proteins. Nucleic Acids Res 15, 2927–2944.
Schrijver, R. S., Hensen, E. J., Langedijk, J. P. M., Daus, F., Middel, W. G. J., Kramps, J. A. & van Oirschot, J. T. (1997). Antibody responses against epitopes on the F protein of bovine respiratory syncytial virus differ in infected or vaccinated cattle. Arch Virol 142, 2195–2210.[CrossRef][Medline]
Shibuta, H., Nozawa, A., Shioda, T. & Kanda, T. (1983). Neuraminidase activity and syncytial formation in variants of parainfluenza 3 virus. Infect Immun 41, 780–788.
Snowder, G. D., Van Vlek, L. D., Cundiff, L. V. & Bennett, G. L. (2006). Bovine respiratory disease in feedlot cattle: environmental, genetic, and economic factors. J Anim Sci 84, 1999–2008.
Stevenson, R. G. & Hore, D. E. (1970). Comparative pathology of lambs and calves infected with parainfluenza virus type 3. J Comp Pathol 80, 613–618.[CrossRef][Medline]
Stokes, A., Tierney, E. L., Murphy, B. R. & Hall, S. L. (1992). The complete nucleotide sequence of the JS strain of human parainfluenza virus type 3: comparison with the Wash/47885/57 prototype strain. Virus Res 25, 91–103.[CrossRef][Medline]
Swierkosz, E. M., Erdman, D. D., Bonnot, T., Schneiderheinze, C. & Waner, J. L. (1995). Isolation and characterisation of a naturally occurring parainfluenza 3 virus variant. J Clin Microbiol 33, 1839–1841.[Abstract]
Thorsen, J. & Henderson, J. P. (1971). Survey for antibody to infectious bovine rhinotracheitis (IBR), bovine virus diarrhea (BVD) and parainfluenza 3(PI3) in moose sera. J Wildl Dis 7, 93–95.
Yener, Z., Saglam, Y. S., Timurkaan, N. & Ilhan, F. (2005). Immunohistochemical detection of parainfluenza type 3 virus antigens in paraffin sections of pneumonic caprine lungs. J Vet Med A Physiol Pathol Clin Med 52, 268–271.[Medline]
Zarnke, R. L. & Erickson, G. A. (1990). Serum antibody prevalence of parainfluenza 3 virus in a free-ranging bison (Bison bison) herd from Alaska. J Wildl Dis 26, 416–419.[Abstract]
Received 1 January 2008;
accepted 28 February 2008.
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