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Comparison of sampling methods for the detection of human rhinovirus RNA
Journal of Clinical Virology 58 (2013) 200–204
Contents lists available at SciVerse ScienceDirect
Journal of Clinical Virology journal homepage: www.elsevier.com/locate/jcv
Comparison of sampling methods for the detection of human rhinovirus RNA Matti Waris a,∗ , Riikka Österback a , Elina Lahti b , Tytti Vuorinen a , Olli Ruuskanen b , Ville Peltola b a b
Department of Virology, University of Turku, Kiinamyllynkatu 13, 20520 Turku, Finland Department of Paediatrics, Turku University Hospital, PB 52, 20521 Turku, Finland
a r t i c l e
i n f o
Article history: Received 5 November 2012 Received in revised form 20 April 2013 Accepted 1 June 2013 Keywords: Rhinovirus Diagnosis PCR Swab Sputum
a b s t r a c t Background: Obtaining a nasal swab (NS) from a child for human rhinovirus (HRV) RNA detection is simple and well tolerated even for repeated sampling, but only few studies have compared them qualitatively and quantitatively with other sampling methods. Objectives: Real-time PCR was used to study the stability of HRV genomes in swabs, and to compare different swabs and induced sputum specimens with nasopharyngeal aspirates (NPAs). Study design: Replicate swabs in a dry test tube were stored at room temperature or mailed to the laboratory before freezing, and compared to freshly frozen specimens. To compare sampling methods, paediatric patients had NPA, NS and throat swab collected. In paired sputum and NPA specimens, viral load was correlated to the amount of -actin mRNA. Results: Specimens were stable at room temperature for at least 4 days and survived mailing without loss of HRV detectability. As compared to NPA, NS had an equal diagnostic sensitivity, with no significant quantitative difference using flocked nylon swabs and a 2.2-fold drop in the average copy number using cotton swabs. The diagnostic sensitivity of cotton swab-collected throat specimens was 97%, with a 26fold lower mean copy number. Sputum specimens had higher HRV RNA (2.3-fold) and -actin mRNA (1.6-fold) copy numbers than NPAs, but there was a poor correlation between HRV RNA and -actin mRNA. Conclusion: HRV remains well detectable by PCR in specimens mailed to the laboratory. The diagnostic efficacy of NPA can be obtained with NS, quantitative comparison and patient comfort favouring flocked nylon-tipped over cotton-tipped swabs. © 2013 Elsevier B.V. All rights reserved.
1. Background Human rhinovirus (HRV) is one of the major findings in respiratory infections including common cold, otitis media, sinusitis, bronchiolitis, asthma exacerbation, pneumonia, and severe respiratory illness in the newborn [1–8]. Thus, HRV seems to be frequently involved in the inflammatory process that leads to these manifestations, with or without bacterial or multiviral co-infection. On the other hand, a literature review found a 15% average detection rate of HRV RNA in asymptomatic subjects, especially in young children [9].
Abbreviations: HRV, human rhinovirus; RT-qPCR, quantitative reverse transcription-PCR; 5 NCR, 5 non-coding region; NPA, nasopharyngeal aspirate; NS, nasal swab. ∗ Corresponding author. Tel.: +358 2 333 7465; fax: +358 2 251 3303. E-mail address: matti.waris@utu.fi (M. Waris). 1386-6532/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jcv.2013.06.002
Nucleic acid testing has proved to be much more effective than virus culture for the detection of HRV [10–12]. This was partially due to strains that could not be amplified in cell culture but only by PCR assays [13–15]. Phylogenetic analysis revealed their divergence from the traditionally defined 75 species A and 25 species B prototype strains, suggesting the definition of species C [16,17]. The success of nucleic acid amplification methods in HRV detection rely on the highly conserved sequence regions at the 5 NCR of human rhino- and enterovirus (HEV) genomes. Different primer combinations and modifications targeting this site have been used in a number of assays described in the literature [10,11,18,19]. Several different methods are used for collecting samples from the respiratory tract for the detection of HRV RNA, but only few studies have compared the sampling methods. 2. Objectives Here, we tested samples collected by different ways and addressed the stability of the samples in ambient conditions. We
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used a quantitative reverse transcription-PCR (RT-qPCR) assay targeting the 5 non-coding region (5 NCR), allowing direct comparison of the amount of HRV RNA between samples. 3. Study design 3.1. Patients and samples Specimens were collected from children attending the Department of Paediatrics of Turku University Hospital and manifesting symptoms of viral respiratory infection (nasal discharge and stuffiness, cough, sore throat, or fever), and from their accompanying parents if they also had acute respiratory symptoms. For stability testing, a nasopharyngeal aspirate (NPA) was collected and 6 replicate cotton swabs were dipped into the aspirate. They were placed into dry, sterile tubes, and tested immediately for HRV as described below or frozen at −70 ◦ C on day 0, 1, 2, 3, or 4 after collection. Series from HRV positive individuals (n = 8) were included in the study. To compare sampling methods, following samples were collected from 39 children in the indicated order: (1) nasal swab (NS) using wood-shafted cotton swab; (2) NS using Copan Pernasal flocked swab (Copan Diagnostics, Brescia, Italy); (3) throat swab using wood-shafted cotton swab; (4) NPA. Each NS was collected from the opposite nostrils at the depth of 2–3 cm (adults, 3–5). NPA was collected using a disposable mucus extractor (Vycon, Ecouen, France) by inserting the catheter through the nostril until the resistance of the nasopharynx was felt, and pulling it back while applying suction from an electronic device. Two portions were separated from the nasopharyngeal aspirate by using cotton swabs. All specimens were placed into dry, sterile test tubes. One portion of the aspirate was sent to the laboratory from different locations around the city using standard mail. In addition, 14 symptomatic parents to the children donated nasal specimens collected from the opposite nostrils using cotton swabs; one of them was mailed to the laboratory. All samples were frozen at −70 ◦ C either immediately or in the laboratory upon receipt. Paired induced sputum and NPA specimens were originally collected for a study investigating their usefulness in the diagnosis of pneumonia [20]. 3.2. Nucleic acid extraction Each sample was suspended into 1 ml of phosphate buffered saline by vortex mixing. Stability study specimens were extracted with E.Z.N.A. Viral RNA Isolation Kit (Omega Bio-Tek, Doraville, GA) according to the manufacturer’s protocol using 150 l of the suspension. Other specimens were processed with Nuclisense easyMag automated nucleic acid extractor (BioMerieux, Boxtel, The Netherlands) using 550 l of the suspension and an elution volume of 55 l. An extraction control of HRV1b with a target copy number of 106 /ml was included in each easyMag run. Two negative controls (water) were included in each extraction run of 24 samples; no carryover contamination was detected in any of the runs.
Fig. 1. Variation of HRV copy numbers measured by RT-qPCR in cotton swabs stored in a dry test tube at room temperature. For each series (n = 8, different symbols; means, grey line), 6 swabs were dipped into NPAs from different donors. Reference sample at day 0 (Ref-0) was analysed immediately, other replicates were frozen at −70 ◦ C on indicated time points and each series was analysed in the same assay.
Amplifications were performed in 25-l reactions with QuantiTect SYBR Green master mix (Qiagen, Hilden, Germany), and 600 nM ENRI3+ (CGGCCCCTGAATGCGGCTAA) and ENRI4− primers, and 5 l of the cDNA using Rotor Gene 3000 (Corbett Research, Mortlake, Australia). Amplification included the following steps: 15 min 95 ◦ C; 45 cycles of 15 s 94 ◦ C, 30 s 65–56 ◦ C (touch-down 1 ◦ C/cycle for the first 10 cycles), 40 s 72 ◦ C; melt 72–95 ◦ C with 1 ◦ C/5 s increments. Melting curve analysis of the 108-bp amplicons was used for initial differentiation between HRVs and HEVs [21]. The threshold cycle (Ct ) values were compared to those of the HRV16 RNA standard for copy number calculation using pre-determined amplification efficiency (93.5%) of serially diluted standard RNA. Negative control (water) was included in each PCR run. Differentiation between rhino- and enteroviruses was confirmed by sequencing a longer 397-bp amplicon produced with primers ENRIu2+ (CAAGCACTTCTGTTTCCCC) and ENRI4−. Amplicons were cleaned with QIAquick PCR purification kit (Qiagen) and sequenced in the DNA Sequencing Service laboratory at the Turku Centre for Biotechnology. To obtain a relative measure of cellular material in the pneumonia study specimens, a qPCR for -actin mRNA was performed using oligo-dT primer in RT and probe-based qPCR as described earlier [22]. 3.4. Statistical analysis IBM SPSS Statistics 20 for Mac was used for descriptive statistics and to calculate significances with paired two-sample t-test for means of log copy numbers, using general linear model with Bonferroni adjustment of P values for multiple comparisons. Microsoft Excel 14.2.0 for Mac was used for linear regression analyses. Specimens with a negative result were calculated with log value = 2.0. 4. Results
3.3. RT-qPCRs 4.1. Stability of HRV RNA in the swabs RNA was reverse transcribed with 20 U of MMLV RNase Htranscriptase (Promega, Madison, WI), 4 U of Rnasin ribonuclease inhibitor, 500 nM dNTP, 1.2 M ENRI4- primer (GAAACACGGACACCCAAAGTA), RT buffer, and 10 l of nucleic acid extract in a total volume of 40 l [19]. The cDNA synthesis was carried out at 42 ◦ C for 1 h. No RT template control (water), and standard preparation of HRV16 RNA (106 copies/l) was included in each run of RT-reactions.
The amount of viral genome was not obviously affected by storage of the swab at room temperature for at least up to 4 days (Fig. 1). There were no significant differences (P = 1.000) between the calculated mean amounts of HRV RNA in the specimens at any single time point. The mean log copy number ± SD at day 0 was 8.0 ± 1.9 (Ct 25.0 ± 5.9), and at day 4 7.6 ± 1.7 (Ct 26.2 ± 5.5). The mean difference between the lowest and highest log copy number ± SD in each
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The highest total virus copy numbers from the 3 children with HEV infection or co-infection were measured in the throat swabs (5.2, 8.3, and 8.8 log copies/sample), much higher than that in the corresponding NPA or NS specimens (0.0–3.3, 4.0–6.3, and 7.0–7.8, respectively). 4.3. HRV in induced sputum and NPA There was a positive correlation of HRV copy numbers between induced sputum and NPA specimens from children with pneumonia (Fig. 3.). In direct comparison, the HRV copy number tended to be higher (5.3 ± 1.9 vs. 5.0 ± 1.7, P = 0.079) in induced sputum than in NPA, while no difference was observed for -actin mRNA (4.3 ± 1.7 vs. 4.5 ± 1.6, P = 0.369). The relative virus loads in sputum and NPA were similar (P = 0.184), when the ratios of HRV RNA/-actin mRNA were compared. However, the overall correlation between HRV RNA and -actin mRNA copy numbers was poor as illustrated in Fig. 3. Fig. 2. Boxplots of HRV copy number variation in different specimen types. Distribution of values in other specimen types were compared pairwise (pairs not shown) with those of the reference NPA, and P values indicate significance levels by t-test with Bonferroni correction. Filled circle represent a negative result. Specimens were frozen at −70 ◦ C immediately after collection, except NPA/mailed were first transported to the laboratory by standard mail. Separation of NPA portions and collection of NS and TS (throat swab) was done with cotton tipped swabs, collection of NS/Copan with flocked nylon swabs.
test series (excluding the freshly tested specimens) was 1.3 ± 0.6 (range 0.6–2.1). 4.2. Detection of HRV in different specimen types Of the 39 children in the specimen type study, 32 tested positive, 29 for HRV, 1 for HEV, and 2 for both. Of the 14 adults, 7 had a sole HRV infection and 1, like her child, had also a co-infection with HEV. Median age of the children with HRV only was 1.4 years (range 8 weeks to 2.7 years, plus one 13.4 year old child); other children’s ages were in the same range. NPA, cotton swab from the nose, and throat swab was obtained from all 29, mailed NPA from 28, and flocked swab from the nose of 25 children with a sole HRV infection. The HRV1b positive control (6.0 log copies/ml) included in the determinations through extraction, RT, and PCR, was used as a measure of test-to-test variation. The observed copy number ± SD for the positive control in 14 runs was 5.9 ± 0.1 log copies/ml (Ct ± SD, 26.7 ± 0.4), with a CV of 23%. HRV yield (mean log copies/sample ± SD) in the other specimen types was compared against that in the freshly frozen NPAs (7.3 ± 1.1), including only paired specimens from children with a sole HRV infection in each comparison (Fig. 2). There was a slight, 2.2-fold reduction in average HRV copy number when the specimen was taken directly with a cotton swab (6.9 ± 1.2, P = 0.313) as compared to aspiration. A 2.6-fold mean drop was observed when the NPA specimen was mailed (6.9 ± 1.3, P = 0.009), which caused a delay of 3.2 ± 1.8 days in ambient temperature. In contrast, 26-fold lower average HRV copy numbers were measured in the corresponding throat swabs (5.9 ± 1.3, P < 0.0001), but only one of them was found negative. Nasal specimens with flocked swabs (7.3 ± 1.0) yielded similar (P = 1.000) quantitative results as the NPA. Comparing only complete sets of specimens from children (n = 24), the highest HRV copy number was observed in 15 flocked swabs, 6 NPAs, 2 cotton swabs, and 1 throat swab. Virus positive adults had much lower copy numbers than children. There was no significant difference (P = 0.461) between freshly frozen (4.4 ± 1.4 log copies/sample) and mailed (4.0 ± 1.2) NS from parents with a sole HRV infection (7 of 14); in each group 6 of 7 specimens were positive for the virus.
5. Discussion Our study shows that a swab with nasal discharge has an excellent stability in a dry test tube at room temperature allowing the use of self-sampling and standard mail to transport the specimen to the laboratory. When compared with freshly frozen specimens, insignificantly smaller copy numbers were obtained in specimens that were first stored at room temperature for 1–4 days, or in mailed specimens from adults, whereas a small but statistically significant drop was observed in mailed specimens from children. The diagnostic sensitivity was the same in either way. Obviously, most specimens had a relatively high virus load and it cannot be excluded that specimens with a low load might turn negative in prolonged storage. Still, we confirm the rationale of instructing parents how to collect NS from their child or themselves and mailing the specimens to the laboratory for follow-up studies, as was done in our earlier study of HRV transmission within families [21]. Surprisingly, 28 of 29 (97%) throat swabs from HRV infected children tested positive for the virus. Average copy numbers were still 10–26 -fold lower in throat swabs than in other specimen types, suggesting that in an adult population with lower overall copy numbers, throat swabs would be suboptimal for HRV detection. On the other hand, our results in children may not be extrapolatable to adults beacause of differences in anatomical structures of the upper respiratory tract. When HEV was found, the highest copy numbers were obtained in the throat swabs, but the number of cases was too few to allow firm conclusions. Nevertheless, our findings support the method of Lambert et al. combining nasal and throat swabs for improved rate of HRV and HEV detections [23]. From the children’s point of view, the best result was the excellent performance of the flocked swab. The soft brush and flexible shaft allows much more pleasant sampling than wood shafted cotton swab, and was also preferable to NPA. Majority of children also prefer flocked swabs over washes with bulb suction [24]. Concentration of the virus in a flocked swab was similar as in the corresponding NPA. Others have reported good results using flocked swabs from the same manufacturer in detection of other respiratory viruses [23,25–28]. Using cotton swabs, Heikkinen et al. found equal sensitivity of NS and NPA specimens for HRV detection by culture [29]. Spyridaki et al. used conventional PCR and found higher HRV detection rate in nasal washes than in cotton-tipped swabs, but they removed cells from the suspended swab before nucleic acid extraction [30]. Moore et al. demonstrated good stability for RNA detection of the relatively labile respiratory syncytial virus in dry swabs, both cotton and flocked [27]. In the current study, the stability tests were limited to the use of cotton swabs,
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Fig. 3. Correlations between HRV RNA, -actin mRNA, sputum, and NPA. Pearson’s correlation coefficients (R2) of the linear regressions were: 0.586 for HRV RNA in NPA vs. sputum, 0.647 for -actin mRNA in NPA vs. sputum and 0.181 for -actin mRNA vs. HRV RNA in all specimens. Specimens with a negative result were given log value = 2.0.
but one would expect similar stability of HRV RNA in the flocked swabs made of nylon. In contrast to this report, a study in adult haematological patients found flocked swabs to have poor sensitivity as compared to NPA, 65% overall (n = 20) and 78% for HRV (n = 9) [31]. It is noteworthy, that they collected the swab from the anterior nostril and used culture medium for storage of the swab. We have already reported that induced sputum is a useful specimen for the detection of respiratory pathogens in pneumonia [2,20]. In this study, we extended the analyses of paired NPA and induced sputum specimens with quantitative comparison of the amount of HRV and mRNA of a cellular gene, -actin, in the same specimens. One could expect that the virus load would have a clear association with the amount of cellular transcripts representing exfoliated epithelial cells. According to our results, the statistical significance of higher HRV copy numbers in induced sputum specimens disappears when the results were normalised with the -actin mRNA copy numbers. However, -actin mRNA copy numbers correlated poorly with those of HRV in the total sample material. Similarly, comparing flocked swabs and NPAs and using -actin gene copies as the reference, Ohrmalm et al. found higher average copy numbers of both -actin gene and virus genomes in NPAs, but a poor correlation between them [31]. Thus, in both studies, specimen with a low number of cellular transcripts or genes could have at least a moderate viral load, suggesting a high number of shed virus but only few shed cells. In conclusion, a NS had the same diagnostic sensitivity for HRV RNA detection as NPA. The diagnostic efficacy of the swab was not significantly affected by storage at room temperature or transport by standard mail. Quantitatively, the flocked nylon swab was equal to NPA and more effective than the cotton swab. Induced sputum specimens from paediatric patients with pneumonia had slightly higher HRV RNA and -actin mRNA copy numbers than corresponding NPAs, but there was a poor correlation between the levels of HRV RNA and -actin mRNA. Funding This study was supported by a special government grant from the Turku University Hospital, the Academy of Finland (no. 140251), and Foundation for Paediatric Research (Finland).
Competing interests None declared. Ethical approval The Ethics Committee of the Turku University Hospital approved the study protocols. Informed consent was obtained from the study subjects or the parents of the children. Acknowledgements We thank Anne Nurmi and Kaisu Kaistinen for assistance at the clinic, and Outi Rauta and Tiina Ylinen for technical assistance in the laboratory. References [1]. Broberg E, Niemela J, Lahti E, Hyypia T, Ruuskanen O, Waris M. Human rhinovirus C-associated severe pneumonia in a neonate. J Clin Virol 2011;51:79–82. [2]. Honkinen M, Lahti E, Osterback R, Ruuskanen O, Waris M. Viruses and bacteria in sputum samples of children with community-acquired pneumonia. Clin Microbiol Infect 2012;18:300–7. [3]. Jartti T, Waris M, Niesters HGM, Allander T, Ruuskanen O. Respiratory viruses and acute asthma in children. JACI 2007;120:216. [4]. Kotaniemi-Syrjanen A, Vainionpaa R, Reijonen TM, Waris M, Korhonen K, Korppi M. Rhinovirus-induced wheezing in infancy-the first sign of childhood asthma? JACI 2003;111:66–71. [5]. Miller EK, Bugna J, Libster R, Shepherd BE, Scalzo PM, Acosta PL, et al. Human rhinoviruses in severe respiratory disease in very low birth weight infants. Pediatrics 2012;129:E60–7. [6]. Peltola V, Jartti T, Putto-Laurila A, Mertsola J, Vainionpaa R, Waris M, et al. Rhinovirus infections in children: a retrospective and prospective hospital-based study. J Med Virol 2009;81:1831–8. [7]. Ruohola A, Waris M, Allander T, Ziegler T, Heikkinen T, Ruuskanen O. Viral Etiology of Common Cold in Children, Finland. Emerg Infect Dis 2009;15:344–6. [8]. Wiertsema SP, Chidlow GR, Kirkham LAS, Corscadden KJ, Mowe EN, Vijayasekaran S, et al. High detection rates of nucleic acids of a wide range of respiratory viruses in the nasopharynx and the middle ear of children with a history of recurrent acute otitis media. J Med Virol 2011;83:2008–17. [9]. Jartti T, Jartti L, Peltola V, Waris M, Ruuskanen O. Identification of respiratory viruses in asymptomatic subjects asymptomatic respiratory viral infections. Pediatr Infect Dis J 2008;27:1103–7. [10]. Hyypia T, Puhakka T, Ruuskanen O, Makela M, Arola A, Arstila P. Molecular diagnosis of human rhinovirus infections: comparison with virus isolation. J Clin Microbiol 1998;36:2081–3.
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[11]. Loens K, Goossens H, de Laat C, Foolen H, Oudshoorn P, Pattyn S, et al. Detection of rhinoviruses by tissue culture and two independent amplification techniques, nucleic acid sequence-based amplification and reverse transcription-PCR in children with acute respiratory infections during a winter season. J Clin Microbiol 2006;44:166–71. [12]. Vuorinen T, Vainionpaa R, Hyypia T. Five years’ experience of reversetranscriptase polymerase chain reaction in daily diagnosis of enterovirus and rhinovirus infections. Clin Infect Dis 2003;37:452–5. [13]. Lau SKP, Yip CCY, Tsoi HW, Lee RA, So LY, Lau YL, et al. Clinical features and complete genome characterization of a distinct human rhinovirus (HRV) genetic previously undetected HRV cluster, probably representing a species, HRV-C, associated with acute respiratory illness in children. J Clin Microbiol 2007;45:3655–64. [14]. McErlean P, Shackelton LA, Lambert SB, Nissen MD, Sjoots TP, Mackay IM. Characterisation of a newly identified human rhinovirus, HRV-QPM, discovered in infants with bronchiolitis. J Clin Virol 2007;39:67–75. [15]. Renwick N, Schweiger B, Kapoor V, Liu Z, Villari J, Bullmann R, et al. A recently identified rhinovirus genotype is associated with severe respiratory-tract infection in children in Germany. J Infect Dis 2007;196:1754–60. [16]. Briese T, Renwick N, Venter M, Jarman RG, Ghosh D, Kondgen S, et al. Global distribution of novel rhinovirus genotype. Emerg Infect Dis 2008;14:944–7. [17]. Simmonds P, McIntyre C, Savolainen-Kopra C, Tapparel C, Mackay IM, Hovi T. Proposals for the classification of human rhinovirus species C into genotypically assigned types. J Gen Virol 2010;91:2409–19. [18]. Deffernez C, Wunderli W, Thomas Y, Yerly S, Perrin L, Kaiser L. Amplicon sequencing and improved detection of human rhinovirus in respiratory samples. J Clin Microbiol 2004;42:3212–8. [19]. Santti J, Vainionpaa R, Hyypia T. Molecular detection and typing of human picornaviruses. Virus Res 1999;62:177–83. [20]. Lahti E, Peltola V, Waris M, Virkki R, Rantakokko-Jalava K, Jalava J, et al. Induced sputum in the diagnosis of childhood community-acquired pneumonia. Thorax 2009;64:252–7. [21]. Peltola V, Waris M, Osterback R, Susi P, Ruuskanen O, Hyypia T. Rhinovirus transmission within families with children: incidence of symptomatic and asymptomatic infections. J Infect Dis 2008;197:382–9.
[22]. Makela M, Oling V, Marttila J, Waris M, Knip M, Simell O, et al. Rotavirusspecific T cell responses and cytokine mRNA expression in children with diabetes-associated autoantibodies and type 1 diabetes. Clin Exp Immunol 2006;145:261–70. [23]. Lambert SB, Whiley DM, O‘Neill NT, Andrews EC, Canavan FM, Bletchly C, et al. Comparing nose-throat swabs and nasopharyngeal aspirates collected from children with symptoms for respiratory virus identification using real-time polymerase chain reaction. Pediatrics 2008;122:E615–20. [24]. Walsh P, Nguyen TA, Higashida K, Michaelson S, Pham K, Nguyen P, et al. Toddlers prefer nasal swabs or washes for specimen collection? Pediatr Infect Dis J 2010;29:1156–7. [25]. Abu-Diab A, Azzeh M, Ghneim R, Zoughbi M, Turkuman S, Rishmawi N, et al. Comparison between pernasal flocked swabs and nasopharyngeal aspirates for detection of common respiratory viruses in samples from children. J Clin Microbiol 2008;46:2414–7. [26]. Chan KH, Peiris JSM, Lim W, Nicholls JM, Chiu SS. Comparison of nasopharyngeal flocked swabs and aspirates for rapid diagnosis of respiratory viruses in children. J Clin Virol 2008;42:65–9. [27]. Moore C, Corden S, Sinha J, Jones R. Dry cotton or flocked respiratory swabs as a simple collection technique for the molecular detection of respiratory viruses using real-time NASBA. J Virol Methods 2008;153:84–9. [28]. Munywoki PK, Hamid F, Mutunga M, Welch S, Cane P, Nokes DJ. Improved detection of respiratory viruses in pediatric outpatients with acute respiratory illness by real-time PCR using nasopharyngeal flocked swabs. J Clin Microbiol 2011;49:3365–7. [29]. Heikkinen T, Marttila J, Salmi AA, Ruuskanen O. Nasal swab versus nasopharyngeal aspirate for isolation of respiratory viruses. J Clin Microbiol 2002;40: 4337–9. [30]. Spyridaki IS, Christodoulou I, de Beer L, Hovland V, Kurowski M, OlszewskaZiaber A. Comparison of four nasal sampling methods for the detection of viral pathogens by RT-PCR – a GALEN project. J Virol Methods 2009;156:102–6. [31]. Ohrmalm L, Wong M, Rotzen-Ostlund M, Norbeck O, Broliden K, Tolfvenstam T. Flocked nasal swab versus nasopharyngeal aspirate for detection of respiratory tract viruses in immunocompromised adults: a matched comparative study. BMC Infect Dis 2010;10:340.
Contents lists available at SciVerse ScienceDirect
Journal of Clinical Virology journal homepage: www.elsevier.com/locate/jcv
Comparison of sampling methods for the detection of human rhinovirus RNA Matti Waris a,∗ , Riikka Österback a , Elina Lahti b , Tytti Vuorinen a , Olli Ruuskanen b , Ville Peltola b a b
Department of Virology, University of Turku, Kiinamyllynkatu 13, 20520 Turku, Finland Department of Paediatrics, Turku University Hospital, PB 52, 20521 Turku, Finland
a r t i c l e
i n f o
Article history: Received 5 November 2012 Received in revised form 20 April 2013 Accepted 1 June 2013 Keywords: Rhinovirus Diagnosis PCR Swab Sputum
a b s t r a c t Background: Obtaining a nasal swab (NS) from a child for human rhinovirus (HRV) RNA detection is simple and well tolerated even for repeated sampling, but only few studies have compared them qualitatively and quantitatively with other sampling methods. Objectives: Real-time PCR was used to study the stability of HRV genomes in swabs, and to compare different swabs and induced sputum specimens with nasopharyngeal aspirates (NPAs). Study design: Replicate swabs in a dry test tube were stored at room temperature or mailed to the laboratory before freezing, and compared to freshly frozen specimens. To compare sampling methods, paediatric patients had NPA, NS and throat swab collected. In paired sputum and NPA specimens, viral load was correlated to the amount of -actin mRNA. Results: Specimens were stable at room temperature for at least 4 days and survived mailing without loss of HRV detectability. As compared to NPA, NS had an equal diagnostic sensitivity, with no significant quantitative difference using flocked nylon swabs and a 2.2-fold drop in the average copy number using cotton swabs. The diagnostic sensitivity of cotton swab-collected throat specimens was 97%, with a 26fold lower mean copy number. Sputum specimens had higher HRV RNA (2.3-fold) and -actin mRNA (1.6-fold) copy numbers than NPAs, but there was a poor correlation between HRV RNA and -actin mRNA. Conclusion: HRV remains well detectable by PCR in specimens mailed to the laboratory. The diagnostic efficacy of NPA can be obtained with NS, quantitative comparison and patient comfort favouring flocked nylon-tipped over cotton-tipped swabs. © 2013 Elsevier B.V. All rights reserved.
1. Background Human rhinovirus (HRV) is one of the major findings in respiratory infections including common cold, otitis media, sinusitis, bronchiolitis, asthma exacerbation, pneumonia, and severe respiratory illness in the newborn [1–8]. Thus, HRV seems to be frequently involved in the inflammatory process that leads to these manifestations, with or without bacterial or multiviral co-infection. On the other hand, a literature review found a 15% average detection rate of HRV RNA in asymptomatic subjects, especially in young children [9].
Abbreviations: HRV, human rhinovirus; RT-qPCR, quantitative reverse transcription-PCR; 5 NCR, 5 non-coding region; NPA, nasopharyngeal aspirate; NS, nasal swab. ∗ Corresponding author. Tel.: +358 2 333 7465; fax: +358 2 251 3303. E-mail address: matti.waris@utu.fi (M. Waris). 1386-6532/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.jcv.2013.06.002
Nucleic acid testing has proved to be much more effective than virus culture for the detection of HRV [10–12]. This was partially due to strains that could not be amplified in cell culture but only by PCR assays [13–15]. Phylogenetic analysis revealed their divergence from the traditionally defined 75 species A and 25 species B prototype strains, suggesting the definition of species C [16,17]. The success of nucleic acid amplification methods in HRV detection rely on the highly conserved sequence regions at the 5 NCR of human rhino- and enterovirus (HEV) genomes. Different primer combinations and modifications targeting this site have been used in a number of assays described in the literature [10,11,18,19]. Several different methods are used for collecting samples from the respiratory tract for the detection of HRV RNA, but only few studies have compared the sampling methods. 2. Objectives Here, we tested samples collected by different ways and addressed the stability of the samples in ambient conditions. We
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used a quantitative reverse transcription-PCR (RT-qPCR) assay targeting the 5 non-coding region (5 NCR), allowing direct comparison of the amount of HRV RNA between samples. 3. Study design 3.1. Patients and samples Specimens were collected from children attending the Department of Paediatrics of Turku University Hospital and manifesting symptoms of viral respiratory infection (nasal discharge and stuffiness, cough, sore throat, or fever), and from their accompanying parents if they also had acute respiratory symptoms. For stability testing, a nasopharyngeal aspirate (NPA) was collected and 6 replicate cotton swabs were dipped into the aspirate. They were placed into dry, sterile tubes, and tested immediately for HRV as described below or frozen at −70 ◦ C on day 0, 1, 2, 3, or 4 after collection. Series from HRV positive individuals (n = 8) were included in the study. To compare sampling methods, following samples were collected from 39 children in the indicated order: (1) nasal swab (NS) using wood-shafted cotton swab; (2) NS using Copan Pernasal flocked swab (Copan Diagnostics, Brescia, Italy); (3) throat swab using wood-shafted cotton swab; (4) NPA. Each NS was collected from the opposite nostrils at the depth of 2–3 cm (adults, 3–5). NPA was collected using a disposable mucus extractor (Vycon, Ecouen, France) by inserting the catheter through the nostril until the resistance of the nasopharynx was felt, and pulling it back while applying suction from an electronic device. Two portions were separated from the nasopharyngeal aspirate by using cotton swabs. All specimens were placed into dry, sterile test tubes. One portion of the aspirate was sent to the laboratory from different locations around the city using standard mail. In addition, 14 symptomatic parents to the children donated nasal specimens collected from the opposite nostrils using cotton swabs; one of them was mailed to the laboratory. All samples were frozen at −70 ◦ C either immediately or in the laboratory upon receipt. Paired induced sputum and NPA specimens were originally collected for a study investigating their usefulness in the diagnosis of pneumonia [20]. 3.2. Nucleic acid extraction Each sample was suspended into 1 ml of phosphate buffered saline by vortex mixing. Stability study specimens were extracted with E.Z.N.A. Viral RNA Isolation Kit (Omega Bio-Tek, Doraville, GA) according to the manufacturer’s protocol using 150 l of the suspension. Other specimens were processed with Nuclisense easyMag automated nucleic acid extractor (BioMerieux, Boxtel, The Netherlands) using 550 l of the suspension and an elution volume of 55 l. An extraction control of HRV1b with a target copy number of 106 /ml was included in each easyMag run. Two negative controls (water) were included in each extraction run of 24 samples; no carryover contamination was detected in any of the runs.
Fig. 1. Variation of HRV copy numbers measured by RT-qPCR in cotton swabs stored in a dry test tube at room temperature. For each series (n = 8, different symbols; means, grey line), 6 swabs were dipped into NPAs from different donors. Reference sample at day 0 (Ref-0) was analysed immediately, other replicates were frozen at −70 ◦ C on indicated time points and each series was analysed in the same assay.
Amplifications were performed in 25-l reactions with QuantiTect SYBR Green master mix (Qiagen, Hilden, Germany), and 600 nM ENRI3+ (CGGCCCCTGAATGCGGCTAA) and ENRI4− primers, and 5 l of the cDNA using Rotor Gene 3000 (Corbett Research, Mortlake, Australia). Amplification included the following steps: 15 min 95 ◦ C; 45 cycles of 15 s 94 ◦ C, 30 s 65–56 ◦ C (touch-down 1 ◦ C/cycle for the first 10 cycles), 40 s 72 ◦ C; melt 72–95 ◦ C with 1 ◦ C/5 s increments. Melting curve analysis of the 108-bp amplicons was used for initial differentiation between HRVs and HEVs [21]. The threshold cycle (Ct ) values were compared to those of the HRV16 RNA standard for copy number calculation using pre-determined amplification efficiency (93.5%) of serially diluted standard RNA. Negative control (water) was included in each PCR run. Differentiation between rhino- and enteroviruses was confirmed by sequencing a longer 397-bp amplicon produced with primers ENRIu2+ (CAAGCACTTCTGTTTCCCC) and ENRI4−. Amplicons were cleaned with QIAquick PCR purification kit (Qiagen) and sequenced in the DNA Sequencing Service laboratory at the Turku Centre for Biotechnology. To obtain a relative measure of cellular material in the pneumonia study specimens, a qPCR for -actin mRNA was performed using oligo-dT primer in RT and probe-based qPCR as described earlier [22]. 3.4. Statistical analysis IBM SPSS Statistics 20 for Mac was used for descriptive statistics and to calculate significances with paired two-sample t-test for means of log copy numbers, using general linear model with Bonferroni adjustment of P values for multiple comparisons. Microsoft Excel 14.2.0 for Mac was used for linear regression analyses. Specimens with a negative result were calculated with log value = 2.0. 4. Results
3.3. RT-qPCRs 4.1. Stability of HRV RNA in the swabs RNA was reverse transcribed with 20 U of MMLV RNase Htranscriptase (Promega, Madison, WI), 4 U of Rnasin ribonuclease inhibitor, 500 nM dNTP, 1.2 M ENRI4- primer (GAAACACGGACACCCAAAGTA), RT buffer, and 10 l of nucleic acid extract in a total volume of 40 l [19]. The cDNA synthesis was carried out at 42 ◦ C for 1 h. No RT template control (water), and standard preparation of HRV16 RNA (106 copies/l) was included in each run of RT-reactions.
The amount of viral genome was not obviously affected by storage of the swab at room temperature for at least up to 4 days (Fig. 1). There were no significant differences (P = 1.000) between the calculated mean amounts of HRV RNA in the specimens at any single time point. The mean log copy number ± SD at day 0 was 8.0 ± 1.9 (Ct 25.0 ± 5.9), and at day 4 7.6 ± 1.7 (Ct 26.2 ± 5.5). The mean difference between the lowest and highest log copy number ± SD in each
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The highest total virus copy numbers from the 3 children with HEV infection or co-infection were measured in the throat swabs (5.2, 8.3, and 8.8 log copies/sample), much higher than that in the corresponding NPA or NS specimens (0.0–3.3, 4.0–6.3, and 7.0–7.8, respectively). 4.3. HRV in induced sputum and NPA There was a positive correlation of HRV copy numbers between induced sputum and NPA specimens from children with pneumonia (Fig. 3.). In direct comparison, the HRV copy number tended to be higher (5.3 ± 1.9 vs. 5.0 ± 1.7, P = 0.079) in induced sputum than in NPA, while no difference was observed for -actin mRNA (4.3 ± 1.7 vs. 4.5 ± 1.6, P = 0.369). The relative virus loads in sputum and NPA were similar (P = 0.184), when the ratios of HRV RNA/-actin mRNA were compared. However, the overall correlation between HRV RNA and -actin mRNA copy numbers was poor as illustrated in Fig. 3. Fig. 2. Boxplots of HRV copy number variation in different specimen types. Distribution of values in other specimen types were compared pairwise (pairs not shown) with those of the reference NPA, and P values indicate significance levels by t-test with Bonferroni correction. Filled circle represent a negative result. Specimens were frozen at −70 ◦ C immediately after collection, except NPA/mailed were first transported to the laboratory by standard mail. Separation of NPA portions and collection of NS and TS (throat swab) was done with cotton tipped swabs, collection of NS/Copan with flocked nylon swabs.
test series (excluding the freshly tested specimens) was 1.3 ± 0.6 (range 0.6–2.1). 4.2. Detection of HRV in different specimen types Of the 39 children in the specimen type study, 32 tested positive, 29 for HRV, 1 for HEV, and 2 for both. Of the 14 adults, 7 had a sole HRV infection and 1, like her child, had also a co-infection with HEV. Median age of the children with HRV only was 1.4 years (range 8 weeks to 2.7 years, plus one 13.4 year old child); other children’s ages were in the same range. NPA, cotton swab from the nose, and throat swab was obtained from all 29, mailed NPA from 28, and flocked swab from the nose of 25 children with a sole HRV infection. The HRV1b positive control (6.0 log copies/ml) included in the determinations through extraction, RT, and PCR, was used as a measure of test-to-test variation. The observed copy number ± SD for the positive control in 14 runs was 5.9 ± 0.1 log copies/ml (Ct ± SD, 26.7 ± 0.4), with a CV of 23%. HRV yield (mean log copies/sample ± SD) in the other specimen types was compared against that in the freshly frozen NPAs (7.3 ± 1.1), including only paired specimens from children with a sole HRV infection in each comparison (Fig. 2). There was a slight, 2.2-fold reduction in average HRV copy number when the specimen was taken directly with a cotton swab (6.9 ± 1.2, P = 0.313) as compared to aspiration. A 2.6-fold mean drop was observed when the NPA specimen was mailed (6.9 ± 1.3, P = 0.009), which caused a delay of 3.2 ± 1.8 days in ambient temperature. In contrast, 26-fold lower average HRV copy numbers were measured in the corresponding throat swabs (5.9 ± 1.3, P < 0.0001), but only one of them was found negative. Nasal specimens with flocked swabs (7.3 ± 1.0) yielded similar (P = 1.000) quantitative results as the NPA. Comparing only complete sets of specimens from children (n = 24), the highest HRV copy number was observed in 15 flocked swabs, 6 NPAs, 2 cotton swabs, and 1 throat swab. Virus positive adults had much lower copy numbers than children. There was no significant difference (P = 0.461) between freshly frozen (4.4 ± 1.4 log copies/sample) and mailed (4.0 ± 1.2) NS from parents with a sole HRV infection (7 of 14); in each group 6 of 7 specimens were positive for the virus.
5. Discussion Our study shows that a swab with nasal discharge has an excellent stability in a dry test tube at room temperature allowing the use of self-sampling and standard mail to transport the specimen to the laboratory. When compared with freshly frozen specimens, insignificantly smaller copy numbers were obtained in specimens that were first stored at room temperature for 1–4 days, or in mailed specimens from adults, whereas a small but statistically significant drop was observed in mailed specimens from children. The diagnostic sensitivity was the same in either way. Obviously, most specimens had a relatively high virus load and it cannot be excluded that specimens with a low load might turn negative in prolonged storage. Still, we confirm the rationale of instructing parents how to collect NS from their child or themselves and mailing the specimens to the laboratory for follow-up studies, as was done in our earlier study of HRV transmission within families [21]. Surprisingly, 28 of 29 (97%) throat swabs from HRV infected children tested positive for the virus. Average copy numbers were still 10–26 -fold lower in throat swabs than in other specimen types, suggesting that in an adult population with lower overall copy numbers, throat swabs would be suboptimal for HRV detection. On the other hand, our results in children may not be extrapolatable to adults beacause of differences in anatomical structures of the upper respiratory tract. When HEV was found, the highest copy numbers were obtained in the throat swabs, but the number of cases was too few to allow firm conclusions. Nevertheless, our findings support the method of Lambert et al. combining nasal and throat swabs for improved rate of HRV and HEV detections [23]. From the children’s point of view, the best result was the excellent performance of the flocked swab. The soft brush and flexible shaft allows much more pleasant sampling than wood shafted cotton swab, and was also preferable to NPA. Majority of children also prefer flocked swabs over washes with bulb suction [24]. Concentration of the virus in a flocked swab was similar as in the corresponding NPA. Others have reported good results using flocked swabs from the same manufacturer in detection of other respiratory viruses [23,25–28]. Using cotton swabs, Heikkinen et al. found equal sensitivity of NS and NPA specimens for HRV detection by culture [29]. Spyridaki et al. used conventional PCR and found higher HRV detection rate in nasal washes than in cotton-tipped swabs, but they removed cells from the suspended swab before nucleic acid extraction [30]. Moore et al. demonstrated good stability for RNA detection of the relatively labile respiratory syncytial virus in dry swabs, both cotton and flocked [27]. In the current study, the stability tests were limited to the use of cotton swabs,
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Fig. 3. Correlations between HRV RNA, -actin mRNA, sputum, and NPA. Pearson’s correlation coefficients (R2) of the linear regressions were: 0.586 for HRV RNA in NPA vs. sputum, 0.647 for -actin mRNA in NPA vs. sputum and 0.181 for -actin mRNA vs. HRV RNA in all specimens. Specimens with a negative result were given log value = 2.0.
but one would expect similar stability of HRV RNA in the flocked swabs made of nylon. In contrast to this report, a study in adult haematological patients found flocked swabs to have poor sensitivity as compared to NPA, 65% overall (n = 20) and 78% for HRV (n = 9) [31]. It is noteworthy, that they collected the swab from the anterior nostril and used culture medium for storage of the swab. We have already reported that induced sputum is a useful specimen for the detection of respiratory pathogens in pneumonia [2,20]. In this study, we extended the analyses of paired NPA and induced sputum specimens with quantitative comparison of the amount of HRV and mRNA of a cellular gene, -actin, in the same specimens. One could expect that the virus load would have a clear association with the amount of cellular transcripts representing exfoliated epithelial cells. According to our results, the statistical significance of higher HRV copy numbers in induced sputum specimens disappears when the results were normalised with the -actin mRNA copy numbers. However, -actin mRNA copy numbers correlated poorly with those of HRV in the total sample material. Similarly, comparing flocked swabs and NPAs and using -actin gene copies as the reference, Ohrmalm et al. found higher average copy numbers of both -actin gene and virus genomes in NPAs, but a poor correlation between them [31]. Thus, in both studies, specimen with a low number of cellular transcripts or genes could have at least a moderate viral load, suggesting a high number of shed virus but only few shed cells. In conclusion, a NS had the same diagnostic sensitivity for HRV RNA detection as NPA. The diagnostic efficacy of the swab was not significantly affected by storage at room temperature or transport by standard mail. Quantitatively, the flocked nylon swab was equal to NPA and more effective than the cotton swab. Induced sputum specimens from paediatric patients with pneumonia had slightly higher HRV RNA and -actin mRNA copy numbers than corresponding NPAs, but there was a poor correlation between the levels of HRV RNA and -actin mRNA. Funding This study was supported by a special government grant from the Turku University Hospital, the Academy of Finland (no. 140251), and Foundation for Paediatric Research (Finland).
Competing interests None declared. Ethical approval The Ethics Committee of the Turku University Hospital approved the study protocols. Informed consent was obtained from the study subjects or the parents of the children. Acknowledgements We thank Anne Nurmi and Kaisu Kaistinen for assistance at the clinic, and Outi Rauta and Tiina Ylinen for technical assistance in the laboratory. References [1]. Broberg E, Niemela J, Lahti E, Hyypia T, Ruuskanen O, Waris M. Human rhinovirus C-associated severe pneumonia in a neonate. J Clin Virol 2011;51:79–82. [2]. Honkinen M, Lahti E, Osterback R, Ruuskanen O, Waris M. Viruses and bacteria in sputum samples of children with community-acquired pneumonia. Clin Microbiol Infect 2012;18:300–7. [3]. Jartti T, Waris M, Niesters HGM, Allander T, Ruuskanen O. Respiratory viruses and acute asthma in children. JACI 2007;120:216. [4]. Kotaniemi-Syrjanen A, Vainionpaa R, Reijonen TM, Waris M, Korhonen K, Korppi M. Rhinovirus-induced wheezing in infancy-the first sign of childhood asthma? JACI 2003;111:66–71. [5]. Miller EK, Bugna J, Libster R, Shepherd BE, Scalzo PM, Acosta PL, et al. Human rhinoviruses in severe respiratory disease in very low birth weight infants. Pediatrics 2012;129:E60–7. [6]. Peltola V, Jartti T, Putto-Laurila A, Mertsola J, Vainionpaa R, Waris M, et al. Rhinovirus infections in children: a retrospective and prospective hospital-based study. J Med Virol 2009;81:1831–8. [7]. Ruohola A, Waris M, Allander T, Ziegler T, Heikkinen T, Ruuskanen O. Viral Etiology of Common Cold in Children, Finland. Emerg Infect Dis 2009;15:344–6. [8]. Wiertsema SP, Chidlow GR, Kirkham LAS, Corscadden KJ, Mowe EN, Vijayasekaran S, et al. High detection rates of nucleic acids of a wide range of respiratory viruses in the nasopharynx and the middle ear of children with a history of recurrent acute otitis media. J Med Virol 2011;83:2008–17. [9]. Jartti T, Jartti L, Peltola V, Waris M, Ruuskanen O. Identification of respiratory viruses in asymptomatic subjects asymptomatic respiratory viral infections. Pediatr Infect Dis J 2008;27:1103–7. [10]. Hyypia T, Puhakka T, Ruuskanen O, Makela M, Arola A, Arstila P. Molecular diagnosis of human rhinovirus infections: comparison with virus isolation. J Clin Microbiol 1998;36:2081–3.
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