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Journal of Clinical Microbiology, May 1998, p. 1388-1391, Vol. 36, No. 5
Respiratory and Enteric Viruses Branch,
Received 9 October 1997/Returned for modification 17 December
1997/Accepted 13 February 1998
Reverse transcription (RT)-PCR assays have been widely described
for use in the diagnosis of human parainfluenza viruses (HPIVs) and
other respiratory virus pathogens. However, these assays are mostly
monospecific, requiring separate amplifications for each HPIV type. In
the present work, we describe multiplex RT-PCR assays that detect and
differentiate HPIV serotypes 1, 2, and 3 in a combined reaction.
Specifically, a mixture of three pairs of primers to conserved regions
of the hemagglutinin-neuraminidase gene of each HPIV serotype was used
for primary amplification, yielding amplicons with similar sizes. For
typing, a second amplification was performed with a mixture of nested
primers, yielding amplicons with sizes easily differentiated by agarose
gel electrophoresis. A modified single-amplification RT-PCR assay with
fluorescence-labeled nested primers, followed by analysis of the
labeled products on an automated sequencing gel, was also evaluated.
Fifteen temporally and geographically diverse HPIV isolates from the
Centers for Disease Control and Prevention archives and 26 of 30 (87%)
previously positive nasopharyngeal specimens (8 of 10 positive for HPIV
serotype 1 [HPIV1], 9 of 10 positive for HPIV2, and 9 of 10 positive
for HPIV3) were positive and were correctly typed by both assays. Negative results were obtained with naso- or oropharyngeal specimens and/or culture isolates of 33 unrelated respiratory tract pathogens, including HPIV4, enterovirus, rhinovirus, respiratory syncytial virus,
adenovirus, influenza virus, and Streptococcus pneumoniae. Our multiplex RT-PCR assays provide sensitive, specific, and simplified tools for the rapid diagnosis of HPIV infections.
Human parainfluenza viruses (HPIVs)
are medically important respiratory pathogens and are second only to
respiratory syncytial virus (RSV) as a major cause of lower respiratory
tract (LRT) illness in infants and young children (4, 5, 15, 17, 18, 22). Although repeat infections in healthy older children and
adults are typically less severe, serious LRT illness caused by HPIVs
has been reported among immunocompromised individuals (1, 2, 12,
13, 26-28) and institutionalized elderly individuals (7).
Of the four recognized serotypes of HPIV (HPIV serotype 1 [HPIV1],
HPIV2, HPIV3, and HPIV4) HPIV3 is most commonly associated with serious
LRT illness, followed by HPIV1 and HPIV2; HPIV4 is rarely associated
with serious illness (9). The use of classic diagnostic
methods like viral isolation and serology can often result in a delay
of several weeks before the results are available, and these methods
are therefore less useful for making therapeutic decisions
(4). Direct antigen detection methods are widely used for
the rapid diagnosis of HPIV infections (11, 20, 21), but
results can be variable (10), and some HPIV strains may be
missed entirely by assays with specific monoclonal antibodies (24). To address these problems, reverse transcription
(RT)-PCR assays that have been shown to provide rapid and sensitive
detection of HPIV1 and HPIV3 have been developed (6, 14,
15). However, all of these methods are monospecific, requiring
separate amplifications for each virus. As an alternative, multiplex
PCR assays permit simultaneous amplification of several viruses in a
single reaction (25). In this study, we describe novel
multiplex RT-PCR assays for the detection and identification of HPIV1,
-2, and -3.
Virus isolates.
Cell culture isolates of reference strains
of virus obtained from the Centers for Disease Control and Prevention
archives were used to develop and standardize the RT-PCR assays. These included (i) HPIV prototype strains C-35 (HPIV1), Greer (HPIV2), and
C-243 (HPIV3); (ii) 12 geographically and temporally diverse HPIV
isolates from the United States, Canada, and Argentina (HPIV1 from
Córdoba, Argentina [1967], Massachusetts [1985], Oregon [1974], and Nevada [1985]; HPIV2 from California [1980],
Massachusetts [1971], Connecticut [1977], and Kansas [1982];
HPIV3 from Idaho [1977], Alabama [1979], Oregon [1981], and
Manitoba, Canada [1981]); and (iii) individual isolates of HPIV4, RSV
subtypes A and B, influenza A and B viruses, adenovirus, enterovirus,
and rhinovirus.
Clinical samples.
The clinical samples used to evaluate the
efficacies of the RT-PCR assays included 45 throat or nasopharyngeal
swabs collected from pediatric patients at the Cardinal Glennon
Children's Hospital, St. Louis, Mo., and placed in viral transport
medium; 10 specimens each were previously culture positive for HPIV1,
-2, and -3, and the remaining 15 specimens were positive for other
respiratory viruses, including 3 each for RSV, adenovirus, influenza A
virus, enterovirus, and rhinovirus. Ten nasopharyngeal specimens from adults culture positive for Streptococcus pneumoniae and
culture negative for respiratory viruses were also tested. All clinical samples had been stored at Primer design and preparation.
Previously published
sequences of the HPIV hemagglutinin-neuraminidase gene (from 27 HPIV1
isolates, 2 HPIV2 isolates, and 10 HPIV3 isolates) obtained from
GenBank were aligned by using the Wisconsin Analysis Package (version
8), Genetics Computer Group, Madison, Wis. External and nested primer
sequences were chosen from conserved regions of the
hemagglutinin-neuraminidase gene and were designed to achieve optimal
performance in a multiplex reaction (Table
1). All primers were prepared on a model
394 ABI DNA synthesizer (Perkin-Elmer). Fluorescence-tagged primers were labeled at the 5' end with 6-phosphoramidite (FAM; Glen Research). FAM-labeled primers were purified by reversed-phase high-performance liquid chromatography. All other primers were used without further purification.
RNA extraction.
RNA was extracted from the clinical samples
and virus isolates as described previously (3). Briefly, 50 µl of each sample was treated with 200 µl of extraction buffer (4 M
guanidinium thiocyanate, 0.5% N-lauryl sarcosine, 1 mM
dithiothreitol, 25 mM sodium citrate, and 0.1 mg of glycogen per ml),
followed by isopropanol and 70% ethanol precipitations.
RT.
The vacuum-dried RNA pellets were resuspended in 10 µl
of hybridization buffer (300 mM NaCl, 5 mM Tris-HCl [pH 7.5], 1 mM EDTA) containing HPIV1F, HPIV2F, and HPIV3F each at a concentration of
2 µM and were denatured for 3 min at 94°C. The RNA template and
primers were then allowed to hybridize for 45 min at 50°C. A 40-µl
volume of RT buffer (10 mM Tris-HCl [pH 8.3]; 6 mM magnesium chloride; 1 mM dithiothreitol; dATP, dGTP, dCTP, and dTTP at a concentration of 1 mM each; and 40 U of RNase inhibitor [RNAsin; Boehringer Mannheim]) containing 50 U of avian myeloblastosis reverse
transcriptase (Boehringer Mannheim) was then added, and the mixture was
incubated for 1 h at 42°C. The enzyme was finally inactivated by
incubation for 5 min at 92°C.
PCR amplification and product detection, two-step method.
For primary PCR amplification, 5 µl of cDNA prepared in the RT
reaction was added to a PCR mixture containing 10 mM Tris-HCl (pH 8.3);
50 mM KCl; 3 mM MgCl; dATP, dCTP, dGTP, and dTTP each at a
concentration of 200 µM; the primary amplification primers (Table 1)
each at a concentration of 2 µM; and 1.25 U of Taq polymerase (AmpliTaq; Perkin-Elmer Cetus) in a final volume of 50 µl
and the mixture was overlaid with mineral oil. Amplification was
performed on a model 480 DNA Thermal Cycler (Perkin-Elmer Cetus)
programmed for 35 cycles of 1 min of denaturation at 94°C, 1 min of
annealing at 50°C, and 1 min of elongation at 72°C; an additional 2 min of denaturation preceded the first cycle, and elongation was
extended to 5 min in the last cycle. For nested PCR, 1 µl of the
primary amplification products was added to 49 µl of a new PCR
mixture containing nested instead of primary reaction primers (Table
1), and the thermal cycle program annealing temperature was changed to
58°C. The PCR products were sized by gel electrophoresis on 2%
agarose containing 0.5 g of ethidium bromide per ml in TBE (Tris-borate-EDTA) buffer and were visualized under UV light. Standard
precautions were taken to avoid carryover contamination. Pipetting was
performed with aerosol-resistant tips, and different biosafety cabinets
were used for sample extraction and first amplification or nested
amplification. Amplicon detection was performed in a different room.
PCR amplification and product detection, one-step method.
Primary amplification was performed with the nested primers directly
with the RT products, but substituting FAM-labeled forward primers for
the unlabeled forward primers. The labeled products were subjected to
6% acrylamide gel electrophoresis under denaturing conditions on an
ABI PRISM 377 DNA sequencer (Perkin-Elmer Corp.) and were analyzed with
GeneScan Analysis Software (version 2.0.2).
Viral isolates.
PCR products of the expected size were
obtained with the three HPIV prototype strains when they were tested
alone or in a mixed reaction (Fig. 1).
Serial dilution experiments showed that the nested PCR was about
100-fold more sensitive than cell culture isolation for all three
types. All reference HPIV isolates were detected and were correctly
identified by both the one-step and the two-step procedures. All
non-HPIV isolates were PCR negative.
0095-1137/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
Simultaneous Detection and Identification of Human
Parainfluenza Viruses 1, 2, and 3 from Clinical Samples by
Multiplex PCR
ABSTRACT
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
70°C.
TABLE 1.
Multiplex primers for HPIVs
RESULTS
Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
View larger version (21K):
[in a new window]
FIG. 1.
Detection and typing of prototype HPIV1, HPIV2, and
HPIV3 strains. (A) Lane M, molecular weight markers; lanes 1, 2, and 3, HPIV1, HPIV2, and HPIV3, respectively; lane A, mixture of all three
HPIV types. (B) GeneScan Analysis Software profile of the mixture in
lane A.
Clinical samples. Of the clinical samples from which HPIVs had previously been isolated, 8 of 10 (80%) samples containing HPIV1, 9 of 10 (90%) samples containing HPIV2, and 9 of 10 (90%) samples containing HPIV3 were RT-PCR positive and were correctly typed by both methods (Fig. 2). Only three of them were detected after the first amplification in the two-step method. HPIV was reisolated from one of the four clinical samples that were negative by RT-PCR. The isolate was positive for HPIV-1 by RT-PCR, consistent with the original isolate identification. None of the 25 control specimens containing other respiratory viruses or bacterial pathogens were positive. No discrepancies were found between the one-step and two-step procedures.
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DISCUSSION |
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Top
Abstract
Introduction
Materials & Methods
Results
Discussion
References
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RT-PCR assays for the detection of HPIVs have been described, but they have been limited to the detection of HPIV3 (14, 15) or HPIV1 and HPIV3 (6), without distinguishing between the two serotypes. In contrast, our multiplex assays both detect and differentiate all three medically important HPIV serotypes and provide a sensitive and specific means of identifying HPIVs directly from clinical specimens; 87% (26 of 30) of the clinical samples culture positive for HPIVs were correctly identified, whereas none of the specimens containing a diverse group of other respiratory virus and bacterial pathogens were positive.
Although our primers were designed to react with temporally and geographically diverse HPIV isolates, published HPIV sequence data were limited, and therefore unanticipated strain sequence variation in the primer regions could result in occasional false-negative results. This may account for our failure to amplify virus from three of the initially culture-positive clinical samples. However, we believe that the most likely explanation for our false-negative results with these specimens was that the samples were subjected to at least two freeze-thaw cycles between the initial isolation and RT-PCR procedures, possibly causing a decrease in virus titer and RNA degradation. Further testing of diverse fresh clinical samples by RT-PCR will be required to provide a better estimate of the sensitivity and specificity of this assay in a diagnostic setting.
By combining the reactions into fewer assays, the multiplex design is able to achieve substantial savings in time and reagent costs compared with those required for monospecific methods. The two-step multiplex procedure can be readily adapted to laboratories familiar with RT-PCR procedures by conventional DNA detection methods. The one-step procedure offers several important advantages for laboratories with access to the required equipment. PCR products labeled by incorporation of fluorescence-labeled primers can be detected and sized (or typed) with sensitivity comparable to that of nested PCR, but without the added risk of DNA contamination associated with nested PCR procedures; moreover, the GeneScan Analysis Software makes it possible to size bands with single-base-pair accuracy (19), minimizing the chance of detecting an incorrect but similarly sized product.
In summary, the multiplex RT-PCR assays described here proved to be very effective for the detection and rapid identification of HPIVs. Since primers for other respiratory viruses can be added to the reaction mixture, the efficiencies of these assays can potentially be further improved. However, additional studies will be required to determine the extent to which these assays can complement or replace conventional diagnostic methods.
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ACKNOWLEDGMENT |
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This study was performed at the Centers for Disease Control and Prevention with funding support from the Spanish Ministry of Science and Education (grant PR95-32; Estancias de Investigadores Españoles en Centros de Investigación Extranjeros).
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FOOTNOTES |
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* Corresponding author. Mailing address: Centro Nacional de Microbiología, Instituto de Salud Carlos III, Carretera Majadahonda-Pozuelo s/n, 28220 Majadahonda, Madrid, Spain. Phone: 34-1-5097901. Fax: 34-1-5097966. E-mail: jeecheva{at}isciii.es.
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REFERENCES |
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Abstract
Introduction
Materials & Methods
Results
Discussion
References
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Journal of Clinical Microbiology, May 1998, p. 1388-1391, Vol. 36, No. 5
0095-1137/98/$04.00+0
Copyright © 1998, American Society for Microbiology. All rights reserved.
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