Abstract
Laboratory diagnosis has played a critical role in the Global Polio Eradication Initiative since 1988, by isolating and identifying poliovirus (PV) from stool specimens by using cell culture as a highly sensitive system to detect PV. In the present study, we aimed to develop a molecular method to detect PV directly from stool extracts, with a high efficiency comparable to that of cell culture. We developed a method to efficiently amplify the entire capsid coding region of human enteroviruses (EVs) including PV. cDNAs of the entire capsid coding region (3.9 kb) were obtained from as few as 50 copies of PV genomes. PV was detected from the cDNAs with an improved PV-specific real-time reverse transcription-PCR system and nucleotide sequence analysis of the VP1 coding region. For assay validation, we analyzed 84 stool extracts that were positive for PV in cell culture and detected PV genomes from 100% of the extracts (84/84 samples) with this method in combination with a PV-specific extraction method. PV could be detected in 2/4 stool extract samples that were negative for PV in cell culture. In PV-positive samples, EV species C viruses were also detected with high frequency (27% [23/86 samples]). This method would be useful for direct detection of PV from stool extracts without using cell culture.
INTRODUCTION
Laboratory diagnosis has played a critical role in the Global Polio Eradication Initiative since 1988, by detecting and identifying poliovirus (PV). To date, PV has been detected by isolating the virus, using cell culture systems, from stool specimens from acute flaccid paralysis (AFP) cases, including polio and other paralysis cases, in the World Health Organization (WHO) Global Polio Laboratory Network (1, 2). The advantages of the cell culture-based procedure are (i) minimal equipment requirements, (ii) high sensitivity (detection limit of 1 infectious unit containing 50 to 1,000 virions) (3), and, importantly, (iii) biological amplification of PV to high titers (about 106 50% cell culture infectious dose [CCID50] or 108 to 109 viral genome copies per μl of cell lysate) for subsequent nucleotide sequence analysis of the capsid protein VP1 coding region, which is required for final identification and molecular epidemiological analysis of PV strains. Sequencing is essential for classifying PV isolates into vaccine strains, wild-type strains, and circulating vaccine-derived PV (VDPV) strains and for determining the necessity of additional vaccination plans by identifying virus reservoirs, along with molecular epidemiological methods (4). A major disadvantage of the cell culture system is the timeliness of reporting; it takes 10 days to confirm that a sample is negative for PV (2). To improve the efficiency of PV detection and identification, including the timeliness, the development of methods for direct detection of PV has been encouraged by WHO (http://www.polioeradication.org/Research/Grantsandcollaboration.aspx). Such direct detection methods must provide efficient PV detection, comparable to that of cell culture, and also provide material to allow sequencing of the entire VP1 coding region.
One promising approach for direct detection and identification of PV is real-time reverse transcription (rRT)-PCR, with practical affordability (5, 6). However, the sensitivity of pan-PV PCR has been a major challenge [e.g., 25,000 copies for detection of PV1(Sabin)]. This low sensitivity might reflect intrinsic properties of degenerate primers and probes that use inosine and modified nucleotides to detect diverse nucleotide sequences conserved among the 3 types of PV in the VP1 coding region (7–10). With nondegenerate primers, highly sensitive rRT-PCR systems to detect PV vaccine strains have been developed (6, 11). VDPV strains could also be detected with a recently developed rRT-PCR system (12). However, alternative methodologies remain to be developed to obtain the nucleotide sequence of the entire VP1 coding region from quite small amounts of PV in stool extracts, which could be <100.5 CCID50 or about 1,000 viral genome copies in 50 μl of stool extract (5, 6).
Previously, we developed magnetic nanoparticles sensitized with soluble PV receptor (PVR) (i.e., PVR magnetic beads [MB]) for extraction of PV from stool extracts, to increase the amounts of PV available for direct detection assays (6). During that study, we noticed that some stool extracts contained substantial amounts of noninfectious PV that could not be extracted with PVR MB (about 70 to 80% of total PV in the extracts). This suggested that untreated stool extracts could contain substantial amounts of viral RNA, greater than amounts expected from viable viruses, and could serve as a promising source for direct detection and identification of PV by molecular methods.
In the present study, we developed an efficient entire-capsid-coding-region amplification (ECRA) method to amplify the entire capsid coding region of PV (about 3.9 kb) from PV genomic RNA at as little as 50 copies, for direct detection and identification of PV from stool extracts. We found that application of this ECRA method combined with improved PV rRT-PCR, VP1 sequencing, and PV extraction with PVR MB was effective for direct detection of PV from stool extracts with high efficiency, comparable to that of cell culture.
MATERIALS AND METHODS
Cells and clinical samples.
RD cells (a human rhabdomyosarcoma cell line) and L20B cells (a mouse L cell line expressing the PV receptor) were cultured as monolayers in Dulbecco's modified Eagle medium (DMEM) supplemented with 10% fetal calf serum (FCS) and were used for titration of viruses. PV-positive stool extracts from AFP cases, which were obtained in part from a national polio laboratory in Maiduguri and stored at the US Centers for Disease Control and Prevention (CDC) and were positive for type 1 or 3 wild-type strains, vaccine strains, or VDPV strains (total of 88 samples), were used in this study. Virus isolation was performed according to current PV isolation procedures (2). Briefly, RD cells and L20B cells were inoculated with 100 μl of the stool extracts and then incubated at 35°C. The cells were observed for cytopathic effect (CPE) for 5 days. After 5 days, if the samples showed no CPE, then the supernatants of the samples were passaged to the same cell line after freezing and thawing, followed by additional observation for 5 days. The virus isolates from RD cells were subjected to reinoculation into L20B cells to confirm CPE in L20B cells.
Entire-capsid-coding-region amplification method.
Purified viral genomic RNA solution (50 μl) was prepared from 140 μl of stool extracts or PV extracted from 100 μl of stool extracts with 5 μl of PVR MB, as described previously (6), by using a QIAamp viral RNA kit (Qiagen). For reverse transcription (RT) reactions, 13 μl purified viral genomic RNA solution was used for RT with random hexamers using a PrimeScript II High Fidelity RT-PCR kit, according to the manufacturer's instructions (total of 20 μl of RT reaction solution; TaKaRa). For the first RT-PCR, the entire capsid coding region was amplified with 8 μl of RT reaction solution by using a KOD FX Neo kit (Toyobo) with primers 5′NTR-529–552+ (5′-TGGCGGAACCGACTACTTTGGGTG-3′) and Cre-4485–4460− (5′-TCAATACGGTGTTTGCTCTTGAACTG-3′), according to the manufacturer's instructions (total of 50 μl of the first RT-PCR solution). PCR conditions consisted of a denaturing step at 94°C for 2 min and 45 cycles of thermal cycling at 98°C for 10 s and 68°C for 4 min. The first RT-PCR products were purified by using a Wizard SV gel and PCR clean-up system kit (Promega) prior to PV intratypic differentiation (ITD) real-time RT-PCR.
Nucleotide sequence analysis of VP1 coding region.
cDNA of the VP1 coding region was amplified with 1.0 μl of unpurified first RT-PCR product as the template by using a Qiagen OneStep RT-PCR kit (Qiagen) with primers Y7 (5′-GGGTTTGTGTCAGCCTGTAATGA-3′) and Q8 (5′-AAGAGGTCTCTRTTCCACAT-3′) or with primers Y7 and PV1A (5′-TTIAIIGCRTGICCRTTRTT-3′), according to the manufacturer's instructions (total of 50 μl of the second PCR solution) (10). PCR conditions consisted of a denaturing step at 95°C for 15 min, 35 cycles of 94°C for 30 s, 50°C for 30 s, and 72°C for 1 min, and an additional elongation step at 72°C for 7 min. The second PCR products were purified by using a Wizard SV gel and PCR clean-up system kit (Promega) for sequence reactions with primer Y7 or PV1A. DNA sequencing was performed by using a BigDye Terminator v3.1 cycle sequencing ready reaction kit (Applied Biosystems) and an ABI Prism 3130 genetic analyzer (Applied Biosystems).
Conventional and improved PV ITD rRT-PCR assays.
Conventional PV ITD rRT-PCR was performed by using a poliovirus diagnostic rRT-PCR kit provided by the US CDC, according to the kit instructions. One microliter of the purified first RT-PCR product was used for PV ITD rRT-PCR. The fluorescence emission of the probe was monitored and analyzed by using a 7500 Fast real-time PCR system (Applied Biosystems). Improved PV ITD rRT-PCR was performed by using a qScript XLT One-Step RT-qPCR ToughMix kit (Quanta) with the same primer/probe sets as provided in the poliovirus diagnostic rRT-PCR kit (US CDC) and dual-stage thermal cycling (RT reaction/inactivation at 42°C for 45 min and 95°C for 3 min and then 15 cycles of PCR at 95°C for 24 s, 44°C for 30 s, and 60°C for 24 s, followed by 40 cycles of PCR at 95°C for 24 s, 47°C for 30 s, and 65°C for 24 s) (12).
RESULTS
Development of an efficient ECRA method.
To efficiently amplify the entire capsid coding region of PV, we designed a nondegenerate primer set for conserved nucleotide sequences in the 5′ nontranslated region (NTR) and in the cis-acting replication element (Cre). We found that the nucleotide sequences in these regions were well conserved among Enterovirus A (EV-A), EV-B, and EV-C species, but there are no specific nucleotide sequences for PV or EV-C types closely related to PV (coxsackievirus A11 [CVA11], CVA13, CVA17, and CVA20 strains) (Fig. 1A).
FIG 1.
Development of an efficient ECRA method. (A) Primer design for the efficient ECRA method. Primers were designed for the 5′ NTR and Cre regions of human EV. (B) Sensitivity of the efficient ECRA method. Indicated amounts of PV1(Sabin) cDNA were subjected to RT-PCR with the indicated DNA polymerases. PCR products were subjected to 1% agarose gel electrophoresis and then stained with ethidium bromide. (C) Flow chart of PV detection and identification of PV by using the current PV ITD rRT-PCR system and VP1 nucleotide sequence analysis in combination with the ECRA method and the PVR MB PV-specific extraction method.
With a primer set designed for these regions (primers 5′NTR-529–552+ and Cre-4485–4460−), we examined RT-PCR systems. With RT-PCR systems with a conventional highly processive polymerase, the limit of amplification and detection was approximately 5 × 104 copies with viral genomic RNA of the PV1(Sabin) strain (Fig. 1B). We found that the use of KOD FX Neo polymerase drastically improved the amplification efficiency of RT-PCR, with a limit of amplification and detection of ≤50 copies, suggesting about 1,000-fold higher sensitivity than that of the conventional RT-PCR systems. This suggested the potential of this ECRA method to amplify the entire PV capsid coding region from quite small amounts of viral RNA in PV-positive stool extracts.
Validation of direct PV detection using the ECRA method with PV-positive stool extracts.
We performed the ECRA method according to the flow chart presented in Fig. 1C, with 88 PV-positive stool extracts stored in the US CDC. We confirmed the presence of PV by using the standard cell culture system, and we found that 4 of 88 samples (samples 40, 67, 73, and 86) were negative for PV. We performed the ECRA method for all 88 samples, and we successfully amplified the expected product for 100% of the PV-positive samples (84 of 84 samples) and 50% of the PV-negative samples (2 of 4 samples [samples 40 and 67]) (Table 1; also see Table S1 in the supplemental material). To facilitate VP1 sequencing, we performed the second PCR (nested) by using the first RT-PCR products as the template with primers Y7, Q8, and PV1A, and we obtained corresponding cDNA for 99% of PV-positive samples (83/84 samples) and 50% of PV-negative samples (2 of 4 samples [samples 40 and 67]). This suggested that the ECRA method provides efficient nested amplification and sequencing of VP1 directly from PV-positive stool extract RNA.
TABLE 1.
Detection and identification of PV from stool extracts without PV extraction with the PVR MB method
| Analysis type and primers/probes | No. (%) of PV-positive stool extracts (of 84 samples) | No. of PV-negative stool extracts (of 4 samples) | Total no. (%) (of 88 samples) |
|---|---|---|---|
| First RT-PCR (ECRA method) | 84 (100) | 2 | 86 (98) |
| Second PCR (template DNA preparation for VP1 nucleotide sequence analysis) | 83 (99) | 2 | 85 (97) |
| Conventional ITD rRT-PCR (detection of PV from ECRA products)a with pan-PV primers/probe (detection of PV) | 75 (89) | 2 | 77 (88) |
| Improved PV ITD rRT-PCR (detection/ITD of PV from ECRA products)a | |||
| With pan-PV primers/probe (detection of PV) | 81 (96) | 2 | 83 (94) |
| With PV1, PV2, PV3, and Sabin 1, 2, and 3 primers/probes (ITD of PV) | 79 (90) | 2 | 81 (92) |
| VP1 nucleotide sequence analysis (detection of PV VP1) | 78 (93) | 1 | 79 (94) |
Detection of PV from ECRA products was evaluated with conventional and improved real-time PCR kits.
For detection of PV in the stool extracts, we first performed conventional and improved PV ITD rRT-PCR by using the first RT-PCR products with a pan-PV primer/probe set. PV was detected by conventional rRT-PCR for 89% of the PV-positive samples (75/84 samples) and 50% of the PV-negative samples (2/4 samples) and by improved rRT-PCR for 96% of the PV-positive samples (81/84 samples) and 50% of the PV-negative samples (2/4 samples). We also performed type-specific PV ITD rRT-PCR and detected each type of PV (90% for the PV-positive samples [79/84 samples] and 50% for the PV-negative samples [2/4 samples]). Next, we performed nucleotide sequence analysis of the VP1 coding region starting with the second PCR with the Y7/Q8 primer set, which is currently used for PV nucleotide sequence analysis in the Global Polio Laboratory Network (10). With second PCR products obtained with the Y7/Q8 primers, PV VP1 sequences were obtained with the Y7 primer for only 68% of the samples (60/88 samples). With sequencing with the PV1A primer, the PV VP1 sequence was obtained from an additional 8 samples but still the overall detection rate was low (78% [68/88 samples]). We also performed nucleotide sequence analysis with the first RT-PCR products and the second PCR products obtained with Y7/PV1A primers. In these analyses, the PV VP1 sequence was obtained from an additional 11 samples (94% [79/88 samples]). In addition to PV, EV-C viruses were detected in PV-positive samples with a high frequency (27% [23/86 samples]). We found discordant results for 3 samples (samples 15, 70, and 81) in the results of PV ITD rRT-PCR and nucleotide sequence analysis, compared with those of the original virus isolation. These results suggested that PV could be efficiently detected by using improved PV ITD rRT-PCR with the first RT-PCR products obtained with the ECRA method (96% detection with the pan-PV primer/probe set). The nucleotide sequence of the VP1 coding region could be obtained from the first RT-PCR products or from the second PCR products (93% of PV-positive samples).
Validation of direct PV detection using the ECRA method with PV extracted from PV-positive stool extracts with PVR MB.
Next, we used the ECRA method with PV extracted with PVR MB from the stool extracts that were negative for PV in improved and/or conventional PV ITD rRT-PCR assays (3 and 8 samples, respectively [samples 11, 29, and 41 and samples 13, 46, 47, 64, 65, 24, 73, and 86, respectively]) or in PV VP1 nucleotide sequence analysis (3 samples [samples 40, 61, and 68]) or samples that showed discordant results with original virus isolation records (3 samples [samples 15, 70, and 81]). Three samples (samples 24, 73, and 86) were not available due to the lack of remaining sample volume; thus, a total of 14 samples were examined for PV extraction with PVR MB (Table 2). By using PV extracted with PVR MB, PV was successfully detected in 3 samples that were negative for PV with the improved PV ITD rRT-PCR (3 of 3 samples [samples 11, 29, and 41]). Therefore, with untreated stool extracts and PV extracted with PVR MB, PV was detected in all PV-positive stool extracts with the improved PV ITD real-time RT-PCR in combination with the ECRA method (84/84 samples [100%]). Among 5 samples that were negative for PV by conventional PV ITD rRT-PCR, PV could be detected in 3 samples (3 of 5 samples [samples 46, 64, and 65]). We detected the PV VP1 nucleotide sequence after PV extraction from 4 of 6 PV VP1-negative samples (samples 11, 29, 41, and 68), thus resulting in PV VP1 detection from 82/84 PV-positive samples (samples 47 and 61 were negative for PV in this VP1 nucleotide sequence analysis). For 3 samples with discordant results, PV was detected and showed results consistent with those of virus isolation (3 of 3 samples [samples 15, 70, and 81]). These results suggest that PV extraction by PVR MB could complement direct detection of PV from untreated stool extracts by current systems with the ECRA method.
TABLE 2.
Detection and identification of PV extracted from stool extracts with the PVR MB methoda
| Analysis type and primers/probes | No. (%) of PV-positive stool extracts (of 13 samples) | No. of PV-negative stool extracts (of 1 sample) | Total no. (%) (of 14 samples) |
|---|---|---|---|
| First RT-PCR (ECRA method) | 12 (92) | 1 | 13 (93) |
| Second PCR (template DNA preparation for VP1 nucleotide sequence analysis) | 12 (92) | 1 | 13 (93) |
| Conventional ITD rRT-PCR (detection/ITD of PV from ECRA products) | |||
| With pan-PV primers/probe (detection of PV) | 9 (69) | 0 | 10 (71) |
| With PV1, PV2, PV3, and Sabin 1, 2, and 3 primers/probes (ITD of PV) | 9b (100) | 0 | 9 (64) |
| VP1 nucleotide sequence analysis (detection of PV VP1) | 10 (77) | 0 | 10 (71) |
Stool extracts that were negative in the conventional PV ITD rRT-PCR assay or in the PV VP1 nucleotide sequence analysis or showed discordant results with the original virus isolation records were examined, except for 3 samples that were not available because of lack of sample volume (total of 14 samples examined).
The total number of extracts in this analysis was 9.
DISCUSSION
In the present study, we developed a method to efficiently amplify the entire capsid coding region (about 3.9 kb) of human enteroviruses, including PV, from as few as 50 copies of the viral genome (ECRA method). One of the essential factors for this high efficiency of amplification seemed to be the intrinsic compatibility of the primer set and the polymerase; we could not directly amplify the VP1 coding region with a degenerate primer set (Y7/Q8 primers) under conditions similar to those used for the ECRA method (data not shown). Another factor would be the elongation enhancer in the buffer system, which allowed sustained elongation of polymerase after 30 cycles of PCR. In the polio eradication program, both sensitive detection of PV and nucleotide sequence analysis of the VP1 region (about 900 nucleotides [nt]) are essential for correct identification of PV isolates. It should be noted that the virus isolates are essential for study of phenotypes but not for routine PV surveillance and that the virus isolates are unavailable in the detection system using the ECRA method, in contrast to cell culture. In case a need arises for any further diagnosis or study, all AFP stool samples are stored by reference laboratories for 1 year. For sensitive real-time PCR, short nucleotide sequences (<200 nt) in the target regions were generally used as the targets for amplification, for optimized sensitivity. The existence of a target gene detected by sensitive real-time PCR does not necessarily indicate the availability of the entire nucleotide sequence of the gene; thus, different methods are required for this purpose. The ECRA method would serve as a bridge between sensitive PV detection and VP1 nucleotide sequence analysis with currently implemented rRT-PCR and VP1-sequencing systems in the Global Polio Laboratory Network.
A major challenge in PV detection in stool by rRT-PCR is low sensitivity. Modification of the conditions and reagents have improved the sensitivity about 100-fold, compared to conventional systems (detection limits of about 100 to 1,000 copies of PV viral RNA) (12). Consistently, the PV detection rate for the improved PV ITD rRT-PCR method was increased over that for the conventional method (96% versus 89%, without PV-specific extraction with PVR MB) (Table 1). Some PV-negative stool extracts from PV-positive AFP patients could contain 150 to 260 copies of PV viral RNA in 5 μl of purified viral RNA solution or in 10 μl of original stool extracts (6). The amounts of PV genome in the stool extracts were at or below the detection limits for the improved PV ITD rRT-PCR but could be amplified with the ECRA method. The precise amounts of PV in the PV-positive stool extracts could not be determined, in part because of coexisting EVs, but most of the samples caused complete CPE in the inoculated L20B cells at 3 days postinoculation (p.i.) (71/88 samples), and 15 samples caused complete CPE at 5 days p.i. or later (see Table S1 in the supplemental material). Since about 10 CCID50 of PV could cause complete CPE in the inoculated cells within 2 days p.i (6), the observations described above suggested that most of the stool extracts examined in this study contained minimal infectivity of less than 10 CCID50 (about 103 to 104 viral genome copies) in 100 μl stool extract. Nevertheless, PVs were detected from all of these samples with the improved real-time PCR system in combination with the ECRA and PVR MB extraction methods, suggesting high sensitivity of this system, comparable to that of cell culture.
There were 3 PV-positive stool extract samples (samples 11, 29, and 41) from which PVs were detected only after PV extraction by using PVR MB. In these samples, non-PV EV-C strains (CVA11, CVA13, and CVA17) were the predominant populations, as detected in nucleotide sequence analysis. This suggested that EVs coexisting in the same stool extracts could interfere or compete with amplification of the PV capsid coding region with the ECRA method, possibly because of conservation of the primer sites among EV-C members (primers 5′NTR-529–552+ and Cre-4485–4460−) (Fig. 1A). PV extraction with PVR MB prior to ECRA seems essential for some samples that contain large amounts of non-PV EV-C species. We also found one sample (sample 13) from which PV was detected before PV extraction but not after extraction. This suggested that quite small amounts of PV might not be efficiently extracted by using PVR MB. Therefore, with the current specificity of the ECRA method and VP1 sequencing and the efficiency of PV extraction with PVR MB, along with the sensitivity of real-time PCR systems, analysis of viral RNAs extracted from both PVR MB-treated and non-MB-treated stool extracts might be essential to attain a high PV detection rate.
The PV ITD rRT-PCR system with the pan-PV primer/probe set could detect PV from all of the PV-positive stool extracts with cDNAs obtained by the ECRA method in combination with the PVR MB method. However, further improvement of the sensitivity with the pan-PV primer/probe set will be required to detect PV without PV extraction. A new type-specific PV ITD RT-PCR assay currently under development at the US CDC might also be useful to type pan-PV-positive samples (samples 47, 61, and 68) that could not be typed with the current method.
We found discordant results of PV ITD rRT-PCR analysis and virus isolation for 3 samples (samples 15, 70, and 81). Interestingly, after PV extraction with PVR MB, concordant results were obtained. In our previous study, we found the presence of noninfectious PV strains that could not interact with PVR MB in some stool extracts (6). This observation indicated that noninfectious PV could be the predominant source of viral RNA in some stool extracts and that PV extraction with PVR MB is effective for detecting “viable” PV strains in the samples. Although the importance of noninfectious PV in the circulation is currently unknown, this suggested higher sensitivity of this new detection method than cell culture.
To detect the nucleotide sequence of the PV VP1 coding region, the second PCR products (with the Y7/Q8 and Y7/PV1A primer sets) and the first RT-PCR products were used as the templates with 2 sequencing primers (Y7 and PV1A primers) (10). By using these PCR products with PV1A sequencing primers in combination with the PVR MB extraction method, we managed to detect PV VP1 nucleotide sequences from 82 of 84 PV-positive stool extracts, except for 2 samples (samples 47 and 61). PV was detected in these PV VP1-negative samples by PV ITD real-time RT-PCR, suggesting low specificity/sensitivity of the sequencing primers to detect PV in the context of the current ECRA method, which instead amplified VP1 cDNAs of CVA13, CVA17, and CVA20. In 2014, wild-type PV of serotype 1 continues to circulate (13), type 2 wild-type PV strains have been eliminated (the last isolation was in India in 1999), and type 3 wild-type PV strains have not been isolated since 2012. We found some PV strains that were detected only by nucleotide sequence analysis and not by cell culture (samples 14, 15, and 81), suggesting the potential for higher sensitivity of nucleotide sequence analysis after ECRA than cell culture.
In summary, we developed an efficient ECRA method for sensitive detection of PV with the currently available PV ITD rRT-PCR and VP1-sequencing systems. The combined use with the PVR MB PV-specific extraction method was effective in excluding non-PV EV types. This ECRA method would be useful for further development of a highly sensitive direct detection system for PV in stool extracts without the need for virus isolation in cell culture.
Supplementary Material
ACKNOWLEDGMENTS
We are grateful to Junko Wada for her excellent technical assistance.
This report is based on research funded by a grant from the World Health Organization for a collaborative research project of the Global Polio Eradication Initiative.
Footnotes
Supplemental material for this article may be found at http://dx.doi.org/10.1128/JCM.02384-14.
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