Detection of Severe Acute Respiratory Syndrome Coronavirus in Stool Sp
http://www.100md.com
《微生物临床杂志》
Sunnybrook Health Sciences Centre, Toronto, Ontario
St. Joseph's Healthcare, Hamilton, Ontario
Hospital for Sick Children, Toronto, Ontario
the Toronto Medical Laboratories and Mount Sinai Hospital, Toronto, Ontario
St. Michael's Hospital
the Central Public Health Laboratory, Toronto, Ontario
the University of Toronto, Toronto, Ontario
the Provincial Laboratory for Public Health and the University of Calgary, Calgary, Alberta, Canada
ABSTRACT
Three commercially available real-time reverse transcriptase PCR assays (the Artus RealArt HPA coronavirus LightCycler, the Artus RealArt HPA coronavirus Rotor-Gene, and the EraGen severe acute respiratory syndrome coronavirus POL assay) and three RNA extraction methodologies were evaluated for the detection of severe acute respiratory syndrome coronavirus RNA from 91 stool specimens. The assays' sensitivities were highest (58% to 75%) for specimens obtained 8 to 21 days after symptom onset. The assays were less sensitive when specimens were obtained less than 8 days or more than 21 days after the onset of symptoms. All assays were 100% specific.
TEXT
The discovery of a novel coronavirus (CoV) as the causative agent of severe acute respiratory syndrome (SARS) has led to the development of reverse transcriptase (RT) PCR-based assays for detecting this virus (1, 7, 9, 15). Although SARS is associated primarily with fever and involvement of the respiratory tract, gastrointestinal symptoms may also occur (2, 10). The presence of active viral replication within the large and small intestines, as demonstrated by biopsy samples obtained from colonoscopies or autopsies, suggests that stool may be a useful specimen for SARS-CoV detection (11). The rate of detection of SARS-CoV from stools has been at least as high as that from other clinical specimens by using conventional nested RT-PCR (16), and stool specimens have been a useful alternative to respiratory samples for the detection of SARS-CoV (14, 18). Although there have been evaluations of both conventional and real-time RT-PCR-based assays (12, 19), few studies have evaluated newer, second-generation assays for the detection of SARS-CoV RNA from stool specimens (5). In an outbreak that is short-lived and involves an emerging pathogen, the evaluation of newer diagnostic tests is difficult. While the specimens used were not ideal, we sought to overcome this problem with the use of frozen, archived stool specimens. In this study, we evaluated three RNA extraction methods and three commercially available second-generation, real-time RT-PCR-based assays for the detection of SARS-CoV by using a collection of archived stool samples. We compared these results to those obtained using a conventional nested RT-PCR assay performed at the time of receipt of these same specimens in 2003. The results of this evaluation may be useful, should human infection due to SARS-CoV recur.
Clinical specimens. A total of 91 stool specimens from 57 patients were obtained between April and August 2003 during an outbreak of SARS in Toronto, Ontario, Canada. Thirty-four patients met the WHO case definition for probable or suspected SARS (17) and were seropositive for SARS-CoV antibodies during the convalescent phase of illness, as determined by the Euroimmun immunofluorescent assay (immunoglobulin G; Euroimmun, Lübeck, Germany) (8, 13, 14). Twenty-three patients with other respiratory illnesses did not meet the case definition for SARS and were seronegative for SARS-CoV antibodies. There were 56 stool specimens available for testing from the seropositive individuals and 35 specimens from those who were seronegative.
RNA extraction. RNA was extracted from fresh stool samples upon receipt in the laboratory in 2003 using a viral RNA minikit (QIAGEN Inc., Mississauga, Ontario, Canada) according to the manufacturer's instructions, using 140 μl of a 10% (wt/vol) stool suspension made in 5 ml of sterile diethylpyrocarbonate-treated water (Invitrogen, Burlington, Ontario, Canada). RNA was eluted in a final volume of 50 μl of elution buffer. Stool specimens were then frozen at –80°C until they were retested for this evaluation.
For the present evaluation, stool samples were thawed, and a 10% (wt/vol) stool suspension was made in sterile diethylpyrocarbonate-treated water. RNA was extracted with three different extraction methods, each performed in accordance with the manufacturer's instructions: the viral RNA minikit, the miniMag (bioMerieux, St. Laurent, Quebec, Canada), and the MagaZorb RNA miniprep kit (Cortex Biochem, Inc., San Leandro, CA). A sample of 140 μl was used for both the viral RNA minikit and the miniMag extractions, while 200 μl was used for the MagaZorb extraction. RNA was eluted in a final volume of 50 μl elution buffer.
RT-PCR assays. Conventional, nested RT-PCR was performed as described by Drosten et al. (6). Real-time RT-PCR with the RealArt HPA coronavirus (HPAC) LightCycler and RealArt HPAC Rotor-Gene kits (Artus GmbH, Hamburg, Germany) and the SARS coronavirus POL assay (EraGen Biosciences, Madison, WI) were performed according to the manufacturers' instructions. The target for the real-time RT-PCR assays was the viral polymerase gene, although details regarding the target for each assay are not available as the test kits evaluated are proprietary. All the RNA eluates were tested using the EraGen and Artus kits. The EraGen and the Artus HPAC LightCycler assays were performed with a LightCycler version 1.2 (Roche Diagnostics, Laval, Quebec), and the Artus HPAC Rotor-Gene assay was performed with a Rotor-Gene model 3000 (Corbett Life Science, Sydney, Australia). Only the Artus kits provided positive controls with which to generate a standard curve for viral load quantitation, but this feature of the assays was not evaluated.
All methods were compared, and their clinical sensitivities and specificities were calculated using serologic evidence for the presence of SARS-CoV antibodies as the gold standard.
A summary of all test results is shown in Table 1. Of the 91 specimens tested, 67 were negative and 17 were positive for SARS-CoV RNA by all seven methods. Twenty-three specimens were found positive for viral RNA by using three or more RT-PCR assays. One specimen was positive by only one of the assay methods (EraGen). Although this outcome may have represented a false-positive test result, the patient met the case definition for SARS and was seropositive for SARS-CoV antibodies.
Although all the assays had 100% specificity, the sensitivities were low, ranging from 34 to 40% (Table 2). There were no statistically significant differences between the sensitivities or specificities of the assays evaluated. However, when the assays were analyzed according to the time of specimen collection from the onset of symptoms, there were certain times that were associated with an improved ability to detect viral RNA (Fig. 1). The sensitivities of RT-PCR for the 13 specimens collected 7 days from the onset of symptoms ranged from 46 to 62%, whereas between 8 and 21 days after symptom onset, the sensitivities ranged from 58 to 75% (12 specimens). For the 31 specimens obtained >21 days after symptom onset, the sensitivities were only 13 to 19%. When individual RT-PCR assay results were compared to a modified gold standard of three or more assays yielding a positive result, the sensitivities improved (range, 83% to 96%; data not shown). None of the 35 specimens collected from the 23 non-SARS patient samples was positive for SARS-CoV RNA.
The newer assays performed as well as a conventional nested RT-PCR assay for the detection of SARS-CoV RNA from stool samples, even after sample storage at –80°C for approximately 2 years. This is consistent with a previous report showing that SARS-CoV could withstand freezing and thawing and was stable for many weeks in clinical specimens maintained at 4°C or at –70°C (3). The newer real-time PCR assays produced more rapid results than the conventional method, which was labor-intensive and required 5 h to complete.
The results of our RT-PCR testing for SARS-CoV in stool samples are similar to those previously reported (4, 14, 16). The negative RT-PCR results for serologically confirmed SARS patients obtained in this study could have been due to suboptimal timing of specimen collection (4, 14). Stool samples collected during the first week of illness or more than 21 days after symptom onset often yielded false-negative results, possibly due to a decreased viral load. More than half (55%) of the specimens from SARS patients that were available for testing had been collected more than 21 days after the onset of symptoms. An additional limitation of this study was the lack of availability of multiple samples from a large number of patients collected over an extended period of time during the course of illness. In the majority of cases, only one stool sample was available for testing. The use of archived stool samples may also have affected the yield, although our data do not appear to suggest that freezing affected RT-PCR assay results.
The results of this evaluation indicate that these commercially available real-time RT-PCR assays with newer extraction methodologies are highly specific and at least as sensitive as conventional nested RT-PCR assays for the detection of SARS-CoV. Major advantages of the newer tests are the ease with which they are performed and their rapidity. Any of these assays would be suitable for use, should a need to detect SARS-CoV arise in the future. However, it is clear that should SARS recur in humans, there is still a need for more knowledge about optimal diagnostic test strategies (appropriate specimens and their timing) and for improved diagnostic assays.
ACKNOWLEDGMENTS
Additional members of the Ontario Laboratory Working Group for the Rapid Diagnosis of Emerging Pathogens include Ilene Guglielmi and Aimin Li, Central Public Health Laboratory, Toronto, Ontario; Donald Low and Allison McGeer, Mount Sinai Hospital, Toronto, Ontario; Anne Matlow, Hospital for Sick Children, Toronto, Ontario; Kevin Katz, Mona Loutfy, and Barbara Mederski, North York General Hospital, Toronto, Ontario; Marvin Gerson, Reena Lovinsky, David Rose, and Zev Shainhouse, The Scarborough Hospital, Toronto, Ontario; Sigmund Krajden, St. Joseph's Health Centre, Toronto, Ontario; Roslyn Devlin, St. Michael's Hospital, Toronto, Ontario; Anita Rachlis and Mary Vearncombe, Sunnybrook Health Sciences Centre, Toronto, Ontario; James Downey and Roland Skrastins, Toronto East General Hospital, Toronto, Ontario; Wayne Gold, University Health Network, Toronto, Ontario; and Elizabeth Phillips, St. Paul's Hospital, Vancouver, British Columbia, Canada.
We thank Catherine Harlton-Strezov for database management.
This study was funded in part by the Ontario Ministry of Health and Long-Term Care and by the Canadian Institutes for Health Research.
FOOTNOTES
Corresponding author. Mailing address: Department of Microbiology, Sunnybrook Health Sciences Centre, B121-2075 Bayview Ave., Toronto, Ontario, Canada M4N 3M5. Phone: 416-480-4549. Fax: 416-480-6845. E-mail: lisa.louie@sunnybrook.ca.
Published ahead of print on 30 August 2006.
REFERENCES
Adachi, D., G. Johnson, R. Draker, M. Ayers, T. Mazzulli, P. J. Talbot, and R. Tellier. 2004. Comprehensive detection and identification of human coronaviruses, including the SARS-associated coronavirus, with a single RT-PCR assay. J. Virol. Methods 122:29-36.
Booth, C. M., L. M. Matukas, G. A. Tomlinson, A. R. Rachlis, D. B. Rose, H. A. Dwosh, S. L. Walmsley, T. Mazzulli, M. Avendano, P. Derkach, I. E. Ephtimios, I. Kitai, B. D. Mederski, S. B. Shadowitz, W. L. Gold, L. A. Hawryluck, E. Rea, J. S. Chenkin, D. W. Cescon, S. M. Poutanen, and A. S. Detsky. 2003. Clinical features and short-term outcomes of 144 patients with SARS in the greater Toronto area. JAMA 289:2801-2809.
Chan, K. H., L. L. Poon, V. C. Cheng, Y. Guan, I. F. Hung, J. Kong, L. Y. Yam, W. H. Seto, K. Y. Yuen, and J. S. Peiris. 2004. Detection of SARS coronavirus in patients with suspected SARS. Emerg. Infect. Dis. 10:294-299.
Chan, P. K., W. K. To, K. C. Ng, R. K. Lam, T. K. Ng, R. C. Chan, A. Wu, W. C. Yu, N. Lee, D. S. Hui, S. T. Lai, E. K. Hon, C. K. Li, J. J. Sung, and J. S. Tam. 2004. Laboratory diagnosis of SARS. Emerg. Infect. Dis. 10:825-831.
Drosten, C., L.-L. Chiu, M. Panning, H. N. Leong, W. Preiser, J. S. Tam, S. Günther, S. Kramme, P. Emmerich, W. L. Ng, H. Schmitz, and E. S. Koay. 2004. Evaluation of advanced reverse transcription-PCR assays and an alternative PCR target region for detection of severe acute respiratory syndrome-associated coronavirus. J. Clin. Microbiol. 42:2043-2047.
Drosten, C., S. Gunther, W. Preiser, S. van der Werf, H. R. Brodt, S. Becker, H. Rabenau, M. Panning, L. Kolesnikova, R. A. Fouchier, A. Berger, A. M. Burguiere, J. Cinatl, M. Eickmann, N. Escriou, K. Grywna, S. Kramme, J. C. Manuguerra, S. Muller, V. Rickerts, M. Sturmer, S. Vieth, H. D. Klenk, A. D. Osterhaus, H. Schmitz, and H. W. Doerr. 2003. Identification of a novel coronavirus in patients with severe acute respiratory syndrome. N. Engl. J. Med. 348:1967-1976.
Emery, S. L., D. D. Erdman, M. D. Bowen, B. R. Newton, J. M. Winchell, R. F. Meyer, S. Tong, B. T. Cook, B. P. Holloway, K. A. McCaustland, P. A. Rota, B. Bankamp, L. E. Lowe, T. G. Ksiazek, W. J. Bellini, and L. J. Anderson. 2004. Real-time reverse transcription-polymerase chain reaction assay for SARS-associated coronavirus. Emerg. Infect. Dis. 10:311-316.
He, Q., I. Manopo, L. Lu, B. P. Leung, H. H. Cheng, A. E. Ling, L. L. Chee, S. W. Chan, E. E. Ooi, Y. L. Sin, B. Ang, and J. Kwang. 2005. Novel immunofluorescence assay using recombinant nucleocapsid-spike fusion protein as antigen to detect antibodies against severe acute respiratory syndrome coronavirus. Clin. Diagn. Lab. Immunol. 12:321-328.
Juang, J. L., T. C. Chen, S. S. Jiang, C. A. Hsiung, W. C. Chen, G. W. Chen, S. M. Lin, J. H. Lin, S. C. Chiu, and Y. K. Lai. 2004. Coupling multiplex RT-PCR to a gene chip assay for sensitive and semiquantitative detection of severe acute respiratory syndrome-coronavirus. Lab. Investig. 84:1085-1091.
Lee, N., D. Hui, A. Wu, P. Chan, P. Cameron, G. M. Joynt, A. Ahuja, M. Y. Yung, C. B. Leung, K. F. To, S. F. Lui, C. C. Szeto, S. Chung, and J. J. Sung. 2003. A major outbreak of severe acute respiratory syndrome in Hong Kong. N. Engl. J. Med. 348:1986-1994.
Leung, W. K., K. F. To, P. K. Chan, H. L. Chan, A. K. Wu, N. Lee, K. Y. Yuen, and J. J. Sung. 2003. Enteric involvement of severe acute respiratory syndrome-associated coronavirus infection. Gastroenterology 125:1011-1017.
Mahony, J. B., A. Petrich, L. Louie, X. Song, S. Chong, M. Smieja, M. Chernesky, M. Loeb, S. Richardson, and the Ontario Laboratory Working Group for the Rapid Diagnosis of Emerging Infections. 2004. Performance and cost evaluation of one commercial and six in-house conventional and real-time reverse transcription-PCR assays for detection of severe acute respiratory syndrome coronavirus. J. Clin. Microbiol. 42:1471-1476.
Manopo, I., L. Lu, Q. He, L. L. Chee, S. W. Chan, and J. Kwang. 2005. Evaluation of a safe and sensitive Spike protein-based immunofluorescence assay for the detection of antibody responses to SARS-CoV. J. Immunol. Methods 296:37-44.
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Poon, L. L., K. H. Chan, O. K. Wong, T. K. Cheung, I. Ng, B. Zheng, W. H. Seto, K. Y. Yuen, Y. Guan, and J. S. Peiris. 2004. Detection of SARS coronavirus in patients with severe acute respiratory syndrome by conventional and real-time quantitative reverse transcription-PCR assays. Clin. Chem. 50:67-72.
Tang, P., M. Louie, S. E. Richardson, M. Smieja, A. E. Simor, F. Jamieson, M. Fearon, S. M. Poutanen, T. Mazzulli, R. Tellier, J. Mahony, M. Loeb, A. Petrich, M. Chernesky, A. McGeer, D. E. Low, E. Phillips, S. Jones, N. Bastien, Y. Li, D. Dick, A. Grolla, L. Fernando, T. F. Booth, B. Henry, A. R. Rachlis, L. M. Matukas, D. B. Rose, R. Lovinsky, S. Walmsley, W. L. Gold, and S. Krajden. 2004. Interpretation of diagnostic laboratory tests for severe acute respiratory syndrome: the Toronto experience. Can. Med. Assoc. J. 170:47-54.
World Health Organization. 2006. Case definitions for surveillance of severe acute respiratory syndrome (SARS). [Online.] http://www. who. int/csr/sars/casedefinition/en. Accessed 24 February 2006.
Wu, H. S., S. C. Chiu, T. C. Tseng, S. F. Lin, J. H. Lin, Y. H. Hsu, M. C. Wang, T. L. Lin, W. Z. Yang, T. L. Ferng, K. H. Huang, L. C. Hsu, L. L. Lee, J. Y. Yang, H. Y. Chen, S. P. Su, S. Y. Yang, S. Y. Lin, T. H. Lin, and I. S. Su. 2004. Serologic and molecular biologic methods for SARS-associated coronavirus infection, Taiwan. Emerg. Infect. Dis. 10:304-310.
Yam, W. C., K. H. Chan, L. L. M. Poon, Y. Guan, K. Y. Yuen, W. H. Seto, and J. S. M. Peiris. 2003. Evaluation of reverse transcription-PCR assays for rapid diagnosis of severe acute respiratory syndrome associated with a novel coronavirus. J. Clin. Microbiol. 41:4521-4524.(L. Louie, A. E. Simor, S. Chong, K. Luin)
St. Joseph's Healthcare, Hamilton, Ontario
Hospital for Sick Children, Toronto, Ontario
the Toronto Medical Laboratories and Mount Sinai Hospital, Toronto, Ontario
St. Michael's Hospital
the Central Public Health Laboratory, Toronto, Ontario
the University of Toronto, Toronto, Ontario
the Provincial Laboratory for Public Health and the University of Calgary, Calgary, Alberta, Canada
ABSTRACT
Three commercially available real-time reverse transcriptase PCR assays (the Artus RealArt HPA coronavirus LightCycler, the Artus RealArt HPA coronavirus Rotor-Gene, and the EraGen severe acute respiratory syndrome coronavirus POL assay) and three RNA extraction methodologies were evaluated for the detection of severe acute respiratory syndrome coronavirus RNA from 91 stool specimens. The assays' sensitivities were highest (58% to 75%) for specimens obtained 8 to 21 days after symptom onset. The assays were less sensitive when specimens were obtained less than 8 days or more than 21 days after the onset of symptoms. All assays were 100% specific.
TEXT
The discovery of a novel coronavirus (CoV) as the causative agent of severe acute respiratory syndrome (SARS) has led to the development of reverse transcriptase (RT) PCR-based assays for detecting this virus (1, 7, 9, 15). Although SARS is associated primarily with fever and involvement of the respiratory tract, gastrointestinal symptoms may also occur (2, 10). The presence of active viral replication within the large and small intestines, as demonstrated by biopsy samples obtained from colonoscopies or autopsies, suggests that stool may be a useful specimen for SARS-CoV detection (11). The rate of detection of SARS-CoV from stools has been at least as high as that from other clinical specimens by using conventional nested RT-PCR (16), and stool specimens have been a useful alternative to respiratory samples for the detection of SARS-CoV (14, 18). Although there have been evaluations of both conventional and real-time RT-PCR-based assays (12, 19), few studies have evaluated newer, second-generation assays for the detection of SARS-CoV RNA from stool specimens (5). In an outbreak that is short-lived and involves an emerging pathogen, the evaluation of newer diagnostic tests is difficult. While the specimens used were not ideal, we sought to overcome this problem with the use of frozen, archived stool specimens. In this study, we evaluated three RNA extraction methods and three commercially available second-generation, real-time RT-PCR-based assays for the detection of SARS-CoV by using a collection of archived stool samples. We compared these results to those obtained using a conventional nested RT-PCR assay performed at the time of receipt of these same specimens in 2003. The results of this evaluation may be useful, should human infection due to SARS-CoV recur.
Clinical specimens. A total of 91 stool specimens from 57 patients were obtained between April and August 2003 during an outbreak of SARS in Toronto, Ontario, Canada. Thirty-four patients met the WHO case definition for probable or suspected SARS (17) and were seropositive for SARS-CoV antibodies during the convalescent phase of illness, as determined by the Euroimmun immunofluorescent assay (immunoglobulin G; Euroimmun, Lübeck, Germany) (8, 13, 14). Twenty-three patients with other respiratory illnesses did not meet the case definition for SARS and were seronegative for SARS-CoV antibodies. There were 56 stool specimens available for testing from the seropositive individuals and 35 specimens from those who were seronegative.
RNA extraction. RNA was extracted from fresh stool samples upon receipt in the laboratory in 2003 using a viral RNA minikit (QIAGEN Inc., Mississauga, Ontario, Canada) according to the manufacturer's instructions, using 140 μl of a 10% (wt/vol) stool suspension made in 5 ml of sterile diethylpyrocarbonate-treated water (Invitrogen, Burlington, Ontario, Canada). RNA was eluted in a final volume of 50 μl of elution buffer. Stool specimens were then frozen at –80°C until they were retested for this evaluation.
For the present evaluation, stool samples were thawed, and a 10% (wt/vol) stool suspension was made in sterile diethylpyrocarbonate-treated water. RNA was extracted with three different extraction methods, each performed in accordance with the manufacturer's instructions: the viral RNA minikit, the miniMag (bioMerieux, St. Laurent, Quebec, Canada), and the MagaZorb RNA miniprep kit (Cortex Biochem, Inc., San Leandro, CA). A sample of 140 μl was used for both the viral RNA minikit and the miniMag extractions, while 200 μl was used for the MagaZorb extraction. RNA was eluted in a final volume of 50 μl elution buffer.
RT-PCR assays. Conventional, nested RT-PCR was performed as described by Drosten et al. (6). Real-time RT-PCR with the RealArt HPA coronavirus (HPAC) LightCycler and RealArt HPAC Rotor-Gene kits (Artus GmbH, Hamburg, Germany) and the SARS coronavirus POL assay (EraGen Biosciences, Madison, WI) were performed according to the manufacturers' instructions. The target for the real-time RT-PCR assays was the viral polymerase gene, although details regarding the target for each assay are not available as the test kits evaluated are proprietary. All the RNA eluates were tested using the EraGen and Artus kits. The EraGen and the Artus HPAC LightCycler assays were performed with a LightCycler version 1.2 (Roche Diagnostics, Laval, Quebec), and the Artus HPAC Rotor-Gene assay was performed with a Rotor-Gene model 3000 (Corbett Life Science, Sydney, Australia). Only the Artus kits provided positive controls with which to generate a standard curve for viral load quantitation, but this feature of the assays was not evaluated.
All methods were compared, and their clinical sensitivities and specificities were calculated using serologic evidence for the presence of SARS-CoV antibodies as the gold standard.
A summary of all test results is shown in Table 1. Of the 91 specimens tested, 67 were negative and 17 were positive for SARS-CoV RNA by all seven methods. Twenty-three specimens were found positive for viral RNA by using three or more RT-PCR assays. One specimen was positive by only one of the assay methods (EraGen). Although this outcome may have represented a false-positive test result, the patient met the case definition for SARS and was seropositive for SARS-CoV antibodies.
Although all the assays had 100% specificity, the sensitivities were low, ranging from 34 to 40% (Table 2). There were no statistically significant differences between the sensitivities or specificities of the assays evaluated. However, when the assays were analyzed according to the time of specimen collection from the onset of symptoms, there were certain times that were associated with an improved ability to detect viral RNA (Fig. 1). The sensitivities of RT-PCR for the 13 specimens collected 7 days from the onset of symptoms ranged from 46 to 62%, whereas between 8 and 21 days after symptom onset, the sensitivities ranged from 58 to 75% (12 specimens). For the 31 specimens obtained >21 days after symptom onset, the sensitivities were only 13 to 19%. When individual RT-PCR assay results were compared to a modified gold standard of three or more assays yielding a positive result, the sensitivities improved (range, 83% to 96%; data not shown). None of the 35 specimens collected from the 23 non-SARS patient samples was positive for SARS-CoV RNA.
The newer assays performed as well as a conventional nested RT-PCR assay for the detection of SARS-CoV RNA from stool samples, even after sample storage at –80°C for approximately 2 years. This is consistent with a previous report showing that SARS-CoV could withstand freezing and thawing and was stable for many weeks in clinical specimens maintained at 4°C or at –70°C (3). The newer real-time PCR assays produced more rapid results than the conventional method, which was labor-intensive and required 5 h to complete.
The results of our RT-PCR testing for SARS-CoV in stool samples are similar to those previously reported (4, 14, 16). The negative RT-PCR results for serologically confirmed SARS patients obtained in this study could have been due to suboptimal timing of specimen collection (4, 14). Stool samples collected during the first week of illness or more than 21 days after symptom onset often yielded false-negative results, possibly due to a decreased viral load. More than half (55%) of the specimens from SARS patients that were available for testing had been collected more than 21 days after the onset of symptoms. An additional limitation of this study was the lack of availability of multiple samples from a large number of patients collected over an extended period of time during the course of illness. In the majority of cases, only one stool sample was available for testing. The use of archived stool samples may also have affected the yield, although our data do not appear to suggest that freezing affected RT-PCR assay results.
The results of this evaluation indicate that these commercially available real-time RT-PCR assays with newer extraction methodologies are highly specific and at least as sensitive as conventional nested RT-PCR assays for the detection of SARS-CoV. Major advantages of the newer tests are the ease with which they are performed and their rapidity. Any of these assays would be suitable for use, should a need to detect SARS-CoV arise in the future. However, it is clear that should SARS recur in humans, there is still a need for more knowledge about optimal diagnostic test strategies (appropriate specimens and their timing) and for improved diagnostic assays.
ACKNOWLEDGMENTS
Additional members of the Ontario Laboratory Working Group for the Rapid Diagnosis of Emerging Pathogens include Ilene Guglielmi and Aimin Li, Central Public Health Laboratory, Toronto, Ontario; Donald Low and Allison McGeer, Mount Sinai Hospital, Toronto, Ontario; Anne Matlow, Hospital for Sick Children, Toronto, Ontario; Kevin Katz, Mona Loutfy, and Barbara Mederski, North York General Hospital, Toronto, Ontario; Marvin Gerson, Reena Lovinsky, David Rose, and Zev Shainhouse, The Scarborough Hospital, Toronto, Ontario; Sigmund Krajden, St. Joseph's Health Centre, Toronto, Ontario; Roslyn Devlin, St. Michael's Hospital, Toronto, Ontario; Anita Rachlis and Mary Vearncombe, Sunnybrook Health Sciences Centre, Toronto, Ontario; James Downey and Roland Skrastins, Toronto East General Hospital, Toronto, Ontario; Wayne Gold, University Health Network, Toronto, Ontario; and Elizabeth Phillips, St. Paul's Hospital, Vancouver, British Columbia, Canada.
We thank Catherine Harlton-Strezov for database management.
This study was funded in part by the Ontario Ministry of Health and Long-Term Care and by the Canadian Institutes for Health Research.
FOOTNOTES
Corresponding author. Mailing address: Department of Microbiology, Sunnybrook Health Sciences Centre, B121-2075 Bayview Ave., Toronto, Ontario, Canada M4N 3M5. Phone: 416-480-4549. Fax: 416-480-6845. E-mail: lisa.louie@sunnybrook.ca.
Published ahead of print on 30 August 2006.
REFERENCES
Adachi, D., G. Johnson, R. Draker, M. Ayers, T. Mazzulli, P. J. Talbot, and R. Tellier. 2004. Comprehensive detection and identification of human coronaviruses, including the SARS-associated coronavirus, with a single RT-PCR assay. J. Virol. Methods 122:29-36.
Booth, C. M., L. M. Matukas, G. A. Tomlinson, A. R. Rachlis, D. B. Rose, H. A. Dwosh, S. L. Walmsley, T. Mazzulli, M. Avendano, P. Derkach, I. E. Ephtimios, I. Kitai, B. D. Mederski, S. B. Shadowitz, W. L. Gold, L. A. Hawryluck, E. Rea, J. S. Chenkin, D. W. Cescon, S. M. Poutanen, and A. S. Detsky. 2003. Clinical features and short-term outcomes of 144 patients with SARS in the greater Toronto area. JAMA 289:2801-2809.
Chan, K. H., L. L. Poon, V. C. Cheng, Y. Guan, I. F. Hung, J. Kong, L. Y. Yam, W. H. Seto, K. Y. Yuen, and J. S. Peiris. 2004. Detection of SARS coronavirus in patients with suspected SARS. Emerg. Infect. Dis. 10:294-299.
Chan, P. K., W. K. To, K. C. Ng, R. K. Lam, T. K. Ng, R. C. Chan, A. Wu, W. C. Yu, N. Lee, D. S. Hui, S. T. Lai, E. K. Hon, C. K. Li, J. J. Sung, and J. S. Tam. 2004. Laboratory diagnosis of SARS. Emerg. Infect. Dis. 10:825-831.
Drosten, C., L.-L. Chiu, M. Panning, H. N. Leong, W. Preiser, J. S. Tam, S. Günther, S. Kramme, P. Emmerich, W. L. Ng, H. Schmitz, and E. S. Koay. 2004. Evaluation of advanced reverse transcription-PCR assays and an alternative PCR target region for detection of severe acute respiratory syndrome-associated coronavirus. J. Clin. Microbiol. 42:2043-2047.
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