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MassTag polymerase chain reaction for differential diagnosis of viral hemorrhagic fevers.

Viral hemorrhagic fevers are associated with high rates of illness and death. Although therapeutic options are limited, early differential diagnosis has implications for containment and may aid in clinical management. We describe a diagnostic system for rapid, multiplex polymerase chain reaction identification of 10 different causes of viral hemorrhagic fevers.

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Increasing international travel, trafficking in wildlife, political instability, and terrorism have made emerging infectious diseases a global concern. Viral hemorrhagic fevers (VHF) warrant specific emphasis because of their high rates of illness and death, and the potential for rapid dissemination by human-to-human transmission. The term "viral hemorrhagic fever" characterizes a severe multisystem syndrome associated with fever, shock, and bleeding diathesis caused by infection with any of several RNA viruses, including Ebola virus and Marburg virus (MARV) (family Filoviridae); Lassa virus (LASV) and the South American hemorrhagic fever viruses Guanarito virus, Junin virus, Machupo virus, and Sabifi virus (Arenaviridae); Rift Valley fever virus (RVFV), Crimean-Congo hemorrhagic fever virus (CCHFV), and hantaviruses (Bunyaviridae); and Kyasanur Forest disease virus (KFDV), Omsk hemorrhagic fever virus, yellow fever virus (YFV), and dengue viruses (Flaviviridae) (1,2). Although clinical management of VHF is primarily supportive, early diagnosis is needed to contain the contagion and implement public health measures, especially if agents are encountered out of their natural geographic context.

Vaccines have been developed for YFV, RVFV, Junin virus, KFDV, and hantaviruses (3-7), but only YFV vaccine is widely available. Early treatment with immune plasma was effective in Junin virus infection (8). The nucleoside analog ribavirin may be helpful if given early in the course of Lassa fever (9), Crimean-Congo hemorrhagic fever (10), or hemorrhagic fever with renal syndrome (11) and is recommended in postexposure prophylaxis and early treatment of arenavirus and bunyavirus infections (12).

Methods for direct detection of nucleic acids of microbial pathogens in clinical specimens are rapid, sensitive, and obviate the need for high-level biocontainment. Numerous systems are described for nucleic acid detection of VHF agents; however, none are multiplex (13). Although geographic location or travel history of suspected patients usually restricts the number of agents to be considered, diagnosis of VHF may be difficult in case of an intentional release (12). Symptoms of VHF are initially nonspecific and may include fever, headache, myalgia, and gastrointestinal or upper respiratory tract complaints (1); thus, assays that allow simultaneous consideration of multiple agents are needed.

We recently described the application of MassTag polymerase chain reaction (PCR) in the context of differential diagnosis of respiratory disease (14). MassTag PCR is a multiplex assay in which microbial gene targets are coded by a library of 64 distinct mass tags. Nucleic acids (RNA or DNA) are amplified by multiplex (reverse transcription-) PCR using up to 64 primers, each labeled by a photo-cleavable link with a different molecular weight tag. After separation of the amplification products from unincorporated primers and release of the mass tags from the amplicons by UV irradiation, tag identity is analyzed by mass spectrometry. The identity of the microbe in the clinical sample is determined by the presence of its 2 cognate tags, 1 from each primer.

The Study

To facilitate rapid differential diagnosis of VHF agents, we established the Greene MassTag Panel VHF version 1.0, which comprises the following targets: Ebola Zaire virus (ZEBOV), Ebola Sudan virus (SEBOV), MARV, LASV, RVFV, CCHFV, Hantaan virus (HNTV), Seoul virus (SEOV), YFV, and KFDV. Oligonucleotide primers were designed in conserved genomic regions to detect the broadest number of members for a given pathogen species. We developed a software program that culls sequence information from GenBank, performs multiple alignments with ClustalW, and designs primers optimized for multiplex PCR. The program uses a greedy algorithm to identify conserved sequences and create the minimum set of primers for amplification of all sequences in the alignment. Primers are selected within standard design constraints whenever possible (melting temperature 55[degrees]C-65[degrees]C, guanine-cytosine content 40%-60%, no hairpins); degenerate positions are introduced in cases where template divergence requires more flexibility. Although degeneracy is not tolerated in the five 3' nucleotides, MassTag PCR allows up to 4 nonneighboring variable positions per primer. Primers are checked by the basic local alignment search tool for potential hybridization to sequenced vertebrate genomes (Table 1).

Because only released mass tags are analyzed, staggering the size of amplification products created in multiplex reactions is unnecessary; thus, primers are selected for efficient and consistent performance irrespective of amplicon size (typically 80-200 bp). Before committing to synthesis of tagged primers, the functionality of candidate multiplex primer panels is examined in a series of amplification reactions that use prototype templates representing individual microbial targets. Primers that fail to yield a single, specific product band in agarose gel analysis are replaced. Target sequence standards for evaluation are cloned into pCR2.1-TOPO (Invitrogen, Carlsbad, CA, USA) by using PCR amplification of cDNA templates obtained by reverse transcription (RT) of extracts from infected, cultured cells or by assembly of overlapping synthetic polynucleotides.

The agents assayed in the VHF panel have RNA genomes; thus, assay sensitivity was determined by using synthetic RNA standards. Synthetic RNA standards were generated from linearized target sequence plasmids by using T7 polymerase (mMessage mMachine, Invitrogen). After quantitation by UV spectrometry, RNA was serially diluted in 2.5 [micro]g/mL yeast tRNA (Sigma, St. Louis, MO, USA), reverse transcribed with random hexamers by using Superscript II (Invitrogen), and analyzed by MassTag PCR as previously described (14). QIAquick 96 PCR purification cartridges (Qiagen, Hilden, Germany, with modified binding and wash buffers) were used to remove unincorporated primers before tags were decoupled from amplification products by UV photolysis in a flow cell and analyzed in a quadrapole mass spectrometer by using positive-mode atmospheric pressure chemical ionization (APCI-MS, Agilent Technologies, Palo Alto, CA, USA). The sensitivity of the 10-plex VHF panel with synthetic RNA standards was [less than or equal to] 50 RNA copies per assay (Table 2). Sensitivity and specificity of multiplex primer panels is assessed empirically by using calibrated synthetic standards as well as tissue culture-derived viral nucleic acid for each assembled panel.

Tissue culture extracts were used to examine assay specificity. Random primed cDNA obtained from cultures of ZEBOV, SEBOV, MARV, YFV isolates from the Gambia and Cote d'Ivoire, RVFV, CCHFV, HTNV, SEOV, and LASV strains Josiah, NL, and AV were subjected to mass tag analysis. In all instances, only the appropriate cognate mass tags were detected (data not shown). No spurious signal was identified in assays with water or RNA controls.

Performance with clinical materials was tested by using blood, sera, or oral swabs from 24 human patients of VHF

previously diagnosed through virus isolation, RT-PCR, or antigen detection enzyme-linked immunosorbent assay. Differential diagnosis by blinded MassTag PCR analysis was accurate in all cases (Table 3). For the samples from the 2005 Angola Marburg outbreak the result of MassTag PCR was similar to that of diagnostic single-plex PCR. ZEBOV sample 5004, obtained on day 17 of illness when serologic test results were positive for immunoglobulin M (IgM) and IgG, was negative by viral culture but positive in MassTag PCR.

Conclusions

These results confirm earlier work in respiratory diseases that show that MassTag PCR offers a rapid, sensitive, specific, and economic approach to differential diagnosis of infectious diseases. Small, low-cost, or mobile APCI-MS units extend the applicability of this technique beyond selected reference laboratories. Given the capacity of the method to code for up to 32 genetic targets, we are expanding the hemorrhagic fever panel to include additional viruses (dengue and South American hemorrhagic fever viruses) and are exploring the inclusion of bacterial and parasitic agents that may result in similar clinical signs and symptoms and, thus, have to be considered in differential diagnosis.

References

(1.) Peters CJ, Zaki SR. Role of the endothelium in viral hemorrhagic fevers. Crit Care Med. 2002;30(5 Suppl):S268-73.

(2.) Geisbert TW, Jahrling PB. Exotic emerging viral diseases: progress and challenges. Nat Med. 2004; 10(12 Suppl): S110-21.

(3.) Pugachev KV, Guirakhoo F, Monath TP. New developments in flavivirus vaccines with special attention to yellow fever. Curr Opin Infect Dis. 2005; 18:387-94.

(4.) Pittman PR, Liu CT, Cannon TL, Makuch RS, Mangiafico JA, Gibbs PH, et al. Immunogenicity of an inactivated Rift Valley fever vaccine in humans: a 12-year experience. Vaccine. 1999;18:181-9.

(5.) Enria DA, Barrera Oro JG. Junin virus vaccines. Curr Top Microbiol Immunol. 2002;263:239-61.

(6.) Hooper JW, Li D. Vaccines against hantaviruses. Curr Top Microbiol Immunol. 2001;256:171-91.

(7.) Dandawate CN, Desai GB, Achar TR, Banerjee K. Field evaluation of formalin inactivated Kyasanur forest disease virus tissue culture vaccine in three districts of Karnataka state. Indian J Med Res. 1994;99:152-8.

(8.) Enria DA, Maiztegui JI. Antiviral treatment of Argentine hemorrhagic fever. Antiviral Res. 1994;23:23-31.

(9.) McCormick JB, King IJ, Webb PA, Scribner CL, Craven RB, Johnson KM, et al. Lassa fever. Effective therapy with ribavirin. N Engl J Med. 1986;314:20-6.

(10.) Ozkurt Z, Kiki I, Erol S, Erdem F, Yilmaz N, Parlak M, et al. Crimean-Congo hemorrhagic fever in eastern Turkey: clinical features, risk factors and efficacy of ribavirin therapy. J Infect. Epub 2005 Jun 13.

(11.) Huggins JW, Hsiang CM, Cosgriff TM, Guang MY, Smith JI, Wu ZO, et al. Prospective, double-blind, concurrent, placebo-controlled clinical trial of intravenous ribavirin therapy of hemorrhagic fever with renal syndrome. J Infect Dis. 1991;164:1119-27.

(12.) Borio L, Inglesby T, Peters CJ, Schmaljohn AL, Hughes JM, Jahrling PB, et al. Hemorrhagic fever viruses as biological weapons: medical and public health management. JAMA. 2002;287:2391-405.

(13.) Drosten C, Kummerer BM, Schmitz H, Gunther S. Molecular diagnostics of viral hemorrhagic fevers. Antiviral Res. 2003;57:61-87.

(14.) Briese T, Palacios G, Kokoris M, Jabado O, Liu Z, Renwick N, et al. Diagnostic system for rapid and sensitive differential detection of pathogens. Emerg Infect Dis. 2005;11:310-3.

(15.) Bowen MD, Rollin PE, Ksiazek TG, Hustad HL, Bausch DG, Demby AH, et al. Genetic diversity among Lassa virus strains. J Virol. 2000;74:6992-7004.

Gustavo Palacios, * (1) Thomas Briese, * (1) Vishal Kapoor, * Omar Jabado, * Zhiqiang Liu, * Marietjie Venter, ([dagger]) Junhui Zhai, * Neil Renwick, * Allen Grolla, ([double dagger]) Thomas W. Geisbert, ([section]) Christian Drosten, ([paragraph] Jonathan Towner, (#) Jingyue Ju, * Janusz Paweska, ** Stuart T. Nichol, ([double dagger]) ([dagger][dagger]) Robert Swanepoel, ** Heinz Feldmann, ([double dagger]) ([dagger][dagger]) Peter B. Jahrling, ([double dagger][double dagger]) and W. Ian Lipkin *

(1) These authors contributed equally to this article.

* Columbia University, New York, New York, USA; ([dagger]) University of Pretoria and National Health Laboratory Services, Pretoria, South Africa; ([double dagger]) Public Health Agency of Canada, Winnipeg, Manitoba, Canada; ([section]) United States Army Medical Research Institute of Infectious Diseases, Fort Detrick, Frederick, Maryland, USA; ([paragraph]) Bernhard-Nocht-Institute of Tropical Medicine, Hamburg, Germany; (#) Centers for Disease Control and Prevention, Atlanta, Georgia, USA; ** National Institute for Communicable Diseases, Sandringham, South Africa; ([dagger][dagger]) University of Manitoba, Winnipeg, Manitoba, Canada; and ([double dagger][double dagger]) National Institutes of Allergy and Infectious Diseases Integrated Research Facility, Fort Detrick, Frederick, Maryland, USA

Address for correspondence: Thomas Briese, Jerome L. and Dawn Greene Infectious Disease Laboratory, Mailman School of Public Health, Columbia University, 722 W 168th St, Rm 1801, New York, NY 10032, USA; fax: 212-342-9044; email: thomas.briese@columbia.edu

Use of trade names is for identification only and does not imply endorsement by the Public Health Service or by the U.S. Department of Health and Human Services.

This work was supported by National Institutes of Health awards AI51292, AI056118, AI55466, and U54AI57158 (Northeast Biodefense Center-Lipkin) and the Ellison Medical Foundation.

Dr Palacios is an associate research scientist in the Jerome L. and Dawn Greene Infectious Disease Laboratory at the Columbia University Mailman School of Public Health. His research focuses on the molecular epidemiology of viruses, virus interactions with their hosts, and innovative pathogen detection methods.
Table 1. Greene MassTaq panel VHF version 1.0 *

Target MassTag Name Sequence

ZEBOV 718 (fwd) EboZA-U234 AACACCGGGTCTTAATTCTTATATCAA
 646 (rev) EboZA-1_319 GGTGGTAAAATTCCCATAGTAGTTCTTT
SEBOV 503 (fwd) EboSU-U416 CGAGCCTAACGTTTTGGGC
 630 (rev) EboSU-L489 GCTCCAGGAATTGTTCGGGTA
MARV 654 (fwd) MARV-U12816C CCCTCCATATCTTAGACAACATATTGTG
 395 (rev) MARV-L12994 CCCAACACTCCTGGTTCACAGC
LASVT 558 (fwd) Las4-U92 ACTGCATTYTCATACTTYCTRGAATC
 686 (rev) Las4-L257 CCRGGYTTGACCAGTGCTGT
RVFV 658 (fwd) RVF-U578 GGATTGACCTGTGCCTGTTGC
 495 (rev) RVF-L660 GCATTAGAAATGTCCTCTTTTGCTGC
CCHFV 499 (fwd) CCHV-U4 AGAAACACGTGCCGCTTACGCCCA
 710 (rev) CCHV-L120 CCATTTCCYTTYTTRAACTCYTCAAACCA
HNTV 479 (fwd) HAN-U179 AYACAGCAGCAGTTAGCCTCCT
 702 (rev) HAN-L245 GCT GCC GTA RGT AGT CCC TGTT
SEOV 455 (fwd) SEO-U243 CAGGATTGCAGCAGGGAAGA
 602 (rev) SEOUL-L309 ATGATCACCAGGYTCTACCCC
YFV 467 (fwd) YF-U186 GCTGGGAGCGCGGTATC
 670 (rev) YF-L249 GGAAGCCCAATGGTCCTCAT
KFDV 483 (fwd) KYF-U170 TGGAAGCCTGGCTGAAAGAG
 614 (rev) KYF-L233 TCATCCCCACTGACCAGCAT

Target Gene

ZEBOV L
SEBOV L
MARV L
LASVT NP
RVFV N
CCHFV N
HNTV N
SEOV N
YFV NS5
KFDV NS5

* ZEBOV, Ebola Zaire virus, SEBOV, Ebola Sudan virus, MARV, Marburg
virus; LASV, Lassa virus, RVFV, Rift Valley fever virus; CCHV,
Crimean-Congo hemorrhagic fever virus; HNTV, Hantaan virus; SEOV, Seoul
virus; YFV, yellow fever virus; KFDV, Kyasanur Forest disease virus;
fwd, forward; rev, reverse.

([dagger]) Primers were designed on Lassa lineage IV sequences (15) and
the recently identified outlier sequence Lassa AV (AF256121).

Table 2. Sensitivity of detection with synthetic RNA standards

Pathogen * Detection threshold (RNA copies) ([dagger])

ZEBOV 20
SEBOV 20
MARV 20
LASV 20
RVFV 20
CCHFV 50
HNTV 20
SEOV 50
YFV 20
KFDV 20

* ZEBOV, Ebola Zaire virus; SEBOV, Ebola Sudan virus; MARV, Marburg
virus; LASV, Lassa virus; RVFV, Rift Valley fever virus; CCHV,
Crimean-Congo hemorrhagic fever virus; HNTV, Hantaan virus; SEOV, Seoul
virus; YFV, yellow fever virus; KFDV, Kyasanur Forest disease virus.
tRNA copies refers to the number of molecules subjected to reverse
transcription; half of the reverse transcription reaction was then used
for polymerase chain reaction amplification.

Table 3. MassTag polymerase chain reaction analysis of clinical
specimens from viral hemorrhaqic fever patients *

Previous diagnosis Sample identification Sample type

ZEBOV 5015 Serum
ZEBOV 5014 Serum
ZEBOV 5004 Serum
ZEBOV 6317 Serum
ZEBOV 6313 Serum
MARV 246-00-5 Hemolyzed whole blood
MARV 226-00-4 Hemolyzed whole blood
MARV 246-00-7 Hemolyzed whole blood
MARV 98-00-2 Hemolyzed whole blood
MARV 461 Blood
MARV 462 Oral swab
MARV 475 Blood
MARV 476 Oral swab
LASV 98-04-1 Serum
LASV 98-04 Serum
LASV 98-045 Serum
LASV 80-041 Serum
RVFV 98002009 Serum
RVFV H6061989 Serum
RVFV 98002019 Serum
RVFV 77-04 Serum
CCHFV 187-86 Serum
CCHFV 30-93 Serum
CCHFV 465-88 Serum
CCHFV 407-89 Serum
CCHFV 215-90 Serum

Previous diagnosis Year/origin MassTag result ([dagger])

ZEBOV 1995/Kikwit, DRC +++, ZEBOV
ZEBOV 1995/Kikwit, DRC +++, ZEBOV
ZEBOV 1995/Kikwit, DRC +++, ZEBOV
ZEBOV 1995/Kikwit, DRC +++, ZEBOV
ZEBOV 1995/Kikwit, DRC +++, ZEBOV
MARV 2000/Durba, DRC +, MARV
MARV 2000/Durba, DRC ++, MARV
MARV 2000/Durba, DRC +, MARV
MARV 2000/Durba, DRC +++, MARV
MARV 2005/Uige, Angola +++, MARV
MARV 2005/Uige, Angola +++, MARV
MARV 2005/Uige, Angola ++, MARV
MARV 2005/Uige, Angola +, MARV
LASV 2004/Sierra Leone +++, LASV
LASV 2004/Sierra Leone ++, LASV
LASV 2004/Sierra Leone +, LASV
LASV 2004/Sierra Leone +++, LASV
RVFV 1998/Kenya +, RVFV
RVFV 1998/Kenya +, RVFV
RVFV 1998/Kenya ++, RVFV
RVFV 2004/Namibia ++, RVFV
CCHFV 1986/South Africa +, CCHFV
CCHFV 1993/South Africa +++, CCHFV
CCHFV 1988/South Africa +++, CCHFV
CCHFV 1989/South Africa +++, CCHFV
CCHFV 1990/South Africa ++, CCHFV

* ZEBOV, Ebola Zaire virus; MARV, Marburg virus; LASV, Lassa virus;
RVFV, Rift Valley fever virus; CCHV, Crimean-Congo hemorrhagic fever
virus, DRC, Democratic Republic of Congo.

([dagger]) Relative ranking of results: +, signal-to-noise ratio
[less than or equal to] 4, ++, signal-to-noise ratio >4 and <8; +++,
signal-to-noise ratio [greater than or equal to] 8.
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Author:Lipkin, W. Ian
Publication:Emerging Infectious Diseases
Geographic Code:1USA
Date:Apr 1, 2006
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