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8 Methods for Detection and Diagnosis I n the pre-eradication era, smallpox was usually diagnosed by its dis- tinct clinical characteristics, particularly the vesicular–pustular rash (see Chapter 2), in the context of a cluster of probable cases with an epidemiologic link. Material from lesions could be analyzed by electron microscopy in reference laboratories, providing morphologic identification of the characteristic brick-shaped virions (Biel and Gelderblom, 1999; see also Chapter 3), but did not distinguish variola from other poxviruses. Recovery of infectious virus from infected persons using tissue culture methods was feasible but was seldom used. Despite the eradication of smallpox, the need remains for robust and safe methods of detection of variola virus and diagnosis of the disease. Diseases caused by poxviruses that can infect the human host, such as monkeypox, continue to circulate and may be confused with smallpox, necessitating precise methods for rapid differential diagnosis. Disseminated vaccinia might also be misdiagnosed as smallpox, although a history of recent vaccination or contact with a recently vaccinated person would usually be obtained. Finally, the classification of smallpox as a category A agent with the potential for aerosolization and broad distribution within the environment requires new approaches to sensitive and specific detection of the virus in nonclinical specimens. The application of contemporary viral diagnostic tools, such as poly- merase chain reaction (PCR) methods, to smallpox diagnosis has received attention because rapid and accurate identification of index cases would be essential for optimal containment of initial spread in a largely unimmunized population in the event of an unintended or intentional release of the virus. 

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 LIVE VARIOLA VIRUS These methods may allow diagnosis in respiratory secretions during the 12–14 day incubation period, which would be quite valuable for controlling transmission. Additionally, recent and forthcoming advances in genomic science mean that new approaches for identification of variola virus in clinical or environmental samples can be developed that involve detecting the presence of genomic DNA or viral proteins. Maximizing the specificity of such tests will require knowledge of the genetic variability of related poxviruses, the background against which variola must be distinguished to maximize the sensitivity of the test, and the variability of variola and viral proteins and their subdomains that are unique to variola. It will also be important to develop new diagnostics that can be used to detect the virus in different types of patient specimens (e.g., lesion material, secretions, organ tissues) and environmental samples (e.g., air, surfaces, fomites). Developing environmental detection and diagnostic methods that do not require the isolation of infectious virus in tissue culture is important because of the risk of human exposure during preparation of specimens to be tested in the laboratory. Such advances in detection and diagnosis would facilitate forensic investigations to determine the source of variola virus in the event of an intentional release. This chapter reviews the current status of methods to detect variola virus and diagnose smallpox, relevant regulatory requirements, and the need for live variola virus to achieve advances in the development of detec- tion and diagnostic capabilities. CURRENT STATUS OF DETECTION AND DIAgNOSTIC METHODS The 1999 committee offered the following conclusion related to detec- tion and diagnosis: If further development of procedures for the environmental detec- tion of variola virus or for diagnostic purposes were to be pursued, more extensive knowledge of the genome variability, predicted pro- tein sequences, virion surface structure, and functionality of variola virus from widely dispersed geographic sources would be needed. Since 1999, substantial work has been done on the development of new techniques for the detection of variola virus and diagnosis of smallpox and for the differentiation of variola virus from other orthopoxviruses that infect humans (e.g., vaccinia, monkeypox, cowpox). Most of these assays have been based on nucleic acid detection by PCR, and some have been validated using clinical samples. Some experience has been reported with the use of multiplex PCR to detect variola and differentiate it from other poxviruses or unrelated viruses in laboratory-created specimens containing

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 METHODS FOR DETECTION AND DIAGNOSIS mixed genomic fragments. Limited experience exists with direct detection of variola virus in stored patient specimens or in specimens from nonhuman primates. Relatively little has been done to create assays that detect variola virus proteins or to refine serologic approaches to smallpox diagnosis. The capacity to carry out seroepidemiologic surveillance with rapid high- throughput serologic assays for variola virus-specific IgG antibodies would be valuable to characterize the extent of the spread of the virus in an out- break setting, and serologic assays for variola virus-specific IgM antibodies would be useful to document recent infection in individuals who were asymptomatic when tested (see Appendix). Polymerase Chain Reaction PCR enables highly sensitive detection of viral nucleic acids to very low copy numbers. PCR products can be sequenced to provide detailed genetic information about the pathogen, and PCR can be performed as a quantita- tive or multiplex assay in which the specimen is tested for multiple patho- gen sequences at the same time. Several different regions of the variola virus genome have been used to design primers that either detect all orthopox- viruses of interest or are specific for individual poxviruses. Real-time PCR for the hemagglutinin gene (J7R) of variola virus was sensitive and specific when tested on variola virus samples from cell culture and infected tissues that contained both viral and cellular DNA (Ibrahim et al., 2003; Aitichou et al., 2008). This assay was evaluated with genomic DNA from 48 differ- ent isolates of variola virus and 25 other poxviruses. Specificity for variola detection was greater than 96 percent; the majority of these samples were derived from virus-infected cell cultures and variola virus-infected tissues. This poxvirus assay was applied successfully to the diagnosis of smallpox from fixed human tissue from one fatal case (Schoepp et al., 2004), even though specimens were obtained and stored under conditions not designed to protect DNA integrity. The assay has been expanded to include other variola virus genes (B9R and B10R) using prepared samples, detecting 12–25 genome copies (Kulesh et al., 2004). It has been adapted for use with dried reagents and for multiplexing with probes for other orthopoxviruses (Aitichou et al., 2008). The hemagglutinin gene has also been used to design primers for detecting all orthopoxviruses for use with a probe that can distinguish variola from other poxviruses by melting curve analysis, and tested on plasmid DNA (Espy et al., 2002) and on tissue and blood spiked with poxvirus DNA (Putkuri et al., 2009). The CrmB (cytokine response modifier B) gene has also served as the target for amplifying orthopoxvirus DNA using consensus primers. Viral (genomic) amplicons may differ in size, but variola and other orthopox- viruses can also be differentiated from each other by analysis of restriction

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 LIVE VARIOLA VIRUS fragment length polymorphism (RFLP) (Loparev et al., 2001). This assay was validated on eight strains of variola virus. In a similar assay, TaqMan probes were designed to be specific for all orthopoxviruses or for variola virus and validated with poxvirus panels and plasmid DNA from the European Network for Imported Viral Diseases (Fedele et al., 2006). A multiplex PCR that distinguished orthopoxviruses from herpesviruses used primers from the CrmB gene for poxvirus identification and RFLP of the PCR product to differentiate one orthopoxvirus from another. This test was developed and validated using plasmid DNA from only a single strain of variola virus (Sias et al., 2007). A real-time PCR assay that combines variola virus-specific and panorthopoxvirus primers targeted to the gene for a 14 Kd protein (A30L) has been developed and validated on genomic DNA from 12 strains of variola virus; variola was differentiated from cowpox, vaccinia, monkeypox, and camelpox viruses (Scaramozzino et al., 2007). A multiplex real-time PCR assay has been developed that includes indi- vidual primers specific for variola (B11R–B12R), vaccinia, monkeypox, and cowpox viruses, plus primers common for all orthopoxviruses, and results in amplicons of different sizes. This assay was validated on DNA from virus grown in culture and on scabs from smallpox skin lesions (Shchelkunov et al., 2005). Another multiplex method targets the 14kD fusion protein (A27L) for amplification from all orthopoxviruses and differentiates variola from other orthopoxviruses by melting curve analysis (Olson et al., 2004). This assay was validated on 14 variola virus samples from tissue culture and from skin lesions in the VECTOR repository and detected four variola genome copies. Multiplex PCR has also been performed using consensus and variola virus-specific primers based on known single nucleotide poly- morphisms (SNPs) in A13L and A36R genes that are different in variola and other poxviruses; these SNPs were identified in PCR products from 43 variola strains but none of 50 other orthopoxviruses (Pulford et al., 2004). These variola virus isolates had been collected over 40 years from diverse geographic locations. A number of PCR assays have been developed and tested for detection and differentiation of variola virus using only plasmid DNA. The genes ana- lyzed include hemagglutinin, RNA polymerase (rpo18), early transcription factor VETF, and small membrane protein p8 (A13L). For each, melting curve analysis was used to distinguish variola from other orthopoxviruses (Nitsche et al., 2004; Panning et al., 2004). PCR has also been combined with immobilization of synthetic oligonucleotides corresponding to variola and other poxvirus genes on nylon membranes to allow direct visualization of products that hybridize to specific oligonucleotides as a simplification, but a PCR apparatus is still required (Fitzgibbon and Sagripanti, 2006).

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 METHODS FOR DETECTION AND DIAGNOSIS In Situ Hybridization In situ hybridization was used to examine sections of tissue speci- mens for the presence of variola virus DNA in skin lesion biopsies from two South American smallpox cases. Specific molecular probes differenti- ated skin cells containing variola from those caused by herpesviruses in formalin-fixed tissue sections that showed no distinguishable differences by standard histopathology methods (Nuovo et al., 2003). gene Chip Analysis Oligonucleotides specific for orthopoxviruses can be immobilized and used to detect interaction with DNA extracted from samples suspected of containing a poxvirus. Specific hybridization can be detected by fluorescent probes (Lapa et al., 2002; Laassri et al., 2003; Ryabinin et al., 2006) or use of electrochemical sensors (Komarova et al., 2005). Chips have been designed using one or two individual variola virus genes (CrmB, Lapa et al., 2002, and Komarova et al., 2005; C23L/B29R, Laassri et al., 2003; C23L/B29R + B19R, Ryabinin et al., 2006) and the complete genomes of multiple strains as resequencing tiling arrays (Sulaiman et al., 2007). These assays can distinguish variola virus from other poxviruses and from herpes- viruses. The resequencing array was tested on amplified DNA from 14 strains of variola virus and can also identify other human orthopoxviruses (Sulaiman et al., 2008). This technology can be used for rapid identifica- tion of a particular variola genome by comparison with known genomes in sequencing databases. A variation on this approach is the development of primers that span the orthopoxvirus genome followed by RFLP, which is then used to distinguish one orthopoxvirus from another. This assay was validated on genomic DNA from two strains of variola virus and on monkeypox, camelpox, cowpox, tanapox, ectromelia, and vaccinia viruses (Li et al., 2007). These whole-genome approaches would be useful to iden- tify variola genomes that had been altered intentionally. Protein-Based Methods Little work has been done to develop direct protein detection methods for variola. At present, these methods depend on developing antibody reagents that bind specifically to variola proteins that are distinct from those made by other orthopoxviruses. Utilizing ELISAs, Ulaeto and colleagues (2002) have begun to characterize the reactivity of 23 strains/isolates of variola virus, both γ®-ray inactivated and viable (under BSL-4 conditions), with a panel of monoclonal antibodies and polyclonal antisera, raised against either vaccinia or variola virus preparations. Polyclonal antibody

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 LIVE VARIOLA VIRUS reagents displayed more uniform detection of variola virus strains than was obtained with monoclonal antibodies (Ulaeto et al., 2002). One monoclonal antibody has been described that is specific for variola virus and can be used to distinguish variola from other poxviruses (Damon, 2006). However, monoclonal antibodies detect a single epitope in a single viral protein, and most are conformation dependent. Specificity for geographically unrelated variola isolates would depend on defining a fully conserved and stable epitope or using a mix of monoclonals that would recognize epitopes in several unique variola virus proteins having no homologues or differing substantially from the related proteins in the other poxviruses. Even when well-characterized reagents are available, designing antigen detection methods that demonstrate the presence of viral proteins in patient materials has been challenging for many human pathogens. Most successes are achieved when the clinical material is a cutaneous lesion specimen, which would be the case for variola at the symptomatic stage of infection. In one example of a poxvirus detection method applied to respiratory secre- tions, a biosensor technique using cyan-5 dye labeled antivaccinia antibody was used to detect vaccina proteins in human throat swab specimens that had been spiked with vaccinia virus from tissue culture (Donaldson et al., 2004). One would expect such approaches to be feasible for variola detection, but their development currently depends on generating panels of antibodies that are highly specific for variola proteins. Pilot experiments were conducted in which ELISAs were used to detect monkeypox virus during the recent outbreaks in Africa and variola virus in specimens from nonhuman primates (Karem et al., 2007). Nevertheless, although inhibi- tors may be encountered, nucleotide detection methods are generally pre- ferred for viral detection because nucleotides can be extracted from patient materials and concentrated for PCR testing, whereas similar processes to enhance sensitivity are difficult for protein detection in respiratory secre- tions or other clinical specimens that would be available from patients in the pre-eruptive phase of smallpox. Proteomics methods may emerge that can identify a specific sequence of amino acid residues by direct analysis of a sample using mass spectroscopy or other methods that do not require antibody reagents, but these tools are not yet applicable for clinical use. With the exception of measuring antibody titers by plaque reduc- tion neutralization assay, serologic assays for IgG and IgM antibodies to variola and other poxviruses are also protein-based detection techniques. ELISA methods detect antibodies in serum samples through their binding to immobilized viral antigens. The development of such an assay for detecting variola virus IgG and IgM antibodies is feasible, but specificity requires the identification of unique proteins that do not elicit cross-reactive antibodies as a result of exposure to other poxviruses, such as by vaccination with vac- cinia. It is anticipated that most variola infections would be symptomatic;

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 METHODS FOR DETECTION AND DIAGNOSIS however, a panpoxvirus serologic assay could be useful for assessing the extent of asymptomatic infection in a population not previously vaccinated should the need arise. Currently, VECTOR is developing next-generation test kits to detect orthopoxviral protein markers. These immunodiagnostic tests will rely on hybridoma technology and technology for producing recombinant anti- bodies to major neutralizing and protective antigens of variola virus and those of other orthopoxviruses pathogenic for humans (Russian federation Patent #2281327; Razumov et al., 2004, 2005; personal communication, Ilya Drozdov, WHOCC for Orthopoxvirus Diagnosis and Repository for Variola Virus Strains and DNA, March 27, 2009). In parallel, VECTOR is working to develop and improve species-specific diagnostics for viruses such as variola, monkeypox, and cowpox based on multiplex PCR, real-time PCR, and microchip technology (Lapa et al., 2002; Laassri et al., 2003; Ryabinin et al., 2006; personal communication, Ilya Drozdov, WHOCC for Orthopoxvirus Diagnosis and Repository for Variola Virus Strains and DNA, March 27, 2009). Detection in the Environment The technical capacity for environmental detection of variola virus would be important in the event of an intentional release. Widespread distribution of the virus could be achieved because poxviruses are stable in aerosol form and can be lyophilized. The molecular methods for variola virus detection that have been developed since 1999 use PCR and in situ hybridization assays that have proven valuable for the clinical detection of many viral pathogens in patient specimens, and a few of these methods have been validated using archived tissues from variola cases. PCR-based methods are also useful for detecting viruses in environmental samples, including air samples, water, and soil, as well as in swabs taken from potentially contaminated surfaces. These methods could be applied to the identification of variola virus in such specimens with certain modifications in the way the materials are prepared for testing. For example, it would be necessary to take into account the inhibitory effects of detergents and other materials on PCR sensitivity, as shown in experiments with vaccinia virus (Kurth et al., 2008). The specificity of PCR for variola virus detection should be preserved, but sensitivity in such samples is difficult to predict. Ideally, tools for detecting the presence of variola virus in the environ- ment would need to be rapid, portable, and easily deployable. Because pox- virus genome detection methods require relatively complex equipment and reagents, it would be necessary at present to bring materials suspected of containing variola virus to a laboratory facility. A more practical variation of the method for field use would be the use of dried reagents in a dual-

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 LIVE VARIOLA VIRUS probe real-time PCR assay for detection of variola or other orthopoxviruses (Aitichou et al., 2008). Even if PCR or ELISA methods were used that could differentiate variola from other poxviruses in environmental samples, their sensitivity in field testing would need to be established. Criteria for speci- ficity might need to be lowered to ensure that a positive sample was not missed under field conditions, with the assumption that all specimens would need to be retested and results validated in a reference laboratory. This gap may be addressed by the development of tools such as direct electro- chemical DNA sensors that can identify nucleotide sequences without the need for PCR amplification and secondary analysis of the products by RFLP or sequencing (Komarova et al., 2005). Nanotechnology-based tools may be developed that can discriminate viruses based on their particle size and other properties; if so, it would be necessary to have at least inactivated variola particles to assess their sensitivity for environmental detection. REgULATORy REqUIREMENTS Currently available in vitro diagnostic devices (IVDs) for the detection and diagnosis of variola infection are limited to research assays developed by DOD, CDC, and academic laboratories. In the United States, licensure of IVDs for various infectious agents, including variola, is regulated pri- marily by the FDA’s Center for Devices and Radiological Health (CDRH), which assesses benefits and risks according to the IVD’s analytical and clinical performance. Medical devices, including IVDs, are categorized as Class I, II, or III according to risk criteria and requirements listed in 21 Code of Federal Regulations (CFR) 800. Whereas most Class I devices are exempt from premarket notification, most Class II devices do require such notification [510(k)], and most Class III devices require premarket approval (PMA), including submission of clinical data to support marketing claims. The potential classification of IVDs for variola virus detection has not been established, although it appears likely, given the critical impor- tance of accurate detection methods, that premarket notification including both general and special controls (Class II designation) would be required. A new section (513(f)(2)) of the Food, Drug and Cosmetics Act as amended by the FDA Modernization Act of 1997 includes a provision whereby a sponsor can request a so-called “de novo” classification that may not require premarket approval, but the sponsor would have to demonstrate that the device would pose very little or no risk of harm, especially for diagnosing suspected human cases. Finally, the use of a new IVD for variola virus detection may also be approved via Emergency Use Authorization (see Chapter 1).

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 METHODS FOR DETECTION AND DIAGNOSIS NEED FOR LIVE VARIOLA VIRUS The identification and characterization of a series of variola virus- specific genetic markers has paved the way for sensitive and specific multi- plex nucleic acid methods, and further progress on refining these approaches should not require live virus. Methods that detect viral proteins have been pursued to a lesser extent but could also be expanded without the need for live virus. Although not essential, better characterization of the sensitivity and specificity of both nucleic acid and protein methods for variola virus detection in relevant samples could be achieved by additional testing of tissues from nonhuman primates infected with the virus. Preservation of tissues for this purpose should be included in antiviral, vaccine, or patho- genesis studies done in animals infected with variola. Since methods devel- oped using only variola proteins could prove inadequate for their detection in clinical materials from infected individuals, archived clinical specimens could be tested to confirm the sensitivity and specificity of such tests, if possible. Further work on protein-based detection would benefit particu- larly from access to proteins made in variola virus-infected cells instead of proteins made using expression vectors to ensure the reliability of the test and to standardize reagents. Environmental detection methods have seen little progress, but further research in this area would use “mocked-up” specimens, so use of the live virus would not be necessary. High-throughput assays, including serologic methods to identify recently infected individuals, would be needed to test large numbers of samples in a possible outbreak situation. However, the development of most new methods would not require live virus as this research could build on work with other validated methods and be scaled up. Some future approaches that might prove valuable, such as those that detect viral particles, could require access to variola virions made in culture cells for their validation. One caveat related to variola detection and smallpox diagnosis is that genomic sequencing of enough geographically diverse isolates is necessary to ensure that PCR tests have adequate specificity. PCR and sequencing of the amplicons would be the first step in a forensic analysis of the source of a variola isolate should a reintroduction of the virus occur, and would also be accomplished most effectively if background information were available on the complete genome sequence of as many variola isolates as possible. It is expected that use of the live virus would not be necessary for this purpose, assuming that sufficient DNA is still available in stored specimens in the U.S. and Russian stocks. Finally, it is not yet clear whether the FDA will require the use of live variola virus in the evaluation of new diagnostic methods.

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0 LIVE VARIOLA VIRUS REFERENCES Aitichou, M., S. Saleh, P. Kyusung, J. Huggins, M. O’Guinn, P. Jahrling, and S. Ibrahim. 2008. Dual-probe real-time PCR assay for detection of variola or other orthopoxviruses with dried reagents. Journal of Virological Methods 153(2):190–195. Biel, S. S., and H. R. Gelderblom. 1999. Diagnostic electron microscopy is still a timely and rewarding method. Journal of Clinical Virology 13:105–119. Damon, I. K. 2006. Poxviruses. In Fields’ virology, 5th edition, edited by B. N. Fields, D. M. Knipe, P. M. Howley, and D. E. Griffin. Philadelphia, PA: Lippincott Williams & Wilkins. Pp. 2947–2976. Donaldson, K. A., M. F. Kramer, and D. V. Lim. 2004. A rapid detection method for Vaccinia virus, the surrogate for smallpox virus. Biosens Bioelectron 20(2):322–327. Espy, M. J., I. F. Cockerill, R. F. Meyer, M. D. Bowen, G. A. Poland, T. L. Hadfield, and T. F. Smith. 2002. Detection of smallpox virus DNA by LightCycler PCR. Journal of Clinical Microbiology 40(6):1985–1988. Fedele, C. G., A. Negredo, F. Molero, M. P. Sanchez-Seco, and A. Tenorio. 2006. Use of inter- nally controlled real-time genome amplification for detection of variola virus and other orthopoxviruses infecting humans. Journal of Clinical Microbiology 44(12):4464–4470. Fitzgibbon, J. E., and J. L. Sagripanti. 2006. Simultaneous identification of orthopoxviruses and alphaviruses by oligonucleotide macroarray with special emphasis on detection of variola and Venezuelan equine encephalitis viruses. Journal of Virological Methods 131(2):160–167. Ibrahim, S. M., D. A. Kulesh, S. S. Saleh, I. K. Damon, J. J. Esposito, A. L. Schmaljohn, and P. B. Jahrling. 2003. Real-time PCR assay to detect smallpox virus. Journal of Clinical Microbiology 41(8):3835–3839. Karem, K.L., M. Reynolds, C. Hughes, Z. Braden, P. Nigam, S. Crotty, J. Glidewell, R. Ahmed, R. Amara, and I. K. Damon. 2007. Monkeypox-induced immunity and failure of childhood smallpox vaccination to provide complete protection. Clinical & Vaccine Immunology: CVI 14 (10):1318–1327. Komarova, E., M. Aldissi, and A. Bogomolova. 2005. Direct electrochemical sensor for fast reagent-free DNA detection. Biosens Bioelectron 21(1):182–189. Kulesh, D. A., R. O. Baker, B. M. Loveless, D. Norwood, S. H. Zwiers, E. Mucker, C. Hartmann, R. Herrera, D. Miller, D. Christensen, L. P. Wasieloski, Jr., J. Huggins, and P. B. Jahrling. 2004. Smallpox and pan-orthopox virus detection by real-time 3’-minor groove binder TaqMan assays on the roche LightCycler and the Cepheid smart Cycler platforms. Journal of Clinical Microbiology 42(2):601–609. Kurth, A., J. Achenbach, L. Miller, I. M. Mackay, G. Pauli, and A. Nitsche. 2008. Orthopox- virus detection in environmental specimens during suspected bioterror attacks: Inhibitory influences of common household products. Applied and Environmental Microbiology 74(1):32–37. Laassri, M., V. Chizhikov, M. Mikheev, S. Shchelkunov, and K. Chumakov. 2003. Detec- tion and discrimination of orthopoxviruses using microarrays of immobilized oligo- nucleotides. Journal of Virological Methods 112(1-2):67–78. Lapa, S., M. Mikheev, S. Shchelkunov, V. Mikhailovich, A. Sobolev, V. Blinov, I. Babkin, A. Guskov, E. Sokunova, A. Zasedatelev, L. Sandakhchiev, and A. Mirzabekov. 2002. Species-level identification of orthopoxviruses with an oligonucleotide microchip. Journal of Clinical Microbiology 40(3):753–757.

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 METHODS FOR DETECTION AND DIAGNOSIS Li, Y., D. S. Carroll, S. N. Gardner, M. C. Walsh, E. A. Vitalis, and I. K. Damon. 2007. On the origin of smallpox: Correlating variola phylogenics with historical smallpox records. Proceedings of the National Academy of Sciences of the United States of America 104:15787–15792. Loparev, V. N., R. F. Massung, J. J. Esposito, and H. Meyer. 2001. Detection and differentia- tion of Old World orthopoxviruses: Restriction fragment length polymorphism of the crmB gene region. Journal of Clinical Microbiology 39(1):94–100. Nitsche, A., H. Ellerbrok, and G. Pauli. 2004. Detection of orthopoxvirus DNA by real-time PCR and identification of variola virus DNA by melting analysis. Journal of Clinical Microbiology 42(3):1207–1213. Nuovo, G. J., J. A. Plaza, and C. Magro. 2003. Rapid diagnosis of smallpox infection and differentiation from its mimics. Diagnostic Molecular Pathology 12(2):103–107. Olson, V. A., T. Laue, M. T. Laker, I. V. Babkin, C. Drosten, S. N. Shchelkunov, M. Niedrig, I. K. Damon, and H. Meyer. 2004. Real-time PCR system for detection of orthopoxviruses and simultaneous identification of smallpox virus. Journal of Clinical Microbiology 42(5):1940–1946. Panning, M., M. Asper, S. Kramme, H. Schmitz, and C. Drosten. 2004. Rapid detection and differentiation of human pathogenic orthopox viruses by a fluorescence resonance energy transfer real-time PCR assay. Clinical Chemistry 50(4):702–708. Pulford, D. J., A. Gates, S. H. Bridge, J. H. Robinson, and D. Ulaeto. 2004. Differential efficacy of vaccinia virus envelope proteins administered by DNA immunisation in protection of BALB/c mice from a lethal intranasal poxvirus challenge. Vaccine 22:3358–3366. Putkuri, N., H. Piiparinen, A. Vaheri, and O. Vapalahti. 2009. Detection of human orthopox- virus infections and differentiation of smallpox virus with real-time PCR. Journal of Medical Virology 81(1):146–152. Razumov, I. A., M. A. Vasil’eva, O. A. Serova, L. V. Artem’eva, N. I. Bormotov, E. F. Belanov, G. V. Kochneva, E. E. Konovalov, and V. B. Loktev. 2004. A study of neutralizing activity and cross-reactivity of monoclonal antibodies to ectromelia virus with orthopoxviruses pathogenic for man. Vestn Ross Akad Med Nauk (8):19–22. Article in Russian. Razumov, I. A., Gileva, I. P., M. A. Vasil’eva, T. S. Nepomniashchikh, M. N. Mishina, E. F. Belanov, G. V. Kochneva, E. E. Konovalov, S. N. Shchelkunov, and V. B. Loktev. 2005. Neutralizing monoclonal antibodies cross-react with fusion proteins encoded by 129L of the ectromelia virus and A30L of the variola virus. Molecular Biology 39(6):918–925. Ryabinin, V. A., L. A. Shundrin, E. B. Kostina, M. Laassri, V. Chizhikov, S. N. Shchelkunov, K. Chumakov, and A. N. Sinyakov. 2006. Microarray assay for detection and discrimination of Orthopoxvirus species. Journal of Medical Virology 78(10):1325–1340. Scaramozzino, N., A. Ferrier-Rembert, A. L. Favier, C. Rothlisberger, S. Richard, J. M. Crance, H. Meyer, and D. Garin. 2007. Real-time PCR to identify variola virus or other human pathogenic orthopox viruses. Clinical Chemistry 53(4):606–613. Schoepp, R. J., M. D. Morin, M. J. Martinez, D. A. Kulesh, L. Hensley, T. W. Geisbert, D. R. Brady, and P. B. Jahrling. 2004. Detection and identification of variola virus in fixed human tissue after prolonged archival storage. Laboratory Investigation: A Journal of Technical Methods and Pathology 84(1):41–48. Shchelkunov, S. N., E. V. Gavrilova, and I. V. Babkin. 2005. Multiplex PCR detection and species differentiation of orthopoxviruses pathogenic to humans. Molecular & Cellular Probes 19(1):1–8. Sias, C., F. Carletti, M. R. Capobianchi, D. Travaglini, R. Chiappini, D. Horejsh, and A. Di Caro. 2007. Rapid differential diagnosis of orthopoxviruses and herpesviruses based upon multiplex real-time PCR. Infez Med 15(1):47–55.

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 LIVE VARIOLA VIRUS Sulaiman, I. M., K. Tang, J. Osborne, S. Sammons, and R. M. Wohlhueter. 2007. GeneChip resequencing of the smallpox virus genome can identify novel strains: A biodefense appli- cation. Journal of Clinical Microbiology 45(2):358–363. Sulaiman, I. M., S. A. Sammons, and R. M. Wohlhueter. 2008. Smallpox virus resequencing GeneChips can also rapidly ascertain species status for some zoonotic non-variola orthopoxviruses. Journal of Clinical Microbiology 46(4):1507–1509. Ulaeto, D., et al., Poster presentation. XIV International Poxvirus &. Iridovirus Workshop, Lake Placid, New York, September 19–25, 2002.