- Open Access
Rapid and sensitive point-of-care detection of Orthopoxviruses by ABICAP immunofiltration
© The Author(s). 2016
- Received: 1 July 2016
- Accepted: 2 December 2016
- Published: 9 December 2016
The rapid and reliable detection of infectious agents is one of the most challenging tasks in scenarios lacking well-equipped laboratory infrastructure, like diagnostics in rural areas of developing countries. Commercially available point-of-care diagnostic tests for emerging and rare diseases are particularly scarce.
In this work we present a point-of-care test for the detection of Orthopoxviruses (OPV). The OPV ABICAP assay detects down to 1 × 104 plaque forming units/mL of OPV particles within 45 min. It can be applied to clinical material like skin crusts and detects all zoonotic OPV infecting humans, including Vaccinia, Cowpox, Monkeypox, and most importantly Variola virus.
Given the high sensitivity and the ease of handling, the novel assay could be highly useful for on-site diagnostics of suspected Monkeypox virus infections in areas lacking proper laboratory infrastructure as well as rapid on-site testing of suspected bioterrorism samples.
- Variola virus
- Monkeypox virus
- Cowpox virus
- Vaccinia virus
- Rapid detection
OPV belong to the Poxviridae family and are large and complex DNA viruses able to infect humans, causing severe diseases . Despite the successful eradication of smallpox caused by Variola virus (VARV), zoonotic infections with still circulating OPVs, namely Monkeypox virus (MPXV), Cowpox virus (CPXV), and Vaccinia virus (VACV) , remain a threat to an increasing number of unvaccinated individuals . Diagnostics and differentiation of OPV from other rash-inducing agents  are usually done by detection of either viral particles by negative staining electron microscopy or viral DNA by quantitative real-time PCR (qPCR), both requiring dedicated laboratories. However, rapid and highly sensitive point-of-care (POC) diagnostics are needed because of the rising incidence of MPXV infections in rural Africa [5, 6] as well as the risk of a potential deliberate release of Variola virus in the case of a bioterrorist attack [7, 8].
In this work, we describe a rapid and sensitive detection system based on the ABICAP technology that reliably detects particles of different human-pathogenic OPV.
Antibodies and proteins
The diagnostic ABICAP is based on two monoclonal antibodies (mAbs) targeting the OPV surface protein A27: mAb A1/40 and mAb A3/710. Briefly, both antibodies bind to A27 with high affinity (A1/40: 2.3 nM; A3/710: 4.3 nM) and recognize non-overlapping linear epitopes located in close proximity to the N-terminal heparin-binding domain of A27 (A1/40: aa 24–38; A3/710: aa 13–27). Both antibodies have previously been selected in an antigen capture ELISA, representing the best combination to detect native VACV, CMLV, CPXV, ECTV, and MPXV. Production, purification, storage, and biotinylation of antibodies was performed as described previously . The following reagent was obtained through the NIH Biodefense and Emerging Infections Research Resources Repository (BEI Resources), NIAID, NIH: Vaccinia virus (WR) rA27 (NR-2622) with a C-terminal histidine tag, recombinantly expressed from baculovirus.
The virus panel used for development and validation included VACV strain New York City Board of Health (NYCBOH, VR-1536™, ATCC/LGC Standards GmbH, Wesel, Germany) and CPXV strain GuWi . Camelpox virus (CMLV) strain CP-19, Ectromelia virus (ECTV) strain Nü-1, and MPXV strain MSF6 were kindly provided by Prof. Dr. Hermann Meyer (Bundeswehr Institute of Microbiology, Munich, Germany). The OPV ABICAP detection system was additionally validated with two VARV strains, VARV Solaiman 1974 and VARV Niger 1969. Parapoxvirus (PPV) ORF D1701 was kindly provided by Achim Rziha (Friedrich-Loeffler-Institute, Tübingen, Germany) whereas herpes simplex virus 1 (HSV-1) was isolated in our lab from a patient. Propagation and titration of viruses on cell culture was done according to standard procedures . Viruses were used as clarified supernatant (HSV, PPV, MPXV, and VARV) or as semi-purified viral particles by centrifugation through a 40% sucrose cushion . All viruses were used natively except for MPXV which was heat-inactivated at 60 °C for 2 h. All work with live VARV was conducted within a biosafety level 4 laboratory in accordance to guidelines and approvals from the World Health Assembly Advisory Committee on Variola Virus Research.
For final validation, a panel containing inactivated (γ-irradiation) highly pathogenic viruses comprising Yellow fever virus strain 17D, Ebola virus strain Zaire, Marburg virus, VACV, and MPXV in Dulbecco’s Modified Eagle Medium was kindly provided by the P.R.O.B.E. consortium. Viruses in this panel have been quantified by quantitative real-time PCR using published assays [14, 15] and were investigated in a blinded manner to test the suitability of the ABICAP assay to discern highly pathogenic viruses in a bioterrorist setting.
Clinical sample material
Clinical sample material tested in this work had been sent to the Robert Koch Institute for poxvirus diagnostics. Crust material and surface swabs were delivered dry and suspended in 500 μL of phosphate buffered saline (PBS, pH 7.4) without magnesium and calcium. Surface swabs were thoroughly mixed on a vortexer while crusts were homogenized in a Fastprep 24 homogenizer (MP Biomedicals, Eschwege, Germany) after addition of Precellys® ceramic beads (1.4 mm; PEQLAB, Erlangen, Germany), and DNA was prepared from 200 μL of sample using the Qiagen Blood and Tissues Kit (Qiagen, Hilden, Germany) according the manufacturer’s recommendations. Quantitative real-time PCR for OPV diagnostics was performed on 5 μL of isolated DNA, using rpo18 as target as published previously . The number of genome equivalents (GE) contained in the samples was estimated based on the published calibration curve of the rpo18 assay. To test the specificity of the ABICAP assay, clinical samples negative in the OPV PCR were also tested. For differential diagnostics, different PCR assays targeting Molluscipox virus , Parapox virus , Myxoma virus  were tested on isolated DNA from homogenized crusts or swabs. Homogenized samples in PBS were tested in the ABICAP assay after dilution in UCBS casein buffer (SDT, Baesweiler, Germany).
OPV ABICAP assay
The preparation of ABICAP columns was done as described before , using mAb A1/40 immobilized on polyethylene filter frits contained inside the ABICAP columns and biotinylated mAb A3/710 as the detection antibody.
To this aim, polyethylene filter frits (Type 180; Porex, Aachen, Germany) used for the coating of the capture antibodies were activated by successive 10-min washes with 96% ethanol, 50% ethanol, and 3 × coating buffer (0.1 M NaHCO3/ Na2CO3, pH 9.5). All activating and coating steps were done at room temperature under constant stirring and vacuum. The frits were then coated with 7.5 μg/frit of mAb A1/40 in coating buffer (75 μg/mL) overnight and blocked for 25 min with PBS-T BND (PBS + 0.05% Tween-20 + 0.05% BND [5-bromo-5-nitro-1,3-dioxane; SDT, Baesweiler, Germany]) + 0.2% bovine serum albumin (BSA; Carl Roth, Karlsruhe, Germany). Finally, frits were prepared for drying by 1 h of incubation with PBS-T BND + 1% BSA + 5% saccharose and air-dried at 35 °C for 30 min in a fluid bed dryer (FBD2000; Endecotts, London, UK). The columns were equipped with antibody-coated frits between two preblocked separation frits (Type 187; Porex) and stored dry at room temperature until further use.
The principle of ABICAP™ immunofiltration assay (Antibody Immuno Column for Analytical Processes, Senova GmbH, Weimar, Germany) is shown in Fig. 1. 500 μL of pre-diluted sample material (UCBS casein buffer, SDT) were applied per column and incubated for 6 min. After a washing step with 750 μL of washing buffer (PBS-T BND + 0.1% BSA), 500 μL of biotinylated mAb A3/710–20 (in AA1-buffer [SDT]) were added and incubated for 6 min. Columns were washed again and incubated with 500 μL of streptavidin-PolyHRP 40 (SA-pHRP; in SA1-buffer, both SDT) for 6 min. Two washing steps later (1 × 750 μL of washing buffer, 1 × 750 μL of substrate buffer [phosphate/citrate buffer, pH 5.0]) 500 μL of precipitating TMB substrate (epTMB; SDT) were added for 6 min. Finally, columns were washed with 750 μL of substrate buffer, and the extinctions were read at 525 nm with a handheld photometer device (Senova; Fig. 1).
To allow for maximum sensitivity of detection at low background, both SA-pHRP and biotinylated detection antibodies were titrated (0.1, 0.5, 1, and 5 μg/mL) in the absence of antigen. For titration of SA-pHRP, the protocol was performed as described, but instead of biotinylated detection antibody AA1-buffer only was added. For titration of biotinylated detection antibodies, SA-pHRP was added at the optimum concentration determined in the previous experiment. Low background was obtained with mAb A3/710 at 0.5 μg/mL and SA-pHRP at 0.5 μg/mL (Additional file 1: Figure S1).
The cutoff to determine the limits of detection (LOD) and to discern positive from negative samples was calculated as the mean + 3 × standard deviation of negative control columns (buffer only, n = 16). The assay’s sensitivity was determined by measuring the LOD for recombinant A27. The assay’s specificity was determined for various OPV strains by using purified viral particles and by testing clinical crust material from CPXV-infected patients. To determine the test’s specificity, additional viruses causing similar clinical pictures were used, including HSV-1 and PPV. The LOD was determined by measuring serial dilutions of semi-purified virus particles and virus from clarified cell culture supernatant. Finally, the OPV ABICAP assay was validated with a blinded panel of PCR-quantified highly pathogenic viruses, including Yellow Fever virus, Ebola virus, Marburg virus, VACV, and MPXV.
To establish the ABICAP assay, two high-affinity and epitope-matched monoclonal antibodies targeting the orthopoxviral attachment protein A27 were tested: mAb A1/40 and mAb A3/710. Based on previous results from sandwich-ELISAs, mAb A1/40 was immobilized as a capture ELISA while biotinylated mAb A3/710 was used as detection antibody. This combination proved superior to all other combinations tested with 8 monoclonal antibodies targeting A27 .
Using the presented protocol, all OPV strains tested were detected with sensitivities of 3.4 × 104 PFU/mL for VACV, 2.4 × 104 and 8.3 × 103 PFU/mL for VARV (Solaiman and Niger strain, respectively), 6.8 × 103 PFU/mL for CPXV, 1.3 × 103 PFU/mL for MPXV, and 6.8 × 103 PFU/mL for CMLV (Fig. 2b). ECTV showed a significantly increased LOD (1.9 × 106 PFU/mL), which was expected since a previous study showed impaired binding of capture antibody A1/40 to ECTV . As ECTV does not cause zoonotic disease in humans and usually only infects laboratory mice, the lacking sensitivity is irrelevant for clinical application of the OPV ABICAP assay.
Analysis of clinical samples
Validation of the OPV ABICAP assay with clinical samples
Cat, homogenized crust
17.4 (1.6 × 109 GE/mL)
Cat, homogenized skin biopsy
27.5 (1.4 × 106 GE/mL)
Human, homogenized crust
11.7 (8.2 × 1010 GE/mL)
Human, swab of crust
19.6 (3.4 × 108 GE/mL)
Human, homogenized crust
Unknown (OPV + PPV PCR negative)
0.334 ± 0.049
Rat, homogenized crust
Unknown (OPV PCR negative)
0.245 ± 0.007
Rabbit, homogenized crust
0.305 ± 0.092
Human, swab of rash
0.285 ± 0.007
1:10 (2.5 × 105TCID50/ ml)
0.275 ± 0.022
Herpes simplex virus
1:10 (6.9 × 107 PFU/ ml)
0.235 ± 0.065
Application of the ABICAP OPV assay in an external quality assessment for diagnostics of BT samples
Validation of the OPV ABICAP assay in an external quality assessment for diagnostics of BT samples
Genome equivalents tested [GE/mL]
9.2 × 105
1.570 ± 0.001
2.4 × 107
2.032 ± 0.185
6.2 × 104
0.437 ± 0.023
2.3 × 103
0.119 ± 0.016
2.7 × 106
0.169 ± 0.009
8.2 × 106
0.152 ± 0.050
0.194 ± 0.004
Cell culture medium (DMEM)
0.181 ± 0.005
In conclusion, the OPV ABICAP assay was able to detect OPV rapidly from clinical specimens as well as from blinded samples included in a bioterror panel. The assay was sufficiently sensitive to confirm the virus presence in all clinical samples. The observed LOD for real-world samples measured with the OPV ABICAP assay corresponded to approximately 5 × 104 GE/mL.
Molecular diagnostics of infectious diseases has undergone a rapid development over the past years. While there are numerous commercially available tests for clinically relevant infectious diseases, there is an obvious gap regarding tests for rare or emerging diseases, particularly with respect to on-site diagnostic tests. Although unrivalled in sensitivity and specificity, PCR has its drawbacks in field diagnostics, requiring a certain amount of technical equipment, practical skills, and diagnostic knowledge. In contrast, so-called rapid tests like lateral flow assays (LFA) usually circumvent these demands and are easy to perform, but lack sensitivity due to the absence of a signal amplification step. However, they present a promising approach for POC diagnostics.
The established OPV ABICAP immunofiltration assay is the first POC detection assay covering all zoonotic OPV as well as VARV with a satisfactory sensitivity. One previously developed handheld rapid test for poxviruses, the Tetracore Orthopox Biothreat detection assay, displays an LOD of 106 to 107 PFU/mL which is usually sufficient for the diagnosis of skin lesions containing extremely high viral loads . The new OPV ABICAP assay offers an LOD as low as 104 PFU/mL which allows reliable diagnosis even when viral loads in the sample are low because of inappropriate sampling or non-optimal time points in the course of infection. Similarly, an ABICAP assay for the detection of Ebola virus showed an LOD of approximately 1.2 × 104 PFU/mL, indicating that ABICAP systems generally represent sensitive platforms for particle detection . Additionally, the described LODs can be reached within 45 min, which is comparable to the Tetracore assay time.
The higher sensitivity of the ABICAP assay compared to lateral flow assays is also in agreement with previous reports with an ABICAP assay targeting botulinum neurotoxin. Furthermore, in an independent evaluation the ABICAP showed sensitivity comparable to that of an in-house ELISA whereas commercial LFAs either failed completely or lacked sensitivity, even compared to the relatively high LODs according to the manufacturers . Our study also shows that well-characterized antibodies can easily be transferred from ELISA to ABICAP assays. Here, the specific characteristics of the immunofiltration approach offer the clear advantage of relative simplicity and rapid assay times at comparable sensitivity.
Particularly when performing field diagnostics, adequate sampling and sample processing can be impaired dramatically by environmental conditions. Although the ABICAP assay was highly specific for all samples tested in this work, false-positive results remain a risk as in any diagnostics, especially when testing clinical sample material. Independent of the high specificity of the implemented antibodies, matrix effects might cause higher background signals. For an exhaustive validation, more samples, ideally in the field, will be required to be analyzed using this assay.
Finally, as all reagents for an OPV ABICAP test can be stored stably at 4 °C or even ambient temperature, there is no need to freeze reagents to -20 °C, which is usually a limiting factor in the regions in which the diagnosed diseases are endemic. The OPV ABICAP assay is therefore a low-tech alternative, particularly when compared to PCR, and represents an optimal compromise between ease of handling and sensitivity. Most importantly, not only was the ABICAP system able to detect OPV from clinical samples, but also to differentiate OPV from other viruses that can (like OPV) cause skin lesions or potentially be used as bioterror agents.
The panel of different viruses for validation was kindly provided by the P.R.O.B.E. project where it was used for final validation of mobile diagnostic devices. The authors would like to thank Jung-Won Sim-Brandenburg and Silvia Muschter for excellent technical assistance, Andrea Schnartendorff for the pictures of the ABICAP column and reader, and Ursula Erikli for copy-editing.
This work was supported by a grant from the Federal Ministry of Education and Research [grant numbers 13 N9601, 13 N9596].
Availability of data and materials
All data generated or analyzed during this study are included in this published article.
D.S., V.A.O., S.K.S., M.P., L.M., P.M., B.G.D., and A.N. designed the study. D.S., V.A.O., S.K.S, and M.P. performed the experiments. D.S. and A.N. wrote the manuscript with input from the other authors. All authors read and approved the final manuscript.
M.P. and P.M. are affiliated with Senova, the commercial distributor of the ABICAP platform. All other authors report no potential conflicts.
Consent for publication
Ethics approval and consent to participate
The findings and conclusions in this report are those of the author(s) and do not necessarily represent the official position of the Centers for Disease Control and Prevention.
Open AccessThis article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated.
- Damon IK. Poxviruses. In: Knipe DM, Howley PM, editors. Fields virology. 6th ed. Philadelphia: Wolters Kluwer/Lippincott Williams & Wilkins Health; 2013. p. 2160–84.Google Scholar
- Essbauer S, Pfeffer M, Meyer H. Zoonotic poxviruses. Vet Microbiol. 2009;140:229–36.View ArticlePubMedGoogle Scholar
- Shchelkunov SN. An increasing danger of zoonotic orthopoxvirus infections. PLoS Pathog. 2013;9:e1003756.View ArticlePubMedPubMed CentralGoogle Scholar
- MacNeil A, Reynolds MG, Carroll DS, Karem K, Braden Z, Lash R, Moundeli A, Mombouli JV, Jumaan AO, Schmid DS, Damon IK. Monkeypox or varicella? Lessons from a rash outbreak investigation in the Republic of the Congo. Am J Trop Med Hyg. 2009;80:503–7.PubMedGoogle Scholar
- Rimoin AW, Mulembakani PM, Johnston SC, Lloyd Smith JO, Kisalu NK, Kinkela TL, Blumberg S, Thomassen HA, Pike BL, Fair JN, et al. Major increase in human monkeypox incidence 30 years after smallpox vaccination campaigns cease in the Democratic Republic of Congo. Proc Natl Acad Sci U S A. 2010;107:16262–7.View ArticlePubMedPubMed CentralGoogle Scholar
- McCollum AM, Damon IK. Human monkeypox. Clin Infect Dis. 2014;58:260–7.View ArticlePubMedGoogle Scholar
- Arita I, Francis D. Is it time to destroy the smallpox virus? Science. 2014;345:1010.View ArticlePubMedGoogle Scholar
- Henderson DA, Arita I. The smallpox threat: a time to reconsider global policy. Biosecur Bioterror. 2014;12:117–21.View ArticlePubMedGoogle Scholar
- Mabey D, Peeling RW, Ustianowski A, Perkins MD. Diagnostics for the developing world. Nat Rev Microbiol. 2004;2:231–40.View ArticlePubMedGoogle Scholar
- Lucht A, Formenty P, Feldmann H, Gotz M, Leroy E, Bataboukila P, Grolla A, Feldmann F, Wittmann T, Campbell P, et al. Development of an immunofiltration-based antigen-detection assay for rapid diagnosis of Ebola virus infection. J Infect Dis. 2007;196 Suppl 2:S184–92.View ArticlePubMedGoogle Scholar
- Stern D, Pauly D, Zydek M, Miller L, Piesker J, Laue M, Lisdat F, Dorner MB, Dorner BG, Nitsche A. Development of a Genus-Specific Antigen Capture ELISA for Orthopoxviruses-Target Selection and Optimized Screening. PLoS One. 2016;11:e0150110.View ArticlePubMedPubMed CentralGoogle Scholar
- Kurth A, Wibbelt G, Gerber HP, Petschaelis A, Pauli G, Nitsche A. Rat-to-Elephant-to-Human Transmission of Cowpox Virus. Emerg Infect Dis. 2008;14:670–1.View ArticlePubMedPubMed CentralGoogle Scholar
- Kotwal GJ, Abrahams MR. Growing poxviruses and determining virus titer. Methods Mol Biol. 2004;269:101–12.PubMedGoogle Scholar
- Weidmann M, Muhlberger E, Hufert FT. Rapid detection protocol for filoviruses. J Clin Virol. 2004;30:94–9.View ArticlePubMedGoogle Scholar
- Kramski M, Drozd A, Lichtfuss GF, Dabrowski PW, Ellerbrok H. Rapid detection of anti-Vaccinia virus neutralizing antibodies. Virol J. 2011;8:139.View ArticlePubMedPubMed CentralGoogle Scholar
- Nitsche A, Ellerbrok H, Pauli G. Detection of orthopoxvirus DNA by real-time PCR and identification of variola virus DNA by melting analysis. J Clin Microbiol. 2004;42:1207–13.View ArticlePubMedPubMed CentralGoogle Scholar
- Schroeder K, Nitsche A. Multicolour, multiplex real-time PCR assay for the detection of human-pathogenic poxviruses. Mol Cell Probes. 2010;24:110–3.View ArticlePubMedGoogle Scholar
- Nitsche A, Buttner M, Wilhelm S, Pauli G, Meyer H. Real-time PCR detection of parapoxvirus DNA. Clin Chem. 2006;52:316–9.View ArticlePubMedGoogle Scholar
- Albini S, Sigrist B, Guttinger R, Schelling C, Hoop RK, Vogtlin A. Development and validation of a Myxoma virus real-time polymerase chain reaction assay. J Vet Diagn Invest. 2012;24:135–7.View ArticlePubMedGoogle Scholar
- Townsend MB, MacNeil A, Reynolds MG, Hughes CM, Olson VA, Damon IK, Karem KL. Evaluation of the Tetracore Orthopox BioThreat(R) antigen detection assay using laboratory grown orthopoxviruses and rash illness clinical specimens. J Virol Methods. 2013;187:37–42.View ArticlePubMedGoogle Scholar
- Slotved HC, Tanassi JT, Sparding N, Lindqvist A, Steenhard NR, Heegaard NH. Botulinum toxin field assays evaluated using cosmetic botox preparations. Biosecur Bioterror. 2013;11:280–6.View ArticlePubMedGoogle Scholar