Monkeypox Diagnosis Approaches

Monkeypox virus (MPXV) is an adenovirus with a double-stranded DNA genome, belonging to the family Poxviridae, subfamily Chordopoxvirinae, and the genus Orthopoxvirus. MPXV was first reported in 1958 after two pox-like disease outbreaks occurred in monkeys. The original source of MPXV is unknown. Rodents likely harbor the virus, leading to spillover events. The case of human infection by MPXV was first reported in humans in in the Democratic Republic of the Congo in 1970.

monkeypox
real-time PCR
immunoassay
diagnosis

1. Indirect Detection

Indirect detection is based on virus-induced morphological changes to host cells or membranes.

1.1. Monkeypox Diagnosis Based on Virus Culture

Some viruses can induce macroscopic lesions (called pocks) on the chick chorioallantoic membrane (CAM). The pattern of pock formation, the time required for pock formation, and the size of the pock have been explored to differentiate different poxvirus infections, including monkeypox virus (MPXV) ^[1]^[2]. The morphological changes can be observed with a microscope or the naked eye. For instance, when CAM was inoculated with MPXV, the pocks were visible and could be reckoned with the naked eye ^[2]. However, detection solely based on the above-mentioned characteristics may not be sufficient for an accurate diagnosis due to overlapping signs and symptoms with other diseases.

Monkeypox isolates are grown in RK13 cells ^[3], where cytopathic effects are observed within 24–48 h of infection. The major drawback of culture-based virus diagnosis is the prolonged assay time ^[4], which is not suitable for mass testing scenarios. Further, virus culture methods need biosafety level 3 (BSL3) labs and pose a risk of laboratory-acquired infections ^[5]. Shell vial culture (SVC) has been developed as an alternative culture method for the rapid in vitro detection of MPXV and other viruses ^[4]. In this method, a cell monolayer is grown on a cover slip in a shell vial culture tube, and the specimen is inoculated on the monolayer, followed by low-speed centrifugation and immunofluorescence-based detection. The low-speed centrifugation step is introduced to enhance the virus’s infectivity. The mechanical force resulting from low-speed centrifugation is thought to cause cell trauma, which subsequently enhances viral entry into cells, resulting in a reduced cell infection time ^[6].

1.2. Diagnosis Based on Image Analysis

Image digitalization has already gained momentum for infectious disease diagnosis and monitoring. Chatbots have been developed for disease diagnostic evaluation and the recommendation of immediate measures in case a patient contracts SARS-CoV-2 ^[7]. A monkeypox image dataset was constructed comprising 43 original images and 587 images obtained after data augmentation ^[8] (Figure 1). Using the newly developed “Monkeypox 2022” dataset, an image classification model was proposed ^[9]. The study paves the way towards the development of image-analysis-based tools for monkeypox virus detection. The images used in the dataset are from previous outbreaks. The classic MPXV cases were characterized by a generalized rash. In contrast, most of the cases in the current outbreak have localized lesions in anogenital and genitourinary areas ^[10]^[11]. Since many recent MPXV-infected cutaneous images have been reported, the updated dataset may have added value to the above MPXV 2022 image dataset.

Figure 1. Schematic of the image-based MPXV detection workflow. Redrawn from Ref. ^[9] with permission from the author. The source content is licensed under a Creative Commons Attributions 4.0 international license.

2. Direct Detection

In the case of direct detection, nucleic acid and protein components of the virus are detected without the need for a pathogen culture. Molecular detection, immunodiagnostics, and sequencing are widely explored direct detection approaches.

2.1. Monkeypox Immunodiagnostics

The hemagglutination test is a simple and cost-effective approach for virus detection. The test is based on the agglutination of erythrocytes in the presence of a virus ^[4]. The hemagglutination mechanism led to the development of another assay called the hemagglutination inhibition (HI) assay ^[12]. The HI approach relies on virus-specific antibodies to detect viral antigens. The MPXV strains are tested using hemagglutination and HI tests ^[2]. The test cannot differentiate MPXV from the variola and vaccinia viruses but can differentiate cowpox from MPXV and can be used to estimate the evolutionary relationships of viral strains or species.

The enzyme-linked immunosorbent assay is a widely used protein detection method ^[13]. A commercially available Orthopox BioThreat^® Alert Assay for orthopox virus (OPV) detection is a reliable OPV detection method ^[14]. This antibody-based lateral flow assay captures virus antigens and detects the viral load at 10⁴ PFU/mL. The surface protein A27 was found to be the most immunogenic protein for virus particle capture and detection ^[15]. After a comprehensive screening of A27-binding antibodies, an ELISA approach was developed for orthopoxviruses, including MPXV. The method’s detection limit is 1× 10³ PFU/mL. Recently, Ulaeto et al. described the characteristics of an LFA for the detection of orthopoxviruses ^[16]. The assay detects vaccinia virus samples spiked in human saliva and clinical sample buffer with a detection limit of between 10⁴ and 10⁵ PFU/mL within 20 min. Since this assay detects orthopoxviruses, the test can be further explored for MPXV detection in real samples. Combining the clinical presentation of MPXV with the LFA test could provide a rapid MPXV detection tool. All of the above-mentioned immunodetection modalities are suitable for generic orthopox virus detection applications, but none of them are specific for MPXV.

2.2. Whole-Particle Detection

Finding biomarkers for a newly emerged virus is challenging and may hamper the direct implementation of routine diagnostic methods. In this regard, whole-particle detection using electron microscopy (EM) is a powerful alternative ^[17]. Transmission electron microscopy is a good first step for the detection of viruses, as it provides information about the shape and amount of viral load with a small sample volume ^[18]. The use of virus-specific antibodies in immunoelectron microscopy (IEM) further improves the detection accuracy of EM ^[19]. EM has been used to detect monkeypox and other orthopoxviruses ^[20]. Although EM is suitable for the laboratory validation of the virus detection results, the approach has certain limitations, such as the high cost of the instrument, the requirement of highly trained staff, and low sample throughput ^[20].

2.3. Detection by Genome Sequencing

Genome sequencing is the gold standard to identify novel or mutated viruses. Genome sequencing not only identifies the target virus but may pinpoint the presence of other viruses in the sample that can help to create a treatment plan for a particular disease. MPXV detection based on qPCR coupled with genome sequencing has been reported ^[21]. To date, 200 genome sequences of MPXV isolates from recent outbreaks in non-endemic countries have been reported ^[22]. Whole-genome sequencing is a time-consuming process and requires expensive instruments, trained staff, and skilled bioinformaticians for computational analyses. These limitations need to be overcome to harness the potential of genome sequencing approaches.

2.4. Monkeypox Virus Detection Based on PCR

The polymerase chain reaction (PCR) is widely regarded as the gold standard for nucleic acid detection. According to WHO recommendations, PCR (conventional or real-time) is a standard method for MPXV laboratory validation ^[23]. The detection can be combined with sequencing or other orthopox detection assays ^[24]. Conventional PCR-based MPXV detection involves PCR amplification and restriction digestion of the PCR-amplified fragments to identify MPXV based on restriction fragment length polymorphisms.

A hemagglutinin PCR (HA-PCR) assay was developed based on MPXV-specific primers coupled with TaqI restriction digestion ^[25]. The method could not distinguish different MPXV isolates. To improve the detection accuracy of the PCR assay, an A-type inclusion body protein (ATI) gene has been used to detect MPXV and other orthopoxviruses based on PCR-based gene amplification and XbaI digestion ^[26]^[27]. The method can differentiate MPXV strains based on restriction digestion. In another development, the open reading frame (ORF) of the ATI gene was identified, sequenced, and compared with other related poxviruses ^[28]. Unique deletions were found in the OFR of MPXV and were harnessed for the specific detection of the MPXV ATI gene. This PCR method differentiates 19 MPXV strains. The specificity was confirmed by BglII restriction digestion.

Compared to traditional PCR, real-time PCR is rapid and sensitive. Due to the low GC content and almost 90% genome identity with other Eurasian orthopoxviruses, designing an MPXV-specific TaqMan assay is challenging. Li et al. developed a real-time PCR assay where minor-groove-binding protein-based (MGB) probes were developed ^[29]. The use of MGB stabilizes probe–template interactions, enables the use of small probe sequences for single-nucleotide polymorphism (SNP) detection, and enhances assay sensitivity and specificity ^[30]. The method could detect 15 MPXV isolates at a 10 ng concentration. The assay efficiency with freshly diluted DNA is 97%, while it is reduced to 67% after multiple freeze–thaw cycles. These observations indicate that a fresh sample should be used in order to achieve maximum assay efficiency. The detection of MPXV and other orthopoxviruses based on melting-curve analysis (MCA) has also been reported ^[31]^[32]^[33]. Both clades (West African and Congo Basin) of MPXV have 99% sequence identity but are significantly different in terms of virulence ^[34]. It is a big challenge to develop a clade-specific real-time PCR detection approach due to the limited availability of unique sequences.

Multiplex detection can significantly reduce the misidentification of coexisting pathogens ^[35]^[36]. A multicolor, multiplex approach for MPXV detection was reported where MPXV was specifically detected in the presence of the variola virus (VARV) and the varicella-zoster virus (VZV) ^[37]. The target genes harboring unique sequences for MPXV, VARV, and VZV are F3L, B12R, and ORF38, respectively. The specificity of the developed approach is 100%, and LODs of 20 copies per reaction for MPXV and VARV and 50 copies per reaction for VZV were reported. The robustness of the approach was demonstrated by successfully detecting the different combinations of MPXV, VARV, and VZV samples.

The standard poxvirus detection approach combines the disease’s clinical symptoms with a generic poxvirus PCR assay, followed by a poxvirus-specific PCR assay ^[33]. These pan-pox real-time PCR methods are instrumental in the accurate diagnosis of poxvirus infection. Based on the GC content, the chordopoxviruses (poxviruses that infect vertebrates) of the subfamily Chordopoxvirinae have two distinct genome types: one genome type contains high GC content (>60%), while the other genome type is comprised of low GC content (30–40%) ^[38]. GC-content-based pan-pox PCR assays have been developed ^[38]. The assays are termed high-GC PCR and low-GC PCR assays. The developed PCR assays detected DNA samples from more than 150 isolates and strains of chordopoxviruses. The detection approach is based on conventional PCR, and PCR amplicons are evaluated by TaqI RFLP patterns. In a similar line of work, a real-time PCR assay for the universal detection of orthopoxviruses was reported ^[36]. The system was reported to be able to detect poxviruses excluded in a previous study ^[38] as well as those from the subfamily Entomopoxvirinae. This assay targets a 100 bp highly conserved sequence in the D6R gene of poxviruses. The specificity of the assay for vertebrate samples is 99.8%, while it is 99.7% for arthropod samples. The system is 100% sensitive for vertebrate samples and 86.6% sensitive for arthropod samples. The detection limits are reported to be 100 or 1000 copies per reaction, depending on the poxvirus species.

2.5. Detection Based on Isothermal Amplification

More than ten types of different isothermal amplification methods have been reported and demonstrated for nucleic detection ^[39]. Loop-mediated isothermal amplification (LAMP) and recombinase polymerase amplification (RPA) are well-explored isothermal nucleic acid amplification based virus detection methods ^[40]. The LAMP technology relies on two internal primers called the forward internal primer (FIP) and the backward internal primer (BIP), two outer primers known as the forward outer primer (F3) and the backward outer primer (B3), and a DNA polymerase with strand displacement activity ^[40]. The reaction is carried out at 60–65 °C. The amplification reaction is accelerated by using two loop primers, the forward loop (LF) and the backward loop primer (LB) ^[41]. The annealing of the FIP, which has two target sequences (separated by a spacer) complementary to the two different regions of the template, initiates strand synthesis and elongation (Figure 2A). Subsequently, the F3 primer displaces the FIP strand, producing a single-stranded DNA (ssDNA) strand that is used as a template by the BIP (Figure 2B). The BIP, which also has two target sequences complementary to the template DNA at two different regions, starts the strand elongation of the ssDNA template, which is later displaced by the B3 (Figure 2C). The 5′ and 3′ ends of the template DNA have inward complementary sequences, forming a stem-looped DNA that is exponentially amplified by loop primers (Figure 2C,D). LAMP-based MPXV-clade-specific assays have been developed where West African (the assay named W-LAMP) and Congo Basin MPXV (the assay named C-LAMP) clades are selectively detected ^[42]. A turbidimeter is used to analyze the LAMP reaction, and restriction digestion is used to confirm the LAMP products. A LAMP-based method for rapid MPXV detection was recently posted on a preprint server ^[43]. The assay was developed to detect MPXV clades. The method shows satisfactory sensitivity and response times.

Figure 2. Reaction mechanism of LAMP. See text for details. Redrawn from Becherer et al., 2020, Ref. ^[40] © The Royal Society of Chemistry 2020, licensed under a Creative Commons Attributions-Noncommercial 3.0 unported license https://creativecommons.org/licenses/by-nc/3.0/. Accessed on 12 October 2022.

Although promising, the LAMP needs a 60-minute reaction time and six primers. Furthermore, primer design is relatively complex. To overcome these limitations, RPA has been proposed as an attractive alternative ^[44]. The RPA signal is detected by gel electrophoresis, real-time monitoring ^[45], or lateral flow assay ^[45]. In the case of real-time detection, the fluorogenic probe, along with the primers, is added to the reaction system where cleavage of the probe by exonuclease leads to a fluorescent signal. RPA-based MPXV detection shows satisfactory results with reduced assay times and reagent costs ^[46].

Table 1. Summary of MPXV diagnostic methods.

Sr. No	Assay Name	Target Gene	Primers’ Sequences	Probes’ Sequences	Detection Limit	Real-Sample Analysis	References
1	HA-PCR	HA gene	Forward: 5′-CTGATAATGTAGAAG AC -3′ Reverse: 5′-TTGTATTTACGTGGGTG-3′	NA	Not reported	Yes	^[25]
2	ATI-PCR	ATI-gene	Forward: 5′-AATACAAGGAGGATCT-3′ Reverse: 5′-CTTAACTTTTTCTTTTTCTTTCTC-3′	NA	Not reported	Yes	^[26]
3	MPXV PCR assay	ATI-gene	Forward: 5′-GAGAGAATCTCTTGATAT-3′ Reverse: 5′-ATTCTAGATTGTAATC-3′	NA	Not reported	Yes	^[28]
4	Real-time PCR	B6R	Forward: 5′-ATTGGTCATTATTTTTGTCACAGGAACA-3′ Reverse: 5′-AATGGCGTTGACAATTATGGGTG-3′	5′-MGB/DarkQuencher-AGAGATTAGAAATA-3′-FAM	∼10 viral copies (2 fg)	Yes	^[29]
5	Real-time PCR	G2R	Forward: 5′-CACACCGTCTCTTCCACAGA -3′ Reverse: 5′-GATACAGGTTAATTTCCACATCG -3′	5′-FAM AACCCGTCGTAACCAGCAATACATTT-3′-BHQ1	∼8.2 genome copies (1.7 fg)	Yes	^[47]
6	Real-time PCR	G2R	Forward: 5′-TGTCTACCTGGATACAGAAAGCAA-3′ Reverse: 5′-GGCATCTCCGTTTAATACATTGAT -3′	5′-FAM-CCCATATATGCTAAATGTACCGGTACCGGA-3′-BHQ1	∼40.4 copies (9.46 fg)	Yes	^[47]
7	Real-time PCR	F3L	Forward: 5′-CTCATTGATTTTTCG CGGGAT A-3′ Reverse: 5′-GACGATACTCCTCCT CGTTGGT-3′	5′-6FAM-CATCAGAATCTGTAGGCCGT-MGBNFQ-3′	11–55 fg (50–250 copies)	Yes	^[48]
8	Real-time PCR	N3R	Forward: 5′-AACAACCGT CCTACA ATTAAA CAACA-3′ Reverse: 5′-CGCTATCGAACCATT TTTGTAGTCT-3′	5′-6FAM-TAT AAC GGC GAA GAA TAT ACT-MGBNFQ-3′	11–55 fg (50–250 copies)	Rodents	^[48]
9	Real-time PCR	B7R	Forward: 5′-ACGTGTTAAACAATGGGTGATG-3′ Reverse: 5′-AACATTTCCATGAATCGTAGTCC-3′	5′-TAMRA-TGAATGAATGCGATACTGTATGTGTGGG-3′-BHQ2	50 copies per reaction	Yes	^[49]
10	C-LAMP	D14L	FIP-C: 5′-TGGGAGCATTGTAACTTATAGTTGCCCTCCTGAACACATGACA-3′ F3-C: 5′-TGGGTGGATTGGACCATT-3′ BIP-C: 5′-ATCCTCGTATCCGTTATGTCTTCCCACCTATTTGCGAATCTGTT-3′ B3-C: 5′-ATGGTATGGAATCCTGAGG-3′ LOOP-F-C: 5′-GATATTCGTTGATTGGTAACTCTGG-3′ LOOP-C-C: 5′-GTTGGATATAGATGGAGGTGATTGG-3′	N/A	10^2.4 copies per reaction	Yes	^[42]
11	C-LAMP	ATI	FIP-W: 5′-CCGTTACCGTTTTTACAATCGTTAATCAATGCTGATATGGAAAAGAGA-3′ F3-W: 5′-TACAGTTGAACGACTGCG-3′ BIP-W: 5′-ATAGGCTAAAGACTAGAATCAGGGATTCTGATTCATCCTTTGAGAAG-3′ B3-W: 5′-AGTTCAGTTTTATATGCCGAAT-3′ LOOP-F-W: 5′-GATGTCTATCAAGATCCATGATTCT-3′ LOOP-C-W: 5′-TCTTGAACGATCGCTAGAGA-3′	N/A	10³ copies per reaction	Yes	^[42]
12	RPA	G2R	Forward: 5′-AATAAACGGAAGAGATATAGCACCACATGCAC-3′ Reverse: 5′-GTGAGATGTAAAGGTATCCGAACCACACG-3′	5′-ACAGAAGCCGTAATCTATGTTGTCTATCGQTFCCTCCGGGAACTTA-3′	16 DNA molecules/μL	Yes	^[46]

This entry is adapted from the peer-reviewed paper 10.3390/bioengineering9100571

References

Von Magnus, P.; Andersen, E.K.; Petersen, K.B.; Birch-Andersen, A. A pox-like disease in cynomolgus monkeys. Acta Pathol. Microbiol. Scand. 2009, 46, 156–176.
Rondle, C.J.; Sayeed, K.A. Studies on monkeypox virus. Bull World Health Organ 1972, 46, 577–583.
Prier, J.E.; Sauer, R.M. A pox disease of monkeys. Ann. N. Y. Acad. Sci. 1960, 85, 951–959.
Zhu, H.; Fohlerová, Z.; Pekárek, J.; Basova, E.; Neužil, P. Recent advances in lab-on-a-chip technologies for viral diagnosis. Biosens. Bioelectron. 2020, 153, 112041.
Artika, I.M.; Ma’Roef, C.N. Laboratory biosafety for handling emerging viruses. Asian Pac. J. Trop. Biomed. 2017, 7, 483–491.
Jayakeerthi, R.S.; Potula, R.V.; Srinivasan, S.; Badrinath, S. Shell vial culture assay for the rapid diagnosis of Japanese encephalitis, west Nile and dengue-2 viral encephalitis. Virol. J. 2006, 3, 2.
Battineni, G.; Chintalapudi, N.; Amenta, F. AI chatbot design during an epidemic like the novel coronavirus. Healthcare 2020, 8, 20154.
Ahsan, M.M.; Uddin, M.R.; Luna, S.A. Monkeypox image data collection. arXiv 2022, arXiv:2206.01774.
Ahsan, M.M.; Uddin, M.R.; Farjana, M.; Sakib, A.N.; Momin, K.; Luna, S.A. Image data collection and implementation of deep learning-based model in detecting monkeypox disease using modified VGG16. arXiv 2022, arXiv:2206.01862.
Portela-Dias, J.; Sereno, S.; Falcão-Reis, I.; Rasteiro, C. Female monkeypox infection with localized genital lesions. Am. J. Obstet. Gynecol. 2022.
Mailhe, M.; Beaumont, A.-L.; Thy, M.; Le Pluart, D.; Perrineau, S.; Houhou-Fidouh, N.; Deconinck, L.; Bertin, C.; Ferré, V.M.; Cortier, M.; et al. Clinical characteristics of ambulatory and hospitalized patients with monkeypox virus infection: An observational cohort study. Clin. Microbiol. Infect. 2022.
Hierholzer, J.C.; Suggs, M.T.; Hall, E.C. Standardized viral hemagglutination and hemagglutination-inhibition tests II. Description and statistical evaluation. Appl. Microbiol. 1969, 18, 824–833.
Sadeghi, P.; Sohrabi, H.; Hejazi, M.; Jahanban-Esfahlan, A.; Baradaran, B.; Tohidast, M.; Majidi, M.R.; Mokhtarzadeh, A.; Tavangar, S.M.; de la Guardia, M. Lateral flow assays (LFA) as an alternative medical diagnosis method for detection of virus species: The intertwine of nanotechnology with sensing strategies. TrAC Trends Anal. Chem. 2021, 145, 116460.
Townsend, M.B.; MacNeil, A.; Reynolds, M.G.; Hughes, C.M.; Olson, V.A.; Damon, I.K.; Karem, K.L. Evaluation of the tetracore orthopox biothreat® antigen detection assay using laboratory grown orthopoxviruses and rash illness clinical specimens. J. Virol. Methods 2013, 187, 37–42.
Stern, D.; Pauly, D.; Zydek, M.; Miller, L.; Piesker, J.; Laue, M.; Lisdat, F.; Dorner, M.B.; Dorner, B.G.; Nitsche, A. Development of a genus-specific antigen capture ELISA for orthopoxviruses—Target selection and optimized screening. PLoS ONE 2016, 11, e0150110.
Ulaeto, D.O.; Lonsdale, S.G.; Laidlaw, S.M.; Clark, G.C.; Horby, P.; Carroll, M.W. A prototype lateral flow assay for detection of orthopoxviruses. Lancet Infect. Dis. 2022, 22, 1279–1280.
Hughes, L.; Wilkins, K.; Goldsmith, C.S.; Smith, S.; Hudson, P.; Patel, N.; Karem, K.; Damon, I.; Li, Y.; Olson, V.A.; et al. A rapid orthopoxvirus purification protocol suitable for high-containment laboratories. J. Virol. Methods 2017, 243, 68–73.
Richert-Pöggeler, K.R.; Franzke, K.; Hipp, K.; Kleespies, R.G. Electron microscopy methods for virus diagnosis and high resolution analysis of viruses. Front. Microbiol. 2019, 9, 3255.
Lavazza, A.; Tittarelli, C.; Cerioli, M. The use of convalescent sera in immune-electron microscopy to detect non-suspected/new viral agents. Viruses 2015, 7, 2683–2703.
Gelderblom, H.R.; Madeley, D. Rapid viral diagnosis of orthopoxviruses by electron microscopy: Optional or a must? Viruses 2018, 10, 142.
Dumont, C.; Irenge, L.M.; Magazani, E.K.; Garin, D.; Muyembe, J.-J.T.; Bentahir, M.; Gala, J.-L. Simple technique for in field samples collection in the cases of skin rash illness and subsequent PCR detection of orthopoxviruses and varicella zoster virus. PLoS ONE 2014, 9, e96930.
NCBI Monkeypox Genome Sequences. Available online: https://www.ncbi.nlm.nih.gov/labs/virus/vssi/#/virus?SeqType_s=Nucleotide&VirusLineage_ss=Monkeypoxvirus,taxid:10244&CreateDate_dt=2022-04-01T00:00:00.00ZTO2022-12-31T23:59:59.00Z (accessed on 9 August 2022).
WHO Laboratory Testing for the Monkeypox Virus. Available online: https://www.who.int/publications/i/item/WHO-MPX-laboratory-2022.1 (accessed on 16 June 2022).
Saxena, S.K.; Ansari, S.; Maurya, V.K.; Kumar, S.; Jain, A.; Paweska, J.T.; Tripathi, A.K.; Abdel-Moneim, A.S. Re-emerging human monkeypox: A major public-health debacle. J. Med. Virol. 2022.
Ropp, S.L.; Jin, Q.; Knight, J.C.; Massung, R.F.; Esposito, J.J. PCR strategy for identification and differentiation of small pox and other orthopoxviruses. J. Clin. Microbiol. 1995, 33, 2069–2076.
Meyer, H.; Ropp, S.L.; Esposito, J.J. Gene for a-type inclusion body protein is useful for a polymerase chain reaction assay to differentiate orthopoxviruses. J. Virol. Methods 1997, 64, 217–221.
Meyer, H.; Damon, I.K.; Esposito, J.J. Orthopoxvirus diagnostics. In Vaccinia Virus and Poxvirology; Humana Press: Totowa, NJ, USA, 2004; pp. 119–133.
Neubauer, H.; Reischl, U.; Ropp, S.; Esposito, J.; Wolf, H.; Meyer, H. Specific detection of monkeypox virus by polymerase chain reaction. J. Virol. Methods 1998, 74, 201–207.
Li, Y.; Olson, V.A.; Laue, T.; Laker, M.T.; Damon, I.K. Detection of monkeypox virus with real-time PCR assays. J. Clin. Virol. 2006, 36, 194–203.
Afonina, I.; Reed, M.; Lusby, E.; Shishkina, I.; Belousov, Y. Minor groove binder-conjugated DNA probes for quantitative DNA detection by hybridization-triggered fluorescence. BioTechniques 2002, 32, 940–949.
Gelaye, E.; Mach, L.; Kolodziejek, J.; Grabherr, R.; Loitsch, A.; Achenbach, J.E.; Nowotny, N.; Diallo, A.; Lamien, C.E. A novel HRM assay for the simultaneous detection and differentiation of eight poxviruses of medical and veterinary importance. Sci. Rep. 2017, 7, srep42892.
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, 1207–1213.
Olson, V.A.; Laue, T.; Laker, M.T.; Babkin, I.V.; Drosten, C.; Shchelkunov, S.; Niedrig, M.; Damon, I.K.; Meyer, H. Real-time PCR system for detection of orthopoxviruses and simultaneous identification of smallpox virus. J. Clin. Microbiol. 2004, 42, 1940–1946.
Simpson, K.; Heymann, D.; Brown, C.S.; Edmunds, W.J.; Elsgaard, J.; Fine, P.; Hochrein, H.; Hoff, N.A.; Green, A.; Ihekweazu, C.; et al. Human monkeypox—After 40 years, an unintended consequence of smallpox eradication. Vaccine 2020, 38, 5077–5081.
Hughes, C.M.; Liu, L.; Davidson, W.B.; Radford, K.W.; Wilkins, K.; Monroe, B.; Metcalfe, M.G.; Likafi, T.; Lushima, R.S.; Kabamba, J.; et al. A tale of two viruses: Coinfections of monkeypox and varicella zoster virus in the Democratic Republic of Congo. Am. J. Trop. Med. Hyg. 2021, 104, 604–611.
Luciani, L.; Inchauste, L.; Ferraris, O.; Charrel, R.; Nougairède, A.; Piorkowski, G.; Peyrefitte, C.; Bertagnoli, S.; de Lamballerie, X.; Priet, S. A novel and sensitive real-time PCR system for universal detection of poxviruses. Sci. Rep. 2021, 11, 1798.
Maksyutov, R.A.; Gavrilova, E.V.; Shchelkunov, S.N. Species-specific differentiation of variola, monkeypox, and varicella-zoster viruses by multiplex real-time PCR assay. J. Virol. Methods 2016, 236, 215–220.
Li, Y.; Meyer, H.; Zhao, H.; Damon, I.K. GC Content-based pan-pox universal PCR assays for poxvirus detection. J. Clin. Microbiol. 2010, 48, 268–276.
Glökler, J.; Lim, T.S.; Ida, J.; Frohme, M. Isothermal amplifications—A comprehensive review on current methods. Crit. Rev. Biochem. Mol. Biol. 2021, 56, 543–586.
Becherer, L.; Borst, N.; Bakheit, M.; Frischmann, S.; Zengerle, R.; von Stetten, F. Loop-mediated isothermal amplification (LAMP)—Review and classification of methods for sequence-specific detection. Anal. Methods 2020, 12, 717–746.
Nagamine, K.; Hase, T.; Notomi, T. Accelerated reaction by loop-mediated isothermal amplification using loop primers. Mol. Cell Probes 2002, 16, 223–229.
Iizuka, I.; Saijo, M.; Shiota, T.; Ami, Y.; Suzaki, Y.; Nagata, N.; Hasegawa, H.; Sakai, K.; Fukushi, S.; Mizutani, T.; et al. Loop-mediated isothermal amplification-based diagnostic assay for monkeypox virus infections. J. Med. Virol. 2009, 81, 1102–1108.
Bhadra, S.; Ellington, A.D. Portable nucleic acid tests for rapid detection of monkeypox virus. medRxiv 2022.
Lobato, I.M.; O’Sullivan, C.K. Recombinase polymerase amplification: Basics, applications and recent advances. TrAC Trends Anal. Chem. 2017, 98, 19–35.
Fan, X.; Li, L.; Zhao, Y.; Liu, Y.; Liu, C.; Wang, Q.; Dong, Y.; Wang, S.; Chi, T.; Song, F.; et al. Clinical validation of two recombinase-based isothermal amplification assays (RPA/RAA) for the rapid detection of African swine fever virus. Front. Microbiol. 2020, 11, 1696.
Davi, S.D.; Kissenkötter, J.; Faye, M.; Böhlken-Fascher, S.; Stahl-Hennig, C.; Faye, O.; Faye, O.; Sall, A.A.; Weidmann, M.; Ademowo, O.G.; et al. Recombinase polymerase amplification assay for rapid detection of monkeypox virus. Diagn. Microbiol. Infect. Dis. 2019, 95, 41–45.
Li, Y.; Zhao, H.; Wilkins, K.; Hughes, C.; Damon, I.K. Real-time PCR assays for the specific detection of monkeypox virus West African and Congo Basin strain DNA. J. Virol. Methods 2010, 169, 223–227.
Kulesh, D.A.; Loveless, B.M.; Norwood, D.; Garrison, J.; Whitehouse, C.A.; Hartmann, C.; Mucker, E.; Miller, D.; Wasieloski, L.P.; Huggins, J.; et al. Monkeypox virus detection in rodents using real-time 3′-minor groove binder TaqMan® assays on the roche light cycler. Lab. Investig. 2004, 84, 1200–1208.
Shchelkunov, S.N.; Shcherbakov, D.N.; Maksyutov, R.A.; Gavrilova, E.V. Species-specific identification of variola, monkeypox, cowpox, and vaccinia viruses by multiplex real-time PCR assay. J. Virol. Methods 2011, 175, 163–169.

© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.