Review of virological methods for laboratory diagnosis and characteri- zation of monkeypox virus (MPXV): lessons learned from the 2022 Mpox outbreak Katarina Resman Rus 1 , Samo Zakotnik 1 , Martin Sagadin 1 , Marko Kolenc 1 , Lucijan Skubic 1 , Nataša Knap 1 , Misa Korva 1 , Mario Poljak 1 , Tatjana Avšič-Županc 1 ✉ 1 Institute of Microbiology and Immunology, Faculty of Medicine, University of Ljubljana, Ljubljana, Slovenia. 23 2024;33:23-35 doi: 10.15570/actaapa.2024.1 Introduction Monkeypox virus (MPXV) belongs to the genus of Orthopoxvirus- es (OPV) within the family Poxviridae and is the etiological agent of the human disease known as Mpox. In the past, MPXV was en- demic in certain regions in West and Central Africa (1). However, sporadic outbreaks in non-endemic countries such as the United States in 2003, the United Kingdom in 2018, Israel in 2018, and Singapore in 2019 have been linked to recent travel to endemic areas or contact with infected animals (2–5). The current Mpox outbreak, characterized by high human-to-human transmis- sion, has spread in several non-endemic countries in Europe and North America since May 2022 (6). The first cases were detected in the United Kingdom, prompting the World Health Organization (WHO) to declare it a Public Health Emergency of International Concern (PHEIC) in July 2023 due to the rapid increase of cases worldwide (7). As of December 14th, 2023, Mpox cases have been reported in 116 countries, with 91,788 confirmed cases and 167 deaths (8). The MPXV genome is a double-stranded DNA molecule of about 197 kb. Sequences are divided into two main clades in en- demic regions: Clade I (formerly the Central African clade), which includes strains from the Democratic Republic of the Congo, and Clade II (formerly the West African clade) (9). Clade II is further divided into two subclades: Subclade IIa, which includes West Af- rican strains, and Subclade IIb, which contains only genomes as- sociated with the 2022 outbreak. Genomic surveillance conducted throughout the Mpox outbreak indicates that the MPXV sequenc- es from 2022 form a cohesive group but are phylogenetically dis- tinct from the Nigerian strain, leading to their placement in the separate Subclade B.1 (10, 11). To date, several thousand MPXV genome sequences have been deposited in public sequence re- positories, and they have formed several B.1 lineages with unique mutational and evolutionary characteristics (11, 12). The significant increase in Mpox cases in non-endemic coun- tries recorded during the 2022 outbreak, the newly identified transmission route, and constant concerns related to potential resurgence of smallpox (the only officially eradicated human dis- ease) have reinforced the need for rapid and accurate MPXV diag- nostics. This review presents and critically discusses the range of virological methods for laboratory diagnosis and characterization of MPXV as well as related lessons learned from the 2022 Mpox outbreak. In addition to the standard-of-care molecular methods and some innovative molecular approaches (such as digital PCR), the practical experience gained in 2022/2023 with virus isolation in cell culture, detection of MPXV by electron microscopy, and monitoring of the serologic response after MPXV infection and smallpox vaccination are summarized. Abstract Monkeypox virus (MPXV), originally endemic in West Africa (Clade II) and Central Africa (Clade I), has recently emerged worldwide and has reinforced the need for rapid and accurate MPXV diagnostics. This review presents and critically discusses the range of virological methods for laboratory diagnosis and characterization of MPXV as well as related lessons learned and practical experi- ence gained from the 2022 Mpox global outbreak. Real-time PCR is currently considered the diagnostic gold standard and ensures accurate and timely confirmation of suspected Mpox cases based on suspicious skin lesions, and digital PCR improves the preci- sion of MPXV DNA quantification. Whole genome sequencing reveals the diversity within the Clade IIb outbreak and highlights the role of microevolution in the adaptation of the virus to the human host. Continuous genomic surveillance is important for better understanding of human-to-human transmission and prevention of the emergence of variola virus–like strains. Traditional virological methods such as electron microscopy and virus isolation remain essential for comprehensive virus characterization, particularly in the context of vaccine and antiviral drug development. Despite the current challenges, serological tests detecting a range of anti-MPXV antibodies are important adjunct diagnostic and research tools for confirmation of late-presenting or asymp- tomatic MPXV cases, contact tracing, epidemiological studies, seroepidemiological surveys, and better understanding of the role of IgG and neutralizing antibodies in the immune response to infection and vaccination. A multidisciplinary approach combining advanced molecular techniques with traditional virological methods is important for rapid and reliable diagnosis, surveillance, and control of the outbreak. Keywords: monkeypox virus, MPXV diagnostics, whole genome sequencing, isolation of MPXV, electron microscopy, MPXV serology Acta Dermatovenerologica Alpina, Pannonica et Adriatica Acta Dermatovenerol APA Received: 8 December 2023 | Returned for modification: 14 December 2023 | Accepted: 22 December 2023 ✉ Corresponding author: tatjana.avsic@mf.uni-lj.si 24 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. Molecular detection of MPXV and MPOX case confirmation Sample collection for MPXV detection The selection of appropriate samples is crucial for reliable diag- nosis of Mpox. Swabs from lesion exudates, roofs, and crusts or tissue biopsies are most recommended for MPXV detection be- cause testing from such samples has the highest clinical sensitiv- ity (from 91% to 100%) (13, 14). It is advisable to collect several skin/mucosal lesions with the same/similar morphology from different body areas in a single tube. Blood samples are not rec- ommended for diagnostic purposes because they may lead to false negative results due to the brief viremia that usually occurs before the appearance of characteristic skin lesions. For research purposes, additional sample types such as saliva, rectal swabs, nasopharyngeal swabs, semen, urine, fecal samples, and vitre- ous or cerebrospinal fluid samples may provide insight into viral shedding and the role of body fluids in transmission, especially in close contact; for example, sexual activity (13, 15–22). As previously reported, all suspected clinical cases of Mpox in Slovenia were tested in the laboratory for the diagnosis of zo- onoses at the Institute of Microbiology and Immunology, Faculty of Medicine, University of Ljubljana. Suspected cases were those with clinical symptoms and an epidemiological link to exposure to MPXV. Swabs of skin lesions or biopsy samples were collected in commercially available universal transport media (UTM) for MPXV DNA testing. A total of 144 different clinical samples from 129 sus- pected cases were molecularly tested by real-time polymerase chain reaction (PCR), and Mpox was confirmed in 49 patients (23). MPXV molecular detection using real-time PCR Confirmation of an Mpox case involves detection of MPXV DNA by either real-time PCR and/or sequencing. Laboratory protocols usually involve two steps: in the first step, OPV is detected, al- though the species cannot be identified. This is followed by the second step, in which MPXV is specifically detected by real-time PCR or sequencing. Alternatively, some laboratory protocols are based on the initial generic detection of MPXV by real-time PCR followed by specific real-time PCR to differentiate between MPXV clades. Although final Mpox case confirmation usually relies on real-time PCR due to its high accuracy and rapid turnaround time, whole genome sequencing provides important additional com- prehensive information on the molecular characteristics and ori- gin of the virus (13, 24–26). Real-time PCR is considered the “gold standard” laboratory method for virological confirmation of MPXV infection (13, 27). Several validated real-time PCR protocols have been developed for the detection of OPV and specifically MPXV, some of which differentiate between Clade I and Clade II. Various primer and probe sequences for OPV and MPXV detection have been pub- lished, allowing suitably equipped and experienced laboratories to develop their own in-house assays (18, 19, 28–32). Since the last emergence of MPXV, several commercial real-time PCR test kits with high sensitivity and specificity are available on the market for the detection of OPV or specifically MPXV . The tests have been validated using well-characterized clinical samples to ensure the quality and robustness of the tests (16, 33–39). Frequently used targets for MPXV molecular diagnostics are the genes D14L (28), G2R (28), B7R (40), F3L (18, 34, 41, 42), B6R (31), and N3R (41), and the tumor necrosis factor (TNF) receptor gene (43, 44). Most assays have a detection limit of 10 to 250 cop- ies per reaction (27). However, a single nucleotide polymorphism (SNP) observed in 2022 MPXV sequences has significantly reduced the sensitivity of some PCR assays. The SNP was located at the binding site of the reverse primer and caused an approximate in- crease in cycle threshold (Ct) value by 6.8 or 100-fold loss in sen- sitivity (45). The first Mpox case in Slovenia was confirmed on May 23rd, 2022, following the algorithm shown in Figure 1. First, OPV was detected using a previously described real-time PCR protocol (29). In the next step, confirmation of MPXV, particularly the genetic Figure 1 | Laboratory algorithm used for confirmation of the first Mpox case in Slovenia. 25 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus lineage of Clade II (former West Africa), was performed using es- tablished generic and West Africa–specific RT-PCR assays (28). The suspected Mpox case was finally confirmed within 3 hours of receipt of the sample at the laboratory. Subsequently, within 24 hours of confirmation, the complete genome sequence was gener- ated using the ONT GridIon instrument and submitted to the Na- tional Center for Biotechnology Information (NCBI). Our diagnos- tic approach is in line with routine practice in the great majority of other European countries (17). In order to establish a specific, rapid, and cost-effective mo- lecular diagnostic approach for the detection of MPXV in clinical samples, we tested seven PCR assays using lesion swabs obtained from nine Slovenian Mpox patients. In addition to the three in- house RT-PCR protocols already mentioned (28, 29), four commer- cial kits were evaluated for confirmation of OPV and/or MPXV: the LightMix Modular Assay targeting OPV and MPXV-specific Light- Mix Modular Assay (Roche, TIB MolBiol, Berlin, Germany), the RealStar OPV PCR kit (Altona Diagnostics, Hamburg, Germany), the and MPXV Real Time PCR Kit (Bioperfectus, Shanghai, China). The LightMix Modular Assay targets a 113 bp fragment of the 14 kDa gene specific for OPVs, and the MPXV-specific LightMix Mod- ular Assay amplifies a 106 bp fragment of the J2L/J2R gene. Both assays have a limit of detection (LOD) of < 10 copies per reaction. RT-PCR reactions were performed according to the manufacturer’s instructions using the TaqMan Fast Virus 1-Step Master Mix (Ther- moFisher Scientific, Waltham, MA, USA). The RealStar OPV PCR kit uses technology for the simultaneous detection and differentiation of non-variola OPV species (includ- ing MPXV) and variola virus (VARV)–specific DNA. Details of LOD and gene detection region(s) are not available. The testing was performed following the manufacturer’s instructions. In the MPXV Real Time PCR Kit, the specific primers and probes target the F3L gene regions of MPXV. The kit also uses amplifi- cation of the human housekeeping gene (RNase P) to control for sample adequacy and efficiency of nucleic acid extraction. The assay’s LOD is five MPXV copies per reaction. The assay was per- formed according to the manufacturer’s instructions. All seven assays listed above were performed using the Quant- Studio 7 Pro Real-Time PCR System (ThermoFisher Scientific). The assays were compared on the basis of the mean Ct value, which showed clear differences when comparatively evaluated on the same set of samples. The in-house OPV/parapoxvirus (PPV) mul- tiplex real-time PCR assay (29) yielded the highest mean Ct value (24.3), significantly higher than the commercial RealStar OPV PCR kit (Ct 15.7; p = 0.005; one-way ANOVA with Tukey’s multiple com- parison test) and the commercial OPV LightMix Modular assay (Ct 17.0; p = 0.019; one-way ANOVA with Tukey’s multiple com- parison test; Fig. 2). For the assays specifically targeting MPXV , no statistically significant performance differences were observed, with similar mean Ct values obtained for the different methods: Ct 19.5 with the in-house MPXV generic test (28), Ct 20.0 with the in- house MPXV WA–specific test (28), Ct 20.9 with the MPXV-specific LightMix Modular Assay, and Ct 19.8 with the MPXV Real Time PCR Kit. During the 2022 outbreak, a combination of the in-house MPXV generic and the MPXV WA—specific assays (28) was used in Slovenia for detection and final confirmation of Mpox cases. In addition, commercially available OPV and MPXV LightMix Modu- lar assays were used when available. All 49 Mpox cases in Slove- nia were confirmed based on positivity of at least two different real-time PCR tests listed above. Digital PCR quantification Digital PCR technology (dPCR) offers advantages for precise quantification and better comparability between different labo- ratories and platforms. In contrast to conventional quantitative PCR (qPCR), dPCR allows absolute quantification of DNA copies in samples without relying on standard curves. The microfluidic design generates numerous reaction partitions that function like individual reactions. Using the Poisson distribution of positive or negative fractions, dPCR determines the absolute target concen- tration in copies per microliter (46–48). During the Mpox 2022 outbreak, dPCR was used for the abso- lute quantification of MPXV in clinical samples (46), to analyze the kinetics of viral DNA in body fluids during the acute phase of Mpox (49), and to quantify virus growth in vitro (50). A recent study demonstrated a strong correlation between MPXV DNA levels measured by dPCR and Ct levels determined by qPCR (51). However, further studies are needed to determine the utility of dPCR for absolute MPXV quantification in clinical samples (46). We successfully quantified MPXV in clinical samples by pre- paring a MPXV standard by dPCR according to the following pro- tocol. Nanoplate-based microfluidic dPCR was performed using the QIAcuity Probe PCR Kit (Qiagen, Hilden, Germany) in a 40 μl reaction mixture containing a 5 µl sample of DNA, 10 µl of 4× Probe PCR Master Mix, 0.9 µM of primers G2R-G_F and G2R-G_R, 0.25 µM of probe G2R-G_P (28), 0.025 U/µl of restriction enzyme Anza 52 PvuII (Thermo Scientific), and nuclease-free water. Five microlit- ers of nuclease-free water were added in place of a DNA template as a non-template control. The reaction mixture was prepared in a pre-plate, transferred to the QIAcuity Nanoplate 26k 24-well plate (Qiagen), sealed with the QIAcuity Nanoplate Seal (Qiagen), and then loaded into the QIAcuity Four automated digital PCR instru- ment (Qiagen) as specified by the manufacturer. The workflow in- cluded 1) a priming and rolling step with partitioning of the wells, 2) a thermocycling step under the following conditions: initial DNA denaturation and enzyme activation for 2 minutes at 95 °C, followed by 45 amplification cycles of 15 s at 95 °C and 30 s at 58 °C, and 3) an imaging step with image acquisition of the wells on the green channel (exposure time: 500 ms, gain: 6 dB). The data were analyzed using QIAcuity Software Suite v2.2.0.26 (Qiagen). To improve the accuracy of the concentration calculation in each well, Volume Precision Factor v4.0 (Qiagen) was applied accord- ing to the manufacturer’s instructions. The MPXV-positive sam- Figure 2 | Comparison of seven different PCR assays/protocols (three in-house and four commercial) for the detection of OPV or specific MPXV. The mean Ct value of each assay was compared with the mean Ct value of each other assay. One-way ANOVA with Tukey’s multiple comparison test was used to test for sig- nificant differences between the assays. Statistically significant differences are indicated by an asterisk; p < 0.019 (*) and < 0.005 (**). 26 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. ple was tested in three consecutive 10-fold dilutions, the meas- urements of which were then used to determine the MPXV DNA concentration in the original sample. Based on such analysis and calculation of the average value of the measurements, the MPXV DNA concentration in the MPXV2.8 sample was determined to be 2.14 × 10 5 copies/µl (Fig. 3 and Table 1). Using the established MPXV standard, we generated standard curves for both the generic MPXV and the MPXV WA–specific in- house assays (28) (Figs. 4A and 4B). The standard curve for the generic MPXV assay was calculated by five 10-fold standard dilu- tions ranging from 8.39 × 10 5 to 8.39 copies/µl (R² = 0.998; efficien- cy 85.366%), represented by the equation y = −1.62ln(x) + 38.932. In addition, the standard curve for the MPXV WA–specific assay was derived from five 10-fold standard dilutions ranging from 7.34 × 10 5 to 7.34 copies/µl (R² = 0.995; efficiency 88.614%), with the equation y = −1.549ln(x) + 39.359. Therefore, we successfully quantified MPXV in clinical samples using both MPXV generic and MPXV WA–specific in-house assays. The results showed no statistically significant difference between the assays (p = 0.1338). The median amount of MPXV detected by the generic MPXV assay was 1.12 × 10 5 copies/µl (min 1.42 × 10 2 and max 5.46 × 106 copies/µl), and the median amount detected by the specific WA assay was 2.21 × 10 5 copies/µl (min 2.21 × 10 2 and max 7.68 × 10 6 copies/µl) (Fig. 5). Whole genome sequencing Whole genome sequencing is not commonly used in clinical di- agnostics due to the high cost of reagents and infrastructure, the time required, and the need for specialized personnel. However, it is an indispensable tool for tracking the evolution of pathogens and transmission patterns during an outbreak, as the SARS-CoV-2 pandemic has shown (52). Sequencing protocols using metagen- Figure 3 | One-dimensional scatter plots of fluorescence partition intensity when testing three consecutive 10-fold dilutions of MPXV-positive sample MPXV2.8 and the non-template control (NTC) with the digital PCR assay for quantification of MPXV DNA. Blue dots represent positive partitions, gray dots negative partitions, and the red horizontal line the threshold for partition positivity. Table 1 | Quantification of MPXV DNA in three consecutive 10-fold dilutions of MPXV-positive sample MPXV2.8 using digital PCR. Sample dilution Dilution factor CI (95%) Valid partitions Positive partitions λ MPXV DNA concentration in (copies/µl) Sample dilution Original sample MPXV2.8 10 −1 10 1.6% 25,465 22,679 2.213 21,740.80 2.17 × 10 5 MPXV2.8 10 −2 100 2.9% 25,385 4,706 0.205 2,124.00 2.12 × 10 5 MPXV2.8 10 −3 1,000 8.8% 25,447 500 0.020 212.16 2.12 × 10 5 Figure 4 | Plotting of the MPXV standard curve using the MPXV generic PCR in- house real-time assay (A) and the MPXV WA–specific PCR in-house real-time assay (Clade II) (B). Five standard dilutions from 10 5 to 10 1 were used to calcu- late the curve. Figure 5 | MPXV copy number / µl in 49 samples from 49 Slovenian Mpox pa- tients measured with the MPXV generic PCR and the MPXV WA (Clade II)–spe- cific in-house real-time PCR assays. The median value is marked in the graph. For statistically significant differences between the groups, the non-parametric Mann–Whitney test was used. No statistically significant difference between the assays (p = 0.1338) was obtained. 27 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus omics or amplicon approaches and next-generation sequencing tools such as Illumina or MinION were used to generate MPXV whole genome sequences from clinical samples (12, 53–58). On- going genomic surveillance of circulating lineages has proven helpful in supporting public health and government decisions to contain MPXV transmission (27). As of December 14th, 2023, a total of 6,501 MPXV IIb lineage whole genome sequences from 52 countries have been collected in the Gisaid database (https:// gisaid.org/hmpxv-variants/; accessed December 14th, 2023). The greatest number of sequences, 3,187 of them, were contributed by the United States, followed by 786 sequences from Germany and 503 from Portugal (https://gisaid.org/hmpxv-variants/; accessed December 14th, 2023). Other platforms such as Nexstrain, mpox- Spectrum, and MPoxVR also provide a global number of upload- ed MPXV sequences and genomic data (59). As outlined previously, all but one Slovenian MPXV detected in clinical samples have been fully sequenced, and the compre- hensive protocols for this process and subsequent phylogenetic analyses have been described in detail previously (23). The Slo- venian sequences are well positioned in Subclade IIb, where all sequences from the 2022 global outbreak are located. Subclade IIb includes lineages A.1, A.1.1, A.2 (further subdivided into three sublineages), A.3, C.1 (https://nextstrain.org/mpox/clade-IIb; ac- cessed December 14th, 2023), and B.1, which is responsible for the Mpox outbreak in Europe in 2022 and subsequent spread to the Americas, Oceania, and Asia (11). The B.1 lineage has further adapted and diversified during the outbreak into 14 sublineages (https://nextstrain.org/mpox/clade-IIb; accessed December 14, 2023). Although it has more mutations than lineage A relative to the presumed ancestor, the data show greater variability within lineage A than within B.1, with all B.1 isolates being closely re- lated (12, 60). The mutation rate detected in MPXV strains belonging to the current outbreak exceeds previous estimates for poxviruses at one to two nucleotides/genome/year, likely due to the activity of the human APOBEC cytosine deaminase enzyme and adaptation of the virus to the human host (12, 61). Microevolutionary events within a short period of time, such as SNPs, gene loss or duplica- tion, and recombination, particularly at the MPXV genomic ends during the outbreak, may contribute to virus evolution, adapta- tion to the human host, and human-to-human transmission (12, 62–67). According to the Nextstrain classification, MPXV Clade-IIb comprises 24 different lineages and sublineages. In various stud- ies on the MPXV strains derived from the 2022 outbreak in Europe, several unrelated introductions into the continent have been identified, suggesting a chain of transmission within a country (23, 68–71). In the European region, lineages B.1, B.1.7, and B.1.10 were found with significant frequency (69). Five different genetic lineages were identified in Slovenia: B.1, B.1.14, B.1.2, B.1.3, and A.2.1. The divergence of MPXV evolution in the European region was calculated to be about 5.70 × 10 –5 substitutions per site per year (69). Monitoring changes in the MPXV genome is crucial to prevent the emergence of an epigone of variola virus. However, such monitoring requires high-quality genomic data and meticu- lous annotation (63). Although whole genome sequencing and subsequent phyloge- netic analysis significantly contributed to viral characterization at the molecular level, electron microscopy and isolation of viable viruses in cell cultures remain essential for complete viral charac- terization, especially in the context of vaccine and antiviral drug development. Electron microscopy After the invention of transmission electron microscopy (TEM) in the 1930s, vaccinia virus (VACV) was one of the first viruses to be visualized by electron microscopy (EM). Continuous advances in EM have played a critical role in elucidating the morphology, size, ultrastructure, and characterization of the viral replication cycle, Figure 6 | Panel A shows VERO E6 cells infected with MPXV under a thin-section electron microscope. The cell nucleus is labelled “Nu+” and the scale corresponds to 5 μm. Panel B shows a closer view of the mature virions within the highlighted area in the blue box of panel A. A lateral body within a maturing virus particle is indicated by the arrow, and the scale is set at 500 nm. Panel C enlarges the area within the red box in panel A and provides higher magnification. Here the viral inclusion bodies, also known as “virus factories”, consist of spherical, immature virus particles (arrowhead). The scale for panel C is 500 nm. Finally, panel D shows brick-shaped virions that have been negatively stained with 2% phosphotungstic acid at a scale of 200 nm. 28 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. including cellular changes induced by MPXV (72–74). In addition to research, EM has also proven to be a valuable diagnostic tool, with the simple negative contrast method being widely used in smallpox diagnostics due to its speed and ability to reliably dis- tinguish between OPV, PPV, and the more common, less severe herpesviruses (72, 75). However, the high cost of purchasing and maintaining EM equipment, combined with the need for very spe- cialized skills of laboratory personnel and the development of highly sensitive, high-throughput molecular assays, have resulted in reduced use of EM for diagnostic purposes (75–78). Various clinical samples, such as vesicle fluids from skin le- sions or solid tissues such as scabs or biopsies, can be examined with EM for the detection of poxviruses. The practical resolution limit of EM for biological material is 2 nm, so that the fine struc- tures of the viral particles are clearly visible. The ultrastructural analysis contributes to a better understanding of the evolution and spread of MPXV (79). OPVs are assembled in the host cyto- plasm in “viral factories” and exist in intermediate compartments such as the immature, mature, and extracellular envelope virions. Mature OPVs generally have an ovoid or brick-shaped morphol- ogy, ranging from 220 to 450 nm in length and 140 to 260 nm in width, making them among the largest known animal viruses. However, OPVs cannot be distinguished morphologically (75, 80). After confirmation of the first Mpox case in Slovenia, the mor- phological characteristics of MPXV were determined and studied by TEM directly on clinical specimens with negative staining and after isolation of the virus in a cell culture by ultra-thin EM (Fig. 6). Negative staining electron microscopy In negative staining EM, also known as negative contrast, an aqueous solution of heavy metal salts is applied to a sample. When the solution dries in the air on the grid, it forms an electron- tight “glass” around the objects on the grid. Biological materials, which have a lower electron density, appear as lighter structures against the darker background of the stain. The process of nega- tive staining involves three steps: adsorption, washing, and stain- ing (75). The clinical samples were centrifuged in UTM viral transport medium for 10 minutes at low speed (2,000 g) to remove larger particles and cell debris. Formvar-coated grids were exposed to a drop of UTM medium for 5 minutes and then negatively stained with 2% phosphotungstic acid. In addition, the UTM medium was ultracentrifuged (Airfuge; Beckman Coulter, Indianapolis, IN, USA) at high speed (100,000 g) to concentrate the viruses directly on the grid, and negative staining was performed using the same procedure. Examination of the grids was performed using a TEM (JEM-1400 Plus; JEOL, Tokyo, Japan) at 120 kV. The OPVs were identified based on their characteristic brick-shaped morphology and a size of approximately 350 × 270 nm (Fig. 6D). Ultrathin section electron microscopy After successful MPXV isolation in cell culture (see details below), ultrathin sections of MPXV-infected VERO E6 cells were prepared. Two days after inoculation, the cells were fixed with a modified Karnovsky fixative. Fixation was followed by post-fixation in 1% (w/v) osmium tetroxide for 1 h. Samples were afterwards dehy- drated in a graded series of ethanol and embedded in Epon (Serva Electrophoresis, Heidelberg, Germany). Ultrathin sections (60–80 nm) were cut with an ultramicrotome from the resin block em- bedded with samples and contrasted with uranyl acetate and lead citrate using an automatic contrasting system (EM ACE 20; Leica, Wetzlar, Germany). Finally, the grids and stained sections were examined with JEM-1400 Plus TEM at 120 kV . The EM showed both mature and immature virus particles, indicating virus replication in the cytoplasm of the infected cell. Viral inclusion bodies (“viral factories”), composed of round, spherical, and less dense immature virus particles, were visible inside the cytoplasm next to mature poxvirus particles with a typical oval profile and dumbbell core (Fig. 6A, 6B, 6C). Our EM results are in accordance with other published Mpox cases, which have demonstrated the importance of TEM for study of replica- tion and pathogenesis as well as characterization of this emerging pathogen (4, 42, 74, 81–84). Virus isolation MPXV was originally isolated in cell culture in 1958 from pus- tules of cynomolgus monkeys showing non-lethal smallpox-like symptoms (85), and it was later detected in infected humans in the Democratic Republic of the Congo in 1970 (86). Initially, viral culture was considered the MPXV diagnostic gold standard and confirmed the Mpox diagnosis in 67% of suspected cases after the eradication of smallpox in the Democratic Republic of the Congo in the 1980s (87). Lately, molecular assays targeting viral DNA have become the most reliable MPXV diagnostic method due to their rapid turnaround time and higher sensitivity. MPXV isolation: practical considerations Only highly trained personnel should perform in vitro testing with viable MPXV using full personal protection in biosafety level 3 (BSL-3) laboratory facilities. Several mammalian cell lines (VERO, VERO E6, VERO 76, LLC-MK2, BSC-40, BSC-1, HeLa, PEK, HEP-2, A549, MRC-5) have been shown to be susceptible to MPXV infec- tion and support its replication (14, 88–90). The ability of poxvi- ruses to infect cells without specific receptors contributes to this broad susceptibility of cell lines (91–93). However, VERO cells are particularly useful because MPXV replicates in these cell lines with a higher titer compared to HEP-2 and PEK cells (86). Clinical samples with low Ct values when tested with PCR and thus with anticipated high virus concentration (plaque-forming units [PFU]/ml)—for example, from skin lesions—are most appro- priate for virus isolation (14). Viable MPXV has been efficiently isolated from a range of clinical specimens, including semen samples collected 6 days after symptom onset and from anal and urethral swabs (89, 94). Successful virus growth in cell culture is most frequently associated with samples with PCR Ct values up to 30 (14, 88, 95, 96). However, the Ct values can vary significantly between different laboratories and different PCR assays (Fig. 2), thus no single Ct value can be used as an eligibility cut-off for vi- ral culture. Extensive cytopathic effects (CPE) usually occur 1 to 4 days after inoculation, with unique CPE patterns observed in specific cells, often with widespread cellular detachment and de- generation (88). Upon passage of the virus on VERO cultures, CPE is observed as infectious foci (96). During the 2022 Mpox outbreak, samples from six PCR-con- firmed Slovenian Mpox patients (swabs of vesicle fluid from four patients and biopsy samples from two patients) were inoculated into the VERO E6 cell line in the BSL-3 laboratory at our facili- ty. The LightMix modular MPXV PCR Ct values ranged from 16.9 29 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus to 30.4. For each patient, 500 µl of the samples was inoculated directly into the VERO E6 cell growth medium (ATCC-LGC; CRL- 1586) consisting of Dulbecco’s modified eagle medium (DMEM; Gibco, Thermo Fisher Scientific) with 5% fetal bovine serum (FBS; Euroclone, Pero, Italy) and antibiotics (penicillin, streptomycin, amphotericin B). The inoculated cells were incubated in a T25 flask (TPP , Trasadingen, Switzerland) at 37 °C and 5% CO 2 . The in- oculated cells were monitored daily for CPE, and an intense pox- virus-specific effect was observed on the 2nd day (Fig. 7). Isolation of viable MPXV was successful in five out of six samples tested, with the exception of a sample with a PCR Ct value of 30.4. In ad- dition, when we intended to sequence the whole MPXV genome from this sample, we were only able to cover 40% of the genome, suggesting that the MPXV DNA was substantially degraded in this sample and the virus was most probably not infectious (23). Although virus isolation is not recommended or routinely per- formed in MPXV diagnostics, it is essential for the study of viral pathogenesis and for research on future diagnostic methods, anti- viral drugs, and vaccines. Determination of a patient’s infectivity is an important adjunct tool in infection control, patient isolation protocols, public health guidelines, and cases of prolonged viral shedding (88). Viral cultures remain essential for the detection of rare and/or emerging viral pathogens, especially in samples with presumptive high viral concentrations (95). MPXV inactivation Ensuring safe laboratory conditions when handling highly path- ogenic viruses, such as OPVs, requires appropriate and reliable virus inactivation procedures. Heat treatment is a widely used method due to its ability to denature the secondary structures of viruses. Although literature data on inactivation methods were previously only available for VACV and VARV viruses among the OPVs, recent research by Batejat et al. in 2022 on MPXV recom- mends a 30-minute treatment at 60 °C for the Clade II and Clade I genetic lineages using viral transport media (VTM) and FBS (97). Further evaluation of heat treatment under different conditions (30 minutes at 65 °C, 15 minutes at 65 °C, or 15 minutes at 95 °C) by Fischer in 2022 revealed no plaque formation after such a pro- cedure, providing viable protocols for MPXV inactivation (98). Of note, heat treatment at 60, 70, or 95 °C only minimally affected the detection of MPXV DNA by qPCR, indicating that inactivated sam- ples are suitable for molecular diagnostics without a significant decrease in Ct values (97). In addition, evaluations of commonly used reagents in clinical laboratories such as Trizol and commercially available lysis buffer AVL with ethanol were performed, showing inactivation of MPXV after a 10-minute incubation at room temperature (98). A study testing seven commercial diagnostic buffers confirmed inactiva- tion of MPXV at different incubation times, and heat (56 °C and 60 °C) and sodium dodecyl sulfate detergent were also found to be effective, highlighting the importance of matching the protocol to downstream biological processes (99). As a result of the initial absence of literature information on MPXV inactivation, an in-house viral inactivation protocol was developed at our institution. A 500 μl MPXV sample suspended in cell culture medium was heat inactivated at 60 °C for 1 hour. Sub- sequently, three passages of the inactivated virus were performed on VERO E6 cells, where no CPE was observed and the Ct values of an MPXV generic assay (28) performed on all passages increased from 15.8 to 32.6 after three passages, supporting the effectiveness of the heat inactivation protocol developed for the subsequent MPXV PCR (Fig. 8). Prolonged stability of MPXV DNA is crucial for shipping sam- ples between laboratories to be used as positive controls (espe- cially at outbreak onset) and for referral purposes. Room-temper- ature shipment substantially simplifies transport logistics and reduces associated costs, facilitating the establishment of a global Figure 7 | Isolation of viable MPXV in cell culture. Onset of cytopathic effect on the VERO E6 cell line 1, 2, and 3 days after inoculation (d.p.i.) of a swab sample from an Mpox patient compared to the negative control (NC). The images were acquired with the EVOS FL Imaging System (4×; Life Technologies), and the scale is 1,000 nm. 30 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. network of laboratories capable of performing molecular testing. Thus, after virus inactivation, stability of isolated DNA was moni- tored in our laboratory through 5 days of storage at room tempera- ture by daily aliquot testing using the MPXV generic assay (28). The results showed that the MPXV DNA was stable over 5 days of storage with no significant drop in Ct values observed, even when the DNA aliquot was exposed to temperatures above 30 °C for sev- eral hours on day 3. This stability is consistent with the results of other studies (97, 99). In addition, the DNA stability after MPXV inactivation using an autoclave (121 °C, 30 min) was investigated. Despite successful virus inactivation (no CPE on VERO E6 cells were observed), viral DNA was still detectable by MPXV generic assay (28) without a significant decrease in Ct values (isolated vi- rus Ct value: 14.2; autoclaved virus Ct value: 15.0). Detection and quantification of MPXV viability and infectivity Although qPCR can detect viral DNA quickly and accurately and provide an estimate of its concentration, it cannot detect and quantify viral viability and infectivity. This limitation is particu- larly notable in the case of MPXV because it is a very stable DNA virus. Instead, the TCID 50 assay (median tissue culture infectious dose) and the plaque assay are used to quantify infectious viruses (Fig. 9). The plaque assay is considered the gold standard for the quan- tification of replication-capable lytic virions. In this method, the virus is diluted and 10-fold dilutions applied to susceptible cells. After a certain incubation period, the liquid medium is replaced by a solid or semi-solid medium to prevent the virus from spread- ing, resulting in visible plaques within the cell monolayer. The vi- rus titer, expressed as PFU, is determined by staining and count- ing the plaques (100). In contrast, the TCID50 assay determines the point at which 50% of the cultured cells are infected and thus provides an ap- proximate virus titer. In contrast to the plaque assay, in the TCID50 assay the viral inoculum is mixed with a cell medium and inoculated into the cells, which are usually seeded on 96-well plates. The wells are observed and analyzed for the presence or absence of CPE, and the virus dilution at which 50% of the wells show infection is calculated. TCID50 titers are determined using the Reed–Muench or Spearman–Kärber methods (100–102). MPXV plaque assay protocol. During the 2022 Mpox out- break, a plaque assay was developed in our laboratory to quan- tify the infectious virus titer of MPXV isolates on cell cultures. Although MPXV forms plaques on VERO E6 cell monolayers, an overlay medium with agarose was used for the assay. VERO E6 cells were seeded overnight in DMEM cell growth medium sup- plemented with 10% FBS at a concentration of 1 × 10 5 cells / 500 μl / well on a 24-well plate. The next day, the cells were washed with 2× Medium 199 (Gibco, Thermo Fisher Scientific), and 200 µl of a 10-fold serial dilution of the virus in 2× Medium 199 was inocu- lated into the cells in four replicates. Uninfected wells served as negative controls. After 1 hour of incubation at 37 °C and 5% CO 2 , the inoculum was removed and the wells were covered with 500 µl of overlay medium. The overlay medium consisted of supple- mented medium (2× Medium 199 with 6% NaHCO₃ and 4% FBS) and 2.5% CMC agarose (Sigma M0512-100G methylcellulose) at a ratio of 1:1. The plates were then incubated at 37 °C and 5% CO 2 for 5 days. The infected cells were fixed with a 4% formaldehyde solu- tion for 30 minutes. The formaldehyde and overlay medium were removed prior to staining. Plaque formation was visualized with a 1% crystal violet solution. PFU per ml were calculated based on the number of plaques multiplied by the dilution factor. MPXV isolated on VERO E6 cells in Slovenia during the 2022 outbreak reached a mean titer of 3 × 10 7 PFU/ml (between 6.9 × 10 4 and 1.6 × 10 8 PFU/ml) after three passages on cells (Fig. 9B). MPXV TCID 50 method protocol. During the 2022 Mpox out- break, we also implemented the TCID50 method for MPXV quanti- fication because it is faster and easier to perform than the plaque Figure 8 | Ct values measured using MPXV generic PCR after heat inactivation of an MPXV sample suspended in cell culture medium at 60 °C for 1 hour (total virus) and after three passages (p1, p2, and p3) on the VERO E6 cell line. Figure 9 | Quantification of viable MPXV using the TCID 50 method and plaque assay. In the TCID 50 method (Panel A), 10-fold dilutions of the virus in DMEM medium from 10 −1 to 10 −10 were inoculated into columns 1 to 10 of 96-well plates. Columns 11 and 12 served as negative controls (NC), where only DMEM medium without virus was added. For the plaque assay (Panel B), 10-fold virus dilutions were inoculated, from 10 −2 (column 1) to 10 −6 dilutions (up to column 5). Column 6 serves as a negative control (NC), in which only virus-free medium is added to the cells. The TCID 50 and plaque assay plates are stained with crystal violet. 31 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus assay. This method closely follows our previously established pro- tocol for SARS-CoV-2 quantification (103). Tenfold dilutions of the virus in DMEM containing 10% FBS were inoculated onto 24-hour- old VERO E6 cells seeded on a 96-well plate at a concentration of 1 × 10 5 cells / 100 μl / well in eight replicates. The plates were incu- bated at 37 °C and 5% CO 2 for 3 days. At the end of incubation, the cells were fixed with a 4% formaldehyde solution for 20 minutes and stained with a 1% crystal violet solution. Wells in which CPE were observed in at least 50% of the cells per well were identified as MPXV positive. Virus concentration was calculated using the Spearman–Kärber algorithm (102). Slovenian MPXV 2022 isolates on VERO E6 were quantified with a median of 4 × 10 8 TCID 50/ml (between 1 × 10 6 and 2 × 10 9 TCID 50/ml; Fig. 9A). Serological assays Serologic methods measuring immune response against MPXV are not vital for diagnostic confirmation of individual Mpox cases, but they are essential for assessing population seroprevalence and immunity, and for conducting epidemiologic and vaccine- related studies. In the past, the lack of commercial anti-MPXV se- rologic tests was due to the high cross-reactivity between OPVs as a result of > 90% similarity at the amino acid level between MPXV and VARV (40). Consequently, there are no commercially availa- ble serologic tests that could reliably distinguish MPXV from other OPVs such as VACV, VARV, and cowpox virus (CPXV) (104). As a result, such tests may not have been useful in OPV-endemic areas such as Europe (CPXV) (105, 106) and Africa (MPXV) (107, 108). In the traditional endemic regions of MPXV , only limited serological surveillance data are still available (109). Despite these challenges, various in-house immunoassays— immunofluorescence assay (IFA) and enzyme-linked immuno- sorbent assay (ELISA)—have been developed for surveillance, seroprevalence studies, and differentiation between vaccination- induced and naturally acquired immunity (14, 17, 20, 110—112). Following the global Mpox 2022 outbreak, a range of innovative serological methods based on different platforms and technolo- gies were developed, such as the multi-antigen ELISA, which in- cluded 27 poxvirus antigens (24 MPXV and three VACV antigens) (111), a chemiluminescence-based serological assay (113), a robust peptide-based IgG ELISA suitable for serosurveys in endemic ar- eas (104), and a multiplex bead-based microsphere immunoassay with high throughput and high sensitivity (93%) for the detection of anti-MPXV IgG antibodies using a combination of MPXV pep- tides and OPV protein (114, 115). A study conducted during the 2003 Mpox outbreak in the United States provided important insights into the timing of the immune response against MPXV. An IgM response was detected in serum collected more than 4 days after infection, and IgG re- sponse was observed in serum collected more than 14 days after MPXV infection. Of note, previously unvaccinated Mpox cases showed a more rapid and sustained IgM response that lasted up to 18 weeks post-infection, and in individuals previously vaccinated against smallpox the immune response lasted 11 weeks post-in- fection. Conversely, the IgG immune response was long-lasting in previously vaccinated individuals, up to 21 weeks after rash on- set, compared to 19 weeks in previously unvaccinated individuals. This study suggests that MPXV infection induces a strong immune response in the acute phase even in individuals that have been previously vaccinated against smallpox (116). Further serological studies supported these initial results and indicated that IgM an- tibodies can persist for several weeks (peak 2 to 3 weeks after OPV exposure) and IgG antibodies for several years after infection (14, 117–120). Recently, particularly after the COVID-19 pandemic, the im- portance of serological tests for the detection of functional an- tibodies, especially neutralizing antibodies (N-Ab), has become evident. Vaccination with VACV is thought to provide functional cross-protection against MPXV infection, with polyclonal N-Ab playing a crucial role. However, a limited number of studies have investigated the correlation between N-Ab levels and clinical pro- tection (121–124). Traditionally, the detection of N-Ab against OPV has relied on the plaque reduction neutralization test (PRNT). However, recently accumulated evidence suggests that a micro- neutralization assay supplemented with an external complement source increases sensitivity of N-Ab detection (125). Studies dem- onstrated that patients develop a robust immune response pro- ducing IgM, IgG, and N-Abs approximately 2 weeks after the onset of symptoms (112). Of note, N-Ab and IgG titers show a significant negative correlation with the MPXV isolation success rate and a positive correlation with time from symptom onset (112). This suggests that patients are effectively controlling the infection and reducing virus shedding (112). In addition, antibodies that neu- tralize MPXV have also been detected decades after historical smallpox vaccination. However, the exact role of MPXV N-Ab in disease protection and transmissibility is still under investigation (125, 126), especially considering that they are also detected in asymptomatic individuals (127). In-house immunofluorescence assay protocol Although ELISA-based assays have higher sensitivity compared to indirect immunofluorescence assays (IFA), an in-house IFA was initially used in Slovenia to characterize individual Mpox cases (128). After confirmation of the first cases in 2022 and successful isolation of the virus, we soon developed an in-house IFA for de- termination of anti-OPV antibodies in serum samples. In the BSL-3 facility, VERO E6 cells were cultured for 48 hours before inocu- lation with MPXV at a concentration of 10 4 TCID 50 /ml. Five days after inoculation, infected cells were washed twice with saline containing 5% FBS. The cells were removed from the flask with glass beads and the pellet was resuspended in freezing medium consisting of DMEM with 10% FBS and 10% dimethyl sulfoxide (DMSO). The DMSO-treated cells containing MPXV antigen were frozen at −80 °C until further use. To prepare IFT slides, the frozen cells were thawed and centrifuged for 5 minutes at 1,500 rpm and 4 °C. The infected cells were washed twice with PBS and resus- pended in saline containing 5% FBS. For IFT slides, infected cells were mixed with uninfected VERO E6 cells in a 1:3 ratio, and 7 µl of the cell suspension per well was added to the 15-well slides. The slides were fixed in ice-cold acetone for 15 minutes and frozen at −30 °C until further use. For validation of the in-house IFA developed, serum samples were collected from the first three confirmed Mpox Slovenian cases, one sample each per patient: 7, 21, and 40 days after PCR- confirmation, respectively. The smallpox vaccination status of these patients was not available. Serum samples were diluted twofold from 1:16 to 1:1,024 in phosphate-buffered saline (PBS), applied to prepared slides and incubated for 30 minutes at room temperature. After washing, a fluorescein isothiocyanate (FITC)- conjugated anti-human IgG (Vircell, Granada, Spain) was used for detection. After incubation at room temperature for 30 minutes, 32 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. the slides were washed in water and mounted in 3% glycerol/ water. Positive FITC staining was detected in infected cells with a fluorescence microscope at ×20 magnification. The highest IgG antibody titer (1:512) against MPXV was de- tected in the serum collected 21 days after PCR confirmation of the disease. IgG antibodies were also found in the serum collected 40 days after PCR confirmation (1:64), whereas the serum collected 7 days after PCR confirmation was anti-MPXV IgG antibody negative (Fig. 10). These results are consistent with recent studies in which OPV-directed IgG antibodies are typically detected 2 to 4 weeks after the onset of symptoms in titers between 1:40 and 1:320 (17, 20, 110). Neutralization test protocol The neutralization test (NT) is considered the serological gold standard in virology. The test detects N-Abs, which are central players in humoral immunity. N-Abs block the entry of viruses into the host cells and neutralize their biological effect. In NT, vi- rus and patient serum are mixed and inoculated into the cells. The N-Abs present in the serum bind to the free virus and prevent the virus from binding to the receptors of the host cell and infecting the cells (129, 130). During the 2022 Mpox outbreak, the NT was performed at our BSL-3 facility following a previously established procedure with slight modifications (103). In brief, Vero E6 cells were propagated at a density of 10 5 cells per well in a 96-well plate 1 day before the start of the NT assay. Heat-inactivated (60 °C, 30 min) serum samples from Mpox patients were used in a twofold serial dilution starting at 1:10. The diluted samples (100 µl) were then incubated with an equal volume of MPXV (at a concentration of 10 3 TCID 50 / ml). After 1 hour of incubation at 37 °C, 50 µl of each plasma dilu- tion–virus mixture was added to the cells in triplicate and incu- bated at 37 °C and 5% CO 2 for 5 days. After fixing the plates with 4% paraformaldehyde for 30 minutes, crystal violet was used to detect CPE. The neutralizing endpoint titer was determined as the highest dilution causing a 90% reduction in CPE in at least two of three parallels. An NT titer ≥ 1:10 was considered positive. Serum samples from the same patients used for validation of IFT were also tested with NT, and a serum sample from a healthy smallpox-unvaccinated person was included as a negative con- trol. With the NT, relatively low N-Ab titers were detected in two patients, 1:20 and 1:40, in samples collected 21 and 40 days after Mpox confirmation, respectively. Specific N-Abs were absent in the serum sample collected 7 days after Mpox confirmation as well as in the healthy unvaccinated control (Fig. 11). The NT results con- firmed the IFA results and showed that anti-MPXV IgG and N-Abs can be detected at least 2 weeks after Mpox confirmation, which is consistent with previously reported results (112). Conclusions In the last 2 years, MPXV has become a public health problem worldwide. At present, real-time PCR is considered the diagnos- tic gold standard and ensures accurate and timely confirmation of suspected Mpox cases based on suspicious skin lesions, and digital PCR improves the precision of MPXV DNA quantification. Whole genome sequencing reveals the diversity within the Clade IIb outbreak and highlights the role of microevolution in the ad- aptation of the virus to the human host. Traditional virological methods such as EM and virus isolation remain essential for com- prehensive virus characterization. Despite the current challenges, serological tests detecting a range of anti-MPXV antibodies are important adjunct diagnostic and research tools for confirmation of late-presenting or asymptomatic MPXV cases, contact trac- ing, epidemiological studies, and seroepidemiological surveys. A multidisciplinary approach combining advanced molecular tech- Figure 10 | Immunofluorescence assay (IFA) for the detection of IgG antibodies against MPXV in the serum of an MPXV-positive patient. VERO E6 cells infected with MPXV were used as antigen in the IFA. Panel A shows a positive immunofluorescence signal for anti-MPXV IgG antibody detected in a serum sample collected 21 days after PCR-confirmation of the disease, and panel B shows a negative control. The figure shows the immunofluorescence of 1:64 serum dilution. Figure 11 | Neutralization test (NT) for the detection of neutralizing antibodies (NT-Ab) against MPXV in the serum of an Mpox patient. Serum dilutions mixed with MPXV were inoculated in triplicate. A serum sample from an individual that had not previously been vaccinated against smallpox and in whom no OPV infection had previously been detected was used as a negative control. In the positive control (PC) column, only the virus was inoculated without serum, and, in the negative control (NC) columns, only growth medium was added to the cells. NT-Abs were detected in the patient’s serum at a titer of 1:40 (columns marked with a red square), whereby an inhibition of the cytopathic effect was observed in comparison to the unvaccinated control. 33 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus niques with traditional virological methods is important for rapid and reliable diagnosis, surveillance, and control of the outbreak. Acknowledgments We thank Mateja Jelovšek, Kaja Kotnik Koman, Katja Potočnik, Patricija Pozvek, and Jan Slunečko for their excellent laboratory assistance and literature research. Funding The research was funded by the Institute of Microbiology and Im- munology, Faculty of Medicine, University of Ljubljana; by the Slovenian Research and Innovation Agency (grant P3-0083); and by the European Virus Archive–GLOBAL project, which received funding from the European Union’s Horizon 2020 research and in- novation program under grant agreement no. 871029. References 1. Kabuga AI, El Zowalaty ME. A review of the monkeypox virus and a recent out- break of skin rash disease in Nigeria. J Med Virol. 2019;91:533–40. 2. Centers for Disease Control and Prevention. Update: multistate outbreak of mon- keypox—Illinois, Indiana, Kansas, Missouri, Ohio, and Wisconsin, 2003. MMWR Morb Mortal Wkly Rep. 2003;52:642–6. 3. Vaughan A, Aarons E, Astbury J, Balasegaram S, Beadsworth M, Beck CR, et al. Two cases of monkeypox imported to the United Kingdom, September 2018. Euro Surveill. 2018;23:1800509. 4. Erez N, Achdout H, Milrot E, Schwartz Y, Wiener-Well Y, Paran N, et al. Diagnosis of imported monkeypox, Israel, 2018. Emerg Infect Dis. 2019;25:980–3. 5. Ng OT, Lee V, Marimuthu K, Vasoo S, Chan G, Lin RTP, et al. A case of imported monkeypox in Singapore. Lancet Infect Dis. 2019;19:1166. 6. Heskin J, Belfield A, Milne C, Brown N, Walters Y, Scott C, et al. Transmission of monkeypox virus through sexual contact—a novel route of infection. J Infect. 2022;85:334–63. 7. WHO. WHO Director General’s statement following IHR Emergency Committee regarding the multi-country outbreak of monkeypox [Internet]. 2022 Jul 23 [cited 2023 Dec 14]. Available from: https:/ /www.who.int/director-general/speeches/ detail/whodirector-general-s-statement-on-the-press-conference-following- IHRemergency-committee-regarding-the-multi--country-outbreak-ofmonkey- pox--23-july-2022. 8. WHO. 2022–23 Mpox (Monkeypox) outbreak: global trends [Internet]. 2023 Nov [cited 2023 Dec 14]. Available from: https:/ /worldhealthorg.shinyapps.io/ mpx_global/. 9. Nakazawa Y, Mauldin MR, Emerson GL, Reynolds MG, Lash RR, Gao J, et al. A phylogeographic investigation of African monkeypox. Viruses. 2015;7:2168–84. 10. Happi C, Adetifa I, Mbala P , Njouom R, Nakoune E, Happi A, et al. Urgent need for a non-discriminatory and non-stigmatizing nomenclature for monkeypox virus. PLoS Biol. 2022;20:e3001769. 11. Luna N, Munoz M, Bonilla-Aldana DK, Patino LH, Kasminskaya Y, Paniz-Mondolfi A, et al. Monkeypox virus (MPXV) genomics: a mutational and phylogenomic analyses of B.1 lineages. Travel Med Infect Dis. 2023;52:102551. 12. Isidro J, Borges V, Pinto M, Sobral D, Santos JD, Nunes A, et al. Phylogenomic characterization and signs of microevolution in the 2022 multi-country outbreak of monkeypox virus. Nat Med. 2022;28:1569–72. 13. WHO. Diagnostic testing for the monkeypox virus (MPXV). Interim guidance [Internet]. 2023 Nov 9 [cited 2023 Dec 14]. Available from: https://iris.who. int/bitstream/handle/10665/373966/WHO-MPX-Laboratory-2023.1.eng. pdf?sequence=1. 14. Lim CK, Roberts J, Moso M, Liew KC, Taouk ML, Williams E, et al. Mpox diagnos- tics: review of current and emerging technologies. J Med Virol. 2023;95:e28429. 15. Sofi Ibrahim M, Kulesh DA, Saleh SS, Damon IK, Esposito JJ, Schmaljohn AL, et al. Real-time PCR assay to detect smallpox virus. J Clin Microbiol. 2003;41:3835–9. 16. Peiró-Mestres A, Fuertes I, Camprubí-Ferrer D, Marcos MA, Vilella A, Navarro M, et al. Frequent detection of monkeypox virus DNA in saliva, semen, and other clinical samples from 12 patients, Barcelona, Spain, May to June 2022. Euro Sur - veill. 2022;27: 2200503. 17. Noe S, Zange S, Seilmaier M, Antwerpen MH, Fenzl T, Schneider J, et al. Clinical and virological features of first human monkeypox cases in Germany. Infection. 2023;51:265–70. 18. Huo S, Chen Y, Lu R, Zhang Z, Zhang G, Zhao L, et al. Development of two mul- tiplex real-time PCR assays for simultaneous detection and differentiation of monkeypox virus IIa, IIb, and I clades and the B.1 lineage. Biosaf Health. 2022;4: 392–8. 19. Mills MG, Juergens KB, Gov JP , McCormick CJ, Sampoleo R, Kachikis A, et al. Eval- uation and clinical validation of monkeypox (mpox) virus real-time PCR assays. J Clin Virol. 2023;159:105373. 20. Adler H, Gould S, Hine P, Snell LB, Wong W, Houlihan CF, et al. Clinical features and management of human monkeypox: a retrospective observational study in the UK. Lancet Infect Dis. 2022;22:1153–62. 21. Antinori A, Mazzotta V, Vita S, Carletti F, Tacconi D, Lapini LE, et al. Epidemio- logical, clinical and virological characteristics of four cases of monkeypox sup- port transmission through sexual contact, Italy, May 2022. Euro Surveill. 2022; 27:2200421. 22. Algarate S, Bueno J, Crusells MJ, Ara M, Alonso H, Alvarado E, et al. Usefulness of non-skin samples in the PCR diagnosis of Mpox (monkeypox). Viruses. 2023; 15:1107. 23. Resman Rus K, Zakotnik S, Sagadin M, Knap N, Suljic A, Zorec TM, et al. Mo- lecular epidemiology of the 2022 monkeypox virus outbreak in Slovenia. Acta Dermatovenerol Alp Pannonica Adriat. 2023;32:111–7. 24. ECDC. Epidemiological update: Monkeypox multi-country outbreak [Internet]. 2022 May 25 [cited 2023 Dec 14]. Available from: https:/ /www.ecdc.europa.eu/ en/news-events/epidemiological-update-monkeypox-multi-country-outbreak. 25. PAHO. Laboratory guidelines for the detection and diagnosis of monkeypox virus infection [Internet]. 2022 May 23 [cited 2023 Dec 14]. Available from: https:/ /www.paho.org/en/documents/laboratory-guidelines-detection-and- diagnosis-monkeypox-virus-infection-23-may-2022. 26. Nakhaie M, Arefinia N, Charostad J, Bashash D, Haji Abdolvahab M, Zarei M. Mon- keypox virus diagnosis and laboratory testing. Rev Med Virol. 2023;33:e2404. 27. Silva S, Kohl A, Pena L, Pardee K. Clinical and laboratory diagnosis of monkey- pox (mpox): current status and future directions. iScience. 2023;26:106759. 28. Li Y, Zhao H, Wilkins K, Hughes C, Damon IK. Real-time PCR assays for the spe- cific detection of monkeypox virus West African and Congo Basin strain DNA. J Virol Methods. 2010;169:223–7. 29. Schroeder K, Nitsche A. Multicolour, multiplex real-time PCR assay for the detec- tion of human-pathogenic poxviruses. Mol Cell Probes. 2010;24:110–3. 30. Ropp SL, Jin Q, Knight JC, Massung RF, Esposito JJ. PCR strategy for identifica- tion and differentiation of small pox and other orthopoxviruses. J Clin Microbiol. 1995;33:2069–76. 31. Li Y, Olson VA, Laue T, Laker MT, Damon IK. Detection of monkeypox virus with real-time PCR assays. J Clin Virol. 2006;36:194–203. 32. Kasuya F, Negishi A, Kumagai R, Yoshida I, Murakami K, Fujiwara T, et al. Genetic characteristics of the virus detected in the first Mpox imported case in Tokyo, Japan. Jpn J Infect Dis. 2023;76:259–62. 33. Peris MP, Clusa L, Alonso H, Escolar C, Fortuno B, Rezusta A, et al. Clinical per- formance evaluation of a rapid real-time PCR assay for monkeypox diagnosis: a retrospective and comparative study. Microbiol Spectr. 2023;11:e0051023. 34. Michel J, Targosz A, Rinner T, Bourquain D, Brinkmann A, Sacks JA, et al. Evalua- tion of 11 commercially available PCR kits for the detection of monkeypox virus DNA, Berlin, July to September 2022. Euro Surveill. 2022;27:2200816. 35. Velu PD, Sipley J, Marino J, Ghanshani S, Lukose G, Cong L, et al. Evaluation of a zoonotic orthopoxvirus PCR assay for the detection of Mpox virus infection. J Mol Diagn. 2023;25:740–7. 36. Erster O, Levy I, Kabat A, Mannasse B, Levy V, Assraf H, et al. A multi-laboratory evaluation of commercial monkeypox virus molecular tests. Microbiol Spectr. 2023;11:e0022523. 37. Tan NK, Madona CP , Taylor JF, Fourali LH, Sehmi JK, Stone MJ, et al. Performance evaluation of the Viasure PCR assay for the diagnosis of monkeypox: a multicen- tre study. J Clin Virol. 2023;158:105350. 38. Li D, Wilkins K, McCollum AM, Osadebe L, Kabamba J, Nguete B, et al. Evaluation of the GeneXpert for human monkeypox diagnosis. Am J Trop Med Hyg. 2017;96: 405–10. 39. Martinez-Murcia A, Navarro A, Garcia-Sirera A, Perez L, Bru G. Internal validation of a real-time qPCR kit following the UNE/EN ISO/IEC 17025:2005 for detection of the re-emerging monkeypox virus. Diagnostics (Basel). 2023;13:1560. 40. Shchelkunov SN, Shcherbakov DN, Maksyutov RA, Gavrilova EV. Species-specif- ic identification of variola, monkeypox, cowpox, and vaccinia viruses by multi- plex real-time PCR assay. J Virol Methods. 2011;175:163–9. 41. Kulesh DA, Loveless BM, Norwood D, Garrison J, Whitehouse CA, Hartmann C, et al. Monkeypox virus detection in rodents using real-time 3'-minor groove binder TaqMan assays on the Roche LightCycler. Lab Invest. 2004;84:1200–8. 42. Paniz-Mondolfi A, Guerra S, Munoz M, Luna N, Hernandez MM, Patino LH, et al. Evaluation and validation of an RT-PCR assay for specific detection of monkey- pox virus (MPXV). J Med Virol. 2023;95:e28247. 43. Elbaz M, Halutz O, Ali Y , Adler A. Diagnosis of monkeypox infection: validation of two diagnostic kits for viral detection using RT-PCR. J Virol Methods. 2023;312: 114653. 34 Acta Dermatovenerol APA | 2024;33:23-35 K. Resman Rus et al. 44. CDC. Poxvirus & Rabies Branch (PRB). Test procedure: monkeypox virus generic real-time PCR test [Internet]. [cited 2023 Dec 14]. Available from: https:/ /www. cdc.gov/poxvirus/mpox/pdf/PCR-Diagnostic-Protocol-508.pdf. 45. Karagoz A, Tombuloglu H, Alsaeed M, Tombuloglu G, AlRubaish AA, Mahmoud A, et al. Monkeypox (mpox) virus: classification, origin, transmission, genome organization, antiviral drugs, and molecular diagnosis. J Infect Public Health. 2023;16:531–41. 46. Pomari E, Mori A, Accordini S, Donini A, Cordioli M, Tacconelli E, et al. Evaluation of a ddPCR commercial assay for the absolute quantification of the monkeypox virus West Africa in clinical samples. Diagnostics (Basel). 2023;13:1349. 47. Kojabad AA, Farzanehpour M, Galeh HEG, Dorostkar R, Jafarpour A, Bolandian M, et al. Droplet digital PCR of viral DNA/RNA, current progress, challenges, and future perspectives. J Med Virol. 2021;93:4182–97. 48. Chen B, Jiang Y, Cao X, Liu C, Zhang N, Shi D. Droplet digital PCR as an emerging tool in detecting pathogens nucleic acids in infectious diseases. Clin Chim Acta. 2021;517:156–61. 49. Colavita F, Mazzotta V, Rozera G, Abbate I, Carletti F, Pinnetti C, et al. Kinetics of viral DNA in body fluids and antibody response in patients with acute monkey- pox virus infection. iScience. 2023;26:106102. 50. Yang J, Guo R, Li H, Chen G, Lin Y, Wang X, et al. Development of highly accurate digital PCR method and reference material for monkeypox virus detection. Anal Bioanal Chem. 2023;415:1333–7. 51. Specchiarello E, Carletti F, Matusali G, Abbate I, Rozera G, Minosse C, et al. De- velopment and validation of a nanoplate-based digital PCR assay for absolute MPXV quantification. J Virol Methods. 2023;321:114802. 52. Markov PV, Ghafari M, Beer M, Lythgoe K, Simmonds P , Stilianakis NI, et al. The evolution of SARS-CoV-2. Nat Rev Microbiol. 2023;21:361–79. 53. Taouk ML, Steinig E, Taiaroa G, Savic I, Tran T, Higgins N, et al. Intra- and inter- host genomic diversity of monkeypox virus. J Med Virol. 2023;95:e29029. 54. Vandenbogaert M, Kwasiborski A, Gonofio E, Descorps-Declere S, Selekon B, Nkili Meyong AA, et al. Nanopore sequencing of a monkeypox virus strain iso- lated from a pustular lesion in the Central African Republic. Sci Rep. 2022;12: 10768. 55. Forni D, Moltrasio C, Sironi M, Mozzi A, Quattri E, Venegoni L, et al. Whole-ge- nome sequencing of hMPXV1 in five Italian cases confirms the occurrence of the predominant epidemic lineage. J Med Virol. 2023;95:e28493. 56. Howard M, Maki JJ, Connelly S, Hardy DJ, Cameron A. Whole-genome sequences of human monkeypox virus strains from two 2022 global outbreak cases in west- ern New York state. Microbiol Resour Announc. 2022;11:e0084622. 57. Choi CH, No JS, Kim JW, Lee M, Shin H, Choi MM, et al. Complete genome se- quence of monkeypox virus strain MPXV-ROK-P1-2022 isolated from the first monkeypox patient in the Republic of Korea. Microbiol Resour Announc. 2022; 11:e0085322. 58. Chen Y , Wu C, Ruhan A, Zhao L, Zhang Z, Tan W. Perspective on the application of genome sequencing for monkeypox virus surveillance. Virol Sin. 2023;38:327– 33. 59. Nasri F, Kongkitimanon K, Wittig A, Cortes JS, Brinkmann A, Nitsche A, et al. MpoxRadar: a worldwide MPXV genomic surveillance dashboard. Nucleic Acids Res. 2023;51:W331–7. 60. Ndodo N, Ashcroft J, Lewandowski K, Yinka-Ogunleye A, Chukwu C, Ahmad A, et al. Distinct monkeypox virus lineages co-circulating in humans before 2022. Nat Med. 2023;29:2317–24. 61. Firth C, Kitchen A, Shapiro B, Suchard MA, Holmes EC, Rambaut A. Using time- structured data to estimate evolutionary rates of double-stranded DNA viruses. Mol Biol Evol. 2010;27:2038–51. 62. Brinkmann A, Kohl C, Pape K, Bourquain D, Thurmer A, Michel J, et al. Extensive ITR expansion of the 2022 Mpox virus genome through gene duplication and gene loss. Virus Genes. 2023;59:532–40. 63. Pfaff F, Hoffmann D, Beer M. Monkeypox genomic surveillance will challenge les- sons learned from SARS-CoV-2. Lancet. 2022;400:22–3. 64. Esteban DJ, Hutchinson AP. Genes in the terminal regions of orthopoxvirus ge- nomes experience adaptive molecular evolution. BMC Genomics. 2011;12:261. 65. Hughes AL, Friedman R. Poxvirus genome evolution by gene gain and loss. Mol Phylogenet Evol. 2005;35:186–95. 66. Kugelman JR, Johnston SC, Mulembakani PM, Kisalu N, Lee MS, Koroleva G, et al. Genomic variability of monkeypox virus among humans, Democratic Republic of the Congo. Emerg Infect Dis. 2014;20:232–9. 67. Kumar R, Nagar S, Haider S, Sood U, Ponnusamy K, Dhingra GG, et al. Monkey- pox virus: phylogenomics, host-pathogen interactome and mutational cascade. Microb Genom. 2023;9:mgen000987. 68. Ferraro F, Caraglia A, Rapiti A, Cereda D, Vairo F, Mattei G, et al. Letter to the edi- tor: multiple introductions of MPX in Italy from different geographic areas. Euro Surveill. 2022;27:2200456. 69. Chakraborty C, Bhattacharya M, Saikumar G, Alshammari A, Alharbi M, Lee SS, et al. A European perspective of phylogenomics, sublineages, geographical dis- tribution, epidemiology, and mutational landscape of mpox virus: emergence pattern may help to fight the next public health emergency in Europe. J Infect Public Health. 2023;16:1004–14. 70. Gao L, Shi Q, Dong X, Wang M, Liu Z, Li Z. Mpox, caused by the MPXV of the Clade IIb lineage, goes global. Trop Med Infect Dis. 2023;8:76. 71. Sharma SK, Dash PK, Yadav RG, Shrivastava A, Menon R, Kumar JS, et al. Isola- tion and characterization of emerging Mpox virus from India. J Med Virol. 2023; 95:e28911. 72. Nagler FP, Rake G. The use of the electron microscope in diagnosis of variola, vaccinia, and varicella. J Bacteriol. 1948;55:45–51. 73. Schramlova J, Arientova S, Hulinska D. The role of electron microscopy in the rap- id diagnosis of viral infections—review. Folia Microbiol (Praha). 2010;55:88–101. 74. Witt ASA, Trindade GS, Souza FG, Serafim MSM, da Costa AVB, Silva MVF, et al. Ultrastructural analysis of monkeypox virus replication in Vero cells. J Med Virol. 2023;95:e28536. 75. Gelderblom HR, Madeley D. Rapid viral diagnosis of orthopoxviruses by electron microscopy: optional or a must? Viruses. 2018;10:142. 76. Almeida JD. Uses and abuses of diagnostic electron microscopy. Curr Top Micro- biol Immunol. 1983;104:147–58. 77. Laue M, Bannert N. Detection limit of negative staining electron microscopy for the diagnosis of bioterrorism-related micro-organisms. J Appl Microbiol. 2010; 109:1159–68. 78. Hazelton PR, Gelderblom HR. Electron microscopy for rapid diagnosis of infec- tious agents in emergent situations. Emerg Infect Dis. 2003;9:294–303. 79. Paniz-Mondolfi A, Reidy J, Pagani N, Lednicky JA, McGrail JP, Kasminskaya Y, et al. Genomic and ultrastructural analysis of monkeypox virus in skin lesions and in human/animal infected cells reveals further morphofunctional insights into viral pathogenicity. J Med Virol. 2023;95:e28878. 80. Hyun J. Poxvirus under the eyes of electron microscope. Appl Microsc. 2022;52: 11. 81. Moltrasio C, Boggio FL, Romagnuolo M, Cagliani R, Sironi M, Di Benedetto A, et al. Monkeypox: a histopathological and transmission electron microscopy study. Microorganisms. 2023;11:1781. 82. Muller M, Ingold-Heppner B, Stocker H, Heppner FL, Dittmayer C, Laue M. Elec- tron microscopy images of monkeypox virus infection in 24-year-old man. Lan- cet. 2022;400:1618. 83. Rodriguez-Cuadrado FJ, Najera L, Suarez D, Silvestre G, Garcia-Fresnadillo D, Roustan G, et al. Clinical, histopathologic, immunohistochemical, and electron microscopic findings in cutaneous monkeypox: a multicenter retrospective case series in Spain. J Am Acad Dermatol. 2023;88:856–63. 84. Stagles MJ, Watson AA, Boyd JF, More IA, McSeveney D. The histopathology and electron microscopy of a human monkeypox lesion. Trans R Soc Trop Med Hyg. 1985;79:192–202. 85. Magnus von P, Andersen EK, Petersen KB, Birch-Andersen A. A pox-like dis- ease in cynomolgus monkeys. Acta Pathologica Microbiologica Scandinavica. 1959;46: 156–76. 86. Marennikova SS, Seluhina EM, Mal’ceva NN, Cimiskjan KL, Macevic GR. Isolation and properties of the causal agent of a new variola-like disease (monkeypox) in man. Bull World Health Organ. 1972;46:599–611. 87. Hutin YJ, Williams RJ, Malfait P, Pebody R, Loparev VN, Ropp SL, et al. Outbreak of human monkeypox, Democratic Republic of Congo, 1996 to 1997. Emerg Infect Dis. 2001;7:434–8. 88. Ma A, Langer J, Hanson KE, Bradley BT. Characterization of the cytopathic effects of monkeypox virus isolated from clinical specimens and differentiation from common viral exanthems. J Clin Microbiol. 2022;60:e0133622. 89. Moschese D, Pozza G, Mileto D, Giacomelli A, Cutrera M, Cossu MV, et al. Isola- tion of viable monkeypox virus from anal and urethral swabs, Italy, May to July 2022. Euro Surveill. 2022;27:2200675. 90. Rosa RB, Ferreira de Castro E, Vieira da Silva M, Paiva Ferreira DC, Jardim ACG, Santos IA, et al. In vitro and in vivo models for monkeypox. iScience. 2023;26: 105702. 91. Lalani AS, Masters J, Zeng W, Barrett J, Pannu R, Everett H, et al. Use of chemokine receptors by poxviruses. Science. 1999;286:1968–71. 92. Blasco R, Sisler JR, Moss B. Dissociation of progeny vaccinia virus from the cell membrane is regulated by a viral envelope glycoprotein: effect of a point muta- tion in the lectin homology domain of the A34R gene. J Virol. 1993;67:3319–25. 93. Lin CL, Chung CS, Heine HG, Chang W. Vaccinia virus envelope H3L protein binds to cell surface heparan sulfate and is important for intracellular mature virion morphogenesis and virus infection in vitro and in vivo. J Virol. 2000;74:3353–65. 94. Lapa D, Carletti F, Mazzotta V , Matusali G, Pinnetti C, Meschi S, et al. Monkeypox virus isolation from a semen sample collected in the early phase of infection in a patient with prolonged seminal viral shedding. Lancet Infect Dis. 2022;22: 1267–9. 95. Callaby H, Emery K, Killip M, Rampling T, Richards KS, Houlihan CF. Monkeypox virus isolation from longitudinal samples from four patients to infer risk of on- wards transmission: an interim analysis. J Hosp Infect. 2023;135:139–44. 96. Paran N, Yahalom-Ronen Y, Shifman O, Lazar S, Ben-Ami R, Yakubovsky M, et al. Monkeypox DNA levels correlate with virus infectivity in clinical samples, Israel, 2022. Euro Surveill. 2022;27:2200636. 97. Batejat C, Grassin Q, Feher M, Hoinard D, Vanhomwegen J, Manuguerra JC, et al. Heat inactivation of monkeypox virus. J Biosaf Biosecur. 2022;4:121–3. 98. Fischer RJ, Gallogly S, Schulz JE, van Doremalen N, Munster V, Das S. Evaluation of five buffers for inactivation of monkeypox virus and feasibility of virus detec- tion using the Panther Fusion® Open Access System. Viruses. 2022;14:2227. 35 Acta Dermatovenerol APA | 2024;33:23-35 Review of laboratory methods for monkeypox virus 99. Queromes G, Frobert E, Bouscambert-Duchamp M, Oblette A, Valette M, Billaud G, et al. Rapid and reliable inactivation protocols for the diagnostics of emerg- ing viruses: the example of SARS-CoV-2 and monkeypox virus. J Med Virol. 2023; 95:e28126. 100. Zapata-Cardona MI, Florez-Alvarez L, Gomez-Gallego DM, Moncada-Diaz MJ, Hernandez JC, Diaz F, et al. Comparison among plaque assay, tissue culture in- fectious dose (TCID(50)) and real-time RT-PCR for SARS-CoV-2 variants quantifi- cation. Iran J Microbiol. 2022;14:291–9. 101. Lei C, Yang J, Hu J, Sun X. On the calculation of TCID(50) for quantitation of virus infectivity. Virol Sin. 2021;36:141–4. 102. Hierholzer JC. KRA. Virus isolation and quantitation. In: Mahy BWJ, Kangro HO, editors. Virology methods manual. London: Academic Press; 1996. p. 25–46. 103. Resman Rus K, Korva M, Knap N, Avsic Zupanc T, Poljak M. Performance of the rapid high-throughput automated electrochemiluminescence immunoassay targeting total antibodies to the SARS-CoV-2 spike protein receptor binding do- main in comparison to the neutralization assay. J Clin Virol. 2021;139:104820. 104. Taha TY, Townsend MB, Pohl J, Karem KL, Damon IK, Mbala Kingebeni P, et al. Design and optimization of a monkeypox virus specific serological assay. Patho- gens. 2023;12:396. 105. Baxby D, Bennett M, Getty B. Human cowpox 1969–93: a review based on 54 cases. Br J Dermatol. 1994;131:598–607. 106. Pelkonen PM, Tarvainen K, Hynninen A, Kallio ER, Henttonen K, Palva A, et al. Cow- pox with severe generalized eruption, Finland. Emerg Infect Dis. 2003;9:1458– 61. 107. Di Giulio DB, Eckburg PB. Human monkeypox: an emerging zoonosis. Lancet In- fect Dis. 2004;4:15–25. 108. McCollum AM, Damon IK. Human monkeypox. Clin Infect Dis. 2014;58:260–7. 109. Shete AM, Patil DY, Jain R, Sahay RR, Chenayil S, Mohandas S, et al. Assessment of antibody kinetics in mpox cases with indigenously developed IgM and IgG ELISA. J Med Virol. 2023;95:e29073. 110. De Baetselier I, Van Dijck C, Kenyon C, Coppens J, Michiels J, de Block T, et al. Retrospective detection of asymptomatic monkeypox virus infections among male sexual health clinic attendees in Belgium. Nat Med. 2022;28:2288–92. 111. Otter AD, Jones S, Hicks B, Bailey D, Callaby H, Houlihan C, et al. Monkeypox virus-infected individuals mount comparable humoral immune responses as smallpox-vaccinated individuals. Nat Commun. 2023;14:5948. 112. Sammartino JC, Cassaniti I, Ferrari A, Piralla A, Bergami F, Arena FA, et al. Char- acterization of immune response against monkeypox virus in cohorts of infected patients, historic and newly vaccinated subjects. J Med Virol. 2023;95:e28778. 113. Therrien C, Prevost J, Blais AC, Turcotte S, Gendron-Lepage G, Finzi A, et al. De- velopment and validation of a highly specific in-house chemiluminescent-based serological assay for the detection of antibodies directed against the human monkeypox virus. J Virol Methods. 2023;322:114836. 114. Davis I, Payne JM, Olguin VL, Sanders MP , Clements T, Stefan CP , et al. Develop- ment of a specific MPXV antigen detection immunodiagnostic assay. Front Mi- crobiol. 2023;14:1243523. 115. Yates JL, Hunt DT, Kulas KE, Chave KJ, Styer L, Chakravarthi ST, et al. Develop- ment of a novel serological assay for the detection of mpox infection in vacci- nated populations. J Med Virol. 2023;95:e29134. 116. Karem KL, Reynolds M, Braden Z, Lou G, Bernard N, Patton J, et al. Characteriza- tion of acute-phase humoral immunity to monkeypox: use of immunoglobulin M enzyme–linked immunosorbent assay for detection of monkeypox infection dur- ing the 2003 North American outbreak. Clin Diagn Lab Immunol. 2005;12:867– 72. 117. Crotty S, Felgner P, Davies H, Glidewell J, Villarreal L, Ahmed R. Cutting edge: long-term B cell memory in humans after smallpox vaccination. J Immunol. 2003;171:4969–73. 118. Hammarlund E, Lewis MW, Hansen SG, Strelow LI, Nelson JA, Sexton GJ, et al. Du- ration of antiviral immunity after smallpox vaccination. Nat Med. 2003;9:1131–7. 119. Minhaj FS, Singh V, Cohen SE, Townsend M, Scott H, Szumowski J, et al. Preva- lence of undiagnosed monkeypox virus infections during global Mpox outbreak, United States, June–September 2022. Emerg Infect Dis. 2023;29:2307–14. 120. Li E, Guo X, Hong D, Gong Q, Xie W, Li T, et al. Duration of humoral immunity from smallpox vaccination and its cross-reaction with Mpox virus. Signal Transduct Target Ther. 2023;8:350. 121. Kempe CH, Bowles C, Meiklejohn G, Berge TO, St Vincent L, Babu BV, et al. The use of vaccinia hyperimmune gamma-globulin in the prophylaxis of smallpox. Bull World Health Organ. 1961;25:41–8. 122. Belyakov IM, Earl P, Dzutsev A, Kuznetsov VA, Lemon M, Wyatt LS, et al. Shared modes of protection against poxvirus infection by attenuated and conventional smallpox vaccine viruses. Proc Natl Acad Sci U S A. 2003;100:9458–63. 123. Edghill-Smith Y, Golding H, Manischewitz J, King LR, Scott D, Bray M, et al. Smallpox vaccine-induced antibodies are necessary and sufficient for protec- tion against monkeypox virus. Nat Med. 2005;11:740–7. 124. Gilchuk I, Gilchuk P, Sapparapu G, Lampley R, Singh V, Kose N, et al. Cross- neutralizing and protective human antibody specificities to poxvirus infections. Cell. 2016;167:684–94 e9. 125. Manenti A, Solfanelli N, Cantaloni P, Mazzini L, Leonardi M, Benincasa L, et al. Evaluation of monkeypox- and vaccinia virus-neutralizing antibodies in human serum samples after vaccination and natural infection. Front Public Health. 2023;11:1195674. 126. Zaeck LM, Lamers MM, Verstrepen BE, Bestebroer TM, van Royen ME, Gotz H, et al. Low levels of monkeypox virus-neutralizing antibodies after MVA-BN vaccina- tion in healthy individuals. Nat Med. 2023;29:270–8. 127. Moschese D, Giacomelli A, Mileto D, Sacco Mpox Study G. Ongoing sporadic monkeypox cases: neutralising antibody detection in asymptomatic individuals. Lancet Microbe. 2023;4:e765. 128. Huang B, Zhao H, Song J, Zhao L, Deng Y, Wang W, et al. Isolation and char- acterization of monkeypox virus from the first case of monkeypox—Chongqing Municipality, China, 2022. China CDC Wkly. 2022;4:1019–24. 129. Abebe EC, Dejenie TA. Protective roles and protective mechanisms of neutraliz- ing antibodies against SARS-CoV-2 infection and their potential clinical implica- tions. Front Immunol. 2023;14:1055457. 130. Rocha VPC, Quadros HC, Fernandes AMS, Goncalves LP, Badaro R, Soares MBP, et al. An overview of the conventional and novel methods employed for SARS- CoV-2 neutralizing antibody measurement. Viruses. 2023;15:1504.