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Korean J. Vet. Serv. 2022; 45(1): 19-30

Published online March 30, 2022

https://doi.org/10.7853/kjvs.2022.45.1.19

© The Korean Socitety of Veterinary Service

Simple and rapid colorimetric detection of African swine fever virus by loop-mediated isothermal amplification assay using a hydroxynaphthol blue metal indicator

Ji-Hoon Park 1†, Hye-Ryung Kim 1†, Ha-Kyung Chae 1,3, Jonghyun Park 1,3, Bo-Young Jeon 4, Young S. Lyoo 5, Choi-Kyu Park 1*

1College of Veterinary Medicine & Animal Disease Intervention Center, Kyungpook National University, Daegu 41566, Korea
2DIVA Bio, Inc., Daegu 41519, Korea
3Korea Disease Control and Prevention Agency, Gyeongbuk Regional Center for Disease Control and Prevention, Daegu 41061, Korea
4Department of Biomedical Laboratory Science, College of Health Science, Yonsei University, Wonju 26493, Korea
5College of Veterinary Medicine, Konkuk University, Seoul 05029, Korea

Correspondence to : Choi-Kyu Park
E-mail: parkck@knu.ac.kr
https://orcid.org/0000-0002-0784-9061
These first three authors contributed equally to this work.

Received: March 23, 2022; Accepted: March 28, 2022

This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0). which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

In this study, a simple loop-mediated isothermal amplification (LAMP) combined with visual detection method (vLAMP) assay was developed for the rapid and specific detection of African swine fever virus (ASFV), overcoming the shortcomings of previously described LAMP assays that require additional detection steps or pose a cross-contamination risk. The assay results can be directly detected by the naked eye using hydroxynaphthol blue after incubation for 40 min at 62℃. The assay specifically amplified ASFV DNA and no other viral nucleic acids. The limit of detection of the assay was <50 DNA copies/reaction, which was ten times more sensitive than conventional polymerase chain reaction (cPCR) and comparable to real-time PCR (qPCR). For clinical evaluation, the ASFV detection rate of vLAMP was higher than cPCR and comparable to OIE-recommended qPCR, showing 100% concordance, with a κ value (95% confidence interval) of 1 (1.00∼1.00). Considering the advantages of high sensitivity and specificity, no possibility for cross-contamination, and being able to be used as low-cost equipment, the developed vLAMP assay will be a valuable tool for detecting ASFV from clinical samples, even in resource-limited laboratories.

Keywords African swine fever virus, Loop-mediated isothermal amplification, Visual detection

African swine fever (ASF) is a highly contagious transboundary swine disease caused by the ASF virus (ASFV) and is responsible for devastating economic losses in domesticated pigs and wild boar. ASFV is a large, enveloped double-stranded DNA virus, the only member of genus Asfivirus, family Asfarviridae (Alonso et al, 2018). ASFV strains have been divided into 24 genotypes based on their B646L gene, which encodes the major capsid protein p72 (Bastos et al, 2003; Quembo et al, 2018). The African continent is the endemic area with all 24 ASFV genotypes. Outside Africa, only genotype I was found in Europe, South America, and the Caribbean until genotype II ASFV was introduced in Georgia from east Africa in 2007. Since then, genotype II ASFV had quickly spread to the Russian Federation, Ukraine, and Belarus and in 2014 to the European Union Baltic States and Poland. By 2018, the infection had also spread to Belgium, Hungary, the Czech Republic, Romania, Bulgaria, Slovakia, and Serbia. In August 2018, the virus finally emerged in China and rapidly spread to many Asian countries, including Mongolia, Vietnam, Cambodia, Laos and North Korea (Democratic People’s Republic of Korea), the Philippines, and South Korea (Zhou et al, 2018; Gaudreault et al, 2020; Kim et al, 2020a).

On September 17, 2019, the first ASF outbreak in a domestic pig farm was confirmed in South Korea. On April 29, 2019, 14 cases of ASF outbreaks in domestic pig farms were diagnosed in four cities or counties (Kim et al, 2020a). Phylogenetic analysis showed that all of 14 ASFV isolates in South Korea belonged to genotype II and serogroup 8, identical to European and Chinese genotype II ASFV isolates (Kim et al, 2020b). On the other hand, the first wild boar infection case was confirmed on October 2, 2019, in a wild boar carcass found by the military in the Demilitarized Zone (DMZ). Despite implementing national control measures, ASFV-infected wild boars have been continuously identified in the border areas adjacent to DMZ. ASFV infections in wild boars are considered a risk factor for further ASF outbreaks in domestic pig farms and are becoming a major concern for the Korean Government and pig farmers (Jo and Gortázar, 2020; Yoo et al, 2021). Concerns that ASFV, continuously circulating in wild boar populations, could be transmitted into domestic pig farms has been realized with two additional ASFV outbreaks in domestic pig farms located in wild boar outbreak areas in October 2020 and May 2021.

Because there is no vaccine available, the prevention, control, and eradication of ASF are based on the implementation of appropriated surveillance and strict biosecurity measures. The success of surveillance activities depends on the availability of the most appropriate diagnostic tests. Although a number of good validated ASF diagnostic techniques are available, virus detection methods, including viral genome detection by polymerase chain reaction (PCR), viral antigen detection by antigen enzyme-linked immunosorbent assay or a direct immunofluorescence test, and virus detection using virus isolation, are vital for the rapid implementation of control measures, especially in ASFV-free areas (Gallardo et al, 2019). Currently, PCR assays, including conventional PCR (cPCR) and quantitative real-time PCR (qPCR), are considered the gold standard tests for the early detection of the disease due to their superior sensitivity, specificity, robustness, and high-throughput application to detect the ASFV genome in any kind of clinical samples from domestic pigs, wild boar, and ticks (Agüero et al, 2003; King et al, 2003; Zsak et al, 2005; Tignon et al, 2011; Fernández-Pinero et al, 2013). However, these PCR-based assays require sophisticated equipment, specialized labor, and complicated procedures to detect amplified products; thus, such methods are not suitable for on-site diagnosis in field situations or primitive laboratories in developing countries. Therefore, the development of a simple, rapid, and cost-effective assay with reliable specificity and sensitivity is imperative to detect ASFV from suspected cases.

Loop-mediated isothermal amplification (LAMP) is a valuable tool for detecting various pathogens with high sensitivity, specificity, rapidity, and simplicity (Notomi et al, 2000; Mori and Notomi, 2009; Dhama et al, 2014). Several LAMP assays have been developed for ASFV detection (James et al, 2010; Atuhaire et al, 2014; Wu et al, 2016; Woźniakowski et al, 2018; Wang et al, 2020). With regard to detection methods for assay results, previously reported ASFV LAMP assays determined the assay results by gel electrophoresis, turbidity analysis by real-time turbidimeter, DNA-intercalating dye-mediated visual detection (PicoGreen, Evergreen, or SYBR Green 1), or vertical flow visualization strip. However, detection methods using gel electrophoresis and DNA-intercalating dyes may increase the potential risk for cross-contamination of amplified LAMP products, as these methods require opening the tube lid to add reagents or perform further experiments. Also, real-time turbidity analysis, fluorescence dye detection, and vertical flow visualization strip methods require additional instruments, such as real-time turbidimeter, fluorescence detector, and visualization strip cassette, limiting the applicability of LAMP assay as a field diagnostic assay. To facilitate the practical use of LAMP assays, a simple, rapid, and contamination-free detection method must be developed (Mori and Notomi, 2009). Previously, a simple and rapid colorimetric method was applied to detect LAMP results by adding hydroxynaphthol blue (HNB) metal indicators to the pre-reaction solution, which reduced the chances of carryover contamination of the LAMP product and enabled the direct visual detection of LAMP results without additional instrumentations (Goto et al, 2009). A number of LAMP assays using HNB have been developed to detect various animal pathogens (Lim et al, 2018; Park et al, 2018; Park et al, 2019; Chae et al, 2020; Kim et al, 2021; Kim et al, 2022). However, the simple colorimetric detection method using HNB has rarely been applied to ASFV LAMP assays. Therefore, in this study, a visual LAMP (vLAMP) assay combined with visual detection methodology using HNB was developed for the simple and rapid detection of ASFV, which can be used for on-site laboratory diagnoses.

Samples and nucleic acid extraction

To prevent the risk of unexpected transmission of ASFV, non-infectious ASFV DNA samples were obtained through various routes and used to develop and evaluate the vLAMP assay. A DNA standard was synthesized based on the previously reported ASFV sequence (ASFV-SY18 strain; GenBank accession no. MH766894) and used for the development and sensitivity evaluation of the assay. Twelve viral DNA were extracted from different ASFV genotypes, and 22 clinical DNA samples extracted from ASFV-infected pigs were kindly provided by Professor Bo-Young Jeon, who is conducting international joint research with a Russian ASFV research institute (Table 1). Additionally, 20 clinical pig samples (5 blood, 5 spleen, 5 kidney, and 5 lymph node samples) of Korean diseased pigs were collected and used as negative controls for clinical evaluation. DNA samples were allocated in small volumes and stored at −80℃ until use. To assess the specificity of vLAMP, classical swine fever virus (CSFV; strain LOM), type 1 porcine reproductive and respiratory syndrome virus (PRRSV; strain Lelystad virus), type 2 PRRSV (strain LMY), porcine parvovirus (PPV; strain NADL-2), porcine circovirus (PCV) type 1 (PCV1; from infected PK-15 cell culture), PCV type 2 (strain PCK0201; Park et al, 2004), PCV type3 (strain PCK3-1701; Kim et al, 2017), Erysipelothrix rhusiopathiae (ER; strain NL-11), and ST and PK-15 cells were obtained from the Animal and Plant Quarantine Agency (Korea) and the Animal Disease Intervention Center (Korea). The viral titers of all viruses were determined to be at least 104.0 TCID50/mL. All viral samples were allocated and stored at −80℃ until use. Total RNA or DNA was extracted from 200 μL of virus stocks, cultured cells, and clinical samples using the TAN Bead Nucleic Acid Extraction kit for automated extraction (TAN Bead, Taoyuan, Taiwan), according to manufacturer’s instructions. Extracted RNA or DNA samples were eluted in 100 μL nuclease-free water, allocated in small volumes, and stored at −80℃ until use.

Table 1 . Comparison of diagnostic results by cPCR, qPCR and vLAMP assays using ASFV-related DNA samples

No.DNA sampleSample source (ASFV genotype)aResults of different assaysb

cPCRqPCR (Ct value)vLAMP
1Viral DNAChinese isolate SY18 (G II)+29.53+
2Viral DNARussian field isolate in Itkutsk (G II)+26.76+
3Viral DNARussian field isolate 2 in Stavropol (G II)+25.84+
4Viral DNARussian field isolate 1 in Omsk (G II)+26.71+
5Viral DNARussian field isolate 2 in Omsk (G II)+23.67+
6Viral DNARussian field isolate 1 in Nizhny Novgorod (G II)+23.26+
7Viral DNARussian field isolate 2 in Nizhny Novgorod (G II)+21.76+
8Viral DNARussian field isolate 3 in Nizhny Novgorod (G II)+25.73+
9Viral DNARussian field isolate 4 in Nizhny Novgorod (G II)+24.27+
10Viral DNARussian field isolate in Krasnodar (G II)+24.76+
11Viral DNARussian field isolate in Saratov (G II)+27.06+
12Viral DNAAttenuated ASFV KK262 strain (G I)+26.82+
13Viral DNAAttenuated ASFV MK200 strain (G V)+26.04+
14Sample DNABlood of pig 1 infected with V strain, 3 DPC (G II)+29.94+
15Sample DNABlood of pig 1 infected with V strain, 5 DPC (G II)+22.81+
16Sample DNABlood of pig 2 infected with V strain, 3 DPC (G II)+29.42+
17Sample DNABlood of pig 2 infected with V strain, 5 DPC (G II)+23.34+
18Sample DNABlood of pig 3 infected with V strain, 3 DPC (G II)+29.27+
19Sample DNABlood of pig 3 infected with V strain, 4 DPC (G II)+27.23+
20Sample DNABlood of pig 3 infected with V strain, 6 DPC (G II)+21.73+
21Sample DNABlood of pig 4 infected with V strain, 4 DPC (G II)+25.36+
22Sample DNABlood of pig 4 infected with V strain, 5 DPC (G II)+22.56+
23Sample DNABlood of pig 4 infected with V strain, 6 DPC (G II)+25.19+
24Sample DNABlood of pig 5 infected with V strain, 3 DPC (G II)+30.05+
25Sample DNABlood of pig 5 infected with V strain, 4 DPC (G II)-
26Sample DNABlood of pig 5 infected with V strain, 5 DPC (G II)+25.17+
27Sample DNABlood of pig 6 infected with K strain, 3 DPC (G 1)+25.85+
28Sample DNABlood of pig 6 infected with K strain, 5 DPC (G 1)+28.86+
29Sample DNABlood of pig 6 infected with K strain, 7 DPC (G 1)33.35+
30Sample DNABlood of pig 7 infected with K strain, 3 DPC (G 1)+25.74+
31Sample DNABlood of pig 7 infected with K strain, 5 DPC (G 1)+21.45+
32Sample DNABlood of pig 7 infected with K strain, 7 DPC (G 1)+21.06+
33Sample DNABlood of pig 8 infected with K strain, 3 DPC (G 1)+27.18+
34Sample DNABlood of pig 8 infected with K strain, 5 DPC (G 1)+21.85+
35Sample DNABlood of pig 8 infected with K strain, 7 DPC (G 1)+30.92+
36∼40Sample DNABloods of diseased Korean pigs (NC)-
41∼45Sample DNASpleens of diseased Korean pigs (NC)-
46∼50Sample DNAKidneys of diseased Korean pigs (NC)-
51∼55Sample DNALymph nodes of diseased Korean pigs (NC)-

aV strain, Russian field strain Volgograd/wb/2014; K strain, Russian virulent strain KK262; DPC, days of postchallenge; G, genotype of African swine fever virus; NC, negative control.

bThe results of the visual loop-mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) were presented as positive (+) or negative (−), and positive results of the real-time PCR (qPCR) were presented as the threshold cycle (Ct) values for each sample.



Construction of a DNA standard

Because a developing LAMP assay and reference PCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays in this study target the same ASFV B646L gene, the gene was cloned and used as a DNA standard. Based on the sequence of the Chinese genotype II ASFV strain ASFV-SY18 (GenBank accession no. MH766894), 1942-bp of the B646L gene was commercially synthesized and cloned into the pUC57 cloning vector (GenScript, Seoul, Korea) by BIONICS (Seoul, Korea). Recombinant plasmid DNA samples were linearized by digestion with EcoRI (Takara Korea Biomedical, Seoul, Korea) and purified using the Expin CleanUP SV kit (GeneAll Biotechnology, Seoul, Korea), according to the manufacturer’s instructions. DNA concentrations were determined by measuring the absorbance at 260 nm using a NanoDrop Lite spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). DNA transcript copy numbers were quantified using a previously described method (Park et al, 2018). The DNA standard was stored at −80℃.

Primers for the vLAMP assay

Because the B646L gene served as a useful diagnostic marker for OIE-approved reference cPCR and qPCR assays for ASFV, vLAMP primers were designed based on the conserved regions of the B646L gene (Agüero et al, 2003; King et al, 2003). A total of 355 complete or partial B646L gene sequences of 24 ASFV genotypes were retrieved from the National Center for Biotechnology Information (NCBI) GenBank database. Conserved nucleotide sequences within the B646L gene were identified by multiple alignments using the BioEdit Sequence Alignment Editor program (http://www.mbio.ncsu.edu/BioEdit/bioedit.html). Based on these conserved sequences, a set of six primers, including two outer primers (F3 and B3), two inner primers (FIP and BIP), and two loop primers (LF and LB), were manually designed for the vLAMP assay aided by Primer Explorer V5 software (Fujitsu System Solutions Ltd., Tokyo, Japan). The specificity of the primers for vLAMP was confirmed against random nucleotide sequences using a BLAST search of the NCBI GenBank database (http://www.ncbi.nlm.nih.gov/BLAST/). All primers were synthesized by BIONICS, as shown in Table 2.

Table 2 . Primers and probes used in vLAMP and cPCR, and qPCR in this study

MethodPrimer and probeLength (bp)Primer/probe sequence (5’–3’)aGenome positionbReference
vLAMPF318CTCTTCCAGACGCATGTT85893∼85910This study
B318CCGTRGTGATAGACCCCA86196∼86213
LF24TGTAAGAGCTGCAGAACTTTGATG86013∼86036
LB21TTGAARCCACGGGAGGAATAC86108∼86128
FIP (F1c+F2)41GCCTCCGTAGTGRAAGGGTA+86037∼86056+
GTHACTGCTCAYGGTATCAAT85976∼85996
BIP (B1c+B2)44TCCGGGYGCGATGATGATTAC+86080∼86100+
CTTGCTCTGGATACGTTAATATG86141∼86163
PCRPPA121TTCCCAGCGTAGTTGAGATTG84467∼84486Agüero et al (2003)
PPA220AGTTATGGGAAACCCGACCC84705∼84723
qPCRF25CTGCTCATGGTATCAATCTTATCGA85980∼86004King et al (2003)
R25CTGCTCATGGTATCAATCTTATCGA86210∼86229
P25FAM–CCACGGGAGGAATACCAACCCAGTG–BHQ186114∼86138

aBold text in B3, LB, FIP, and BIP sequences used in the vLAMP assay represent a degenerate base: R, A/G; H, A/C/T; Y, C/T. FAM, 6-carboxyfluorescein; BHQ1, Black Hole Quencher 1.

bThe locations of all primer and probe sequences for the visual loop-mediated isothermal amplification (vLAMP), conventional polymerase chain reaction (cPCR) and real-time PCR (qPCR) assays were derived from the complete genome sequence of the Chinese representative African swine fever virus strain ASFV-SY18 (GenBank accession no. MH766894).



Optimization of vLAMP conditions

vLAMP was performed in a reaction mixture containing 20 mM Tris-HCl (pH 8.8), 10mM KCl, 10mM (NH4)2SO4, 0.1% Triton X-100, 1.4 mM dNTPs, 8 mM MgSO4, 0.12 mM HNB (Lemongreen, Shanghai, China), 0.8 M betaine (Sigma–Aldrich, St. Louis, Missouri, USA), 8U Bst DNA polymerase (large fragment; New England Biolabs, Ipswich, MA, USA), 1.6 μM inner primers (FIP and BIP), 0.2 μM outer primers (F3 and B3), 0.8 μM loop primers (LF and LB), 5 μL template RNA, and dH2O to a final volume of 25 μL. The reaction conditions were optimized by performing test amplifications at temperatures ranging from 56℃ to 68℃ and reaction times of 30∼60 min. Reactions were terminated by heating at 80℃ for 5 min. All experiments were performed in triplicate. The assay results were interpreted by visual detection of a color change from purple to sky blue due to the presence of the metal ion indicator HNB (Goto et al, 2009). Amplicons were also detected by observing LAMP-specific ladder-like DNA bands using an ultraviolet light transilluminator (Bio-Rad, Hercules, CA, USA) after subjecting the DNA to 1.5% agarose gel electrophoresis and staining with NEO green dye (Neoscience, Suwon, Korea).

cPCR and qPCR assays

cPCR was performed with ASFV B646L gene-specific primers using an Excel TB 2X Taq premix (Inclone, Yongin, Korea) and an Applied Biosystems thermal cycler, as described previously (Agüero et al, 2003). The expected size of the cPCR product using the primer set (PPA1 and PPA2) was 257 bp (Table 2). The cPCR conditions were as follows: initial denaturation at 95℃ for 2 min, 40 cycles of PCR (95℃ for 20 s, 62℃ for 40 s, and 72℃ for 30 s), and final extension at 72℃ for 5 min. The cPCR products were subjected to electrophoresis on 1.5% agarose gel. The target bands were visualized by staining with NEO green dye (Neoscience) and imaged under blue light illumination (Neoscience). qPCR was performed using a commercial qPCR kit (Premix Ex Taq™ Probe qPCR, Takara Korea Biomedical) with ASFV B646L gene-specific primers and probe (Table 2), according to previously described methods (King et al, 2003). In brief, qPCR was performed in a 25 μL reaction mixture containing 12.5 μL of 2×reaction buffer, 0.5 μL of each ASFV primers (0.2 μM and probe 0.2 μM), 5 μL of DNA template, and 6 μL of water. The reaction was carried out in a CFX96 Touch™ Real-time PCR Detection System (Bio-Rad) under the following conditions: initial denaturation at 95℃ for 30 s, 40 cycles of denaturation at 95℃ for 5 s, and annealing and extension at 58℃ for 30 s. Real-time fluorescence values of the FAM-labeled probe were measured in ongoing reactions at the end of each annealing step. To interpret the qPCR results, samples producing a cycle threshold (Ct) of <40 were considered positive, whereas those with a higher Ct value (>40) were considered negative (King et al, 2003).

Specificity and sensitivity of the vLAMP assay

To test its specificity, the vLAMP assay was performed with DNA or RNA templates extracted from a genotype II ASFV strain (ASFV-SY18), seven control virus cultures (CSFV, type 1 and 2 PRRSVs, PPV, PCV1, PCV2, and PCV3), a bacterial culture (ER), and two uninfected cell cultures (ST and PK-15 cells) as negative controls. The limit of detection (LOD) of the vLAMP assay was determined using 10-fold serial dilutions of an ASFV DNA standard, ranging from 106 to 1 copies/μL. Subsequently, the LOD of the vLAMP assay was compared to cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays using the same DNA templates described above.

Comparative evaluation of the vLAMP assay

For evaluation of the vLAMP assay, 55 DNA samples including a DNA standard (103 copies/μL), 12 viral DNA extracted different ASFV strains, 22 clinical DNA samples extracted from ASFV-infected pigs, and 20 clinical DNA samples extracted from ASFV-negative pigs, were tested using the developed vLAMP assay, and the results were compared to cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays (Table 1). The concordance between the vLAMP and cPCR or qPCR results was analyzed using Cohen’s κ statistics at 95% confidence interval (CI; Kwiecien et al, 2011). When the calculated κ coefficient value (κ) was ≥0.81, the results from these assays were almost 100% concordant.

Optimization of the vLAMP assay

For vLAMP reaction with ASFV DNA templates, a positive color change from purple to sky blue was visually detected in the reaction tubes, and a ladder pattern of products was generated using electrophoresis at temperatures ranging from 56℃∼68℃ (Fig. 1). The color changes and electrophoretic bands were clearer at 58℃, 60℃, 62℃, and 64℃ than at 56℃, 66℃, and 68℃. Subsequently, the vLAMP assay was performed with three dilutions of DNA transcript samples from the cloned ASFV B646L gene with different copy numbers (102, 101, and 100 copies/μL) for reaction times ranging from 30 to 60 min. The color change and electrophoresis results indicated that amplification was initiated within 30 min; however, a definite positive color change and strongly amplified DNA bands were observed at 40 min for reactions with the DNA template containing 101 copies/μL of standard DNA (Fig. 2). Based on these results, the optimal reaction temperature and time for the vLAMP assay were determined to be 62℃ and 40 min, respectively. Therefore, all subsequent experiments were performed under these reaction conditions.

Fig. 1.Optimal reaction temperature for the vLAMP assay to detect African swine fever virus. (A) Visual detection of vLAMP results. A color change in the reaction solution from purple to sky blue indicated a positive reaction. (B) Electrophoretic analysis of vLAMP amplification products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 7, vLAMP results at seven different reaction temperatures (56℃, 58℃, 60℃, 62℃, 64℃, 66℃, and 68℃), respectively. NC, negative control (nuclease-free water).
Fig. 2.Optimal reaction time for African swine fever virus (ASFV) detection using vLAMP (A) Visual detection of vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 3, vLAMP results for ASFV DNAs (from 5×102 to 5×100 copies/reaction) at four different reaction times (30, 40, 50, and 60 min), respectively. NC, negative control (nuclease-free water).

Specificity of the vLAMP assay

After optimization of the vLAMP conditions, an ASFV strain (ASFV-SY18) and seven control virus cultures (CSFV, type 1 and 2 PRRSVs, PPV, PCV1, PCV2, and PCV3), a bacterial culture (ER), and two cell cultures (ST and PK-15 cells) were tested using vLAMP to evaluate primer specificity. vLAMP yielded positive results for ASFV strains but negative for all other porcine pathogens, cell cultures, and negative controls. Therefore, the primer set used for this assay showed high specificity for the ASFV B646L gene (Fig. 3).

Fig. 3.Specificity of the vLAMP assay for African swine fever virus (ASFV). (A) Visual detection of the vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 11, vLAMP results for ASFV strains ASFV-SY18, classical swine fever virus (strain LOM), Erysipelothrix rhusiopathiae (strain NL-11), type 1 porcine reproductive and respiratory syndrome virus (PRRSV, strain Lelystad virus), type 2 PRRSV (strain LMY), porcine parvovirus (strain NADL-2), type 1 porcine circovirus (PCV1, field strain), PCV2 (strain PCK0201), PCV3 (strain PCK3-1701), ST cell and PK-15 cells, respectively. NC, negative control (nuclease-free water).

Sensitivity of vLAMP, cPCR, and qPCR assays

The sensitivity of the vLAMP assay was determined using standard DNA dilutions ranging from 106 to 100 copies/μL and compared to cPCR and qPCR assays. The LOD was determined as the lowest dilution factor in which positive reactions were obtained in all three replicates. The LOD of vLAMP (101 copies/μL) was 10-fold lower than cPCR (102 copies/μL) and comparable to qPCR (101 copies/μL), as shown in Fig. 4. These results indicated that the developed vLAMP assay was highly sensitive and will be an alternative diagnostic tool for the OIE-recommended qPCR assay (King et al, 2003).

Fig. 4.Comparative sensitivities between vLAMP, cPCR, and qPCR assays for African swine fever virus (ASFV) standard DNA. (A) Amplification curves of the qPCR assay. (B) Electrophoretic analysis of cPCR amplicons. (C, D) Visualization and electrophoretic analysis of vLAMP. Lines and tubes 1 to 7, in (A∼D) represent serial 10-fold dilutions of the ASFV DNA standard (from 5×106 to 5×100 copies). Lane M, 1 kb plus DNA marker; tube and lane NC, negative control (nuclease-free water).

Comparative diagnostic performance of the vLAMP assay

To evaluate the ability of the vLAMP assay to detect ASFV DNA from viral cultures and clinical samples, 55 DNA samples (13 viral DNA and 42 clinical DNA) were tested by the vLAMP assay, and the results were compared to cPCR and qPCR assays. The detection rate of ASFV DNA from 13 viral DNA samples was 100.0% (13/13) for all detection methods. The detection rate from clinical DNA samples by vLAMP, cPCR, and qPCR were 50.0% (21/42), 47.6% (20/42), and 50.0% (21/42), respectively. A clinical DNA sample positive by vLAMP and qPCR (Ct value of 33.35) was tested by cPCR as negative. The total detection rates of vLAMP and qPCR (61.8%) were higher than cPCR (60.0%; Table 3). For ASFV detection from DNA samples, the percentages of positive and negative results and the overall agreement for the vLAMP assay relative to the cPCR assay were 97.1% (33/34), 100% (21/21), and 98.2% (54/55), with a κ value (95% CI) of 0.96 (0.92∼1.00). Relative to the qPCR assay, these values were 100% (34/34), 100% (21/21), and 100% (55/55), with a κ value (95% CI) of 1 (1.00∼1.00), indicating that the diagnostic vLAMP results were 100% concordant with the qPCR assay.

Table 3 . Comparison of vLAMP, cPCR, and qPCR results for the detection of African swine fever virus in different DNA samples

SampleNo. testedNo positive (%)

cPCRqPCRvLAMP
Viral DNA from different ASFVs1313 (100)13 (100)13 (100)
Sample DNA from infected pigs2220 (90.9)21 (95.5)21 (95.5)
Sample DNA from uninfected pigs20000
Total5533 (60.0)34 (61.8)34 (61.8)

The calculated κ coefficient value (95% confidence interval) between visual loop–mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) or real-time PCR (qPCR) were 0.96 (0.92∼1.0), and 1 (1.0∼1.0), respectively.


Since 2019, when outbreaks in domestic pigs and wild boars were confirmed in Korea, despite national control measures executed, ASFV infections in wild boars continue to spread geographically, thereby increasing the risk of further outbreaks in domestic pig farms in wild boar outbreak areas (Jo and Gortázar, 2020; Kim et al, 2020a; Yoo et al, 2021). Considering the continuous spread of ASFV in wild boar populations and the risk of ASFV transmission associated with the movement of ASF-suspected wild boar samples, it is necessary to develop a diagnosis method that can be applied at the site where suspected domestic pigs or wild boar are found. LAMP has been recognized as a valuable tool for detecting of various animal pathogens due to its high sensitivity and specificity, rapidity, simplicity, and field applicability (Mori and Notomi, 2009; Dhama et al, 2014). For ASFV diagnosis, some LAMP assays have been reported so far, but these assays have limitations in their use as an on-field diagnostic assay mainly due to the monitoring methods of LAMP results that are difficult to apply in field situations (James et al, 2010; Atuhaire et al, 2014; Wu et al, 2016; Woźniakowski et al, 2018; Wang et al, 2020). Therefore, a vLAMP assay with high sensitivity and specificity, simplicity, rapidity, and field applicability was developed in this study.

The vLAMP assay using six primers that recognize eight regions of the ASFV B646L gene specifically detected ASFV but did not amplify any other porcine pathogens, as shown in Fig. 3. The LOD of the vLAMP assay was 10-fold lower than cPCR and comparable to qPCR (Fig. 4). Furthermore, the vLAMP reaction was completed in 40 min at 62℃, and the results were directly confirmed by the naked eye using HNB as a colorimetric indicator without additional detection equipment or processes (Fig. 1, 2). Moreover, HNB can be added to the reaction mixture before vLAMP reaction, and the results can be monitored immediately after the reaction is over without opening the lid of the reaction tube, thereby preventing DNA cross-contamination via the incorporation of pre-amplified products (Goto et al, 2009). Taken together, the developed vLAMP assay is more convenient and useful as an on-site diagnostic method than previously reported ASFV LAMP assays.

Recently, Dokphut et al (2021) also reported a colorimetric LAMP assay using the same HNB indicator as vLAMP developed in this study. The sensitivity of the Dokphut LAMP assay was 3.68×102 copies/μL, and the reaction time was 90 min. However, our newly developed vLAMP assay has much higher sensitivity (101 copies/μL) and a shorter reaction time of 40 min. These results indicated that the vLAMP assay developed in this study is superior to the Dokphut LAMP assay for use as an on-site diagnostic method requiring a rapid and sensitive ASFV diagnosis. It is unknown what caused the difference in reaction time and diagnostic sensitivity between two LAMP assays. Although various factors can be pointed out, it is most likely due to the difference in the designed primers and reaction components used in each LAMP assay. However, further analysis is needed to determine the exact cause of the difference between the assays.

To increase the global utility of the developed vLAMP assay, the primers were carefully designed by including degenerated sequences to match the target viral gene sequences of all ASFV genotypes (Table 2). The developed vLAMP assay in this study successfully detected genotype II ASFVs as well as genotype 1 and V ASFVs as shown in Table 1, 3. Considering that genotype 1 and II ASFV strains are circulating in most countries, except African countries and genotype II ASFV strains are circulating in Asian countries, including Korea, the developed vLAMP assay will be useful for ASFV detection in these countries (Quembo et al, 2018; Zhou et al, 2018; Gaudreault et al, 2020; Kim et al, 2020a). However, further studies are needed to determine whether the vLAMP assay can detect other ASFV genotypes not evaluated in this study.

In this study, the diagnostic performance of the vLAMP assay was compared to previously described cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays, which were OIE-recommended diagnostic assays and have been widely used for ASFV detection. The comparative evaluation results demonstrated that the diagnostic sensitivity of the developed vLAMP assay was higher than cPCR and was comparable to qPCR (Table 3). Taken together, the newly developed vLAMP assay will be useful for ASFV diagnosis. The percentages of positive, negative, and overall agreement of the vLAMP assay compared to the OIE-recommended qPCR assay were 100%, 100%, and 100%, respectively, with a κ value of 1.0 (Table 3), indicating that the newly developed vLAMP assay can serve as an alternative to qPCR assay for the diagnosis of ASFV infections in clinical pig samples. However, this study has a limitation in that a sufficient number of clinical samples were not provided for clinical evaluation of the vLAMP assay. Therefore, further evaluation will be needed to determine whether this assay will be useful for detecting ASFV in various clinical samples from suspected animal cases. In conclusion, the established vLAMP assay was highly specific and sensitive for detecting ASFV DNA, and amplification can be rapidly accomplished using a conventional, constant-temperature water bath without a specialized thermocycler. Furthermore, vLAMP results can be observed by the naked eye immediately after the reaction, enabling simple and direct monitoring of test results while avoiding the need for electrophoresis or other monitoring apparatuses. Taken together, these advantages suggested that the developed vLAMP assay will be a rapid, sensitive, and reliable diagnosis method for ASFV, even in resource-limited laboratories.

This work was supported by the Commercializations Promotion Agency for R&D Outcomes (COMPA) grant funded by the Korean Government(Ministry of Science and ICT) (2022).

No potential conflict of interest relevant to this article was reported.

  1. Aguero M, Fernadez J, Romero L, Sanchez C, Sanchez-Vizcaino JM. 2003. Highly sensitive PCR assay for the routine diagnosis of African swine fever virus in clinical samples. J Clin Microbiol. 41(9):4431-4434.
    Pubmed KoreaMed CrossRef
  2. Alonso C, Borca M, Dixon L, Revilla Y, Escribano JM; ICTV Report Consortium. 2018. ICTV virus taxonomy profile: Asfarviridae. J Gen Virol. 99:613-614.
    Pubmed CrossRef
  3. Atuhaire DK, Afayoa M, Ochwo S, Katiti D, Mwiine FN, Nanteza A, Mugasa CM, Matovu E, Okuni JB, Ojok L. 2014. Comparative detection of African swine fever virus by loop-mediated isothermal amplification assay and polymerase chain reaction in domestic pigs in Uganda. African J Microbiol Res. 8(23):2322-2328.
    CrossRef
  4. Bastos AD, Penrith ML, Crucière C, Edrich JL, Hutchings G, Roger F, Thomson GR. 2003. Genotyping field strains of African swine fever virus by partial p72 gene characterisation. Arch Virol. 148(4):693-706.
    Pubmed CrossRef
  5. Chae HG, Lim DR, Kim HR, Park CK. 2020. An advanced loop-mediated isothermal amplification assay for the rapid detection of beak and feather disease virus in psittacine birds. J Virol Methods 277:113819.
    Pubmed CrossRef
  6. Dhama K, Karthik K, Chakraborty S, Tiwari R, Kapoor S, Thomas P. 2014. Loop-mediated isothermal amplification of DNA (LAMP): a new diagnostic tool lights the world of diagnosis of animal and human pathogens: a review. Pak J Biol Sci. 17:151-166.
    Pubmed CrossRef
  7. Dokphut A, Boonpornprasert P, Tangdee S. 2021. Development of a loop-mediated isothermal amplification assay for rapid detection of African swine fever. Vet Integr Sci. 19(1):87-100.
    CrossRef
  8. Fernández-Pinero J, Gallardo C, Elizalde M, Robles A, Gómez C, Bishop R, Heath L, Couacy-Hymann E, Fasina FO, Pelayo V, Arias M. 2013. Molecular diagnosis of African swine fever by a new real-time PCR using universal probe library. Transbound Emerg Dis. 60(1):48-58.
    Pubmed CrossRef
  9. Gallardo C, Arias M. 2019. African swine fever (ASF) diagnosis, an essential tool in the epidemiological investigation. Virus Res. 271:197676.
    Pubmed CrossRef
  10. Gaudreault NN, Madden DW, Wilson WC, Richt JA. 2020. African swine fever virus: an emerging DNA Arbovirus. Front Vet Sci. 7:215.
    Pubmed KoreaMed CrossRef
  11. Goto M, Honda E, Ogura A, Hanaki KI. 2009. Colorimetric detection of loop-mediated isothermal amplification reaction by using hydroxy naphthol blue. Biotechniques 46:167-172.
    Pubmed CrossRef
  12. James HE, Ebert K, McGonigle R, Reid SM, Boonham N, Tomlinson JA, Hutchings GH, Denyer M, Oura CA, King DP. 2010. Detection of African swine fever virus by loop-mediated isothermal amplification. J Virol Methods 164(1-2):68-74.
    Pubmed CrossRef
  13. Jo YS and Gortázar C. 2021. African swine fever in wild boar: Assessing interventions in South Korea. Transbound Emerg Dis. 68(5):2878-2889.
    Pubmed CrossRef
  14. Kim DY, Kim HR, Park JH, Kwon NY, Kim JM, Kim JK, Park JH, Lee KK, Kim SH, Kim WI, Park CK. 2022. Detection of a novel porcine circovirus 4 in Korean pig herds using a loop-mediated isothermal amplification assay. J Virol Methods 299:114350.
    Pubmed CrossRef
  15. Kim HJ, Cho KH, Lee SK, Kim DY, Nah JJ, Kim HJ, Kim HJ, Hwang JY, Sohn HJ, Choi JG, Kim YJ. 2020a. Outbreak of African swine fever in South Korea, 2019. Transbound Emerg Dis. 67(2):473-475.
    Pubmed CrossRef
  16. Kim HJ, Cho KH, Ryu JH, Jang MK, Chae HG, Choi JD, Nah JJ, Kang HE. 2020b. Isolation and genetic characterization of African swine fever virus from domestic pig farms in South Korea, 2019. Viruses 12(11):1237.
    Pubmed KoreaMed CrossRef
  17. Kim HR, Park YR, Lim DR, Park MJ, Park JY, Kim SH, Lee KK, Park CK. 2017. Multiplex real-time polymerase chain reaction for the differential detection of porcine circovirus 2 and 3. J Virol Methods 250:11-16.
    Pubmed CrossRef
  18. Kim JK, Kim HR, Kim DY, Kim JM, Kwon NY, Park JH, Park JY, Kim SH, Lee KK, Lee C, Joo HD, Park CK. 2021. A simple colorimetric detection of porcine epidemic diarrhea virus by reverse transcription loop-mediated isothermal amplification assay using hydroxynaphthol blue metal indicator. J Virol Methods 298:114289.
    Pubmed CrossRef
  19. King DP, Reid SM, Hutchings GH, Grierson SS, Wilkinson PJ, Dixon LK, Drew TW. 2003. Development of a TaqMan PCR assay with internal amplification control for the detection of African swine fever virus. J Virol Methods. 107(1):53-61.
    Pubmed CrossRef
  20. Kwiecien R, Blettner M. 2011. Concordance analysis: Part 16 of a series on evaluation of scientific publications. Dtsch Arztebl Int. 108(30):515-521.
    Pubmed KoreaMed CrossRef
  21. Lim DR, Kim HR, Park MJ, Chae HG, Ku BK, Nah JJ, Ryoo SY, Wee SH, Park YR, Jeon HS, Kim JJ, Jeon BY, Lee HW, Park CK. 2018. An improved reverse transcription loop-mediated isothermal amplification for sensitive and specific detection of serotype O foot-and-mouth disease virus. J Virol Methods 260:6-13.
    Pubmed CrossRef
  22. Mori Y and Notomi T. 2009. Loop-mediated isothermal amplification (LAMP): a rapid, accurate, and cost-effective diagnostic method for infectious diseases. J Infect Chemother. 15:62-69.
    Pubmed KoreaMed CrossRef
  23. Notomi T, Okayama H, Masubuchi H, Yonekawa T, Watanabe K, Hase T. 2000. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res. 28:e63.
    Pubmed KoreaMed CrossRef
  24. Park CK, Kim HS. 2004. Genetic characterization of porcine circovirus 2 Korean isolates. Korean J Vet Res. 44:571-579.
  25. Park MJ, Kim HR, Chae HG, Lim DR, Kwon OD, Park CK. 2019. Development of a colorimetric loop-mediated isothermal amplification assay for rapid and specific detection of Aves polyomavirus 1 from psittacine birds. J Virol Methods 273:113687.
    Pubmed CrossRef
  26. Park YR, Kim HR, Kim SH, Lee KK, Lyoo YS, Park CK. 2018. Loop-mediated isothermal amplification assay for the rapid and visual detection of novel porcine circovirus 3. J Virol Methods 53:26-30.
    Pubmed CrossRef
  27. Quembo CJ, Jori F, Heath L. 2018. Genetic characterization of African swine fever virus isolates from soft ticks at the wildlife/domestic interface in Mozambique and identification of a novel genotype. Transbound Emerg Dis. 65(2):420-431.
    Pubmed KoreaMed CrossRef
  28. Tignon M, Gallardo C, Iscaro C, Hutet E, Van der Stede Y, Kolbasov D, De Mia GM, Le Potier MF, Bishop RP, Koenen F. 2011. Development and inter-laboratory validation study of an improved new real-time PCR assay with internal control for detection and laboratory diagnosis of African swine fever virus. J Virol Methods 178(1-2):161-170.
    Pubmed CrossRef
  29. Wang D, Yu J, Wang Y, Zhang M, Li P, Liu Y. 2020. Development of a real-time loop-mediated isothermal amplification (LAMP) assay and visual LAMP assay for detection of African swine fever virus (ASFV). J Virol Methods 276:113775.
    Pubmed CrossRef
  30. Woźniakowski G, Mazur N. 2018. Comparison of loop-mediated isothermal amplification (LAMP) and cross-priming amplification (CPA) for detection of African swine fever virus. Polish J Vet Sci. 21(4):827-830.
    Pubmed CrossRef
  31. Wu X, Xiao L, Wang Y, Yang Z, Peng B. 2016. Development of a rapid and sensitive method for detection of African swine fever virus using loop-mediated isothermal amplification. Brazilian Arch Biol Technol. 59:e16160500.
    CrossRef
  32. Yoo DS, Kim Y, Lee ES, Lim JS, Hong SK, Lee IS, Jung CS, Yoon HC, Wee SH, Fournié G. 2021. Transmission dynamics of African swine fever virus, South Korea, 2019. Emerg Infect Dis. 27(7):1909-1918.
    Pubmed KoreaMed CrossRef
  33. Zhou X, Li N, Luo Y, Liu Y, Miao F, Chen T, Zhang S, Cao P, Li X, Tian K, Hu R. 2018. Emergence of African swine fever in China, 2018. Transbound Emerg Dis. 65(6):1482-1484.
    Pubmed CrossRef
  34. Zsak L, Borca MV, Risati GR, Zsak A, French RA, Lu Z, Kutish GF, Neilan JG, Callahan JD, Rock DL. 2005. Preclinical diagnosis of African swine fever in contact-exposed swine by a real-Time PCR Assay. J Clin Microbiol. 43(1):112-119.
    Pubmed KoreaMed CrossRef

Article

Original Article

Korean J. Vet. Serv. 2022; 45(1): 19-30

Published online March 30, 2022 https://doi.org/10.7853/kjvs.2022.45.1.19

Copyright © The Korean Socitety of Veterinary Service.

Simple and rapid colorimetric detection of African swine fever virus by loop-mediated isothermal amplification assay using a hydroxynaphthol blue metal indicator

Ji-Hoon Park 1†, Hye-Ryung Kim 1†, Ha-Kyung Chae 1,3, Jonghyun Park 1,3, Bo-Young Jeon 4, Young S. Lyoo 5, Choi-Kyu Park 1*

1College of Veterinary Medicine & Animal Disease Intervention Center, Kyungpook National University, Daegu 41566, Korea
2DIVA Bio, Inc., Daegu 41519, Korea
3Korea Disease Control and Prevention Agency, Gyeongbuk Regional Center for Disease Control and Prevention, Daegu 41061, Korea
4Department of Biomedical Laboratory Science, College of Health Science, Yonsei University, Wonju 26493, Korea
5College of Veterinary Medicine, Konkuk University, Seoul 05029, Korea

Correspondence to:Choi-Kyu Park
E-mail: parkck@knu.ac.kr
https://orcid.org/0000-0002-0784-9061
These first three authors contributed equally to this work.

Received: March 23, 2022; Accepted: March 28, 2022

This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/4.0). which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

Abstract

In this study, a simple loop-mediated isothermal amplification (LAMP) combined with visual detection method (vLAMP) assay was developed for the rapid and specific detection of African swine fever virus (ASFV), overcoming the shortcomings of previously described LAMP assays that require additional detection steps or pose a cross-contamination risk. The assay results can be directly detected by the naked eye using hydroxynaphthol blue after incubation for 40 min at 62℃. The assay specifically amplified ASFV DNA and no other viral nucleic acids. The limit of detection of the assay was <50 DNA copies/reaction, which was ten times more sensitive than conventional polymerase chain reaction (cPCR) and comparable to real-time PCR (qPCR). For clinical evaluation, the ASFV detection rate of vLAMP was higher than cPCR and comparable to OIE-recommended qPCR, showing 100% concordance, with a κ value (95% confidence interval) of 1 (1.00∼1.00). Considering the advantages of high sensitivity and specificity, no possibility for cross-contamination, and being able to be used as low-cost equipment, the developed vLAMP assay will be a valuable tool for detecting ASFV from clinical samples, even in resource-limited laboratories.

Keywords: African swine fever virus, Loop-mediated isothermal amplification, Visual detection

INTRODUCTION

African swine fever (ASF) is a highly contagious transboundary swine disease caused by the ASF virus (ASFV) and is responsible for devastating economic losses in domesticated pigs and wild boar. ASFV is a large, enveloped double-stranded DNA virus, the only member of genus Asfivirus, family Asfarviridae (Alonso et al, 2018). ASFV strains have been divided into 24 genotypes based on their B646L gene, which encodes the major capsid protein p72 (Bastos et al, 2003; Quembo et al, 2018). The African continent is the endemic area with all 24 ASFV genotypes. Outside Africa, only genotype I was found in Europe, South America, and the Caribbean until genotype II ASFV was introduced in Georgia from east Africa in 2007. Since then, genotype II ASFV had quickly spread to the Russian Federation, Ukraine, and Belarus and in 2014 to the European Union Baltic States and Poland. By 2018, the infection had also spread to Belgium, Hungary, the Czech Republic, Romania, Bulgaria, Slovakia, and Serbia. In August 2018, the virus finally emerged in China and rapidly spread to many Asian countries, including Mongolia, Vietnam, Cambodia, Laos and North Korea (Democratic People’s Republic of Korea), the Philippines, and South Korea (Zhou et al, 2018; Gaudreault et al, 2020; Kim et al, 2020a).

On September 17, 2019, the first ASF outbreak in a domestic pig farm was confirmed in South Korea. On April 29, 2019, 14 cases of ASF outbreaks in domestic pig farms were diagnosed in four cities or counties (Kim et al, 2020a). Phylogenetic analysis showed that all of 14 ASFV isolates in South Korea belonged to genotype II and serogroup 8, identical to European and Chinese genotype II ASFV isolates (Kim et al, 2020b). On the other hand, the first wild boar infection case was confirmed on October 2, 2019, in a wild boar carcass found by the military in the Demilitarized Zone (DMZ). Despite implementing national control measures, ASFV-infected wild boars have been continuously identified in the border areas adjacent to DMZ. ASFV infections in wild boars are considered a risk factor for further ASF outbreaks in domestic pig farms and are becoming a major concern for the Korean Government and pig farmers (Jo and Gortázar, 2020; Yoo et al, 2021). Concerns that ASFV, continuously circulating in wild boar populations, could be transmitted into domestic pig farms has been realized with two additional ASFV outbreaks in domestic pig farms located in wild boar outbreak areas in October 2020 and May 2021.

Because there is no vaccine available, the prevention, control, and eradication of ASF are based on the implementation of appropriated surveillance and strict biosecurity measures. The success of surveillance activities depends on the availability of the most appropriate diagnostic tests. Although a number of good validated ASF diagnostic techniques are available, virus detection methods, including viral genome detection by polymerase chain reaction (PCR), viral antigen detection by antigen enzyme-linked immunosorbent assay or a direct immunofluorescence test, and virus detection using virus isolation, are vital for the rapid implementation of control measures, especially in ASFV-free areas (Gallardo et al, 2019). Currently, PCR assays, including conventional PCR (cPCR) and quantitative real-time PCR (qPCR), are considered the gold standard tests for the early detection of the disease due to their superior sensitivity, specificity, robustness, and high-throughput application to detect the ASFV genome in any kind of clinical samples from domestic pigs, wild boar, and ticks (Agüero et al, 2003; King et al, 2003; Zsak et al, 2005; Tignon et al, 2011; Fernández-Pinero et al, 2013). However, these PCR-based assays require sophisticated equipment, specialized labor, and complicated procedures to detect amplified products; thus, such methods are not suitable for on-site diagnosis in field situations or primitive laboratories in developing countries. Therefore, the development of a simple, rapid, and cost-effective assay with reliable specificity and sensitivity is imperative to detect ASFV from suspected cases.

Loop-mediated isothermal amplification (LAMP) is a valuable tool for detecting various pathogens with high sensitivity, specificity, rapidity, and simplicity (Notomi et al, 2000; Mori and Notomi, 2009; Dhama et al, 2014). Several LAMP assays have been developed for ASFV detection (James et al, 2010; Atuhaire et al, 2014; Wu et al, 2016; Woźniakowski et al, 2018; Wang et al, 2020). With regard to detection methods for assay results, previously reported ASFV LAMP assays determined the assay results by gel electrophoresis, turbidity analysis by real-time turbidimeter, DNA-intercalating dye-mediated visual detection (PicoGreen, Evergreen, or SYBR Green 1), or vertical flow visualization strip. However, detection methods using gel electrophoresis and DNA-intercalating dyes may increase the potential risk for cross-contamination of amplified LAMP products, as these methods require opening the tube lid to add reagents or perform further experiments. Also, real-time turbidity analysis, fluorescence dye detection, and vertical flow visualization strip methods require additional instruments, such as real-time turbidimeter, fluorescence detector, and visualization strip cassette, limiting the applicability of LAMP assay as a field diagnostic assay. To facilitate the practical use of LAMP assays, a simple, rapid, and contamination-free detection method must be developed (Mori and Notomi, 2009). Previously, a simple and rapid colorimetric method was applied to detect LAMP results by adding hydroxynaphthol blue (HNB) metal indicators to the pre-reaction solution, which reduced the chances of carryover contamination of the LAMP product and enabled the direct visual detection of LAMP results without additional instrumentations (Goto et al, 2009). A number of LAMP assays using HNB have been developed to detect various animal pathogens (Lim et al, 2018; Park et al, 2018; Park et al, 2019; Chae et al, 2020; Kim et al, 2021; Kim et al, 2022). However, the simple colorimetric detection method using HNB has rarely been applied to ASFV LAMP assays. Therefore, in this study, a visual LAMP (vLAMP) assay combined with visual detection methodology using HNB was developed for the simple and rapid detection of ASFV, which can be used for on-site laboratory diagnoses.

MATERIALS AND METHODS

Samples and nucleic acid extraction

To prevent the risk of unexpected transmission of ASFV, non-infectious ASFV DNA samples were obtained through various routes and used to develop and evaluate the vLAMP assay. A DNA standard was synthesized based on the previously reported ASFV sequence (ASFV-SY18 strain; GenBank accession no. MH766894) and used for the development and sensitivity evaluation of the assay. Twelve viral DNA were extracted from different ASFV genotypes, and 22 clinical DNA samples extracted from ASFV-infected pigs were kindly provided by Professor Bo-Young Jeon, who is conducting international joint research with a Russian ASFV research institute (Table 1). Additionally, 20 clinical pig samples (5 blood, 5 spleen, 5 kidney, and 5 lymph node samples) of Korean diseased pigs were collected and used as negative controls for clinical evaluation. DNA samples were allocated in small volumes and stored at −80℃ until use. To assess the specificity of vLAMP, classical swine fever virus (CSFV; strain LOM), type 1 porcine reproductive and respiratory syndrome virus (PRRSV; strain Lelystad virus), type 2 PRRSV (strain LMY), porcine parvovirus (PPV; strain NADL-2), porcine circovirus (PCV) type 1 (PCV1; from infected PK-15 cell culture), PCV type 2 (strain PCK0201; Park et al, 2004), PCV type3 (strain PCK3-1701; Kim et al, 2017), Erysipelothrix rhusiopathiae (ER; strain NL-11), and ST and PK-15 cells were obtained from the Animal and Plant Quarantine Agency (Korea) and the Animal Disease Intervention Center (Korea). The viral titers of all viruses were determined to be at least 104.0 TCID50/mL. All viral samples were allocated and stored at −80℃ until use. Total RNA or DNA was extracted from 200 μL of virus stocks, cultured cells, and clinical samples using the TAN Bead Nucleic Acid Extraction kit for automated extraction (TAN Bead, Taoyuan, Taiwan), according to manufacturer’s instructions. Extracted RNA or DNA samples were eluted in 100 μL nuclease-free water, allocated in small volumes, and stored at −80℃ until use.

Table 1 . Comparison of diagnostic results by cPCR, qPCR and vLAMP assays using ASFV-related DNA samples.

No.DNA sampleSample source (ASFV genotype)aResults of different assaysb

cPCRqPCR (Ct value)vLAMP
1Viral DNAChinese isolate SY18 (G II)+29.53+
2Viral DNARussian field isolate in Itkutsk (G II)+26.76+
3Viral DNARussian field isolate 2 in Stavropol (G II)+25.84+
4Viral DNARussian field isolate 1 in Omsk (G II)+26.71+
5Viral DNARussian field isolate 2 in Omsk (G II)+23.67+
6Viral DNARussian field isolate 1 in Nizhny Novgorod (G II)+23.26+
7Viral DNARussian field isolate 2 in Nizhny Novgorod (G II)+21.76+
8Viral DNARussian field isolate 3 in Nizhny Novgorod (G II)+25.73+
9Viral DNARussian field isolate 4 in Nizhny Novgorod (G II)+24.27+
10Viral DNARussian field isolate in Krasnodar (G II)+24.76+
11Viral DNARussian field isolate in Saratov (G II)+27.06+
12Viral DNAAttenuated ASFV KK262 strain (G I)+26.82+
13Viral DNAAttenuated ASFV MK200 strain (G V)+26.04+
14Sample DNABlood of pig 1 infected with V strain, 3 DPC (G II)+29.94+
15Sample DNABlood of pig 1 infected with V strain, 5 DPC (G II)+22.81+
16Sample DNABlood of pig 2 infected with V strain, 3 DPC (G II)+29.42+
17Sample DNABlood of pig 2 infected with V strain, 5 DPC (G II)+23.34+
18Sample DNABlood of pig 3 infected with V strain, 3 DPC (G II)+29.27+
19Sample DNABlood of pig 3 infected with V strain, 4 DPC (G II)+27.23+
20Sample DNABlood of pig 3 infected with V strain, 6 DPC (G II)+21.73+
21Sample DNABlood of pig 4 infected with V strain, 4 DPC (G II)+25.36+
22Sample DNABlood of pig 4 infected with V strain, 5 DPC (G II)+22.56+
23Sample DNABlood of pig 4 infected with V strain, 6 DPC (G II)+25.19+
24Sample DNABlood of pig 5 infected with V strain, 3 DPC (G II)+30.05+
25Sample DNABlood of pig 5 infected with V strain, 4 DPC (G II)-
26Sample DNABlood of pig 5 infected with V strain, 5 DPC (G II)+25.17+
27Sample DNABlood of pig 6 infected with K strain, 3 DPC (G 1)+25.85+
28Sample DNABlood of pig 6 infected with K strain, 5 DPC (G 1)+28.86+
29Sample DNABlood of pig 6 infected with K strain, 7 DPC (G 1)33.35+
30Sample DNABlood of pig 7 infected with K strain, 3 DPC (G 1)+25.74+
31Sample DNABlood of pig 7 infected with K strain, 5 DPC (G 1)+21.45+
32Sample DNABlood of pig 7 infected with K strain, 7 DPC (G 1)+21.06+
33Sample DNABlood of pig 8 infected with K strain, 3 DPC (G 1)+27.18+
34Sample DNABlood of pig 8 infected with K strain, 5 DPC (G 1)+21.85+
35Sample DNABlood of pig 8 infected with K strain, 7 DPC (G 1)+30.92+
36∼40Sample DNABloods of diseased Korean pigs (NC)-
41∼45Sample DNASpleens of diseased Korean pigs (NC)-
46∼50Sample DNAKidneys of diseased Korean pigs (NC)-
51∼55Sample DNALymph nodes of diseased Korean pigs (NC)-

aV strain, Russian field strain Volgograd/wb/2014; K strain, Russian virulent strain KK262; DPC, days of postchallenge; G, genotype of African swine fever virus; NC, negative control..

bThe results of the visual loop-mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) were presented as positive (+) or negative (−), and positive results of the real-time PCR (qPCR) were presented as the threshold cycle (Ct) values for each sample..



Construction of a DNA standard

Because a developing LAMP assay and reference PCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays in this study target the same ASFV B646L gene, the gene was cloned and used as a DNA standard. Based on the sequence of the Chinese genotype II ASFV strain ASFV-SY18 (GenBank accession no. MH766894), 1942-bp of the B646L gene was commercially synthesized and cloned into the pUC57 cloning vector (GenScript, Seoul, Korea) by BIONICS (Seoul, Korea). Recombinant plasmid DNA samples were linearized by digestion with EcoRI (Takara Korea Biomedical, Seoul, Korea) and purified using the Expin CleanUP SV kit (GeneAll Biotechnology, Seoul, Korea), according to the manufacturer’s instructions. DNA concentrations were determined by measuring the absorbance at 260 nm using a NanoDrop Lite spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). DNA transcript copy numbers were quantified using a previously described method (Park et al, 2018). The DNA standard was stored at −80℃.

Primers for the vLAMP assay

Because the B646L gene served as a useful diagnostic marker for OIE-approved reference cPCR and qPCR assays for ASFV, vLAMP primers were designed based on the conserved regions of the B646L gene (Agüero et al, 2003; King et al, 2003). A total of 355 complete or partial B646L gene sequences of 24 ASFV genotypes were retrieved from the National Center for Biotechnology Information (NCBI) GenBank database. Conserved nucleotide sequences within the B646L gene were identified by multiple alignments using the BioEdit Sequence Alignment Editor program (http://www.mbio.ncsu.edu/BioEdit/bioedit.html). Based on these conserved sequences, a set of six primers, including two outer primers (F3 and B3), two inner primers (FIP and BIP), and two loop primers (LF and LB), were manually designed for the vLAMP assay aided by Primer Explorer V5 software (Fujitsu System Solutions Ltd., Tokyo, Japan). The specificity of the primers for vLAMP was confirmed against random nucleotide sequences using a BLAST search of the NCBI GenBank database (http://www.ncbi.nlm.nih.gov/BLAST/). All primers were synthesized by BIONICS, as shown in Table 2.

Table 2 . Primers and probes used in vLAMP and cPCR, and qPCR in this study.

MethodPrimer and probeLength (bp)Primer/probe sequence (5’–3’)aGenome positionbReference
vLAMPF318CTCTTCCAGACGCATGTT85893∼85910This study
B318CCGTRGTGATAGACCCCA86196∼86213
LF24TGTAAGAGCTGCAGAACTTTGATG86013∼86036
LB21TTGAARCCACGGGAGGAATAC86108∼86128
FIP (F1c+F2)41GCCTCCGTAGTGRAAGGGTA+86037∼86056+
GTHACTGCTCAYGGTATCAAT85976∼85996
BIP (B1c+B2)44TCCGGGYGCGATGATGATTAC+86080∼86100+
CTTGCTCTGGATACGTTAATATG86141∼86163
PCRPPA121TTCCCAGCGTAGTTGAGATTG84467∼84486Agüero et al (2003)
PPA220AGTTATGGGAAACCCGACCC84705∼84723
qPCRF25CTGCTCATGGTATCAATCTTATCGA85980∼86004King et al (2003)
R25CTGCTCATGGTATCAATCTTATCGA86210∼86229
P25FAM–CCACGGGAGGAATACCAACCCAGTG–BHQ186114∼86138

aBold text in B3, LB, FIP, and BIP sequences used in the vLAMP assay represent a degenerate base: R, A/G; H, A/C/T; Y, C/T. FAM, 6-carboxyfluorescein; BHQ1, Black Hole Quencher 1..

bThe locations of all primer and probe sequences for the visual loop-mediated isothermal amplification (vLAMP), conventional polymerase chain reaction (cPCR) and real-time PCR (qPCR) assays were derived from the complete genome sequence of the Chinese representative African swine fever virus strain ASFV-SY18 (GenBank accession no. MH766894)..



Optimization of vLAMP conditions

vLAMP was performed in a reaction mixture containing 20 mM Tris-HCl (pH 8.8), 10mM KCl, 10mM (NH4)2SO4, 0.1% Triton X-100, 1.4 mM dNTPs, 8 mM MgSO4, 0.12 mM HNB (Lemongreen, Shanghai, China), 0.8 M betaine (Sigma–Aldrich, St. Louis, Missouri, USA), 8U Bst DNA polymerase (large fragment; New England Biolabs, Ipswich, MA, USA), 1.6 μM inner primers (FIP and BIP), 0.2 μM outer primers (F3 and B3), 0.8 μM loop primers (LF and LB), 5 μL template RNA, and dH2O to a final volume of 25 μL. The reaction conditions were optimized by performing test amplifications at temperatures ranging from 56℃ to 68℃ and reaction times of 30∼60 min. Reactions were terminated by heating at 80℃ for 5 min. All experiments were performed in triplicate. The assay results were interpreted by visual detection of a color change from purple to sky blue due to the presence of the metal ion indicator HNB (Goto et al, 2009). Amplicons were also detected by observing LAMP-specific ladder-like DNA bands using an ultraviolet light transilluminator (Bio-Rad, Hercules, CA, USA) after subjecting the DNA to 1.5% agarose gel electrophoresis and staining with NEO green dye (Neoscience, Suwon, Korea).

cPCR and qPCR assays

cPCR was performed with ASFV B646L gene-specific primers using an Excel TB 2X Taq premix (Inclone, Yongin, Korea) and an Applied Biosystems thermal cycler, as described previously (Agüero et al, 2003). The expected size of the cPCR product using the primer set (PPA1 and PPA2) was 257 bp (Table 2). The cPCR conditions were as follows: initial denaturation at 95℃ for 2 min, 40 cycles of PCR (95℃ for 20 s, 62℃ for 40 s, and 72℃ for 30 s), and final extension at 72℃ for 5 min. The cPCR products were subjected to electrophoresis on 1.5% agarose gel. The target bands were visualized by staining with NEO green dye (Neoscience) and imaged under blue light illumination (Neoscience). qPCR was performed using a commercial qPCR kit (Premix Ex Taq™ Probe qPCR, Takara Korea Biomedical) with ASFV B646L gene-specific primers and probe (Table 2), according to previously described methods (King et al, 2003). In brief, qPCR was performed in a 25 μL reaction mixture containing 12.5 μL of 2×reaction buffer, 0.5 μL of each ASFV primers (0.2 μM and probe 0.2 μM), 5 μL of DNA template, and 6 μL of water. The reaction was carried out in a CFX96 Touch™ Real-time PCR Detection System (Bio-Rad) under the following conditions: initial denaturation at 95℃ for 30 s, 40 cycles of denaturation at 95℃ for 5 s, and annealing and extension at 58℃ for 30 s. Real-time fluorescence values of the FAM-labeled probe were measured in ongoing reactions at the end of each annealing step. To interpret the qPCR results, samples producing a cycle threshold (Ct) of <40 were considered positive, whereas those with a higher Ct value (>40) were considered negative (King et al, 2003).

Specificity and sensitivity of the vLAMP assay

To test its specificity, the vLAMP assay was performed with DNA or RNA templates extracted from a genotype II ASFV strain (ASFV-SY18), seven control virus cultures (CSFV, type 1 and 2 PRRSVs, PPV, PCV1, PCV2, and PCV3), a bacterial culture (ER), and two uninfected cell cultures (ST and PK-15 cells) as negative controls. The limit of detection (LOD) of the vLAMP assay was determined using 10-fold serial dilutions of an ASFV DNA standard, ranging from 106 to 1 copies/μL. Subsequently, the LOD of the vLAMP assay was compared to cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays using the same DNA templates described above.

Comparative evaluation of the vLAMP assay

For evaluation of the vLAMP assay, 55 DNA samples including a DNA standard (103 copies/μL), 12 viral DNA extracted different ASFV strains, 22 clinical DNA samples extracted from ASFV-infected pigs, and 20 clinical DNA samples extracted from ASFV-negative pigs, were tested using the developed vLAMP assay, and the results were compared to cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays (Table 1). The concordance between the vLAMP and cPCR or qPCR results was analyzed using Cohen’s κ statistics at 95% confidence interval (CI; Kwiecien et al, 2011). When the calculated κ coefficient value (κ) was ≥0.81, the results from these assays were almost 100% concordant.

RESULTS

Optimization of the vLAMP assay

For vLAMP reaction with ASFV DNA templates, a positive color change from purple to sky blue was visually detected in the reaction tubes, and a ladder pattern of products was generated using electrophoresis at temperatures ranging from 56℃∼68℃ (Fig. 1). The color changes and electrophoretic bands were clearer at 58℃, 60℃, 62℃, and 64℃ than at 56℃, 66℃, and 68℃. Subsequently, the vLAMP assay was performed with three dilutions of DNA transcript samples from the cloned ASFV B646L gene with different copy numbers (102, 101, and 100 copies/μL) for reaction times ranging from 30 to 60 min. The color change and electrophoresis results indicated that amplification was initiated within 30 min; however, a definite positive color change and strongly amplified DNA bands were observed at 40 min for reactions with the DNA template containing 101 copies/μL of standard DNA (Fig. 2). Based on these results, the optimal reaction temperature and time for the vLAMP assay were determined to be 62℃ and 40 min, respectively. Therefore, all subsequent experiments were performed under these reaction conditions.

Figure 1. Optimal reaction temperature for the vLAMP assay to detect African swine fever virus. (A) Visual detection of vLAMP results. A color change in the reaction solution from purple to sky blue indicated a positive reaction. (B) Electrophoretic analysis of vLAMP amplification products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 7, vLAMP results at seven different reaction temperatures (56℃, 58℃, 60℃, 62℃, 64℃, 66℃, and 68℃), respectively. NC, negative control (nuclease-free water).
Figure 2. Optimal reaction time for African swine fever virus (ASFV) detection using vLAMP (A) Visual detection of vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 3, vLAMP results for ASFV DNAs (from 5×102 to 5×100 copies/reaction) at four different reaction times (30, 40, 50, and 60 min), respectively. NC, negative control (nuclease-free water).

Specificity of the vLAMP assay

After optimization of the vLAMP conditions, an ASFV strain (ASFV-SY18) and seven control virus cultures (CSFV, type 1 and 2 PRRSVs, PPV, PCV1, PCV2, and PCV3), a bacterial culture (ER), and two cell cultures (ST and PK-15 cells) were tested using vLAMP to evaluate primer specificity. vLAMP yielded positive results for ASFV strains but negative for all other porcine pathogens, cell cultures, and negative controls. Therefore, the primer set used for this assay showed high specificity for the ASFV B646L gene (Fig. 3).

Figure 3. Specificity of the vLAMP assay for African swine fever virus (ASFV). (A) Visual detection of the vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 11, vLAMP results for ASFV strains ASFV-SY18, classical swine fever virus (strain LOM), Erysipelothrix rhusiopathiae (strain NL-11), type 1 porcine reproductive and respiratory syndrome virus (PRRSV, strain Lelystad virus), type 2 PRRSV (strain LMY), porcine parvovirus (strain NADL-2), type 1 porcine circovirus (PCV1, field strain), PCV2 (strain PCK0201), PCV3 (strain PCK3-1701), ST cell and PK-15 cells, respectively. NC, negative control (nuclease-free water).

Sensitivity of vLAMP, cPCR, and qPCR assays

The sensitivity of the vLAMP assay was determined using standard DNA dilutions ranging from 106 to 100 copies/μL and compared to cPCR and qPCR assays. The LOD was determined as the lowest dilution factor in which positive reactions were obtained in all three replicates. The LOD of vLAMP (101 copies/μL) was 10-fold lower than cPCR (102 copies/μL) and comparable to qPCR (101 copies/μL), as shown in Fig. 4. These results indicated that the developed vLAMP assay was highly sensitive and will be an alternative diagnostic tool for the OIE-recommended qPCR assay (King et al, 2003).

Figure 4. Comparative sensitivities between vLAMP, cPCR, and qPCR assays for African swine fever virus (ASFV) standard DNA. (A) Amplification curves of the qPCR assay. (B) Electrophoretic analysis of cPCR amplicons. (C, D) Visualization and electrophoretic analysis of vLAMP. Lines and tubes 1 to 7, in (A∼D) represent serial 10-fold dilutions of the ASFV DNA standard (from 5×106 to 5×100 copies). Lane M, 1 kb plus DNA marker; tube and lane NC, negative control (nuclease-free water).

Comparative diagnostic performance of the vLAMP assay

To evaluate the ability of the vLAMP assay to detect ASFV DNA from viral cultures and clinical samples, 55 DNA samples (13 viral DNA and 42 clinical DNA) were tested by the vLAMP assay, and the results were compared to cPCR and qPCR assays. The detection rate of ASFV DNA from 13 viral DNA samples was 100.0% (13/13) for all detection methods. The detection rate from clinical DNA samples by vLAMP, cPCR, and qPCR were 50.0% (21/42), 47.6% (20/42), and 50.0% (21/42), respectively. A clinical DNA sample positive by vLAMP and qPCR (Ct value of 33.35) was tested by cPCR as negative. The total detection rates of vLAMP and qPCR (61.8%) were higher than cPCR (60.0%; Table 3). For ASFV detection from DNA samples, the percentages of positive and negative results and the overall agreement for the vLAMP assay relative to the cPCR assay were 97.1% (33/34), 100% (21/21), and 98.2% (54/55), with a κ value (95% CI) of 0.96 (0.92∼1.00). Relative to the qPCR assay, these values were 100% (34/34), 100% (21/21), and 100% (55/55), with a κ value (95% CI) of 1 (1.00∼1.00), indicating that the diagnostic vLAMP results were 100% concordant with the qPCR assay.

Table 3 . Comparison of vLAMP, cPCR, and qPCR results for the detection of African swine fever virus in different DNA samples.

SampleNo. testedNo positive (%)

cPCRqPCRvLAMP
Viral DNA from different ASFVs1313 (100)13 (100)13 (100)
Sample DNA from infected pigs2220 (90.9)21 (95.5)21 (95.5)
Sample DNA from uninfected pigs20000
Total5533 (60.0)34 (61.8)34 (61.8)

The calculated κ coefficient value (95% confidence interval) between visual loop–mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) or real-time PCR (qPCR) were 0.96 (0.92∼1.0), and 1 (1.0∼1.0), respectively..


DISCUSSION

Since 2019, when outbreaks in domestic pigs and wild boars were confirmed in Korea, despite national control measures executed, ASFV infections in wild boars continue to spread geographically, thereby increasing the risk of further outbreaks in domestic pig farms in wild boar outbreak areas (Jo and Gortázar, 2020; Kim et al, 2020a; Yoo et al, 2021). Considering the continuous spread of ASFV in wild boar populations and the risk of ASFV transmission associated with the movement of ASF-suspected wild boar samples, it is necessary to develop a diagnosis method that can be applied at the site where suspected domestic pigs or wild boar are found. LAMP has been recognized as a valuable tool for detecting of various animal pathogens due to its high sensitivity and specificity, rapidity, simplicity, and field applicability (Mori and Notomi, 2009; Dhama et al, 2014). For ASFV diagnosis, some LAMP assays have been reported so far, but these assays have limitations in their use as an on-field diagnostic assay mainly due to the monitoring methods of LAMP results that are difficult to apply in field situations (James et al, 2010; Atuhaire et al, 2014; Wu et al, 2016; Woźniakowski et al, 2018; Wang et al, 2020). Therefore, a vLAMP assay with high sensitivity and specificity, simplicity, rapidity, and field applicability was developed in this study.

The vLAMP assay using six primers that recognize eight regions of the ASFV B646L gene specifically detected ASFV but did not amplify any other porcine pathogens, as shown in Fig. 3. The LOD of the vLAMP assay was 10-fold lower than cPCR and comparable to qPCR (Fig. 4). Furthermore, the vLAMP reaction was completed in 40 min at 62℃, and the results were directly confirmed by the naked eye using HNB as a colorimetric indicator without additional detection equipment or processes (Fig. 1, 2). Moreover, HNB can be added to the reaction mixture before vLAMP reaction, and the results can be monitored immediately after the reaction is over without opening the lid of the reaction tube, thereby preventing DNA cross-contamination via the incorporation of pre-amplified products (Goto et al, 2009). Taken together, the developed vLAMP assay is more convenient and useful as an on-site diagnostic method than previously reported ASFV LAMP assays.

Recently, Dokphut et al (2021) also reported a colorimetric LAMP assay using the same HNB indicator as vLAMP developed in this study. The sensitivity of the Dokphut LAMP assay was 3.68×102 copies/μL, and the reaction time was 90 min. However, our newly developed vLAMP assay has much higher sensitivity (101 copies/μL) and a shorter reaction time of 40 min. These results indicated that the vLAMP assay developed in this study is superior to the Dokphut LAMP assay for use as an on-site diagnostic method requiring a rapid and sensitive ASFV diagnosis. It is unknown what caused the difference in reaction time and diagnostic sensitivity between two LAMP assays. Although various factors can be pointed out, it is most likely due to the difference in the designed primers and reaction components used in each LAMP assay. However, further analysis is needed to determine the exact cause of the difference between the assays.

To increase the global utility of the developed vLAMP assay, the primers were carefully designed by including degenerated sequences to match the target viral gene sequences of all ASFV genotypes (Table 2). The developed vLAMP assay in this study successfully detected genotype II ASFVs as well as genotype 1 and V ASFVs as shown in Table 1, 3. Considering that genotype 1 and II ASFV strains are circulating in most countries, except African countries and genotype II ASFV strains are circulating in Asian countries, including Korea, the developed vLAMP assay will be useful for ASFV detection in these countries (Quembo et al, 2018; Zhou et al, 2018; Gaudreault et al, 2020; Kim et al, 2020a). However, further studies are needed to determine whether the vLAMP assay can detect other ASFV genotypes not evaluated in this study.

In this study, the diagnostic performance of the vLAMP assay was compared to previously described cPCR (Agüero et al, 2003) and qPCR (King et al, 2003) assays, which were OIE-recommended diagnostic assays and have been widely used for ASFV detection. The comparative evaluation results demonstrated that the diagnostic sensitivity of the developed vLAMP assay was higher than cPCR and was comparable to qPCR (Table 3). Taken together, the newly developed vLAMP assay will be useful for ASFV diagnosis. The percentages of positive, negative, and overall agreement of the vLAMP assay compared to the OIE-recommended qPCR assay were 100%, 100%, and 100%, respectively, with a κ value of 1.0 (Table 3), indicating that the newly developed vLAMP assay can serve as an alternative to qPCR assay for the diagnosis of ASFV infections in clinical pig samples. However, this study has a limitation in that a sufficient number of clinical samples were not provided for clinical evaluation of the vLAMP assay. Therefore, further evaluation will be needed to determine whether this assay will be useful for detecting ASFV in various clinical samples from suspected animal cases. In conclusion, the established vLAMP assay was highly specific and sensitive for detecting ASFV DNA, and amplification can be rapidly accomplished using a conventional, constant-temperature water bath without a specialized thermocycler. Furthermore, vLAMP results can be observed by the naked eye immediately after the reaction, enabling simple and direct monitoring of test results while avoiding the need for electrophoresis or other monitoring apparatuses. Taken together, these advantages suggested that the developed vLAMP assay will be a rapid, sensitive, and reliable diagnosis method for ASFV, even in resource-limited laboratories.

ACKNOWLEDGEMENTS

This work was supported by the Commercializations Promotion Agency for R&D Outcomes (COMPA) grant funded by the Korean Government(Ministry of Science and ICT) (2022).

CONFLICT OF INTEREST

No potential conflict of interest relevant to this article was reported.

Fig 1.

Figure 1.Optimal reaction temperature for the vLAMP assay to detect African swine fever virus. (A) Visual detection of vLAMP results. A color change in the reaction solution from purple to sky blue indicated a positive reaction. (B) Electrophoretic analysis of vLAMP amplification products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 7, vLAMP results at seven different reaction temperatures (56℃, 58℃, 60℃, 62℃, 64℃, 66℃, and 68℃), respectively. NC, negative control (nuclease-free water).
Korean Journal of Veterinary Service 2022; 45: 19-30https://doi.org/10.7853/kjvs.2022.45.1.19

Fig 2.

Figure 2.Optimal reaction time for African swine fever virus (ASFV) detection using vLAMP (A) Visual detection of vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 3, vLAMP results for ASFV DNAs (from 5×102 to 5×100 copies/reaction) at four different reaction times (30, 40, 50, and 60 min), respectively. NC, negative control (nuclease-free water).
Korean Journal of Veterinary Service 2022; 45: 19-30https://doi.org/10.7853/kjvs.2022.45.1.19

Fig 3.

Figure 3.Specificity of the vLAMP assay for African swine fever virus (ASFV). (A) Visual detection of the vLAMP results. The color of the reaction solution changes from purple (negative result) to sky blue (positive result). (B) Electrophoretic analysis of vLAMP-amplified products. Lane M, 1 kb plus DNA ladder. LAMP-specific ladder-like electrophoresis patterns were observed in vLAMP-positive reactions. Tubes/lanes 1 to 11, vLAMP results for ASFV strains ASFV-SY18, classical swine fever virus (strain LOM), Erysipelothrix rhusiopathiae (strain NL-11), type 1 porcine reproductive and respiratory syndrome virus (PRRSV, strain Lelystad virus), type 2 PRRSV (strain LMY), porcine parvovirus (strain NADL-2), type 1 porcine circovirus (PCV1, field strain), PCV2 (strain PCK0201), PCV3 (strain PCK3-1701), ST cell and PK-15 cells, respectively. NC, negative control (nuclease-free water).
Korean Journal of Veterinary Service 2022; 45: 19-30https://doi.org/10.7853/kjvs.2022.45.1.19

Fig 4.

Figure 4.Comparative sensitivities between vLAMP, cPCR, and qPCR assays for African swine fever virus (ASFV) standard DNA. (A) Amplification curves of the qPCR assay. (B) Electrophoretic analysis of cPCR amplicons. (C, D) Visualization and electrophoretic analysis of vLAMP. Lines and tubes 1 to 7, in (A∼D) represent serial 10-fold dilutions of the ASFV DNA standard (from 5×106 to 5×100 copies). Lane M, 1 kb plus DNA marker; tube and lane NC, negative control (nuclease-free water).
Korean Journal of Veterinary Service 2022; 45: 19-30https://doi.org/10.7853/kjvs.2022.45.1.19

Table 1 . Comparison of diagnostic results by cPCR, qPCR and vLAMP assays using ASFV-related DNA samples.

No.DNA sampleSample source (ASFV genotype)aResults of different assaysb

cPCRqPCR (Ct value)vLAMP
1Viral DNAChinese isolate SY18 (G II)+29.53+
2Viral DNARussian field isolate in Itkutsk (G II)+26.76+
3Viral DNARussian field isolate 2 in Stavropol (G II)+25.84+
4Viral DNARussian field isolate 1 in Omsk (G II)+26.71+
5Viral DNARussian field isolate 2 in Omsk (G II)+23.67+
6Viral DNARussian field isolate 1 in Nizhny Novgorod (G II)+23.26+
7Viral DNARussian field isolate 2 in Nizhny Novgorod (G II)+21.76+
8Viral DNARussian field isolate 3 in Nizhny Novgorod (G II)+25.73+
9Viral DNARussian field isolate 4 in Nizhny Novgorod (G II)+24.27+
10Viral DNARussian field isolate in Krasnodar (G II)+24.76+
11Viral DNARussian field isolate in Saratov (G II)+27.06+
12Viral DNAAttenuated ASFV KK262 strain (G I)+26.82+
13Viral DNAAttenuated ASFV MK200 strain (G V)+26.04+
14Sample DNABlood of pig 1 infected with V strain, 3 DPC (G II)+29.94+
15Sample DNABlood of pig 1 infected with V strain, 5 DPC (G II)+22.81+
16Sample DNABlood of pig 2 infected with V strain, 3 DPC (G II)+29.42+
17Sample DNABlood of pig 2 infected with V strain, 5 DPC (G II)+23.34+
18Sample DNABlood of pig 3 infected with V strain, 3 DPC (G II)+29.27+
19Sample DNABlood of pig 3 infected with V strain, 4 DPC (G II)+27.23+
20Sample DNABlood of pig 3 infected with V strain, 6 DPC (G II)+21.73+
21Sample DNABlood of pig 4 infected with V strain, 4 DPC (G II)+25.36+
22Sample DNABlood of pig 4 infected with V strain, 5 DPC (G II)+22.56+
23Sample DNABlood of pig 4 infected with V strain, 6 DPC (G II)+25.19+
24Sample DNABlood of pig 5 infected with V strain, 3 DPC (G II)+30.05+
25Sample DNABlood of pig 5 infected with V strain, 4 DPC (G II)-
26Sample DNABlood of pig 5 infected with V strain, 5 DPC (G II)+25.17+
27Sample DNABlood of pig 6 infected with K strain, 3 DPC (G 1)+25.85+
28Sample DNABlood of pig 6 infected with K strain, 5 DPC (G 1)+28.86+
29Sample DNABlood of pig 6 infected with K strain, 7 DPC (G 1)33.35+
30Sample DNABlood of pig 7 infected with K strain, 3 DPC (G 1)+25.74+
31Sample DNABlood of pig 7 infected with K strain, 5 DPC (G 1)+21.45+
32Sample DNABlood of pig 7 infected with K strain, 7 DPC (G 1)+21.06+
33Sample DNABlood of pig 8 infected with K strain, 3 DPC (G 1)+27.18+
34Sample DNABlood of pig 8 infected with K strain, 5 DPC (G 1)+21.85+
35Sample DNABlood of pig 8 infected with K strain, 7 DPC (G 1)+30.92+
36∼40Sample DNABloods of diseased Korean pigs (NC)-
41∼45Sample DNASpleens of diseased Korean pigs (NC)-
46∼50Sample DNAKidneys of diseased Korean pigs (NC)-
51∼55Sample DNALymph nodes of diseased Korean pigs (NC)-

aV strain, Russian field strain Volgograd/wb/2014; K strain, Russian virulent strain KK262; DPC, days of postchallenge; G, genotype of African swine fever virus; NC, negative control..

bThe results of the visual loop-mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) were presented as positive (+) or negative (−), and positive results of the real-time PCR (qPCR) were presented as the threshold cycle (Ct) values for each sample..


Table 2 . Primers and probes used in vLAMP and cPCR, and qPCR in this study.

MethodPrimer and probeLength (bp)Primer/probe sequence (5’–3’)aGenome positionbReference
vLAMPF318CTCTTCCAGACGCATGTT85893∼85910This study
B318CCGTRGTGATAGACCCCA86196∼86213
LF24TGTAAGAGCTGCAGAACTTTGATG86013∼86036
LB21TTGAARCCACGGGAGGAATAC86108∼86128
FIP (F1c+F2)41GCCTCCGTAGTGRAAGGGTA+86037∼86056+
GTHACTGCTCAYGGTATCAAT85976∼85996
BIP (B1c+B2)44TCCGGGYGCGATGATGATTAC+86080∼86100+
CTTGCTCTGGATACGTTAATATG86141∼86163
PCRPPA121TTCCCAGCGTAGTTGAGATTG84467∼84486Agüero et al (2003)
PPA220AGTTATGGGAAACCCGACCC84705∼84723
qPCRF25CTGCTCATGGTATCAATCTTATCGA85980∼86004King et al (2003)
R25CTGCTCATGGTATCAATCTTATCGA86210∼86229
P25FAM–CCACGGGAGGAATACCAACCCAGTG–BHQ186114∼86138

aBold text in B3, LB, FIP, and BIP sequences used in the vLAMP assay represent a degenerate base: R, A/G; H, A/C/T; Y, C/T. FAM, 6-carboxyfluorescein; BHQ1, Black Hole Quencher 1..

bThe locations of all primer and probe sequences for the visual loop-mediated isothermal amplification (vLAMP), conventional polymerase chain reaction (cPCR) and real-time PCR (qPCR) assays were derived from the complete genome sequence of the Chinese representative African swine fever virus strain ASFV-SY18 (GenBank accession no. MH766894)..


Table 3 . Comparison of vLAMP, cPCR, and qPCR results for the detection of African swine fever virus in different DNA samples.

SampleNo. testedNo positive (%)

cPCRqPCRvLAMP
Viral DNA from different ASFVs1313 (100)13 (100)13 (100)
Sample DNA from infected pigs2220 (90.9)21 (95.5)21 (95.5)
Sample DNA from uninfected pigs20000
Total5533 (60.0)34 (61.8)34 (61.8)

The calculated κ coefficient value (95% confidence interval) between visual loop–mediated isothermal amplification (vLAMP) and conventional polymerase chain reaction (cPCR) or real-time PCR (qPCR) were 0.96 (0.92∼1.0), and 1 (1.0∼1.0), respectively..


References

  1. Aguero M, Fernadez J, Romero L, Sanchez C, Sanchez-Vizcaino JM. 2003. Highly sensitive PCR assay for the routine diagnosis of African swine fever virus in clinical samples. J Clin Microbiol. 41(9):4431-4434.
    Pubmed KoreaMed CrossRef
  2. Alonso C, Borca M, Dixon L, Revilla Y, Escribano JM; ICTV Report Consortium. 2018. ICTV virus taxonomy profile: Asfarviridae. J Gen Virol. 99:613-614.
    Pubmed CrossRef
  3. Atuhaire DK, Afayoa M, Ochwo S, Katiti D, Mwiine FN, Nanteza A, Mugasa CM, Matovu E, Okuni JB, Ojok L. 2014. Comparative detection of African swine fever virus by loop-mediated isothermal amplification assay and polymerase chain reaction in domestic pigs in Uganda. African J Microbiol Res. 8(23):2322-2328.
    CrossRef
  4. Bastos AD, Penrith ML, Crucière C, Edrich JL, Hutchings G, Roger F, Thomson GR. 2003. Genotyping field strains of African swine fever virus by partial p72 gene characterisation. Arch Virol. 148(4):693-706.
    Pubmed CrossRef
  5. Chae HG, Lim DR, Kim HR, Park CK. 2020. An advanced loop-mediated isothermal amplification assay for the rapid detection of beak and feather disease virus in psittacine birds. J Virol Methods 277:113819.
    Pubmed CrossRef
  6. Dhama K, Karthik K, Chakraborty S, Tiwari R, Kapoor S, Thomas P. 2014. Loop-mediated isothermal amplification of DNA (LAMP): a new diagnostic tool lights the world of diagnosis of animal and human pathogens: a review. Pak J Biol Sci. 17:151-166.
    Pubmed CrossRef
  7. Dokphut A, Boonpornprasert P, Tangdee S. 2021. Development of a loop-mediated isothermal amplification assay for rapid detection of African swine fever. Vet Integr Sci. 19(1):87-100.
    CrossRef
  8. Fernández-Pinero J, Gallardo C, Elizalde M, Robles A, Gómez C, Bishop R, Heath L, Couacy-Hymann E, Fasina FO, Pelayo V, Arias M. 2013. Molecular diagnosis of African swine fever by a new real-time PCR using universal probe library. Transbound Emerg Dis. 60(1):48-58.
    Pubmed CrossRef
  9. Gallardo C, Arias M. 2019. African swine fever (ASF) diagnosis, an essential tool in the epidemiological investigation. Virus Res. 271:197676.
    Pubmed CrossRef
  10. Gaudreault NN, Madden DW, Wilson WC, Richt JA. 2020. African swine fever virus: an emerging DNA Arbovirus. Front Vet Sci. 7:215.
    Pubmed KoreaMed CrossRef
  11. Goto M, Honda E, Ogura A, Hanaki KI. 2009. Colorimetric detection of loop-mediated isothermal amplification reaction by using hydroxy naphthol blue. Biotechniques 46:167-172.
    Pubmed CrossRef
  12. James HE, Ebert K, McGonigle R, Reid SM, Boonham N, Tomlinson JA, Hutchings GH, Denyer M, Oura CA, King DP. 2010. Detection of African swine fever virus by loop-mediated isothermal amplification. J Virol Methods 164(1-2):68-74.
    Pubmed CrossRef
  13. Jo YS and Gortázar C. 2021. African swine fever in wild boar: Assessing interventions in South Korea. Transbound Emerg Dis. 68(5):2878-2889.
    Pubmed CrossRef
  14. Kim DY, Kim HR, Park JH, Kwon NY, Kim JM, Kim JK, Park JH, Lee KK, Kim SH, Kim WI, Park CK. 2022. Detection of a novel porcine circovirus 4 in Korean pig herds using a loop-mediated isothermal amplification assay. J Virol Methods 299:114350.
    Pubmed CrossRef
  15. Kim HJ, Cho KH, Lee SK, Kim DY, Nah JJ, Kim HJ, Kim HJ, Hwang JY, Sohn HJ, Choi JG, Kim YJ. 2020a. Outbreak of African swine fever in South Korea, 2019. Transbound Emerg Dis. 67(2):473-475.
    Pubmed CrossRef
  16. Kim HJ, Cho KH, Ryu JH, Jang MK, Chae HG, Choi JD, Nah JJ, Kang HE. 2020b. Isolation and genetic characterization of African swine fever virus from domestic pig farms in South Korea, 2019. Viruses 12(11):1237.
    Pubmed KoreaMed CrossRef
  17. Kim HR, Park YR, Lim DR, Park MJ, Park JY, Kim SH, Lee KK, Park CK. 2017. Multiplex real-time polymerase chain reaction for the differential detection of porcine circovirus 2 and 3. J Virol Methods 250:11-16.
    Pubmed CrossRef
  18. Kim JK, Kim HR, Kim DY, Kim JM, Kwon NY, Park JH, Park JY, Kim SH, Lee KK, Lee C, Joo HD, Park CK. 2021. A simple colorimetric detection of porcine epidemic diarrhea virus by reverse transcription loop-mediated isothermal amplification assay using hydroxynaphthol blue metal indicator. J Virol Methods 298:114289.
    Pubmed CrossRef
  19. King DP, Reid SM, Hutchings GH, Grierson SS, Wilkinson PJ, Dixon LK, Drew TW. 2003. Development of a TaqMan PCR assay with internal amplification control for the detection of African swine fever virus. J Virol Methods. 107(1):53-61.
    Pubmed CrossRef
  20. Kwiecien R, Blettner M. 2011. Concordance analysis: Part 16 of a series on evaluation of scientific publications. Dtsch Arztebl Int. 108(30):515-521.
    Pubmed KoreaMed CrossRef
  21. Lim DR, Kim HR, Park MJ, Chae HG, Ku BK, Nah JJ, Ryoo SY, Wee SH, Park YR, Jeon HS, Kim JJ, Jeon BY, Lee HW, Park CK. 2018. An improved reverse transcription loop-mediated isothermal amplification for sensitive and specific detection of serotype O foot-and-mouth disease virus. J Virol Methods 260:6-13.
    Pubmed CrossRef
  22. Mori Y and Notomi T. 2009. Loop-mediated isothermal amplification (LAMP): a rapid, accurate, and cost-effective diagnostic method for infectious diseases. J Infect Chemother. 15:62-69.
    Pubmed KoreaMed CrossRef
  23. Notomi T, Okayama H, Masubuchi H, Yonekawa T, Watanabe K, Hase T. 2000. Loop-mediated isothermal amplification of DNA. Nucleic Acids Res. 28:e63.
    Pubmed KoreaMed CrossRef
  24. Park CK, Kim HS. 2004. Genetic characterization of porcine circovirus 2 Korean isolates. Korean J Vet Res. 44:571-579.
  25. Park MJ, Kim HR, Chae HG, Lim DR, Kwon OD, Park CK. 2019. Development of a colorimetric loop-mediated isothermal amplification assay for rapid and specific detection of Aves polyomavirus 1 from psittacine birds. J Virol Methods 273:113687.
    Pubmed CrossRef
  26. Park YR, Kim HR, Kim SH, Lee KK, Lyoo YS, Park CK. 2018. Loop-mediated isothermal amplification assay for the rapid and visual detection of novel porcine circovirus 3. J Virol Methods 53:26-30.
    Pubmed CrossRef
  27. Quembo CJ, Jori F, Heath L. 2018. Genetic characterization of African swine fever virus isolates from soft ticks at the wildlife/domestic interface in Mozambique and identification of a novel genotype. Transbound Emerg Dis. 65(2):420-431.
    Pubmed KoreaMed CrossRef
  28. Tignon M, Gallardo C, Iscaro C, Hutet E, Van der Stede Y, Kolbasov D, De Mia GM, Le Potier MF, Bishop RP, Koenen F. 2011. Development and inter-laboratory validation study of an improved new real-time PCR assay with internal control for detection and laboratory diagnosis of African swine fever virus. J Virol Methods 178(1-2):161-170.
    Pubmed CrossRef
  29. Wang D, Yu J, Wang Y, Zhang M, Li P, Liu Y. 2020. Development of a real-time loop-mediated isothermal amplification (LAMP) assay and visual LAMP assay for detection of African swine fever virus (ASFV). J Virol Methods 276:113775.
    Pubmed CrossRef
  30. Woźniakowski G, Mazur N. 2018. Comparison of loop-mediated isothermal amplification (LAMP) and cross-priming amplification (CPA) for detection of African swine fever virus. Polish J Vet Sci. 21(4):827-830.
    Pubmed CrossRef
  31. Wu X, Xiao L, Wang Y, Yang Z, Peng B. 2016. Development of a rapid and sensitive method for detection of African swine fever virus using loop-mediated isothermal amplification. Brazilian Arch Biol Technol. 59:e16160500.
    CrossRef
  32. Yoo DS, Kim Y, Lee ES, Lim JS, Hong SK, Lee IS, Jung CS, Yoon HC, Wee SH, Fournié G. 2021. Transmission dynamics of African swine fever virus, South Korea, 2019. Emerg Infect Dis. 27(7):1909-1918.
    Pubmed KoreaMed CrossRef
  33. Zhou X, Li N, Luo Y, Liu Y, Miao F, Chen T, Zhang S, Cao P, Li X, Tian K, Hu R. 2018. Emergence of African swine fever in China, 2018. Transbound Emerg Dis. 65(6):1482-1484.
    Pubmed CrossRef
  34. Zsak L, Borca MV, Risati GR, Zsak A, French RA, Lu Z, Kutish GF, Neilan JG, Callahan JD, Rock DL. 2005. Preclinical diagnosis of African swine fever in contact-exposed swine by a real-Time PCR Assay. J Clin Microbiol. 43(1):112-119.
    Pubmed KoreaMed CrossRef
KJVS
Dec 30, 2022 Vol.45 No.4, pp. 249~342

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