Molecular detection and seroconversion dynamics of lumpy skin disease virus in natural infected cattle herds

Introduction

Lumpy skin disease (LSD) is caused by the lumpy skin disease virus (LSDV), a member of the genus Capripoxvirus within the family Poxviridae and subfamily Chordopoxvirinae. There is only one serotype of LSDV, the Neethling virus (the prototype strain), which is antigenically closely related to the sheep and goat poxviruses (Guyassa, 2022) [9] . In India, LSD is an emerging disease that was first reported in Odisha in 2019 (Kumar et al., 2021) [13] . This initial outbreak was followed by a devastating second wave in 2022, causing the mortality of approximately 1.5 lakh (150,000) animals nationwide and inflicting severe economic losses. Official reports indicate that approximately 29 lakh cattle were affected, with 1.5 lakh deaths recorded in India during 2022 (DAHD, 2022). The significant economic impact stems from severe emaciation, hide damage, infertility in both males and females, mastitis, a drastic drop in milk production, and abortions (Tuppurainen and Oura, 2012) [21] . The LSD virus spreads among clinically and subclinically infected cattle primarily through insect vectors, including mosquitoes (Anopheles stephensi, Aedes aegypti, Culex quinquefasciatus) and biting midges (Culicoides nubeculosus), and less commonly through Ixodid hard ticks like Rhipicephalus appendiculatus (Tuppurainen et al., 2013) [22] . Following its initial emergence in West Bengal and Odisha during the summer of 2019, the disease rapidly expanded its geographical footprint across India, spreading to the northeastern, northern, southern, and western regions between 2019 and 2022. While a preliminary diagnosis of LSD is often based on characteristic nodular skin lesions, accurate diagnostic confirmation is mandatory for implementing effective control measures (EFSA Panel on Animal Health and Welfare, 2015) [4] . Furthermore, mild and subclinical forms of the disease require rapid and reliable laboratory testing. Laboratory diagnosis of LSD encompasses virus isolation in cell cultures, molecular identification techniques such as polymerase chain reaction (PCR)-based assays, detection of viral antigens via fluorescent antibody tests, and the detection of specific antibodies using serological assays (El-Nahas et al., 2011) [6] . Confirmatory testing is primarily achieved through real-time or conventional PCR assays specific to capripoxviruses (Alemayehu et al., 2013) [1] . Recent investigations evaluating the efficacy of these diagnostic tools emphasize that the choice of clinical sample drastically affects detection rates; for instance, PCR analysis reveals significantly higher positivity in tissue samples and scabs compared to nasal swabs or whole blood (Gupta et al., 2025) [8] . Furthermore, combining molecular detection with serological tools like ELISA provides a more comprehensive epidemiological picture of both active infection and prior exposure (Gayal et al., 2022) [7] . Given these diagnostic variables, this study aimed to investigate the effectiveness of molecular and serological methods, as well as the optimal choice of samples, for the confirmation of LSDV outbreaks in cattle and buffaloes.

Material and Methods

Study area and sample collection

Between September 2022 and October 2023, field officers from the Uttar Pradesh Department of Animal Husbandry collected clinical samples from cattle exhibiting symptoms of Lumpy Skin Disease (LSD) across nine districts of the state (Aligarh, Auraiya, Bahraich, Balrampur, Hamirpur Hathras, Mahoba, and Mathura Varanasi). A total of 192 of these archived specimens were retrieved from the Department of Veterinary Microbiology repository for the current investigation. The study was approved with approval no. IAEC/24/I/21-5/03/24 by the institute animal Ethics Committee Uttar Pradesh Pandit Deen dayal Upadhyaya Pashu Chikitsa Vigyan Vishwavidyalaya Evam Go-Anusandhan Sansthan (DUVASU), Mathura UP, India registered with Committee for the purpose of control and Supervision of Experiment on animals (CPCSEA).

Sample processing and DNA extraction

During the defined outbreak period, a total of 273 samples were collected for evaluation, consisting of skin scabs and intranodular saline lavage (n=72), unclotted whole blood (n=78), nasal swabs (n=42), and serum (n=81). Whole blood sample (81) was aseptically collected via jugular venepuncture into EDTA and clot activator vacutainers (Becton-Dickenson). Whole blood samples were initially diluted in PBS at a ratio of 1:1 and later used for DNA extraction. Nasal swabs (n=42) were obtained from affected cattle using sterile swabs. Tubes containing nasal swab samples in PBS were vigorously agitated in a vortex mixer before centrifugation at 3000 x g for 10 min at 4 •C. The clear supernatant fluid was collected and kept at -20 •C for future processing. Nodular tissue samples were homogenized in sterile phosphate buffer saline, subsequently tissue suspension was prepared further centrifuge at 3000 rpm for 10 min and supernatant was collected. Intranodular saline lavage samples were used directly in the DNA extraction procedure. The DNA isolation was carried out using the QIAGEN QIAamp DNA Mini Kit following the manufacturer's instructions. The DNA samples were stored at -20℃ till use in amplification.

Molecular detection of LSDV genome in clinical samples

by PCR: DNA was amplified utilizing capripoxvirus viral attachment protein (p32) gene primer EEV-F-5′-TTTCCTGATTTTTCTTACTAT-3 and EEV-R-5′-AAATTATATACGTAAATAAC-3 (Ireland and Binepal, 1998) [10] . The PCR reactions were prepared in a total volume of 25 μL, consisting of 12.5 μL of 2X EmeraldAmp MAX HS PCR Master Mix (Takara Bio Inc.), 1 μL (10 pmol) of each forward and reverse primer, 5.5 μL of nuclease-free water, and 5 μL of template DNA. The PCR assays were performed in a thermal cycler (Veriti, Applied Biosystems) with the following cycling conditions: an initial denaturation at 94 °C for 5 minutes, followed by 35 cycles of 94 °C for 1 minute, 50 °C for 30 seconds, and 72 °C for 1 minute, with a final extension at 72 °C for 5 minutes. The PCR product were analysed in 1.5 % agarose gel electrophoresis. The gel was visualized under the gel documentation system

Serological detection to estimate anti-LSDV antibodies in serum samples

Anti-LSDV antibodies in serum samples were detected using the ID Screen Capripox Double Antigen Multi-species ELISA kit (Innovative Diagnostics, France) strictly following the manufacturer's instructions. Briefly, 50 μL of test sera and controls were added to microplate wells preloaded with 50 μL of dilution buffer and incubated for 90 minutes at room temperature. Following five washes (300 μL/well), 100 μL of the prepared 1X conjugate was added to each well and incubated for 30 minutes at room temperature. Plates were washed five additional times before the addition of 100 μL of substrate solution. After a 15-minute incubation, the reaction was halted with 100 μL of stop solution. Optical density (OD) was measured at 450 nm. Samples exhibiting a sample-to-positive (S/P) percentage of ≥30% were classified as positive for anti-LSDV antibodies.

Results

A comprehensive screening was performed on a total of 192 samples, which included tissue or nodular lavage samples (n=72), nasal swabs (n=42), and whole blood specimens (n=78). The detection of Lumpy Skin Disease Virus (LSDV) was achieved by targeting the viral attachment protein gene using the OIE-recommended primers in a conventional polymerase chain reaction (PCR) assay. A positive result was confirmed by the specific amplification of a 192 bp amplicon, as illustrated in Fig. 1 The detection of anti-LSDV antibodies was strictly dependent on the clinical matrix evaluated (Fisher's exact test, p<0.0001). The target was detected in 100% of tissue/nodular lavage samples (72/72; 95% CI: 95.0-100%). In stark contrast, no positivity was observed in any of the evaluated non-invasive matrices, including nasal swabs (0/42; 95% CI: 0.0-8.4%) or whole blood samples (0/78; 95% CI: 0.0-4.7%). A total of 31 samples met the criteria for positivity (S/P% ≥ 30%). Stratification of these positive sera by antibody titer revealed that the majority of the cohort mounted a robust humoral response. Specifically, 74.2% of the seropositive samples exhibited either high (S/P% > 100%; n=13, 41.9%) or moderate (S/P% 50-100%; n=10, 32.3%) seroconversion. Only a minority of the samples (n=8, 25.8%) fell into the weak seroconversion bracket (S/P% 30-50%) Fig. 2.

Discussion

Lumpy Skin Disease, by virtue of being a transboundary animal disease, harboured and realised the potential to spread across the globe inflicting substantial losses as well as leading to negative public health outcomes affecting the Indian economy. The present study provides compelling evidence that the successful detection of Lumpy Skin Disease Virus (LSDV) is profoundly dependent on the clinical matrix selected for evaluation. The absolute detection rate (100%) in tissue and nodular lavage samples, contrasted against the complete absence of positivity (0%) in non-invasive matrices such as whole blood and nasal swabs, underscores a critical diagnostic threshold. This stark dichotomy (p<0.0001) aligns with the established pathogenesis of LSDV, which exhibits a strong tropism for cutaneous tissues and endothelial cells of the skin (Babiuk et al. 2008) [2] . While viremia is a known phase of LSDV infection, it is characteristically transient, typically peaking during the early febrile stage before declining rapidly. The failure to detect the target in whole blood and nasal swabs strongly suggests that these matrices are unreliable for field surveillance unless sampling is strictly timed to the acute, early stages of infection. These findings emphasize the suitability of scab/nodular lavage over whole blood samples, as the latter are only effective during the early viremic phase of infection. Previous investigations have highlighted the low occurrence of LSDV detection in whole blood and nasal swabs seen in this research (Khoo et [11,13,18] . LSDV in the bloodstream reaches its zenith at 9 days post-infection, following which the virus often vanishes from the blood (Babiuk et al., 2008) [2] . According to Carn and Kitching (1995) [3] , the viral concentration during viremia is greater in the nodules than it is in the blood hence, skin nodule biopsy samples are preferred for PCR-based detection in clinically infected animals. According to Parvin et al. (2022) [16] , LSD virus DNA was detected in 100% of skin lesions. According to Sanganagouda et al. (2023) [17] , the detection rates were 14.28% in blood samples, 95.04% in skin swabs, and 66.66% in nasal swabs. Previous studies have reported varying detection rates of LSDV in blood samples from suspected animals: 31.81% in India (Sudhakar et al., 2020) [20] , 13% in Russia (Sprygin et al., 2018) [19] , 40% in Azerbaijan (Zeynalova et al., 2016) [25] , and 72.7% in Serbia (Manic et al., 2019). The viremic phase in LSDV-infected animals is relatively short, with blood samples typically testing PCR-positive 4-11 day's post-infection. Zeedan et al. (2024) [24] reported positive detection rates of 56.6% from cattle skin biopsies and 11.11% from blood samples using mPCR and RT-qPCR methods. The findings align with previous studies, which report higher detection rates in tissue samples compared to blood and nasal swabs, due to the short viremic phase in infected animals. The serological data further corroborates the matrixdependent findings and provides crucial context regarding the infection timeline of the sampled population. Among the seropositive cohort, nearly three-quarters of the animals (74.2%) exhibited a robust humoral response, characterized by moderate to high seroconversion (S/P%≥ 50%). The predominance of high-titer antibodies indicates that the majority of these animals had progressed past the acute phase and were well into the post-acute or convalescent stages of infection. This mature IgG response effectively explains the concurrent clearance of the virus from the systemic circulation (resulting in 0% blood positivity), while the virus or viral antigens remained sequestered in the localized cutaneous lesions (resulting in 100% nodular lavage positivity). A previous study by Krešić et al. (2020) [12] found that 85.91% of serum samples tested positive for LSDV antibodies, while 14.09% tested negative, based on a total of 291 bovine serum samples. In the study conducted by Walid et al. 2010 [23] Antibodies against LSDV were identified in 56% of the clinically infected cows using indirect ELISA (iELISA) using the ID Screen® Capripox Double Antigen Multi-species ELISA kit. Its sensitivity in cattle ranges from 35.5% to 75%. Zeedan et al. (2024) [24] reported a 54.8% positive detection rate for LSDV antibodies in cattle using EP-ELISA. The variable detection rate of LSDV antibodies in symptomatic animals can be attributed to the timing of sample collection. Antibody levels might not have risen to detectable levels if samples were taken too early in the course of infection, before a robust immune response developed, or after the immune response had waned. These findings carry significant implications for the optimization of diagnostic protocols during active field outbreaks. Routine surveillance often relies on blood collection or nasal swabbing due to their minimally invasive nature and ease of collection. However, our data unequivocally demonstrate that such non-invasive sampling can lead to a high rate of false-negative diagnostic interpretations, particularly in herds where the infection has progressed beyond the initial viremic window. To minimize diagnostic failures and ensure the accurate molecular and serological confirmation of LSDV, veterinary field personnel must prioritize the collection of skin scabs and intranodular lavage.

References

1. Alemayehu G, Zewde G, Admassu B. Risk assessments of lumpy skin disease in Borena bull market chain and its implication for livelihoods and international trade. Trop Anim Health Prod. 2013;45:1153-1159.
2. Babiuk S, Bowden TR, Boyle DB, Wallace DB, Kitching RP. Capripoxviruses: an emerging worldwide threat to sheep, goats and cattle. Transbound Emerg Dis. 2008;55(7):263-272.
3. Carn VM, Kitching RP. The clinical response of cattle following infection with lumpy skin disease (Neethling) virus. Arch Virol. 1995;140:503-513.
4. EFSA Panel on Animal Health and Welfare (AHAW). Scientific opinion on lumpy skin disease. EFSA J. 2015;13(1):3986.
5. Government of India, Ministry of Fisheries, Animal Husbandry and Dairying, Department of Animal Husbandry and Dairying. F. No. 25-I(08)/2022AHD/AR. New Delhi: Government of India; 2022.
6. El-Nahas EM, El-Habbaa AS, El-Bagoury GF, Radwan ME. Isolation and identification of lumpy skin disease virus from naturally infected buffaloes at Kaluobia, Egypt. Glob Vet. 2011;7(3):234-237.
7. Gayal SD, Sakhare MP, Suman P, Shafi TA, Siddiqui MFMF, Yeotikar PV. Molecular detection and seroprevalence of lumpy skin disease in cattle. Indian J Anim Res. 2022. doi:10.18805/ijar.b-4933.
8. Gupta V, Pravalika A, Pandey MK, Mareddy V, Jain AK, Singh A, et al. Molecular detection, seroprevalence and biochemical analysis of lumpy skin disease virus. Viruses. 2025;17(3):293. doi:10.3390/v17030293.
9. Guyassa C. Epidemiology and diagnostic methods of lumpy skin disease: a short review. Int J Vet Sci Res. 2022;8(2):064-070.
10. Ireland DC, Binepal YS. Improved detection of capripoxvirus in biopsy samples by PCR. J Virol Methods. 1998;74(1):1-7.
11. Khoo CK, Dahlan, et al. Molecular detection of lumpy skin disease virus in Malaysia 2021. Int J Infect Dis. 2022;116:S64.
12. Krešić N, Šimić I, Bedeković T, Acinger-Rogić Ž, Lojkić I. Evaluation of serological tests for detection of International Journal of Advanced Biochemistry Research https://www.biochemjournal.com antibodies against lumpy skin disease virus. J Clin Microbiol. 2020;58(9):e01028-20.
13. Kumar N, Chander, et al. Isolation and characterization of lumpy skin disease virus from cattle in India. PLoS One. 2021;16:e0241022.
14. Li L, Qi C, Li J, et al. Quantitative real-time PCR detection and analysis of a lumpy skin disease outbreak in Inner Mongolia Autonomous Region, China. Front Vet Sci. 2022;9:936581.
15. Manić M, Stojiljković M, Petrović M, Nišavić J, Bacić D, Petrović T, et al. Epizootic features and control measures for lumpy skin disease in south-east Serbia in 2016. Transbound Emerg Dis. 2019;66(5):2087-2099.
16. Parvin R, Chowdhury, et al. Clinical epidemiology, pathology, and molecular investigation of lumpy skin disease outbreaks in Bangladesh during 2020-2021 indicate the re-emergence of an old African strain. Viruses. 2022;14(11):2529.
17. Sanganagouda K, Nagraja, et al. Molecular characterization, phylogenetic analysis and viral load quantification of lumpy skin disease virus in cattle. 2023.
18. Sariya L, Paungpin, et al. Molecular detection and characterization of lumpy skin disease viruses from outbreaks in Thailand in 2021. Transbound Emerg Dis. 2022;69(5):e2145-e2152.
19. Sprygin A, Babin Y, Pestova Y, Kononova S, Wallace DB, Van Schalkwyk A, et al. Analysis and insights into recombination signals in lumpy skin disease virus recovered in the field. PLoS One. 2018;13:e0207480.
20. Sudhakar SB, Mishra N, Kalaiyarasu S, Jhade SK, Hemadri D, Sood R, et al. Lumpy skin disease outbreaks in cattle in Odisha state, India in August 2019: epidemiological features and molecular studies. Transbound Emerg Dis. 2020;67(6):2408-2422.
21. Tuppurainen ESM, Oura CAL. Lumpy skin disease: an emerging threat to Europe, the Middle East and Asia. Transbound Emerg Dis. 2012;59(1):40-48.
22. Tuppurainen ESM, Lubinga JC, Stoltsz WH, Troskie M, Carpenter ST, Coetzer JA, et al. Mechanical transmission of lumpy skin disease virus by Rhipicephalus appendiculatus male ticks. Epidemiol Infect. 2013;141(2):425-430.
23. Walid SA, Adel KI, Khaled M, Khaled MF, Mervet IAM. Evaluation of different diagnostic methods for diagnosis of lumpy skin disease in cows. Trop Anim Health Prod. 2010;42(4):777-783.
24. Zeedan GS, Abdalhamed AM, Allam AM, Abdel-Shafy S. Molecular detection of lumpy skin disease virus in naturally infected cattle and buffaloes: unveiling the role of tick vectors in disease spread. Vet Res Commun. 2024;48(6):3921-3939.
25. Zeynalova S, Asadov K, Guliyev F, Vatani M, Aliyev V. Epizootology and molecular diagnosis of lumpy skin disease among livestock in Azerbaijan. Front Microbiol. 2016;7:1022.