On November 4, 2025, a research team from the National Centre for Exotic Animal Diseases of the Canadian Food Inspection Agency published a new study in the journal *Viruses*, reporting the distribution dynamics of the viral genome in peripheral lymphoid organs of pigs after oral-nasal infection with a moderately virulent African swine fever virus strain.
Research Highlights
* For the first time, this study systematically depicted the temporal-spatial distribution of the viral genome of attenuated African swine fever strains Estonia 2014 (genotype II) and Malta’78 (genotype I) in peripheral lymphoid organs of pigs after oral-nasal infection, comparing the differences between the two strains. Estonia 2014 was detected earlier and resulted in faster pig mortality, while Malta’78 had a longer survival period.
* This study confirmed that the viral genome can be detected in superficial inguinal lymph nodes (SILN) as early as 2-3 days after infection, reaching a peak at 5-9 days, and is highly synchronized with the viral load in the spleen.
* Nine dead pigs were 100% SILN positive, with Ct values differing from spleen samples by less than one cycle. This solved the operational challenges of WOAH-recommended samples (requiring necropsy and prone to contamination), establishing the gold standard for passive monitoring without evisceration.
* Surviving pigs showed a viral clearance trend at 10-18 dpi, with continuously rising SILN Ct values, providing molecular evidence for "recovery assessment."
* A triple diagnostic test combining histopathology, immunohistochemistry (IHC), and in situ hybridization (ISH) revealed that the virus only infects macrophages/dendritic cells → these cells secrete pro-inflammatory cytokines → lymphocytes, without viral infection, undergo apoptosis/necrosis → ultimately leading to lymphoid tissue damage (hemorrhagic necrosis), clarifying the indirect mechanism of immune injury.
This study used two moderately virulent strains, ASFV Estonia 2014 and ASFV Malta’78, to conduct experiments on pigs using a simulated field contact transmission method. The dynamic distribution of the virus in blood, spleen, tonsils, and various superficial lymph nodes was systematically analyzed.
Results showed that the virus could be detected in superficial inguinal lymph nodes 2-3 days after infection, reaching its peak at 5-9 days. The study further confirmed that the viral genome content in the spleen of dead pigs was highly consistent with that in SILN (spleen-intestinal lymph node), supporting SILN as a highly efficient sample type for screening dead pigs for African swine fever.
Introduction
African swine fever (ASF) is rampant globally. Passive monitoring relying on spleen removal is time-consuming, labor-intensive, and carries high biosafety risks. The Canadian Food Inspection Agency's Centre for External Diseases (NCFAD) team proposed the hypothesis that "SILN can replace the spleen" in 2022, but data on its applicability in the early infection stage and whether attenuated strains are similarly distributed remains lacking. This study aims to fill this gap.
Results
1. Differences in virus detection timing:
Viralization in pigs infected with ASFV Estonia 2014 began at 2 days post-infection (dpi), while in pigs infected with ASFV Malta’78 it began at 3 dpi; both viruses were detectable in SILN at 3 dpi, and viral load increased rapidly over time.
Figure 1. Genomic distribution of Estonia 2014 African swine fever virus strain in individual pigs (a-c) and daily average detection results (with standard error of average Ct values) (d-f)
(a, d) show the distribution in whole blood, spleen, and tonsils; (b, e) show the distribution in superficial inguinal lymph nodes (SILN), mandibular lymph nodes (SLN), and superficial cervical lymph nodes (SCLN); (c, f) show the distribution in popliteal lymph nodes (PLN), anterior femoral lymph nodes (PFLN), and gastrohepatic lymph nodes (GHLN). Error bars in the figure represent the standard error (SEM) of the Ct values at each time point and for each sample type.
2. Peak and Clearance Patterns:
The viral load of ASFV Estonia 2014 in lymphoid organs peaked at 7-9 dpi, while that of ASFV Malta’78 peaked at 5-7 dpi. The SILN levels in dead pigs were comparable to those in the spleen, while the viral load in the peripheral lymphoid organs of surviving pigs gradually decreased, indicating viral clearance.
Figure 2. Histopathological observation and virus distribution in superficial inguinal lymph nodes (SILNs) of piglets after oral-nasal inoculation with moderately virulent Estonia 2014 strain African swine fever virus.
On day 3 post-inoculation (3 dpi), no significant histopathological changes were observed: (a) Immunohistochemistry (IHC) showed scattered single cells positive for African swine fever virus (ASFV) antigen (arrow); (b) In situ hybridization (ISH) detected ASFV RNA with a similar distribution (arrow).
On day 4 post-inoculation (4 dpi), multifocal hemorrhage areas were observed at the cortex and corticomedullary junction (d, arrow). IHC showed obvious scattered single-cell positive staining patches (e, arrow), and the viral genomic material detected by ISH (f) showed the same distribution and intensity.
On day 5 post-inoculation (5 dpi), multifocal necrosis with hemorrhage appeared mainly along the corticomedullary junction (g, arrow). Viral antigen (h) and viral RNA (i) were detected in the corresponding areas of necrotic lesions and in scattered macrophage-like cells throughout the tissue.
On day 7 post-inoculation (7 dpi), extensive hemorrhage and necrosis were observed at the corticomedullary junction and throughout the multifocal area of the cortex (j, arrow). A large number of African swine fever virus antigens were present at the corticomedullary junction, and some scattered positive cells were also found in the cortex (k). Compared to previous time points, the detection level of viral nucleic acid was reduced at this time (l).
On day 9 post-inoculation (9 dpi), extensive necrosis and hemorrhage were observed throughout the lymph node (m, arrow), including endothelial cell degeneration (m, inset showing a higher magnification of the necrotic area). The levels of viral antigen (n) and viral nucleic acid (o) detected throughout the multifocal area of the lymph node were lower than those observed at 7 dpi. However, viral antigens were still observed in vascular endothelial cells (n, o inset).
On day 11 post-inoculation (11 dpi), moderate necrosis was observed (p), but immunostaining was significantly reduced (q, r).
3. Application value of SILN:
SILN sample collection does not require dissection, and the virus detection rate in dead pigs is consistent with that in the spleen. It can be stably detected in the early infection stage (after 3 dpi), making it an ideal alternative sample for screening dead pigs.
Figure 3. Genomic distribution of Malta’78 African swine fever virus in individual pigs (a-c) and average daily detection results (with average Ct value standard error) (d-f)
(a,d) show the distribution in whole blood, spleen, and tonsils; (b,e) show the distribution in superficial inguinal lymph nodes (SILN), submandibular lymph nodes (SLN), and superficial cervical lymph nodes (SCLN); (c,f) show the distribution in popliteal lymph nodes (PLN), anterior femoral lymph nodes (PFLN), and gastrohepatic lymph nodes (GHLN). Error bars in the figure represent the standard error (SEM) of Ct values at each time point and for each sample type.
4. Pathological and Molecular Validation:
Immunohistochemical and in situ hybridization results confirmed that the distribution trends of viral antigens and nucleic acids in SILN were consistent with the real-time PCR results. Pathological changes such as lymphoid tissue necrosis and hemorrhage were observed in the later stages of infection, and the viral clearance process was synchronized with the repair of pathological damage.
Figure 4. Histopathological observation and virus distribution in superficial inguinal lymph nodes (SILNs) of piglets after oral-nasal inoculation with moderately virulent Malta’78 strain of African swine fever virus.
On day 4 post-inoculation (4 dpi), no obvious lesions were observed in HE sections (a). Scattered, macrophage-like single cells were observed by immunohistochemistry (IHC) (b, arrow) and in situ hybridization (ISH) (c, arrow).
On day 5 post-inoculation (5 dpi), a small number of small necrotic foci were observed in the medulla (d, arrow). Abundant African swine fever virus (ASFV) antigen (e) and viral RNA (f) were detected in cells with morphology consistent with macrophages and dendritic cells.
On day 7 post-inoculation (7 dpi), areas of medullary necrosis appeared (g, arrow). Compared with 5 dpi, the amount of ASFV antigen (h) and viral RNA (i) detected was reduced.
On day 10 post-inoculation (10 dpi), hemorrhage and necrosis were observed primarily at the cortico-medullary junction (j), accompanied by weak immunostaining signals (k, l).
On day 18 post-inoculation (18 dpi), reactive hyperplasia (m) was observed in the SILN tissue, and no significant staining signals were observed by either IHC (n) or ISH (o).
Conclusion
This study systematically elucidated the distribution dynamics of two moderately virulent ASFV strains in the peripheral lymphoid organs of pigs after oral-nasal infection through animal experiments. The study found that the virus rapidly spread to the superficial inguinal lymph nodes (SILN) after infection, and the viral load in the SILN of all deceased pigs remained highly consistent with that in the spleen, thus confirming from a pathogenic perspective that the SILN has a solid scientific basis as a screening sample for ASF-affected pigs.
It is worth emphasizing that this study also clarified the applicability boundaries of this sampling method: potentially surviving infected pigs will gradually clear the virus from their peripheral lymph nodes, and the viral load and variability in lymph nodes are low in the early stages of infection and during the recovery period. Therefore, SILN is mainly suitable for rapid screening of dead or dying pigs, and is not recommended for routine pathogen surveillance in early-stage infection or surviving animals.
The findings of this study have clear practical significance: SILN sampling does not require carcass dissection, is simple and quick to operate, and can significantly reduce the difficulty of on-site sampling and biosafety risks. It provides key technical support for optimizing ASF passive surveillance programs, especially for improving the ability to detect outbreaks early.
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