Abstract: Transboundary and Emerging DiseasesVolume 67, Issue 5 p. 2086-2092 ORIGINAL ARTICLEOpen Access African swine fever virus survival in buried wild boar carcasses Laura Zani, Corresponding Author Laura Zani [email protected] orcid.org/0000-0001-5263-8389 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, Germany Correspondence Laura Zani, Friedrich-Loeffler-Institut, International Animal Health Team, Suedufer 10, 17493 Greifswald-Insel Riems, Germany. Email: [email protected] for more papers by this authorMarius Masiulis, Marius Masiulis State Food and Veterinary Service, Vilnius, Lithuania Lithuanian University of Health Sciences, Kaunas, LithuaniaSearch for more papers by this authorPaulius Bušauskas, Paulius Bušauskas State Food and Veterinary Service, Vilnius, LithuaniaSearch for more papers by this authorKlaas Dietze, Klaas Dietze orcid.org/0000-0002-6138-6707 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorGediminas Pridotkas, Gediminas Pridotkas National Food and Veterinary Risk Assessment Institute, Vilnius, LithuaniaSearch for more papers by this authorAnja Globig, Anja Globig Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorSandra Blome, Sandra Blome orcid.org/0000-0001-5465-5609 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorThomas Mettenleiter, Thomas Mettenleiter Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorKlaus Depner, Klaus Depner Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorBirutė Karvelienė, Birutė Karvelienė Lithuanian University of Health Sciences, Kaunas, LithuaniaSearch for more papers by this author Laura Zani, Corresponding Author Laura Zani [email protected] orcid.org/0000-0001-5263-8389 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, Germany Correspondence Laura Zani, Friedrich-Loeffler-Institut, International Animal Health Team, Suedufer 10, 17493 Greifswald-Insel Riems, Germany. Email: [email protected] for more papers by this authorMarius Masiulis, Marius Masiulis State Food and Veterinary Service, Vilnius, Lithuania Lithuanian University of Health Sciences, Kaunas, LithuaniaSearch for more papers by this authorPaulius Bušauskas, Paulius Bušauskas State Food and Veterinary Service, Vilnius, LithuaniaSearch for more papers by this authorKlaas Dietze, Klaas Dietze orcid.org/0000-0002-6138-6707 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorGediminas Pridotkas, Gediminas Pridotkas National Food and Veterinary Risk Assessment Institute, Vilnius, LithuaniaSearch for more papers by this authorAnja Globig, Anja Globig Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorSandra Blome, Sandra Blome orcid.org/0000-0001-5465-5609 Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorThomas Mettenleiter, Thomas Mettenleiter Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorKlaus Depner, Klaus Depner Friedrich-Loeffler-Institut, Greifswald-Insel Riems, GermanySearch for more papers by this authorBirutė Karvelienė, Birutė Karvelienė Lithuanian University of Health Sciences, Kaunas, LithuaniaSearch for more papers by this author First published: 26 March 2020 https://doi.org/10.1111/tbed.13554Citations: 24 Funding information Parts of this work have been funded by the FLI internal African swine fever virus research network. AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinkedInRedditWechat Abstract Since the first introduction of African swine fever (ASF) into the European wild boar population in 1957, the question of virus survival in carcasses of animals that succumbed to the disease has been discussed. The causative African swine fever virus (ASFV) is known to be very stable in the environment. Thus, carcasses of infected wild boar could play a major role as ASFV reservoir and thereby help to locally maintain and spread the disease in wild boar populations. To minimize this risk, removal of wild boar carcasses in ASF affected areas is regarded to be crucial for effective disease control. If removal is not feasible, carcasses are usually disposed by burial on the spot to avoid direct contact of wild boar to the infection source. In this study, carcasses of ASFV infected wild boar buried in Lithuania at different time points and locations have been excavated and retested for the presence of infectious ASFV by in vitro assays and for viral genome by qPCR. Soil samples potentially contaminated by body fluids have been additionally tested for viral genome. In seventeen out of twenty burial sites, samples of excavated carcasses were positive for ASFV genome. However, in none of the carcass samples ASFV could be isolated. On seven sites soil samples contained ASF viral DNA. These results unexpectedly negate the long-term persistence of infectious ASFV in wild boar carcasses independent from the burial time. In this context, sensitivity of ASFV isolation from carcass samples versus susceptibility of animals and doses needed for oral inoculation has to be further investigated. Furthermore, research is required to consider alternative ASF infection sources and drivers in the infection cycle among wild boar. 1 INTRODUCTION African swine fever (ASF) affects all members of the Suidae family. While it is asymptomatic in African indigenous wild pigs, for example warthogs (Thomson, Gainaru, & Van Dellen, 1980), it causes a fatal disease in domestic pigs and Eurasian wild boar (Sus scrofa). Due to its extensive spread within the last decade, ASF became a tremendous threat to the global pig industry (Sánchez-Cordón, Montoya, Reis, & Dixon, 2018). Since there is no vaccine and no treatment available, biosecurity and culling of animals are the only tools to fight the disease in pig holdings. As long as ASF affected pig holdings are detected early and all measures are accompanied by proper cleaning and disinfection, the risk for further spread of the disease can be minimized (Zani et al., 2019). If ASF occurs in wild boar, the situation is much more complex. The infected area has to be defined, and ASF control strategies like ban of hunting and/or fencing to limit the wild boar movement must be adapted to the local situation (EFSA Panel on Animal Health & Welfare, 2018). Since carcasses of ASF infected wild boar could act as a source of infection, they have to be detected and removed as soon as possible. Leak-proof collection and transport of infected carcasses to incineration plants are regarded to be the safest way of carcass disposal (Depner et al., 2017). However, carcass removal and transport to rendering facilities can be problematic or even impossible if the terrain is difficult to access or covered by dense vegetation. In most of the countries, carcass disposal by burning on spot is strictly prohibited due to environmental purposes. In the case of Lithuania, where a rendering plant was not available for the disposal of wild boar carcasses, burying of carcasses as suggested by the Food and Agricultural Organization of the United Nations (Miller & Flory, 2018) was applied to reduce the risk for disease spread. Consequently, in Lithuania, with its large and remote forests, but with soil suitable for digging, many wild boar carcasses have been buried since the start of the epidemic in this country in 2014 (Pautienius et al., 2018). In this study, wild boar carcasses buried at different time points and locations in Lithuania in 2017–2018 were excavated and tested for the presence of infectious ASFV and viral DNA to check for virus survival in those carcasses. Virus survival has been analysed in several studies (Table 1) that investigated meat products (Mebus et al., 1997; Petrini et al., 2019), pig excretions (Davies et al., 2017) and different environmental samples (EFSA Panel on Animal Health & Welfare, 2014). However, mainly samples from experimentally infected animals or spiked samples have been tested. To evaluate if the data holds true for samples under field conditions, we investigated specimens taken from wild boar that succumbed to ASF and were buried in Lithuania if proper carcass removal was impossible. Bone marrow, surrounding soil and, if available, residual organ matrix were sampled and tested. TABLE 1. Summary of studies conducted on African swine fever virus tenacity Material Duration and conditions Method Reference Blood 140 days in the dark Bioassay Eustace Montgomery (1921) Blood >6 years at 4–6°C Bioassay (injection) Kovalenko, Sidorov, and Burba (1972) Blood >90 days Virus isolation (high titres) Blome and Dietze, 2011 (unpublished data) Putrefied blood 15 weeks Unknown USDA, 1997 cited by EFSA Panel on Animal Health and Welfare (2010) Spleen 240 days (6–8°C) Bioassay (injection) Kovalenko et al. (1972) Spleen >90 days Virus isolation (high titres) Blome and Dietze, 2011 (unpublished data) Muscle 155 days (6–8°C) Bioassay (injection) Kovalenko et al. (1972) Muscle 183 days Unknown McKercher et al. (1987) Muscle 90 days Virus isolation (low titre) Blome and Dietze, 2011 (unpublished data) Fat 123 days Virus isolation McKercher et al. (1987) Cured pork belly 60 days Virus isolation + bioassay (oral) Petrini et al. (2019) Cured loin 83 days Virus isolation + bioassay (oral) Petrini et al. (2019) 2 MATERIALS AND METHODS 2.1 Study design Wild boar carcass excavation including decontamination measures of the excavation place and surrounding areas has been approved by the Lithuanian Ministry of Environment and the State Food and Veterinary Service of the Republic of Lithuania. Wild boar carcasses, initially found and tested positive for ASFV infection by quantitative polymerase chain reaction (qPCR) as part of disease control measures, have been excavated during 8–11 October 2018 on 20 different locations across Lithuania (see Figure S1). Bone marrow and, if available, residual organ matrix were taken from each excavated carcass. In addition, soil was sampled on each excavation site by pooling three samples randomly taken next to the decomposing bodies. For better comparison, the decomposition status of each sampled carcass was scored. Score ① denotes that the cadaver was generally intact with closed skin and identifiable organs. Carcasses that were dismembered or open but had residual tissue obtained score ② and if only bones were left, the carcasses received score ③ (Figure 1). After excavating and sampling, the carcasses were buried again on the spot. Local soil surfaces and all involved instruments were treated using a broad-spectrum disinfectant containing peroxides, surfactants, organic acids and an inorganic buffer system (Virkon® S, Lanxess). To avoid unintentional virus inactivation of carcass samples, disinfectants were strictly applied after completion of the sampling process. The obtained specimens were subsequently tested by qPCR and virus isolation at the National Food and Veterinary Risk Assessment Institute, harbouring the National Reference Laboratory (NRL) for ASF in Lithuania and at the Friedrich-Loeffler-Institut (FLI), Germany. FIGURE 1Open in figure viewerPowerPoint Exemplary pictures of applied carcass condition score. Score ① skin closed, organs intact; Score ② residual tissue; Score ③ only bones 2.2 Laboratory investigations 2.2.1 Sample processing After collection, samples were stored at −80°C until further use. For qPCR and virus isolation, bone marrow samples were homogenized with a 5 mm stainless steel bead in 1 ml phosphate-buffered saline (PBS) using TissueLyser II (Qiagen). Tissue samples were homogenized by grinding in sterile sand with pestle and mortar. 2.2.2 Genome detection At the Lithuanian NRL, ASF viral nucleic acid was extracted using the QIAamp® RNA Viral Mini Kit (Qiagen). All samples were screened for ASFV genome by qPCR according the protocol of King et al. (2003). At FLI, nucleic acids of soil and decay matrix samples have been extracted by using the DNeasy® PowerSoil® Kit (Qiagen). The obtained eluates were screened for ASFV genome by qPCR (Tignon et al., 2011) with beta-actin gene serving as internal control. In addition, soil samples were screened for swine cytochrome B gene (Forth, 2015) to check for the presence of wild boar DNA. 2.2.3 Virus isolation To detect infectious ASFV, homogenized bone marrow and tissue samples were tested for hemadsorption on peripheral blood mononuclear (PBMC)-derived macrophages by directly adding the homogenate but also by running three blind passages first. Blood for the preparation of PBMC-derived macrophages was collected from healthy domestic donor pigs. In brief, PBMCs were obtained from EDTA-anti-coagulated blood using Pancoll animal density gradient medium (PAN Biotech). PBMCs were grown in RPMI-1,640 cell culture medium with 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) and 10% foetal calf serum (FCS) at 37°C in a humidified atmosphere containing 5% CO2. The medium was supplied with amphotericin B, streptomycin and penicillin to avoid bacterial and fungal growth. To facilitate maturation of macrophages, GM-CSF (granulocyte-macrophage colony-stimulating factor; Biomol) was added to the cell culture medium at 2 ng/ml. For blind passaging, 1 ml of the homogenate was added to PBMC-derived macrophages (1 × 107 cells/ml) in a T 12.5 cell culture flask (Corning™). After an incubation time of 48–72 hr, the cell culture flasks were freeze-thawed before further passaging and testing. The hemadsorption test (HAT) was carried out according to slightly modified standard procedures (Carrascosa, Bustos, & Leon, 2011). In brief, 100 μl of isolated PBMCs per well was seeded into a 96-well microplate (Corning™ Primaria™) at a density of 5 × 106 cells/ml. After 16–24 hr, non-adherent cells were removed and cell culture medium containing GM-CSF was replenished as described above. The culture was then incubated for 24 hr to allow initial maturation of macrophages. Subsequently, 30 µl of the supernatant of the homogenized sample was added to each well. From blind passaged samples, 100 µl of the cell culture supernatant was added to each well. Samples were tested in eight replicates. After 24 hr of incubation, 20 µl of homologous 1% erythrocyte suspension was added to each well. For readout, cultures were checked for hemadsorption over a period of 3 days. When using bone marrow or tissue homogenates directly, cells were washed after 2 hr of adsorption time at 37°C using lukewarm PBS, whereas cell culture supernatant was left on the cells until the final evaluation of the test. ASFV Armenia 2007 virus stocks were used as positive control. Doubtful results were confirmed by an additional passage and qPCR testing (King et al., 2003) of cell supernatant. To exclude the presence of non-hemadsorbing ASFV strains, samples of each carcass were additionally screened by indirect immunofluorescence staining according to the protocol published by Carrascosa et al. (2011). In brief, the formalin-fixated macrophages were incubated with monoclonal antibodies against the viral protein p30 (kindly provided by Linda Dixon, Pirbright Institute, UK) and then stained with commercially available fluorescent antibodies Goat anti-mouse IgG (H + L), Alexa Fluor® 488 (Thermo Fisher). Blood donation from pigs kept for laboratory investigation at FLI, Germany was approved by the competent authority (Landesamt für Landwirtschaft, Lebensmittelsicherheit und Fischerei (LALLF) Mecklenburg-Vorpommern) under reference number LALLF 7221.3-2-041/17. No further animal experiments have been conducted within the present study. Data analysis was performed using Microsoft Excel 2007 (Microsoft) and SigmaPlot for Windows version 11.0 (Systat software, Inc.). 3 RESULTS 3.1 Sample amount and decay conditions of excavated wild boar carcasses In total, 45 bone marrow samples and 18 tissue samples from carcasses excavated at 20 sites have been obtained (Table S1). The sampled carcasses were buried between 18 and 440 days before excavation and were in different stages of decay. The decomposition status varied between the sites and burial times. On two sites, carcasses with score ① were sampled. Eleven carcasses were excavated in condition score ② and from eight carcasses only bones (score ③) could be found and sampled (Figure 2). Only on excavation site #9, two carcasses with different decomposition score could be clearly identified and separate sample panels were obtained. On all other sites, either only one carcass was buried or, if the identification of corresponding body pieces was not feasible, a general carcass scoring was applied. For seven sites, additional information on the carcass decay condition at the time point of burial was available (Figure 2). FIGURE 2Open in figure viewerPowerPoint Overview of burial time and condition of sampled carcasses. Carcass condition score after excavation has been assessed for each location (right side). If more than one carcass was buried and carcasses at the same location showed different level of decay, separate scoring was applied. If the information was available, carcass condition score at time of burial has been added (left side) 3.2 Genome detection in all but three excavation sites and in soil samples of seven sites All sites selected for this study contained wild boar carcasses that were initially tested positive for ASFV genome by qPCR during routine diagnostics. When retested after excavation, in all except three carcass samples, ASFV genome could be detected again by qPCR. The mean ct value of the initial testing was 29.8 with a SD of 3.94. The mean ct value of excavated carcass samples was 31.14 with a SD of 3.32 (Figure 3). On seven sites (Figure 2), soil samples yielded ASFV positive results in qPCR with a mean ct value of 34.12 (SD: 1.29). Traces of swine cytochrome B gene were found in all but six excavation sites with a mean ct value of 31.18 (SD: 3.84). Only on one site (#5) ASFV genome was detected without the detectable presence of swine DNA. All soil samples tested positive for the internal control (beta-actin gene). FIGURE 3Open in figure viewerPowerPoint qPCR results of wild boar carcasses. Black dots indicate ct values of qPCR testing shortly before the carcass was buried, and white dots indicate ct values after carcass excavation. If more than one dot for each category is displayed for a time point, several samples of the same carcasses have been tested or carcasses on different spots have been sampled after the same burial time 3.3 Virus isolation From none of the bone marrow and tissue samples obtained from excavated carcasses, infectious ASFV could be isolated. Neither in samples passaged three times on macrophages nor in those that have been applied without previous passaging, was any hemadsorption observed. Also, the direct immunofluorescence test did not reveal any indication for the presence of infectious ASFV. 4 DISCUSSION Several studies investigating ASFV tenacity in pork products have been conducted. Under favourable conditions, meaning cool and associated with tissue or blood the virus has shown to be rather stable in uncooked pork (Table 1). Nonetheless, there is so far no data available for the survival of ASFV in wild boar carcasses decomposing under field conditions. Due to the high stability of the virus and the fact that pigs dying during the acute phase of the disease usually show high titres of ASFV in blood (Zhao et al., 2019) it has been assumed that carcasses of infected wild boar contain considerable amounts of viable virus (Chenais, Stahl, Guberti, & Depner, 2018). The detection of very stable quantities of ASFV genome (Figure 3) in the obtained samples shows that passive surveillance by sampling dead wild boar is highly reasonable even if the carcass is in advanced stage of decomposition and only bone marrow is available. In our study, only four bone marrow samples (from carcasses after 96, 274, 321 and 440 days of burial) were negative for ASFV genome. However, only on three sites (96, 321 and 440 days of burial) all obtained carcass samples tested negative. The fluctuation of ct values within samples obtained from the same carcass before and after burial shows that viral DNA is rather stable but not homogeneously distributed in body tissues as shown in other studies (Petrov, Forth, Zani, Beer, & Blome, 2018). However, mean ct values after carcass excavation were comparable to previous testing (Figure 3). In soil samples, ASF viral DNA was found at seven out of twenty sites while swine-specific cytochrome B DNA was detected in fourteen samples. This indicates a rather low virus contamination of soil surrounding buried carcasses although host DNA is present. This might be different in case of carcasses decomposing on the surface where the body is exposed to outdoor conditions and insects or vertebrate species have access. The virus load in soil samples could also have been reduced by the natural presence of humic acids that have been shown to reduce infectivity of other pathogens (Kuznetsova, Cullingham, McKenzie, & Aiken, 2018). Wild boar have shown to root in ground soaked with decay liquids (Probst, Globig, Knoll, Conraths, & Depner, 2017). Therefore, the low contamination rate shown in the presented study would decrease the chance of disease spread by this behaviour. However, the number of obtained soil samples is limited and further studies are under way to evaluate the transmission potential of soil. As mentioned before, soil samples were tested by using the protocol of Tignon et al. (2011) while carcass samples have been tested by using the protocol of King et al. (2003). Both protocols are listed in the OIE Manual of Diagnostic Tests and Vaccines for Terrestrial Animals (Sánchez-Vizcaíno, Dixon, & Heath, 2019) and show similar sensitivity and specificity but in our hands the (Tignon et al., 2011) protocol led to less inhibition artefacts with problematic soil samples. Therefore, ct values of carcass and soil samples cannot be directly compared with the ct values of carcass samples in this study but the contamination rate of the obtained soil samples could be reliably assessed. In none of the carcasses sampled during this study, infectious ASFV could be isolated independent from burial time and carcass condition. This result was rather unexpected and might have been influenced by different factors. The obtained sample material is challenging for hemadsorption testing due to advanced decomposition status of sampled tissues and high fat content of white bone marrow. To mitigate the influence of sample quality, blind passaging was performed and antibiotics were added to the cell culture medium. In case of direct testing, the sample was removed after 2 hr from the cell layer to avoid possible damage due to toxic effects. However, a negative influence of sample material on test sensitivity cannot be ruled out. Bioassays, that is testing of infectivity by administration of the sample to live pigs, are known to be more sensitive (Petrini et al. 2019; Table 1). It has been shown that pigs were successfully orally infected with ASFV even if the sample previously tested negative in virus isolation (Niederwerder et al., 2019; Petrini et al., 2019). However, for animal welfare reasons and the low amount of sample material, this was not attempted in our study. Another factor affecting virus stability might be the unknown conditions during the decomposing process of the sampled carcasses. In published tenacity studies, samples are usually stored at a standardized setting which is different from field conditions. Lithuania has a humid continental climate with average air temperatures not above −3°C in the coolest month and up to 22°C in the warmest months. At least for 4 months, the average air temperature is above 10°C (Bukantis, 2001). Compared to air temperature, soil surface is on average 3–6°C warmer during summer and several tenths of a degree colder in winter depending on vegetation and humidity (Juknevičiūtė & Laurinavičius, 2008). All sampled carcasses were buried for at least one summer period, and in case of advanced decomposition status at time of burial (Figure 2) it can be assumed that the exposure to outdoor conditions was even longer. ASFV is known to be heat sensitive (Plowright & Parker, 1967) and inactivated at 37°C after around 20 days (Mazur-Panasiuk, Zmudzki, & Wozniakowski, 2019). Therefore, taking into consideration the additional temperature increase of the carcass during decomposition (above 32°C at an outer temperature of 23°C) it does not seem unlikely that the virus has been inactivated effectively during the decomposition process (Johnson, Mikac, & Wallman, 2013). Nonetheless, as the presence and titres of infectious virus at the time of death of the wild boar included in this study are not known, inactivation is difficult to prove and further research is required. 5 CONCLUSIONS Due to the high stability of ASFV, carcasses of infected wild boar are considered as major driver for local spread and the maintenance of ASF in the habitat. However, the presented study could not demonstrate the presence of infectious ASFV in buried wild boar carcasses at different decomposition status. This result was unexpected but should be taken with care considering a risk of infection for swine coming into contact with dead wild boar which succumbed to ASF. However, it might indicate that there are additional sources of infection that might have been underestimated such as direct and indirect contact between wild boar. Furthermore, proper burial as an alternative tool for wild boar carcass disposal can be regarded as a safe way to mitigate ASFV spread within the habitat if safe collection and transport to rendering plants are not feasible. ACKNOWLEDGEMENTS The authors would like to thank MSc Felix Närmann, University of Greifswald for preparing the map of carcass excavation spots. CONFLICTS OF INTEREST The authors declare no conflict of interest. Open Research DATA AVAILABILITY STATEMENT The data supporting the findings of this study are available in the supplementary material of this article or are available from the corresponding author upon request. Supporting Information Filename Description tbed13554-sup-0001-Figure.pngPNG image, 864.9 KB Figure S1 tbed13554-sup-0002-Table1.xlsxMS Excel, 11.9 KB Table S1 tbed13554-sup-0003-Captions.docxWord document, 13 KB Please note: The publisher is not responsible for the content or functionality of any supporting information supplied by the authors. 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