African swine fever (ASF) is one of the most dangerous and devastating diseases of domestic pigs and wild boar. The disease poses a serious risk to the worldwide trade in pork meat, causing enormous economic losses. Since 2005, 74 countries have confirmed cases of ASF on their territory (29). The causative agent of ASF, African swine fever virus (ASFV), belongs to the
According to the literature, ASFV is among the persistent pathogens in the environment, but the virus can be easily eradicated with most common disinfectants (10, 11, 14). Direct contact plays an important role in the transmission of the disease. Infected animals excrete the virus with oral and nasal fluids, faeces and urine. African swine fever virus can also be transmitted over short distances
Legislation stipulates the procedure for introduction of a new pig herd onto a farm previously affected by ASF; in Poland it permits housing a new herd on a farm after 40 days, counting from the day when cleaning and disinfection processes are completed. Rehoused animals should serve as sentinels and be tested for the presence of specific anti-ASFV antibodies after 45 days. Shortly after the negative results are obtained, a new herd can be introduced safely. Alternatively, pigs may be brought onto the farm again 6 months from the completion of the cleaning and disinfection processes without retesting for antibodies (8). Such a long period ensures the safety of a newly introduced herd; however, in light of recently published studies in this area, from the economic point of view it may be open to discussion if this duration is necessary for the inactive phase (7, 19).
In the present study the molecular contamination of an animal facility was evaluated during and after highly pathogenic ASFV infection, to investigate the virus’ location and its initial amount in the environment. Isolation of infectious virus from collected samples was also attempted, to indicate the risk to animal safety from a contaminated environment. Different routes of transmission were focused upon,
The data presented in this study were gathered during an independent animal trial from a group of six 10-week-old Danbred Duroc domestic pigs. The animal experiment was approved by the Local Ethical Committee for Animal Experiments in Lublin (under approval number 82/2022). All procedures including euthanasia were performed in compliance with current legal regulations.
The virulent ASFV genotype II Georgia 2007 strain at a dose of 1 × 105 50% haemadsorbing doses (HAD50) per animal was used for intranasal infection. The virus was isolated by Dr. Linda Dixon at the Pirbright Institute, Woking, UK, and kindly provided for the present study by the Institut de Recerca i Tecnologia Agroalimentàries – Centre de Recerca en Sanitat Animal, Barcelona, Spain.
Animals were kept in a biosafety level 3 animal facility at the National Veterinary Research Institute (Puławy, Poland) and provided with feed and water
The facility floor was cleaned with tap water on a daily basis during the trial by the facility staff. Seven days from the end of the experiment, the facility was decontaminated with 35% hydrogen peroxide vaporisation (Bioquell Z, Andover, UK).
Oral, nasal and rectal swabs were collected at 0 and 7 days post infection (dpi). Additional samples were taken during necropsy. Swabs were placed into tubes containing 1 mL of phosphate-buffered saline, then incubated at room temperature for 10 min and vortexed. An aliquot of 200 μL of each sample was reserved for DNA extraction and real-time PCR analysis.
Environmental samples were collected when the infection was ongoing at 7 dpi, at 14 dpi (before decontamination) and at 15 dpi (24 h after decontamination). Two millilitres of Roswell Park Memorial Institute 1640 medium (PAN Biotech, Aidenbach, Germany) as a growth medium (GM) supplemented by 1% Antibiotic-Antimycotic Solution (Sigma-Aldrich, St. Louis, MO, USA) and 10% foetal bovine serum (GIBCO, Thermo Fisher Scientific, Waltham, MA, USA) was placed into a plastic tube. Designated spots were thoroughly scrubbed by swabs and immersed in the medium. Swabs from the floor (five spots), feeder (one spot), drinker (one spot), a rubber mat (one spot), a wall at a height of about 30 cm (one spot), the HEPA filter cover (one spot), and the doors at a height of about 120 cm (one spot) were collected in triplicate. Samples were aliquoted and stored at -80°C for real-time PCR and
Extraction of DNA was carried out on the Qiagen DNA Mini Kit protocol (Qiagen), with further use of the Virotype ASFV PCR Kit (Qiagen) for a subsequent real-time PCR reaction in a Rotor-Gene Q thermocycler (Qiagen) according to the manufacturer’s instructions.
Porcine primary pulmonary alveolar macrophages were collected by lung lavage from uninfected donor pigs and seeded at 1 × 105 cells per well in GM supplemented by erythrocytes (1 : 300 v/v) in a 96-well plate. One hundred microlitres of each filtered (0.45 μm) environmental sample was added to the respective wells in triplicate. In parallel, the negative control (medium) and positive control (Georgia 2007, 1 × 103 HAD50/mL) were prepared in the same way. The plates were incubated at 37°C in 5% CO2 and inspected for the presence of haemadsorption and cytopathic effect on the 0, 4th and 7th days of incubation. In addition, cells and medium were collected on days 0 and 7 of incubation for real-time PCR analyses after three freeze-thaw cycles.
The initial virus load was estimated for non-infectious samples based on the threshold cycle (Ct) values, as previously described (27).
Statistical analysis of initial virus load in samples from different time points was performed using the Kruskal–Wallis test in GraphPad Prism 8.4.2 (GraphPad Software, San Diego, CA, USA). The significance for statistical difference was defined as P-value < 0.05.
Fever followed by apathy in infected animals could be seen beginning from 4 dpi. Infection developed and resulted in the deaths of 100% of the animals by 9 dpi. During the disease, three out of the six animals reached the humane endpoint and were euthanised at 8 dpi because they were suffering from bloody diarrhoea, severe dyspnoea and recumbency. The evolution of fever and the survival rate in this group are presented in Fig. 1.
Up to the 7th dpi (the first environmental sampling time point), four out of the six animals remained in the facility, and at this time, the presence of ASFV genetic material was confirmed in the rectal swabs of two out of the four animals and the oral and nasal swabs of all four animals. All swab samples collected from all six animals during necropsy were PCR positive, with noticeably higher initial virus loads. The mean virus loads of swab samples collected at 7 dpi and during necropsy based on the Ct values is presented in Fig. 2.
The presence of ASFV DNA could be confirmed in all tested locations except the doors. However, infectious virus could not be recovered in any of the samples collected at any time point. The numbers of qPCR-positive samples at the three sampling time points are presented in Table 1.
Numbers of positive qPCR-samples in tested animal facility locations and their mean threshold cycle (Ct) values
Time | Number of positive samples (Mean Ct (±SD)) | |||||||
---|---|---|---|---|---|---|---|---|
Floor | Wall | Door | HEPA filter cover | Drain | Feeder | Drinker | Mat | |
T1 | 14/15 |
3/3 |
0/3 | 2/3 |
2/3 |
3/3 |
3/3 |
2/3 |
T2 | 4/15 |
2/3 |
0/3 | 1/3 |
1/3 |
3/3 |
3/3 |
3/3 |
T3 | 5/15 |
2/3 |
0/3 | 1/3 |
1/3 |
3/3 |
3/3 |
3/3 |
SD – standard deviation; T1 – ongoing infection (7 days post infection (dpi)); T2 – after infection (T1 + 7,
The highest estimated initial virus load was found while the infection was ongoing (7 dpi) at the drain (reaching 1.92 × 104 eqTCID50/mL). A noticeably decreasing tendency in estimated virus load over time was found at the floor and drain spots at 15 dpi, when the mean virus load had decreased to 5.5 × 101 50% tissue culture infectious dose equivalents (eqTCID50)/mL and 1.2 × 103 eqTCID50/mL, respectively. These decreases were statistically significant changes, which was shown by a P-value of 0.002 for the floor and one of 0.0393 for the drain. It is noteworthy that the mean initial virus load at feeder and drinker spots was slightly higher than at faeces and urine or aerosol contact spots (
After infection and replication in the host’s lymphatic organs, ASFV spreads through the pig’s body with blood and can soon be found in almost all tissues (1). The virus can be secreted with oral fluid, faeces and urine (4, 27). Shedding of the virus and the presence of dead animals in the pigsty worsen environmental contamination, pose a risk for indirect transmission of ASF and consequently block the sale of animals (2, 23). Presently, the only action to control the disease is the imposition of biosecurity measures (6). Knowledge of ASFV locations and virus load may facilitate cleaning and disinfection processes in the pigsty after culling, leading to effective eradication of the pathogen and fast reintroduction of animals onto a previously affected farm.
In this study, well-established shedding could be seen at 7 dpi. The presence of the virus’ DNA in all tested matrices is in accordance with previously published findings (9, 22, 25). The environmental samples collected in the present study reflected the ongoing infection situation and consequent facility contamination. Genetic traces of ASFV were found on surfaces that may have had contact with faeces and urine (the floor, walls and drain) and with oral and nasal excretions (the feeder and drinker). The latter outcome may suggest that the virus could be transmitted not only
In this study we were not able to isolate infectious virus from environmental samples. Attempts at infectious virus isolation directly from oral fluid were not successful either in the studies of Guinat
The absence of genetic material of ASFV on doors indicates that using personal protective equipment (gloves) and disinfecting hands during standard veterinary procedures prevents further spreading of the virus. It was also noticeable that the estimated initial virus load decreased significantly on surfaces which had been cleaned mechanically with tap water. This showed that even simple maintenance of a pig-farming facility could minimise the risk of the pathogen’s transmission, but only on non-porous surfaces. Porous materials (
Besides its use in proving the restoration of satisfactory hygiene standards, examination of environmental samples could be useful in epidemiological analysis,
To the best of our knowledge, this is the first report on contamination of a facility housing pigs during and after ASF infection. The gathered data provide useful practical knowledge, facilitating processes of cleaning and disinfection, and bring additional insight into the mechanism of indirect transmission of ASF and the virus presence in pig husbandry facilities.