Chapter 3.4.12. – Lumpy skin disease

Chapter 3.4.12. – Lumpy skin disease

Chapter 3.4.12.

lumpy skin disease

SUMMARY

**Description of the disease:**Lumpy skin disease (LSD) is a poxvirus disease of cattle characterised by fever, nodules on the skin, mucous membranes and internal organs, emaciation, enlarged lymph nodes, oedema of the skin, and sometimes death. The disease is of economic importance as it can cause a temporary reduction in milk production, temporary or permanent sterility in bulls, damage to hides and, occasionally, death. Various strains of capripoxvirus are responsible for the disease. These are antigenically indistinguishable from strains causing sheep pox and goat pox yet distinct at the genetic level. LSD has a partially different geographical distribution from sheep and goat pox, suggesting that cattle strains of capripoxvirus do not infect and transmit between sheep and goats. Transmission of LSD virus (LSDV) is thought to be predominantly by arthropods, natural contact transmission in the absence of vectors being inefficient. Lumpy skin disease is endemic in ~~most~~many African and Middle Eastern countries. Between 2012 and ~~2018~~2022, LSD spread into south-east Europe, the Balkans, Russia and ~~the Caucasus~~Asia as part of the Eurasian LSD epidemic.

**Pathology:**the nodules are firm, and may extend to the underlying subcutis and muscle. Acute histological key lesions consist of epidermal vacuolar changes with intracytoplasmic inclusion bodies and dermal vasculitis. Chronic key histological lesions consist of fibrosis and necrotic sequestra~~e sequestrum~~.

**Detection of the agent:** Laboratory confirmation of LSD is most rapid using a real-time or conventional polymerase chain reaction (PCR) method specific for capripoxviruses in combination with a clinical history of a generalised nodular skin disease and enlarged superficial lymph nodes in cattle. Ultrastructurally, capripoxvirus virions are distinct from those of parapoxvirus~~virions~~, which causes bovine papular stomatitis and pseudocowpox, but cannot be distinguished morphologically from orthopoxvirus virions, including cowpox and vaccinia viruses, both of which can cause disease in cattle, although neither causes generalised infection and both are uncommon in cattle. LSDV will grow in tissue culture of bovine, ovine or caprine origin. In cell culture, LSDV causes a characteristic cytopathic effect and intracytoplasmic inclusion bodies that is distinct from infection with Bovine herpesvirus 2, which causes pseudo-lumpy skin disease and produces syncytia and intranuclear inclusion bodies in cell culture. Capripoxvirus antigens can be demonstrated in tissue culture using immunoperoxidase or immunofluorescent staining and the virus can be neutralised using specific antisera.

A variety of conventional and real-time PCR tests as well as isothermal amplification tests using capripoxvirus-specific primers have been published for use on a variety of samples. 

**Serological tests:** The virus neutralisation test (VNT) and enzyme-linked immunosorbent assays (ELISAs) are widely used and have been validated. The agar gel immunodiffusion test and indirect immunofluorescent antibody test are less specific than the VNT due to cross-reactions with antibody to other poxviruses. Western blotting using the reaction between the P32 antigen of LSDV with test sera is both sensitive and specific, but is difficult and expensive to carry out.  

**Requirements for vaccines:**All strains of capripoxvirus examined so far, whether derived from cattle, sheep or goats, are antigenically similar. Attenuated cattle strains, and strains derived from sheep and goats, have been used as live vaccines against LSDV.

A.  introduction

Lumpy skin disease (LSD) was first seen in Zambia in 1929, spreading into Botswana by 1943 (Haig, 1957), and then into South Africa the same year, where it affected over eight million cattle causing major economic loss. In 1957 it entered Kenya, ~~associated with~~at the same time as an outbreak of sheep pox (Weiss, 1968). In 1970 LSD spread north into the Sudan, by 1974 it had spread west as far as Nigeria, and in 1977 was reported from Mauritania, Mali, Ghana and Liberia. Another epizootic of LSD between 1981 and 1986 affected Tanzania, Kenya, Zimbabwe, Somalia and the Cameroon, with reported mortality rates in affected cattle of 20%. The occurrence of LSD north of the Sahara desert and outside the African continent was confirmed for the first time in Egypt and Israel between 1988 and 1989, and was reported again in 2006 (Brenner *et al.,* 2006). In the past decade, LSD occurrences have been reported in the Middle Eastern, European and Asian regions (for up-to-date information, consult WOAH WAHIS interface[footnoteRef:2]). Lumpy skin disease outbreaks tend to be sporadic, depending upon animal movements, immune status, and wind and rainfall patterns affecting vector populations. The principal method of transmission is thought to be mechanical by various arthropod vectors (Tuppurainen *et al*., 2015).  [2:  	https://www.woah.org/en/what-we-do/animal-health-and-welfare/disease-data-collection/] 

Lumpy skin disease virus (LSDV) belongs to the family *Poxviridae*, subfamily *~~Chordopoxviridae~~Chordopoxvirinae*, and genus *Capripoxvirus*. In common with other poxviruses LSDV replicates in the cytoplasm of an infected cell, forming distinct perinuclear viral factories. The LSD virion is large and brick-shaped measuring 293–299nm (length) and 262–273nm (width). The LSDV genome structure is also similar to other poxviruses, consisting of double-stranded linear DNA that is 25% GC-rich, approximately 150,000 bp in length, and encodes around 156 open reading frames (ORFs). An inverted terminal repeat sequence of 2200–2300bp is found at each end of the linear genome. The linear ends of the genome are joined with a hairpin loop. The central region of the LSDV genome contains ORFs predicted to encode proteins required for virus replication and morphogenesis and exhibit a high degree of similarity with genomes of other mammalian poxviruses. The ORFs in the outer regions of the LSDV genome have lower similarity and likely encode proteins involved in viral virulence and host range determinants. 

Phylogenetic analysis shows the majority of LSDV strains group into two monophyletic clusters (cluster 1.1 and 1.2) (Biswas *et al.,*2020; Van Schalkwyk *et al.,* 2021). Cluster 1.1 consists of LSDV Neethling vaccine strains that are based on the LSDV/Neethling/LW-1959 vaccine strain (Kara *et al.,* 2003; Van Rooyen *et al.,* 1959; van Schalkwyk *et al.,* 2020) and historic wild-type strains from South Africa. Cluster 1.2 consists of wild-type strains from southern Africa, Kenya, the northern hemisphere, and the Kenyan KSGP O-240 commercial vaccine. In addition to these two clusters, there have recently been recombinant LSDV strains isolated from clinical cases of LSD in the field in Russia and central Asia (Flannery *et al.,* 2021; Sprygin *et al.,* 2018; 2020; Wang *et al.,* 2021). These recombinant viruses show unique patterns of accessory gene alleles, consisting of sections of both wild-type and “vaccine” LSDV strains.

The severity of the clinical signs of LSD is highly variable and depends on a number of factors, including the strain of ~~capripox~~virus, the age of the host, immunological status and breed. *Bos taurus* is generally more susceptible to clinical disease than *Bos indicus;* the Asian buffalo (*Bubalus spp.*) has also been reported to be susceptible. Within *Bos taurus,* the fine-skinned Channel Island breeds develop more severe disease, with lactating cows appearing to be the most at risk. However, even among groups of cattle of the same breed kept together under the same conditions, there is a large variation in the clinical signs presented, ranging from subclinical infection to death (Carn & Kitching, 1995). There may be failure of the virus to infect the whole group, probably depending on the virulence of the virus isolate, immunological status of the host, host genotype, and vector prevalence.

The incubation period under field conditions has not been reported, but following experimental inoculation is 6–9 days until the onset of fever. In the acutely infected animal, there is an initial pyrexia, which may exceed 41°C and persist for 1 week. All the superficial lymph nodes become enlarged. In lactating cattle there is a marked reduction in milk yield. Lesions develop over the body, particularly on the head, neck, udder, scrotum, vulva and perineum between 7 and 19 days after virus inoculation (Coetzer, 2004). The characteristic integumentary lesions are multiple, well circumscribed to coalescing, 0.5–5 cm in diameter, firm, flat-topped papules and nodules. The nodules involve the dermis and epidermis, and may extend to the underlying subcutis and occasionally to the adjacent striated muscle. These nodules have a creamy grey to white colour on cut section, which may initially exude serum, but over the ensuing 2 weeks a cone-shaped central core or sequestrum of necrotic material/necrotic plug (“sit-fast”) may appear within the nodule. The acute histological lesions consist of epidermal vacuolar changes with intracytoplasmic inclusion bodies and dermal vasculitis. The inclusion bodies are numerous, intracytoplasmic, eosinophilic, homogenous to occasionally granular and they may occur in endothelial cells, fibroblasts, macrophages, pericytes, and keratinocytes. The dermal lesions include vasculitis with fibrinoid necrosis, oedema, thrombosis, lymphangitis, dermal-epidermal separation, and mixed inflammatory infiltrate. The chronic lesions are characterised by an infarcted tissue with a sequestered necrotic core, often rimmed by granulation tissue gradually replaced by mature fibrosis. At the appearance of the nodules, the discharge from the eyes and nose becomes mucopurulent, and keratitis may develop. Nodules may also develop in the mucous membranes of the mouth and alimentary tract, particularly the abomasum and in the trachea and the lungs, resulting in primary and secondary pneumonia. The nodules on the mucous membranes of the eyes, nose, mouth, rectum, udder and genitalia quickly ulcerate, and by then all secretions, ocular and nasal discharge and saliva contain LSD virus (LSDV). The limbs may be oedematous and the animal is reluctant to move. Pregnant cattle may abort, and there is a report of intrauterine transmission (Rouby & Aboulsoudb, 2016). Bulls may become permanently or temporarily infertile and the virus can be excreted in the semen for prolonged periods (Irons *et al*., 2005). Recovery from severe infection is slow; the animal is emaciated, may have pneumonia and mastitis, and the necrotic plugs of skin, which may have been subject to fly strike, are shed leaving deep holes in the hide (Prozesky & Barnard, 1982).

The main differential diagnosis is pseudo-LSD caused by bovine herpesvirus 2 (BoHV-2). This is usually a milder clinical condition, characterised by superficial nodules, resembling only the early stage of LSD. Intra-nuclear inclusion bodies and viral syncytia are histopathological characteristics of BoHV-2 infection not seen in LSD. Other differential diagnoses (for integumentary lesions) include: dermatophilosis, dermatophytosis, bovine farcy, photosensitisation, actinomycosis, actinobacillosis, urticaria, insect bites, besnoitiosis, ~~nocardiasis~~nocardiosis, demodicosis, onchocerciasis, pseudo-cowpox, and cowpox. Differential diagnoses for mucosal lesions include: foot and mouth disease, bluetongue, mucosal disease, malignant catarrhal fever, infectious bovine rhinotracheitis, and bovine papular stomatitis.

LSDV is not transmissible to humans. However, all laboratory manipulations must be performed at an appropriate containment level determined using biorisk analysis (see Chapter 1.1.4 *Biosafety and biosecurity: Standard for managing biological risk in the veterinary laboratory and animal facilities*)*.*

B.  DIAGNOSTIC TECHNIQUES

Table 1. Test methods available for the diagnosis of LSD and their purpose

Key: +++ = recommended for this purpose; ++ recommended but has limitations; + = suitable in very limited circumstances; – = not appropriate for this purpose. PCR = polymerase chain reaction; VNT = virus neutralisation test; IFAT = indirect fluorescent antibody test; ELISA = enzyme-linked immunosorbent assay.

1.	Detection of the agent

1.1.	Specimen collection, submission and preparation

Material for virus isolation and antigen detection should be collected as biopsies or from skin nodules at post-mortem examination. Samples for virus isolation should preferably be collected within the first week of the occurrence of clinical signs, before the development of neutralising antibodies (Davies, 1991; Davies *et al*., 1971); however virus can be isolated from skin nodules for at least 3–4 weeks thereafter. Samples for genome detection using conventional or real-time polymerase chain reaction (PCR) may be collected when neutralising antibody is present. Following the first appearance of the skin lesions, the virus can be isolated for up to 35 days and viral nucleic acid can be demonstrated via PCR for up to 3 months (Tuppurainen *et al*., 2005; Weiss, 1968). Buffy coat from blood collected into heparin or EDTA (ethylene diamine tetra-acetic acid) during the viraemic stage of LSD (before generalisation of lesions or within 4 days of generalisation), can also be used for virus isolation. 

~~Samples for histology should include the lesion and tissue from the surrounding (non-lesion) area, be a maximum size of 2 cm3, and be placed immediately following collection into ten times the sample volume of 10% neutral buffered formal saline.~~

~~Tissues in formalin have no special transportation requirements in regard to biorisks.~~Blood samples with anticoagulant for virus isolation from the buffy coat should be placed immediately on ice after gentle mixing and processed as soon as possible. In practice, the samples may be kept at 4°C for up to 2 days prior to processing, but should not be frozen or kept at ambient temperatures. Tissues for virus isolation and antigen detection should be kept at 4°C, on ice or at –20°C. If it is necessary to transport samples over long distances without refrigeration, the medium should contain 10% glycerol; the samples should be of sufficient size (e.g. 1 g in 10 ml) that the transport medium does not penetrate the central part of the biopsy, which should be used for virus isolation.

Samples for histology should include the lesion and tissue from the surrounding (non-lesion) area, be a maximum size of 2 cm3, and be placed immediately following collection into ten times the sample volume of 10% neutral buffered formaldehyde. Tissues in formalin have no special transportation requirements in regard to biorisks. Material for histology should be prepared using standard techniques and stained with haematoxylin and eosin (H&E) (Burdin, 1959). 

Lesion material for virus isolation and antigen detection is minced using a sterile scalpel blade and forceps and then macerated in a sterile steel ball-bearing mixer mill, or ground with a pestle in a sterile mortar with sterile sand and an equal volume of sterile phosphate buffered saline (PBS) or serum-free modified Eagle’s medium containing sodium penicillin (1000 international units [IU]/ml), streptomycin sulphate (1 mg/ml), mycostatin (100 IU/ml) or fungizone (amphotericin, 2.5 μg/ml) and neomycin (200 IU/ml). The suspension is freeze–thawed three times and then partially clarified using a bench centrifuge at 600 ***g*** for 10 minutes. In cases where bacterial contamination of the sample is expected (such as when virus is isolated from skin samples), the supernatant can be filtered through a 0.45 μm pore size filter after the centrifugation step. Buffy coats may be prepared from unclotted blood using centrifugation at 600 ***g*** for 15 minutes, and the buffy coat carefully removed into 5 ml of cold double-distilled water using a sterile Pasteur pipette. After 30 seconds, 5 ml of cold double-strength growth medium is added and mixed. The mixture is centrifuged at 600 ***g*** for 15 minutes, the supernatant is discarded and the cell pellet is suspended in 5 ml growth medium, such as Glasgow’s modified Eagle’s medium (GMEM). After centrifugation at 600 ***g*** for a further 15 minutes, the resulting pellet is suspended in 5 ml of fresh GMEM. Alternatively, the buffy coat may be separated from a heparinised sample by using a Ficoll gradient.

1.2.	Virus isolation on cell culture

LSDV will grow in tissue culture of bovine, ovine or caprine origin. MDBK (Madin–Darby bovine kidney) cells are often used, as they support good growth of the virus and are well characterised (Fay *et al.,* 2020). Primary cells, such as lamb testis (LT) cells, also support viral growth, but care needs to be taken to ensure they are not contaminated with viruses such as bovine viral diarrhoea virus. One ml of clarified supernatant or buffy coat is inoculated onto a confluent monolayer in a 25 cm2 culture flask at 37°C and allowed to adsorb for 1 hour. The culture is then washed with warm PBS and covered with 10 ml of a suitable medium, such as GMEM, containing antibiotics and 2% fetal calf serum. If available, tissue culture tubes containing appropriate cells and a flying cover-slip, or tissue culture microscope slides, are also infected.

The flasks/tissue culture tubes are examined daily for 7–14 days for evidence of cytopathic effects (CPE). Infected cells develop a characteristic CPE consisting of retraction of the cell membrane from surrounding cells, and eventually rounding of cells and margination of the nuclear chromatin. At first only small areas of CPE can be seen, sometimes as soon as 2 days after infection; over the following 4–6 days these expand to involve the whole cell ~~sheet~~monolayer. If no CPE is apparent by day 14, the culture should be freeze–thawed three times, and clarified supernatant inoculated on to a fresh cell monolayer. At the first sign of CPE in the flasks, or earlier if a number of infected cover-slips are being used, a cover-slip should be removed, fixed in acetone and stained using H&E. Eosinophilic intracytoplasmic inclusion bodies, which are variable in size but up to half the size of the nucleus and surrounded by a clear halo, are diagnostic for poxvirus infection. PCR may be used as an alternative to H&E for confirmation of the diagnosis. The CPE can be prevented or delayed by adding specific anti-LSDV serum to the medium. In contrast, the herpesvirus that causes pseudo-LSD produces a Cowdry type A intranuclear inclusion body. It also forms syncytia.

An ovine testis cell line (OA3.Ts) has been evaluated for the propagation of capripoxvirus isolates (Babiuk *et al.*, 2007), however this cell line has been found to be contaminated with noncytopathic pestivirus and should be used with caution.

1.3.	Polymerase chain reaction

The conventional gel-based PCR method described below is a simple, fast and sensitive method for the detection of capripoxvirus genome in EDTA blood, semen or tissue culture samples  (Tuppurainen *et al*., 2005).

1.3.1.	Test procedure

The extraction method described below can be replaced using commercially available DNA extraction kits.

i)	Freeze and thaw 200 μl of blood in EDTA, semen or tissue culture supernatant and suspend in 100 μl of lysis buffer containing 5 M guanidine thiocyanate, 50 mM potassium chloride, 10 mM Tris/HCl (pH 8); and 0.5 ml Tween 20.

ii)	Cut skin and other tissue samples into fine pieces using a sterile scalpel blade and forceps. Grind with a pestle in a mortar. Suspend the tissue samples in 800 μl of the above mentioned lysis buffer.

iii)	Add 2 μl of proteinase K (20 mg/ml) to blood samples and 10 μl of proteinase K (20 mg/ml) to tissue samples. Incubate at 56°C for 2 hours or overnight, followed by heating at 100°C for 10 minutes. Add phenol:chloroform:isoamylalcohol (25:24:1 [v/v]) to the samples in a 1:1 ratio. Vortex and incubate at room temperature for 10 minutes. Centrifuge the samples at 16,060 ***g*** for 15 minutes at 4°C. Carefully collect the upper, aqueous phase (up to 200 μl) and transfer into a clean 2.0 ml tube. Add two volumes of ice cold ethanol (100%) and 1/10 volume of 3 M sodium acetate (pH 5.3). Place the samples at –20°C for 1 hour. Centrifuge again at 16,060 ***g*****for 15 minutes at 4°C and discard the supernatant. Wash the pellets with ice cold 70% ethanol (100 μl) and centrifuge at 16,060 ***g*****for 1 minute at 4°C. Discard the supernatant and dry the pellets thoroughly. Suspend the pellets in 30 μl of nuclease-free water and store immediately at –20°C (Tuppurainen *et al*., 2005). Alternatively a column-based extraction kit may be used.

iv)	The primers for this PCR assay were developed from the gene encoding the viral attachment protein. The size of the expected amplicon is 192 bp (Ireland & Binepal, 1998). The primers have the following gene sequences:

Forward primer 5’-TCC-GAG-CTC-TTT-CCT-GAT-TTT-TCT-TAC-TAT-3’

Reverse primer 5’-TAT-GGT-ACC-TAA-ATT-ATA-TAC-GTA-AAT-AAC-3’.

v)	DNA amplification is carried out in a final volume of 50 μl containing: 5 μl of 10 × PCR buffer, 1.5 μl of MgCl2 (50 mM), 1 μl of dNTP (10 mM), 1 μl of forward primer, 1 μl of reverse primer, 1 μl of DNA template (~10 ng), 0.5 μl of *Taq* DNA polymerase and 39 μl of nuclease-free water. The volume of DNA template required may vary and the volume of nuclease-free water must be adjusted to the final volume of 50 μl.

vi)	Run the samples in a thermal cycler as follows: 2 minutes at 95°C; then 45 seconds at 95°C, 50 seconds at 50°C and 1 minute at 72°C (34 cycles); 2 minutes at 72°C and hold at 4°C until analysis.

vii)	Mix 10 μl of each sample with loading dye and load onto a 1.5% agarose gel in TAE buffer (Tris/acetate buffer containing EDTA). Load a parallel lane with a 100 bp DNA-marker ladder. Electrophoretically separate the products using approximately 8–10 V/cm for 40–60 minutes and visualise with a suitable DNA stain and transilluminator. 

Quantitative real-time PCR methods have been described that are reported to be faster and have higher sensitivity than conventional PCRs (Balinsky *et al*., 2008; Bowden *et al*., 2008). A real-time PCR method that differentiates between LSDV, sheep pox virus and goat pox virus has been published (Lamien *et al.,* 2011). 

Quantitative real-time PCR assays have been designed to differentiate between Neethling-based LSDV strains, which are often used for vaccination, and wild-type LSDV strains from cluster 1.2 (Agianniotaki *et al.,* 2017; Pestova *et al.,* 2018; Vidanovic *et al.,* 2016). These “DIVA” assays (DIVA: differentiation of infected from vaccinated animals) enable, for example, differentiation of “Neethling response” caused by vaccination with a LSDV Neethling vaccine strain from disease caused by infection with a cluster 1.2 wild-type virus. However, these DIVA PCR assays cannot distinguish between a LSDV Neethling vaccine strain and the novel recombinant LSDV strains recently isolated from disease outbreaks in Asia (Byadovskaya *et al.,* 2021; Flannery *et al.,* 2021). These DIVA assays are also not capable of discriminating between LSDV Neethling vaccine strains and recently characterised (historic) wild-type viruses from South Africa belonging within cluster 1.1 (Van Schalkwyk *et al.,* 2020; 2021). Consequently, in regions where recombinant strains (currently Asia and possibly elsewhere) or wild-type cluster 1.1 strains~~are circulating~~(currently South Africa and possibly elsewhere) are circulating, these DIVA assays are not suitable for distinguishing vaccine and wild-type virus. Thus, in order to overcome these constraints, whole genome sequencing is recommended.

1.4.	Transmission electron microscopy

The characteristic poxvirus virion can be visualised using a negative staining preparation technique followed by examination with an electron microscope. There are many different negative staining protocols, an example of which is given below.

1.4.1.	Test procedure

Before centrifugation, material from the original biopsy suspension is prepared for examination under the transmission electron microscope by floating a 400-mesh hexagonal electron microscope grid, with pioloform-carbon substrate activated by glow discharge in pentylamine vapour, onto a drop of the suspension placed on parafilm or a wax plate. After 1 minute, the grid is transferred to a drop of Tris/EDTA buffer, pH 7.8, for 20 seconds and then to a drop of 1% phosphotungstic acid, pH 7.2, for 10 seconds. The grid is drained using filter paper, air-dried and placed in the electron microscope. The capripox virion is brick shaped, covered in short tubular elements and measures approximately 290 × 270 nm. A host-cell-derived membrane may surround some of the virions, and as many as possible should be examined to confirm their appearance (Kitching & Smale, 1986).

The capripox~~virus~~ virions are indistinguishable from those of orthopoxvirus, but, apart from vaccinia virus and cowpox virus, which are both uncommon in cattle and do not cause generalised infection, no other orthopoxvirus is known to cause lesions in cattle. However, vaccinia virus may cause generalised infection in young immunocompromised calves. In contrast, orthopoxviruses are a common cause of skin disease in domestic buffalo causing buffalo pox, a disease that usually manifests as pock lesions on the teats, but may cause skin lesions at other sites, such as the perineum, the medial aspects of the thighs and the head. Orthopoxviruses that cause buffalo pox cannot be readily distinguished from capripoxvirus by electron microscopy. The parapoxvirus virions that cause bovine papular stomatitis and pseudocowpox are smaller, oval in shape and each is covered in a single continuous tubular element that appears as striations over the virion. Capripox~~virus is~~virions are also distinct from the herpesvirus that causes pseudo-LSD (also known as “Allerton” or bovine herpes mammillitis).

1.5.	Fluorescent antibody tests

Capripoxvirus antigen can be identified on infected cover-slips or tissue culture slides using fluorescent antibody tests. Cover-slips or slides should be washed and air-dried and fixed in cold acetone for 10 minutes. The indirect test using immune cattle sera is subject to high background colour and nonspecific reactions. However, a direct conjugate can be prepared from sera from convalescent cattle (or from sheep or goats convalescing from capripox) or from rabbits hyperimmunised with purified capripoxvirus. Uninfected tissue culture should be included as a negative control as cross-reactions can cause problems due to antibodies to cellular components (pre-absorption of these from the immune serum helps solve this issue).

1.6.	Immunohistochemistry

Immunohistochemistry using F80G5 monoclonal antibody specific for capripoxvirus ORF 057 has been described for detection of LSDV antigen in the skin of experimentally infected cattle (Babiuk *et al.,*2008). 

1.7.	Isothermal genome amplification 

Molecular tests using loop-mediated isothermal amplification to detect capripoxvirus genomes are reported to provide sensitivity and specificity similar to real-time PCR with a simpler method and lower cost (Das *et al.,* 2012; Murray *et al.,* 2013). Field validation of the Das *et al.* method was reported by Omoga *et al.* (2016).

2.	Serological tests

All the viruses in the genus *Capripoxvirus*share a common major antigen for neutralising antibodies and it is thus not possible to distinguish strains of capripoxvirus from cattle, sheep or goats using serological techniques.

2.1.	Virus neutralisation

A test serum can either be titrated against a constant titre of capripoxvirus (100 TCID50 [50% tissue culture infective dose]) or a standard virus strain can be titrated against a constant dilution of test serum in order to calculate a neutralisation index. Because of the variable sensitivity of tissue culture to capripoxvirus, and the consequent difficulty of ensuring the accurate and repeatable seeding of 100 TCID50/well, the neutralisation index is the preferred method in most laboratories, although it does require a larger volume of test sera. The test is described using 96-well flat-bottomed tissue-culture grade microtitre plates, but it can be performed equally well in tissue culture tubes with the appropriate changes to the volumes used, although it is more difficult to read an end-point in tubes. 

2.1.1.	Test procedure

i)	Test sera, including a negative and a positive control, are diluted 1/5 in Eagle’s/HEPES (N-2-hydroxyethylpiperazine, N-2-ethanesulphonic acid) buffer and inactivated at 56°C for 30 minutes.

ii)	Next, 50 μl of the first inactivated serum is added to columns 1 and 2, rows A to H of the microtitre plate. The second serum is placed in columns 3 and 4, the third in columns 5 and 6, the positive control serum is placed in columns 7 and 8, the negative control serum is placed in columns 9 and 10, and 50 μl of Eagle’s/HEPES buffer (without serum) is placed in columns 11 and 12, and to all wells in row H.

iii)	A reference strain of capripoxvirus, usually a vaccine strain known to grow well in tissue culture, with a titre of over log10 6 TCID50 per ml is diluted in Eagle’s/HEPES in bijoux bottles to give a log dilution series of log10 5.0, 4.0, 3.5, 3.0, 2.5, 2.0, 1.5 TCID50 per ml (equivalent to log10 3.7, 2.7, 2.2, 1.7, 1.2, 0.7, 0.2 TCID50 per 50 μl).

iv)	Starting with row G and the most diluted virus preparation, 50 μl of virus is added to each well in that row. This is repeated with each virus dilution, the highest titre virus dilution being placed in row A.

v)	The plates are covered and incubated for 1 hour at 37°C. 

vi)	An appropriate cell suspension (such as MDBK cells) is prepared from pregrown monolayers as a suspension of 105 cells/ml in Eagle’s medium containing antibiotics and 2% fetal calf serum. Following incubation of the microtitre plates, 100 μl of cell suspension is added to all the wells, except wells H11 and H12, which serve as control wells for the medium. The remaining wells of row H are cell and serum controls.

vii)	The microtitre plates are covered and incubated at 37°C for 9 days.

viii)	Using an inverted microscope, the monolayers are examined daily from day 4 for evidence of CPE. There should be no CPE in the cells of row H. Using the 0240 KSGP vaccine strain of capripoxvirus, by way of example, the final reading is taken on day 9, and the titre of virus in each duplicate titration is calculated using the Kärber method. If left longer, there is invariably a ‘breakthrough’ of virus in which virus that was at first neutralised appears to disassociate from the antibody. 

ix)	*Interpretation of the results:* The neutralisation index is the log titre difference between the titre of the virus in the negative serum and in the test serum. An index of ≥1.5 is positive. The test can be made more sensitive if serum from the same animal is examined before and after infection. Because the immunity to capripoxviruses is predominantly cell mediated, a negative result, particularly following vaccination, after which the antibody response may be low, does not imply that the animal from which the serum was taken is not protected.

Antibodies to capripoxvirus can be detected from 1 to 2 days after the onset of clinical signs. These remain detectable for about 7 months.

2.2.	Enzyme-linked immunosorbent assay

Enzyme-linked immunosorbent assays (ELISAs) for the detection of capripoxviral antibodies are widely used and are available in commercial kit form (Milovanovic *et al.,* 2019; Samojlovic *et al.,* 2019). 

2.3.	Indirect fluorescent antibody test

Capripoxvirus-infected tissue culture grown on cover-slips or tissue culture microscope slides can be used for the indirect fluorescent antibody test. Uninfected tissue culture control, and positive and negative control sera, should be included in the test. The infected and control cultures are fixed in acetone at –20°C for 10 minutes and stored at 4°C. Dilutions of test sera are made in PBS, starting at 1/20 or 1/40, and positive samples are identified using an anti-bovine gamma-globulin conjugated with fluorescein isothiocyanate. Antibody titres may exceed 1/1000 after infection. Sera may be screened at 1/50 and 1/500. Cross-reactions can occur with orf virus (contagious pustular dermatitis virus of sheep), bovine papular stomatitis virus and perhaps other poxviruses.

2.4.	Western blot analysis

Western blotting of test sera against capripoxvirus-infected cell lysate provides a sensitive and specific system for the detection of antibody to capripoxvirus structural proteins, although the test is expensive and difficult to carry out.

Capripoxvirus-infected LT cells should be harvested when 90% CPE is observed, freeze–thawed three times, and the cellular debris pelleted using centrifugation. The supernatant should be decanted, and the proteins should be separated using SDS/PAGE (sodium dodecyl sulphate/polyacrylamide gel electrophoresis). A vertical discontinuous gel system, using a stacking gel made up of acrylamide (5%) in Tris (125 mM), pH 6.8, and SDS (0.1%), and a resolving gel made up of acrylamide (10–12.5%) in Tris (560 mM), pH 8.7, and SDS (0.1%), is recommended for use with a glycine running buffer containing Tris (250 mM), glycine (2 M), and SDS (0.1%). Samples of supernatant should be prepared by boiling for 5 minutes with an appropriate lysis buffer prior to loading. Alternatively, purified virus or recombinant antigens may replace tissue-culture-derived antigen.

Molecular weight markers should be run concurrently with the protein samples. The separated proteins in the SDS/PAGE gel should be transferred electrophoretically to a nitrocellulose membrane (NCM). After transfer, the NCM is rinsed thoroughly in PBS and blocked in 3% bovine serum albumin (BSA) in PBS, or 5% skimmed milk powder in PBS, on a rotating shaker at 4C overnight. The NCM can then be separated into strips by employing a commercial apparatus to allow the concurrent testing of multiple serum samples, or may be cut into strips and each strip incubated separately thereafter. The NCM is washed thoroughly with five changes of PBS for 5 minutes on a rotating shaker, and then incubated at room temperature on the shaker for 1.5 hours, with the appropriate serum at a dilution of 1/50 in blocking buffer (3% BSA and 0.05% Tween 20 in PBS; or 5% milk powder and 0.05% Tween 20 in PBS). The membrane is again thoroughly washed and incubated (in blocking buffer) with anti-species immunoglobulin horseradish-peroxidase-conjugated immunoglobulins at a dilution determined using titration. After further incubation at room temperature for 1.5 hours, the membrane is washed and a solution of diaminobenzidine tetrahydrochloride (10 mg in 50 ml of 50 ~~mm~~mM Tris/HCl, pH 7.5, and 20 μl of 30% [v/v] hydrogen peroxide) is added. Incubation is then undertaken for approximately 3–7 minutes at room temperature on a shaker with constant observation, and the reaction is stopped by washing the NCM in PBS before excessive background colour is seen. A positive and negative control serum should be used on each occasion.

Positive test samples and the positive control will produce a pattern consistent with reaction to proteins of molecular weights 67, 32, 26, 19 and 17 kDa – the major structural proteins of capripoxvirus – whereas negative serum samples will not react with all these proteins. Hyperimmune serum prepared against parapoxvirus (bovine papular stomatitis or pseudocowpox virus) will react with some of the capripoxvirus proteins, but not the 32 kDa protein that is specific for capripoxvirus.

C.  REQUIREMENTS FOR VACCINES[This section is under review in the 2023/2024 review cycle]

1.	Background: rationale and intended use of the product

Live attenuated strains of capripoxvirus have been used as vaccines specifically for the control of LSD (Brenner *et al*., 2006; Capstick & Coakley, 1961; Carn, 1993). Capripoxviruses are cross-reactive within the genus. Consequently, it is possible to protect cattle against LSD using strains of capripoxvirus derived from sheep or goats (Coakley & Capstick, 1961). However, it is recommended to carry out controlled trials, using the most susceptible breeds, prior to introducing a vaccine strain not usually used in cattle. The duration of protection provided by LSD vaccination is unknown. 

Capripoxvirus vaccine strains can produce a large local reaction at the site of inoculation in *Bos taurus* breeds (Davies, 1991), which some stock owners find unacceptable. This has discouraged the use of vaccine, even though the consequences of an outbreak of LSD are invariably more severe. Risk–benefit of vaccination should be assessed following stakeholder discussion.

2.	Outline of production and minimum requirements for conventional vaccines

General requirements set for the facilities used for the production of vaccines and for the documentation and record keeping throughout the whole manufacturing process are described in Chapter 1.1.8 *Principles of veterinary vaccine production*. The documentation should include standard operating procedures (SOP) for the method of manufacture and each step for the testing of cells and reagents used in the process, each batch and the final product.

2.1.	Characteristics of the seed

2.1.1.	Biological characteristics

Each seed strain of capripoxvirus used for vaccine production must be accompanied by records clearly and accurately describing its origin, isolation and tissue culture or animal passage history. Preferably, the species and strain of capripoxvirus are characterised using PCR or DNA sequencing techniques.

A quantity of master seed vaccine virus should be prepared, frozen or desiccated and stored at low temperatures such as –80°C and used to produce a consistent working seed for regular vaccine production. 

Each master seed strain must be non-transmissible, remain attenuated after further tissue culture passage, and provide complete protection against challenge with virulent field strains for a minimum of 1 year. It must produce a minimal clinical reaction in cattle when given via the recommended route.

The necessary safety and potency tests are described in Section C.2.2.4 *Final product batch tests*.

2.1.2.	Quality criteria (sterility, purity, freedom from extraneous agents)

Each master seed must be tested to ensure its identity and shown to be free from adventitious viruses, in particular pestiviruses, such as border disease and bovine viral diarrhoea virus, and free from contamination with bacteria, fungi or mycoplasmas.

The general procedures for sterility or purity tests are described in Chapter 1.1.9 *Tests for sterility and freedom from contamination of biological materials intended for veterinary use*.

2.2.	Method of manufacture

The method of manufacture should be documented as the Outline of Production.

2.2.1.	Procedure

Vaccine batches are produced on an appropriate cell line such as MDBK. The required number of vials of seed virus is reconstituted with GMEM or other appropriate medium and inoculated onto a monolayer. Cells should be harvested after 4–8 days when they exhibit 50–70% CPE for maximum viral infectivity, or earlier if CPE is extensive and cells appear ready to detach. Techniques such as sonication or repeated freeze–thawing are used to release the intracellular virus from the cytoplasm. The lysate may then be clarified to remove cellular debris (for example by use of centrifugation at 600 ***g*** for 20 minutes, with retention of the supernatant). A second passage of the virus may be required to produce sufficient virus for a production batch.

An aliquot of the virus suspension is titrated to check the virus titre. The virus-containing suspension is then mixed with a suitable protectant such as an equal volume of sterile, chilled 5% lactalbumin hydrolysate and 10% sucrose (dissolved in double-distilled water or appropriate balanced salt solution), and transferred to individually numbered bottles for storage at low temperatures such as –80°C, or for freeze–drying. A written record of all the procedures followed must be kept for all vaccine batches.

2.2.2.	Requirements for substrates and media

The specification and source of all ingredients used in the manufacturing procedure should be documented and the freedom of extraneous agents (bacteria, fungi, mycoplasma and viruses) should be tested. The detailed testing procedure is described in Chapter 1.1.9. The use of antibiotics must meet the requirements of the licensing authority. 

2.2.3.	In-process control

i)	Cells

Records of the source of the master cell stocks should be maintained. The highest and lowest passage numbers of the cells that can be used for vaccine production must be indicated in the Outline of the Production. The use of a continuous cell line (such as MDBK, etc.) is strongly recommended, unless the virus strain only grows on primary cells. The key advantage of continuous over primary cell lines is that there is less risk of introduction of extraneous agents.

ii)	Serum

Serum used in the growth or maintenance medium must be free from antibodies to capripoxvirus and free from contamination with pestivirus or other viruses, extraneous bacteria, mycoplasma or fungi.

iii)	Medium

Media must be sterile before use.

iv)	Virus

Seed virus and final vaccine must be titrated and pass the minimum release titre set by the manufacturer. For example, the minimum recommended field dose of the South African Neethling strain vaccines (Mathijs *et al.,* 2016) is log10 3.5 TCID50, although the minimum protective dose is log10 2.0 TCID50. Capripoxvirus is highly susceptible to inactivation by sunlight and allowance should be made for loss of activity in the field. 

The recommended field dose of the Romanian sheep pox vaccine for cattle is log10 2.5 sheep infective doses (SID50), and the recommended dose for cattle of the RM65-adapted strain of Romanian sheep pox vaccine is log10 3 TCID50 (Coakley & Capstick, 1961). 

2.2.4.	Final product batch tests

i)	Sterility/purity

Vaccine samples must be tested for sterility/purity. *Tests for sterility and freedom from contamination of biological materials intended for veterinary use* may be found in Chapter 1.1.9.

ii)	Safety and efficacy

The efficacy and safety studies should be demonstrated using statistically valid vaccination–challenge studies using seronegative young LSDV susceptible dairy cattle breeds. The group numbers recommended here can be varied if statistically justified. Fifteen cattle are placed in a high containment level large animal unit and serum samples are collected. Five randomly chosen vials of the freeze-dried vaccine are reconstituted in sterile PBS and pooled. Two cattle are inoculated with 10 times the recommended field dose of the vaccine, and eight cattle are inoculated with the recommended field dose. The remaining five cattle are unvaccinated control animals. The animals are clinically examined daily and rectal temperatures are recorded. On day 21 after vaccination, the animals are again serum sampled and challenged with a known virulent capripoxvirus strain. The challenge virus solution should also be tested free from extraneous viruses. The clinical response is recorded during the following 14 days. Animals in the unvaccinated control group should develop the typical clinical signs of LSD, whereas there should be no local or systemic reaction in the vaccinates other than a  raised area in the skin at the site of vaccination, which should disappear after 4 days. Serum samples are again collected on day 30 after vaccination. The day 21 serum samples are examined for seroconversion to selected viral diseases that could have contaminated the vaccine, and the days 0 and 30 samples are compared to confirm the absence of antibody to pestivirus. Because of the variable response in cattle to LSD challenge, generalised disease may not be seen in all of the unvaccinated control animals, although there should be a large local reaction.

Once the efficacy of the particular strain being used for vaccine production has been determined in terms of minimum dose required to provide immunity, it is not necessary to repeat this on the final product of each batch, provided the titre of virus present has been ascertained.

iii)	Batch potency 

Potency tests in cattle must be undertaken for vaccine strains of capripoxvirus if the minimum immunising dose is not known. This is usually carried out by comparing the titre of a virulent challenge virus on the flanks of vaccinated and control animals. Following vaccination, the flanks of at least three animals and three controls are shaved of hair. Log10 dilutions of the challenge virus are prepared in sterile PBS and six dilutions are inoculated intradermally (0.1 ml per inoculum) along the length of the flank; four replicates of each dilution are inoculated down the flank. An oedematous swelling will develop at possibly all 24 inoculation sites on the control animals, although preferably there will be little or no reaction at the four sites of the most dilute inocula. The vaccinated animals may develop an initial hypersensitivity reaction at sites of inoculation within 24 hours, which should quickly subside. Small areas of necrosis may develop at the inoculation site of the most concentrated challenge virus. The titre of the challenge virus is calculated for the vaccinated and control animals; a difference in titre >log10 2.5 is taken as evidence of protection.

2.3.	Requirements for regulatory approval

2.3.1.	Safety requirements

i)	Target and non-target animal safety

The vaccine must be safe to use in all breeds of cattle for which it is intended, including young and pregnant animals. It must also be non-transmissible and remain attenuated after further tissue culture passage.

Safety tests should be carried out on the final product of each batch as described in Section C.2.2.4.

ii)	Reversion-to-virulence for attenuated/live vaccines

The selected final vaccine should not revert to virulence during further passages in target animals.

iii)	Environmental consideration

Attenuated vaccine should not be able to perpetuate autonomously in a cattle population. Strains of LSDV are not a hazard to human health.

2.3.2.	Efficacy requirements

i)	For animal production

The efficacy of the vaccine must be demonstrated in statistically valid vaccination challenge experiments under laboratory conditions. The group numbers recommended here can be varied if statistically justified. Fifteen cattle are placed in a high containment level large animal unit and serum samples are collected. Five randomly chosen vials of the freeze-dried vaccine are reconstituted in sterile PBS and pooled. Two cattle are inoculated with 10 times the field dose of the vaccine, eight cattle are inoculated with the recommended field dose. The remaining five cattle are unvaccinated control animals. The animals are clinically examined daily and rectal temperatures are recorded. On day 21 after vaccination, the animals are again serum sampled and challenged with a known virulent capripoxvirus strain using intravenous and intradermal inoculation (the challenge virus solution should also be tested and shown to be free from extraneous viruses). The clinical response is recorded during the following 14 days. Animals in the unvaccinated control group should develop the typical clinical signs of LSD, whereas there should be no local or systemic reaction in the vaccinates other than a raised area in the skin at the site of vaccination which should disappear after 4 days. Serum samples are again collected on day 30 after vaccination. The day 21 serum samples are examined for seroconversion to selected viral diseases that could have contaminated the vaccine, and the days 0 and 30 samples are compared to confirm the absence of antibody to pestivirus. Because of the variable response in cattle to challenge with LSDV, generalised disease may not be seen in all of the unvaccinated control animals, although there should be a large local reaction.

Once the potency of the particular strain being used for vaccine production has been determined in terms of minimum dose required to provide immunity, it is not necessary to repeat this on the final product of each batch, provided the titre of virus present has been ascertained.

ii)	For control and eradication

Vaccination is the only effective way to control LSD outbreaks in endemic countries and recent experiences of the disease in Eastern Europe and the Balkans suggests this is also true for outbreaks in non-endemic countries. Unfortunately, currently no marker vaccines allowing a~~the differentiation of infected from vaccinated~~animals (DIVA~~)~~strategy are available, although to a limited extent PCR can be used for certain vaccines. 

The duration of immunity produced by LSDV vaccine strains is currently unknown. 

2.3.3.	Stability

All vaccines are initially given a shelf life of 24 months before expiry. Real-time stability studies are then conducted to confirm the appropriateness of the expiry date. Multiple batches of the vaccine should be re-titrated periodically throughout the shelf-life period to determine the vaccine stability. 

Properly freeze-dried preparations of LSDV vaccine, particularly those that include a protectant, such as sucrose and lactalbumin hydrolysate, are stable for over 25 years when stored at –20°C and for 2–4 years when stored at 4°C. There is evidence that they are stable at higher temperatures, but no long-term controlled experiments have been reported. No preservatives other than a protectant, such as sucrose and lactalbumin hydrolysate, are required for the freeze-dried preparation.

3.	Vaccines based on biotechnology

A new generation of capripox vaccines is being developed that uses the LSDV as a vector for the expression and delivery of immuno-protective proteins of other ruminant pathogens with the potential for providing dual protection (Boshra *et al*., 2013; Wallace & Viljoen, 2005), as well as targeting putative immunomodulatory genes for inducing improved immune responses (Kara *et al.,* 2018). 

REFERENCES

AGIANNIOTAKI E.I., CHAINTOUTIS S.C., HAEGEMAN A., TASIOUDI K.E., DE LEEUW I., KATSOULOS P.D., SACHPATZIDIS A., DE CLERCQ K., ALEXANDROPOULOS T., POLIZOPOULOU Z.S., CHONDROKOUKI E.D. & DOVAS C.I. (2017). Development and validation of a TaqMan probe-based real-time PCR method for the differentiation of wild type lumpy skin disease virus from vaccine virus strains. *J. Virol. Methods*, **249**, 48–57.

Babiuk S., Bowden T.R., Parkyn G., Dalman B., Manning L., Neufeld J., Embury-Hyatt C., Copps J. & Boyle D.B. (2008). Quantification of lumpy skin disease virus following experimental infection in cattle. *Transbound. Emerg. Dis.*, **55**, 299–307.

Babiuk S., Parkyn G., Copps J., Larence J.E., Sabara M.I., Bowden T.R., Boyle D.B. & Kitching R.P. (2007). Evaluation of an ovine testis cell line (OA3.Ts) for propagation of capripoxvirus isolates and development of an immunostaining technique for viral plaque visualization. *J. Vet. Diagn.Invest.*, **19**, 486–491. 

BALINSKY C.A, DELHON G, SMOLIGA G, PRARAT M, FRENCH R.A, GEARY S.J, ROCK D.L & RODRIGUEZ L.L. (2008). Rapid preclinical detection of sheep pox virus by a real-time PCR assay*. J. Clin. Microbiol*., **46**, 438–442.

BYADOVSKAYA O., PESTOVA Y., KONONOV A., SHUMILOVA I., KONONOVA S., NESTEROV A., BABIUK S. & SPRYGIN A. (2021). Performance of the currently available DIVA real-time PCR assays in classical and recombinant lumpy skin disease viruses. *Transbound. Emerg. Dis*., **68**, 3020-3024. doi: 10.1111/tbed.13942. Epub 2021 May 13. PMID: 33253485. 

BISWAS S., NOYCE R.S., BABIUK L.A., LUNG O., BULACH D. M., BOWDEN T.R., BOYLE D. B., BABIUK S. & EVANS D.H. (2020). Extended sequencing of vaccine and wild-type capripoxvirus isolates provides insights into genes modulating virulence and host range. *Transbound. Emerg. Dis.*, **67**, 80–97.

BOSHRA H., TRUONG T., NFON C., GERDTS V., TIKOO S., BABIUK L.A., KARA P., MATHER A., WALLACE D. & BABIUK S. (2013). Capripoxvirus-vectored vaccines against livestock diseases in Africa. *Antiviral Res*., **98**, 217–227. 

Bowden, T.R, Babiuk S.L, Parkyn G.R., Copps J.S. & Boyle D.B. (2008). Capripoxvirus tissue tropism and shedding: A quantitative study in experimentally infected sheep and goats. *Virology*, **371**, 380–393. 

BRENNER J., HAIMOVITZ M., ORON E., STRAM Y., FRIDGUT O., BUMBAROV V., KUZNETZOVA L., OVED Z., WASERMAN A., GARAZZI S., PERL S., LAHAV D., EDERY N. & YADIN H. (2006). Lumpy skin sease (LSD) in a large dairy herd in Israel. *Isr. J. Vet. Med*., **61**, 73–77.

BURDIN M.L. (1959). The use of histopathological examination of skin material for the diagnosis of lumpy skin disease in Kenya. *Bull. Epizoot. Dis. Afr*., **7**, 27–36.

CAPSTICK P.B. & COAKLEY W. (1961). Protection of cattle against lumpy skin disease. Trials with a vaccine against Neethling type infection. *Res. Vet. Sci*., **2**, 362–368

CARN V.M. (1993). Control of capripoxvirus infections. *Vaccine*, **11**, 1275–1279.

CARN V.M. & KITCHING, R.P. (1995). The clinical response of cattle following infection with lumpy skin disease (Neethling) virus. *Arch. Virol.*, **140**, 503–513.

COAKLEY W. & CAPSTICK P.B. (1961). Protection of cattle against lumpy skin disease. Factors affecting small scale production of tissue culture propagated virus vaccine. *Res. Vet. Sci*., **2**, 369–371.

COETZER J.A.W. (2004). Lumpy skin disease. *In:* Infectious Diseases of Livestock, Second Edition Coetzer J.A.W. & Justin R.C., eds. Oxford University Press, Cape Town, South Africa, 1268–1276.

DAS A., BABIUK S. & MCINTOSH M.T. (2012). Development of a loop-mediated isothermal amplification assay for rapid detection of capripoxviruses. *J. Clin. Microbiol.,***50**, 1613–1620.

DAVIES F.G. (1991). Lumpy Skin Disease, a Capripox Virus Infection of Cattle in Africa. FAO, Rome, Italy.

DAVIES F.G., KRAUSS H., LUND L.J. & TAYLOR M. (1971). The laboratory diagnosis of lumpy skin disease. *Res. Vet. Sci*., **12**, 123–127.

FAY P.C., COOK C.G., WIJESIRIWARDANA N., TORE G., COMTET L., CARPENTIER A., SHIH B., FREIMANIS G., HAGA I.R. & BEARD P.M. (2020). Madin-Darby bovine kidney (MDBK) cells are a suitable cell line for the propagation and study of the bovine poxvirus lumpy skin disease virus. *J. Virol. Methods*, **285**, 113943. doi: 10.1016/j.jviromet.2020.113943.

FLANNERY J., SHIH B., HAGA I.R., ASHBY M., CORLA A., KING S., FREIMANIS G., POLO N., TSE A.C., BRACKMAN C.J., CHAN J., PUN P., FERGUSON A.D., LAW A., LYCETT S., BATTEN C. & BEARd P.M. (2021). A novel strain of lumpy skin disease virus causes clinical disease in cattle in Hong Kong. *Transbound. Emerg. Dis*., doi: 10.1111/tbed.14304 

HAIG D. (1957). Lumpy skin disease. *Bull. Epizoot. Dis. Afr.*, **5**, 421–430.

IRELAND D.C. & BINEPAL Y.S. (1998). Improved detection of capripoxvirus in biopsy samples by PCR. *J. Virol. Methods*, **74**, 1–7.

IRONS P.C., TUPPURAINEN E.S.M. & VENTER E.H. (2005). Excretion of lumpy skin disease virus in bull semen. *Theriogenology*, **63**, 1290–1297.

KARA P.D., AFONSO C.L., WALLACE D.B., KUTISH G.F., ABOLNIK C., LU Z., VREEDE F.T., TALJAARD L.C., ZSAK A., VILJOEN G.J. & ROCK D.L. (2003). Comparative sequence analysis of the South African vaccine strain and two virulent field isolates of Lumpy skin disease virus. *Arch. Virol*., **148**, 1335–1356.

KARA P.D., MATHER A.S., PRETORIUS A., CHETTY T., BABIUK S. & WALLACE D.B. (2018). Characterisation of putative immunomodulatory gene knockouts of lumpy skin disease virus in cattle towards an improved vaccine. *Vaccine*, **36**, 4708-4715. doi: 10.1016/j.vaccine.2018.06.017.

KITCHING R.P. & SMALE C. (1986). Comparison of the external dimensions of capripoxvirus isolates. *Res. Vet. Sci*., **41**, 425–427. 

LAMIEN C.E., LELENTA M., GOGER W., SILBER R., TUPPURAINEN E., MATIJEVIC M., LUCKINS A.G. & DIALLO A. (2011). Real time PCR method for simultaneous detection, quantitation and differentiation of capripoxviruses *J. Virol. Methods*, **171**, 134–140. 

MATHIJS E., VANDENBUSSCHE F., HAEGEMAN A., KING A., NTHANGENI B., POTGIETER C., MAARTENS L., VAN BORM S. &DE CLERCQ K. (2016). Complete Genome Sequences of the Neethling-Like Lumpy Skin Disease Virus Strains Obtained Directly from Three Commercial Live Attenuated Vaccines. *Genome Announc*., **4** (6). pii: e01255-16. doi: 10.1128/genomeA.01255-16.

MILOVANOVIC M., DIETZE K., MILICEVIC V., RADOJICIC S., VALCIC M., MORITZ T. & HOFFMANN B. (2019). Humoral immune response to repeated lumpy skin disease virus vaccination and performance of serological tests. *BMC Vet. Res*., **15**, 80. doi: 10.1186/s12917-019-1831-y.

MURRAY L., EDWARDS L., TUPPURAINEN E.S., BACHANEK-BANKOWSKA K., OURA C.A., MIOULET V. & KING D.P. (2013). Detection of capripoxvirus DNA using a novel loop-mediated isothermal amplification assay.***BMC Vet. Res**.*, **9**, 90.

OMOGA D.C.A., MACHARIA M., MAGIRI E., KINYUA J., KASIITI J. & HOLTON T. (2016). Molecular based detection, validation of a LAMP assay and phylogenetic analysis of capripoxvirus in Kenya. *J. Adv. Biol. Biotech*., **7**, 1–12.

PESTOVA Y., BYADOVSKAYA O., KONONOV A. & SPRYGIN A. (2018). A real time high-resolution melting PCR assay for detection and differentiation among sheep pox virus, goat pox virus, field and vaccine strains of lumpy skin disease virus. *Mol. Cell. Probes*, **41**, 57–60.

PROZESKY L. & BARNARD B.J.H. (1982). A study of the pathology of lumpy skin disease in cattle. *Onderstepoort J. Vet. Res.*, **49**, 167–175.

ROUBY S. & ABOULSOUDB E. (2016). Evidence of intrauterine transmission of lumpy skin disease virus. *Vet. J*., **209**, 193–195. 

SAMOJLOVIC M., POLACEK V., GURJANOV V., LUPULOVIC D., LAZIC G., PETROVIĆ T., LAZIC S. (2019). Detection of antibodies against lumpy skin disease virus by virus neutralization test and ELISA methods. *Acta Vet*., **69**, 47–60.

SPRYGIN A., BABIN Y., PESTOVA Y., KONONOVA S., WALLACE D.B., VAN SCHALKWYK A., BYADOVSKAYA O., DIEV V., LOZOVOY D. & KONONOV A. (2018). Analysis and insights into recombination signals in lumpy skin disease virus recovered in the field. *PLoS One*, **13**, e0207480.

SPRYGIN A., VAN SCHALKWYK A., SHUMILOVA I., NESTEROV A., KONONOVA S., PRUTNIKOV P., BYADOVSKAYA O. & KONONOV A. (2020). Full-length genome characterization of a novel recombinant vaccine-like lumpy skin disease virus strain detected during the climatic winter in Russia, 2019. *Arch. Virol*., **165**, 2675–2677.

TUPPURAINEN E.S.M., VENTER E.H. & COETZER J.A.W. (2005). The detection of lumpy skin disease virus in samples of experimentally infected cattle using different diagnostic techniques. *Onderstepoort J. Vet. Res.,***72**, 153–164.

Tuppurainen E.S., Venter E.H., Coetzer J.A. & Bell-Sakyi L. (2015). Lumpy skin disease: attempted propagation in tick cell lines and presence of viral DNA in field ticks collected from naturally-infected cattle. *Ticks Tick Borne Dis*., **6**, 134–140.

Van Rooyen P.J., Komm N.A.L., Weiss K.E. & Alexander R.A. (1959). A preliminary note on the adaptation of a strain of lumpy skin disease virus to propagation in embryonated eggs. *Bull. Epizoot. Dis. Afr*., **7**, 79–85.

Van Schalkwyk A., Byadovskaya O., Shumilova I., Wallace D.B. & Sprygin A. (2021). Estimating evolutionary changes between highly passaged and original parental lumpy skin disease virus strains. *Transbound. Emerg. Dis*., doi: 10.1111/tbed.14326.

Van Schalkwyk A., Kara P., Ebersohn K., Mather A., Annandale C.H., Venter E.H. & Wallace D.B. (2020). Potential link of single nucleotide polymorphisms to virulence of vaccine-associated field strains of lumpy skin disease virus in South Africa. *Transbound. Emerg. Dis*., **67**, 2946–2960.

Vidanovic D., Sekler M., Petrovic T., Debeljak Z., Vaskovic N., Matovic K. & Hoffman B. (2016). Real time PCR assay for the specific detection of field Balkan strain of lumpy skin disease virus. *Acta Vet. Brno*, **66**, 444–454.

Wang Y., Zhao L., Yang J., Shi M., Nie F., Liu S., Wang Z., Huang D., Wu H., Li D., Lin H. & Li Y. (2021). Analysis of vaccine-like lumpy skin disease virus from flies near the western border of China. *Transbound. Emerg. Dis*., doi: 10.1111/tbed.14159

WALLACE D.B. & VILJOEN G.J. (2005). Immune responses to recombinants of the South African vaccine strain of lumpy skin disease virus generated by using thymidine kinase gene insertion. *Vaccine*, **23**, 3061–3067.

WEISS K.E. (1968). Lumpy skin disease. *Virol. Monogr*., **3**, 111–131.

**   *

**NB:** There are WOAH Reference Laboratories for lumpy skin disease (please consult the WOAH Web site: https://www.woah.org/en/what-we-offer/expertise-network/reference-laboratories/#ui-id-3).Please contact WOAH Reference Laboratories for any further information on diagnostic tests, reagents and vaccines for lumpy skin disease

**NB:** FIRST ADOPTED IN 1989. MOST RECENT UPDATES ADOPTED IN 2021.

WOAH *Terrestrial Manual* 2023	1

2	WOAH *Terrestrial Manual* 2023

WOAH *Terrestrial Manual* 2023	15