Maedi-visna infection in Pomeranian Coarsewool Sheep in Germany: seroprevalence, environmental and genetic risk factors

Small ruminant lentiviruses (SRLV) cause maedi-visna (MV), an incurable, wasting disease, affecting sheep worldwide. This study evaluated the seroprevalence of maedi-visna virus (MVV) in the native German sheep breed Pomeranian Coarsewool Sheep (RPL) and environmental and genetic risk factors for individual and flock-level seropositivity. A total of 849 samples from 35 farms across nine German states were analyzed for MVV antibodies with an enzyme-linked immunosorbent assay (ELISA). Individual seroprevalence and clinical prevalence were 3.5% (30/849) and 13.3% (4/30), respectively. Seropositive sheep were detected in six flocks, with prevalences between 5.3 and 37.5%. Flocks in Eastern Germany and sheep older than 3 years showed the highest seroprevalence. Purchase of ewes was the main environmental risk factor for seropositivity. ELISA-based serotyping of SRLV in the MVV-positive sheep identified, besides genotype A, for the first time in Germany genotype B, whereas molecular genotyping, based on Sanger sequencing, identified only genotype A (A1, A3, A21, and A21-like). Apart from the obvious inconsistencies between the results of these two methods, presence of different viral sero- or genotypes suggest multiple virus introduction routes. Sequencing of exons 2 and 3 of the transmembrane protein 154 gene (TMEM154) of 530 sheep revealed non-synonymous variation at codons 35, 44 (known to be in linkage with a frame-shift variant coded in exon 1), and 70. Analysis of association between the putatively protective TMEM154 codon 35 genotype KK and MVV seropositivity showed only a tendency toward significance (p = 0.088), whereas seroprevalence was significantly (p = 0.001) lower in sheep exclusively carrying the K allele at codon 35 and/or the M allele at codon 44. In contrast, MVV seropositivity was significantly higher in sheep with one or two I alleles at codon 70 (p = 0.045). Despite a low seroprevalence in the RPL breed in Germany, regular MVV monitoring is strongly recommended to avoid its spread. An eradication program based on serological testing and removal of seropositive sheep should be implemented. Selection for certain TMEM154 alleles in affected flocks could support eradication measures, but their effect on MVV susceptibility in this breed requires further investigation.

1 Introduction

Maedi-visna (MV), originally described as a sheep-associated slow virus infection, together with caprine arthritis and encephalitis (CAE) in goats, forms the group of small ruminant lentiviruses (SRLV). The genus lentivirus belongs to the non-oncogenic retroviruses, which also include human (HIV) and feline (FIV) immunodeficiency viruses (1). In contrast to these immunodeficiency viruses, maedi-visna virus (MVV) does not infect T-cells and has a tropism for cells of the monocyte–macrophage lineage (2). Replication of the virus in macrophages may lead to inflammatory lesions in the lungs, joints, mammary glands, and brain (3, 4). Clinical signs such as dyspnea, dry cough, chronic weight loss, lameness, or mammary induration appear at the earliest 1–3 years after infection and sometimes do not manifest at all, indicating a subclinical infection (5–7).

MV is a persistent infection and no vaccines or treatment options are available yet (8). The main route of transmission is via the colostrum or milk of an infected ewe to its offspring (9). Horizontal transmission through close contact is another important route and has also been described between sheep and goats (10, 11). To prevent and eliminate the spread of MVV infection, regular monitoring of flocks, early detection, and culling of infected animals are essential (12, 91). In large flocks with high seroprevalences, motherless rearing of lambs is also a successful but costly sanitation option (13). Another cost-effective prevention and eradication strategy could be breeding for less MVV susceptible sheep (14). Some breeds, such as Texel and East Frisian milk sheep, are known to be highly susceptible to MVV, while in other breeds, such as Merinoland sheep, MVV-positive animals were rarely observed (15). These breed differences suggest that genetic factors play an important role in MVV susceptibility. It was reported that variants of the ovine transmembrane protein 154 gene (TMEM154) are associated with the serological MV status (16) and/or provirus concentration (17). Haplotypes 1–4 are the most common TMEM154 haplotypes in sheep, and all four affect MVV susceptibility (16, 18). Sheep with one copy of either haplotype 2 or 3 (ancestral), both encoding glutamate (E) at position 35, were associated with an increased risk of MVV infection, while sheep homozygous for the derived haplotype 1 with lysine (K) at position 35 were less susceptible (16). Also in German sheep flocks with specific breed backgrounds (Texel, Brown Hair Sheep, East Frisian Milk Sheep, and Lacaune), animals with codon 35 genotypes EK and EE showed a significantly higher risk of being seropositive under MVV infection pressure than sheep with the genotype KK (19, 20). A protective effect was also observed for TMEM154 haplotype 4, defined by a frameshift mutation (A4Δ53) in exon 1, representing a naturally occurring knock-out allele, so that alternate homozygotes do not have a functional TMEM154 protein (16, 21, 22). The A4Δ53 variant is in complete linkage with a threonine to methionine exchange (T44M) encoded in exon 2. Haplotype 2 differs from the ancestral haplotype 3 only by a single amino acid substitution from asparagine to isoleucine at position 70 (N70I). Although both haplotypes (2 and 3) were previously shown to be associated with increased susceptibility to MVV, Arcangeli et al. (23) were the first to report a significantly higher risk for sheep carrying one or two copies of haplotype 2 (codon 70 genotypes IN and II) compared to sheep without (NN).

However, not only host genetics but also pathogen genetics is an important risk factor for MVV infection (24, 25). SRLV are divided into four genotypes (A, B, C, and E), which differ from each other by 25–37% in their nucleotide sequences. The fifth putative group D has been reclassified as genotype A, but is still used by many authors (12). Genotype A is a large heterogeneous group with 27 up-to-date subgenotypes containing MVV-like strains. Genotype B, with five subgenotypes, includes CAEV-like strains (26). Both genotypes have been shown to be able to cross the species barrier and to be distributed worldwide in sheep and goats (11, 27). The other two genotypes are geographically restricted to small ruminants in Norway (genotype C) and goats in Italy (genotype E with 2 subgenotypes) (28–31). However, due to high variability of SRLVs, new variants are constantly emerging as more strains are analyzed (32).

Maedi (Icelandic word for dyspnoea) visna (Icelandic word for wasting) is named after the two typical clinical forms that were first described in Iceland in the 1930s after an import of Karakul rams from Germany (33). Progressive pneumonia, named maedi, is the most common form (34). In contrast, visna, a demyelinating leukoencephalomyelitis, is described only rarely (35, 36). Today, Iceland, Australia, and New Zealand are the only MVV-free countries. Except for these islands, the virus is distributed worldwide (33, 37) and is causing major economic losses in many countries (38). In 2005, Graber and Ganter (39) performed a nationwide survey of the epidemiological features of SRLV infections in Germany, including data from sheep and goat breeding associations and livestock health services. Results showed that only 0.21% of the German sheep population (0.93% of sheep flocks) and approximately 4.2% of the goat population had an accredited MV or CAE negative status. Another study by Hüttner et al. (40) observed a herd prevalence of 51.2% in their representative MVV screening study in Mecklenburg-Western Pomerania. These results show that the MVV is also widespread in Germany, particularly in the Texel breed, but can also occur in other breeds such as Merinoland and East Friesian milk sheep (15).

Pomeranian Coarsewool Sheep (in German ´Rauhwolliges Pommersches Landschaf´, RPL) is an old German land sheep breed that originated from Pomerania and was first named RPL in 1860 (41). In 1982, the breed was almost extinct and a conservation-breeding program was started with 46 ewes, 4 yearlings, and 7 rams (42, 43). Later, the S line, which came from a breeder in southern Germany, was added to the seven existing ram lines (44). The number of RPL pedigree sheep is steadily increasing and the Central Documentation of Animal Genetic Resources in Germany registered almost 3,200 RPL sheep in 2022 (45). RPL is a robust and resilient breed that is less susceptible to certain diseases (46, 47), but there are no data available on its susceptibility to MVV. The first MVV-positive RPL flock was detected by Hüttner et al. (40) in 2010. Sporadic serological investigations in other RPL flocks revealed negative results (unpublished data). The aim of this study was to determine the current seroprevalence of MVV in the RPL sheep population in Germany using an enzyme-linked immunosorbent assay (ELISA) and to identify both environmental and genetic risk factors for seropositivity, focusing on SRLV genotypes (pathogen genetics) and TMEM154 genotypes (host genetics).

2 Materials and methods

2.1 Ethics approval statement

Before implementation, all animal procedures, beyond those only included in the voluntary MV monitoring of sheep flocks, were reviewed and approved by the respective state veterinary offices of the different German states involved (approval numbers: Baden-Württemberg 35-9185.82/A-11/23 (8 August 2023), Brandenburg LAVG-V6-2347-19-2023- 23-E (7 August 2023), Hesse V 54 19 c 20 15 h 01 Nr. V 3/2023 (21 August 2023), Lower Saxony 33.19-42502-04-23-00387 (4 August 2023), Mecklenburg-Western Pomerania 7221.3-2-012/23 (5 September 2023), North Rhine-Westphalia 81-02.04.40.2023. VG036 (11 August 2023), Saxony 25-5131/564/17 (9 October 2023), Saxony-Anhalt 203.m-42502-6- 011_TiHo_G (25 September 2023), Schleswig Holstein IX 552-117467/2023 (56-9/23 V) (25 September 2023)). The handling and sampling of the sheep followed European Union guidelines for animal care and handling, as well as Guidelines for Good Veterinary Practices.

2.2 Sample collection and DNA extraction

Between August and December 2023, blood samples were taken by veterinarians using 9 mL EDTA monovettes from jugular vein or vena cava cranialis of RPL sheep. The latter method is especially used in unshorn sheep (48). The study involved 54 flocks from nine German states, which were divided into three different study areas: North-western Germany (Schleswig Holstein, North Rhine-Westphalia, Lower Saxony), Eastern Germany (Saxony, Saxony-Anhalt, Brandenburg, Mecklenburg-Western Pomerania), and Southern Germany (Hesse, Baden-Württemberg) (number of sampled flocks per German state is given in Figure 1). In each flock, all breeding rams over 1 year of age and 1–10 unrelated females per ram line (no first- or second-degree relatedness) were sampled based on pedigree information, resulting in a total of 530 samples for sequence analysis of TMEM154 exon 2 and 3.

Figure 1

Figure 1. Location and numbers of the RPL flocks sampled in nine different states in Germany. Numbers without brackets: TMEM154 genotyped flocks; Numbers in brackets: TMEM154 genotyped and MVV monitored flocks; Blue: Northwestern Germany; Green: Eastern Germany; Orange; Southern Germany.

Additionally, 35 of these flocks participated in the voluntary, but reportable, MV monitoring (Table 1). For this purpose, all RPL sheep over 1 year of age (plus four sheep aged 10 months) were sampled per flock, resulting in a total of 849 samples for MVV serology. The average flock size of the MVV-tested flocks was 52 RPL sheep (range: 7–295) (Table 1).

Combining the data from both analyses, a total of 395 RPL were tested for both TMEM154 genotyping and MVV serology.

Table 1

Table 1. Number and percentage of MVV-tested flocks and sheep in the three study areas.

EDTA-blood samples were stored at <+8 °C after collection for up to 5 days. Blood samples for MVV serology were centrifuged and plasmas were separated before freezing at −20 °C until use. DNA was extracted from whole blood using the NucleoMag Blood 200 μL kit (Machery-Nagel GmbH & Co. KG, Dueren, Germany), which is based on magnetic separation by reversible adsorption of nucleic acids to paramagnetic beads under appropriate buffer conditions. DNA quality and quantity were measured with a Nanodrop ND-1000 spectrophotometer (Thermo Fisher Scientific Inc., Waltham, MA, USA). DNA samples were stored at −20 °C until use.

2.3 Questionnaire

A questionnaire was developed and sent to each RPL breeder of the 54 flocks to be completed before, during, or after the farm visit for blood sampling. The aim of the questionnaire was to obtain flock-level information including flock size, pedigree breeding, management practices (type of housing, mating, lambing, and lamb rearing), sheep purchases, presence of other sheep breeds or goats on the farm, health measures (such as vaccinations and prevention of ecto- and endoparasites), and potentially known MV status. However, only the 35 questionnaires corresponding to flocks tested serologically for MVV were used for statistical risk analysis.

2.4 MVV serology

All plasma samples were tested by using a commercial competitive ELISA test kit (VMRD Inc., Pullman, Washington, USA), according to the manufacturer’s instructions. Based on the optical density (OD) results, the percent inhibition (PI) was calculated as follows:

$\mathit{PI}=100[1-(\frac{\mathit{\text{Sample}}\mathit{OD}}{\mathit{\text{Negative Control}}\mathit{OD}}\left)\right]$

According to the manufacturer’s recommendations, a sample was classified as seropositive when it produced ≥ 35% inhibition.

Although based on the CAEV envelope glycoprotein, the sensitivity and specificity of the assay were 98.6 and 96.9%, respectively, when used with sheep sera (49).

2.5 SRLV serotyping and genotyping

Twenty-nine of the 30 seropositive plasmas were serotyped with the indirect Eradikit ELISA for SRLV genotyping (Eradikit SRLV Genotyping IN3 Diagnostic, Torino, Italy). The genotyping ELISA plates are coated with specific recombinant subunits of genotypes A, B, and E on three separate strips. Genotype-specific antibodies bind to the corresponding antigen and lead to colour development. To determine a positive result for one genotype, the measured OD value must be > 0.4, and 40% above the other OD values. The results were classified as indeterminate if the OD values of all genotypes were < 0.4, or if the difference between the two most reactive antigens was too low (< 40%). In the latter case, a double infection could be possible, according to the manufacturer’s instructions.

To obtain more precise and comprehensive data, a nested end-point PCR amplification of the SRLV gag-pol region was carried out with DNA samples from all seropositive sheep. In the first round, a 1,300 bp fragment was amplified. In the second round, an 800 bp fragment was obtained using the first-round amplified product as template. The following set of primers was used: GAGF1 for 5′-TGGTGARKCTAGMTAGAGACATGG-3′ and POLR1 rev 5′-CATAGGRGGHGCGGACGGCASCA-3′ in the first round, and GAGF2 for 5′-CAAACWGTRGCAATGCAGCATGG-3′ and POLR2 rev 5′-GCGGACGGCASCACACG-3′ in the second round. The first-round of the nested PCR was carried out in 50 μL reaction volume consisting of 5 μL of Buffer 10X, 2 μL of MgCl2 25 mM, 1 μL of dNTPs 10 mM (ThermoFisher Scientific, Massachusetts, USA), 1.5 μL of each primer (20 μM), 5 μL of BSA 10X, 0.2 μL of Taq Platinum DNA Polymerase (Invitrogen Corporation, USA) and 3 μL of cDNA or 10 μL of DNA. The PCR program used had the following thermal profile: Taq polymerase activation at 94 °C for 2 min, followed by 40 cycles with denaturation at 94 °C for 45 s, annealing at 55 °C for 1 min, and extension at 72 °C for 1 min, followed by 10 min at 72 °C. The second round of the nested PCR was carried out in 50 μL reaction volume consisting of the same volumes from the first-round PCR and 3 μL of template. The PCR program used had the following thermal profile: Taq polymerase activation at 94 °C for 2 min, followed by 40 cycles with denaturation at 94 °C for 45 s, annealing at 65 °C for 1 min, and extension at 72 °C for 1 min, followed by 10 min at 72 °C. Both first and second-round mix volumes were adjusted using DEPC treated water. Results were analyzed via electrophoresis on a 2% agarose gel, using Midori Green Advance (NIPPON Genetics Europe GmbH, Düren, Germany) as an intercalating agent. Only samples that yielded sufficient quantities of PCR products were subjected to Sanger sequencing. Bands of the expected size of 800 bp were excised from agarose gel and purified using the QIAquick Gel Extraction kit (Qiagen, Hilden, Germany) following the manufacturer’s protocol. The sequencing reaction was conducted using the BrilliantDye Terminator Cycle Sequencing v3.1 Kit (NimaGen BV, Nijmegen, The Netherlands). The sequencing process was replicated twice for each sample using the forward primer GAG-F2 and the reverse primer POL-R2, both used at a 3.2 μM concentration. The following thermal profile was used for the sequencing reaction: Taq polymerase activation at 96 °C for 45 s, followed by 25 cycles with denaturation at 96 °C for 10 s, annealing at 50 °C for 5 s and extension at 60 °C for 4 min. Precipitation of sequencing reaction products was done with the DyeEx 2.0 Spin Kit (Qiagen, Hilden, Germany) following the manufacturer’s protocol and denatured at 95 °C for 5 min after the addition of HiDi Formammide (Applied Biosystems Life Technologies, Warrington, UK). Following denaturation, the products were loaded into the ABI PRISM 3130 Genetic Analyzer automatic sequencer (Applied Biosystems, Foster City, CA, USA). Consensus sequences for each sample were obtained using DNAStar Navigator software v.15, then compared and aligned with other SRLV sequences downloaded from GenBank, including references for subgenotypes. The alignment was done with Clustal X v.2 and was further refined and trimmed using BioEdit v.7.2.5 (50). The phylogenetic tree was obtained with MEGA X v.10.2.6, using Maximum Likelihood method and 1,000 bootstrap replicates, and further refinement has been done with ITOL online tool (https://itol.embl.de/).

2.6 Determination of TMEM154 genotypes

PCR amplification and Sanger sequencing of TMEM154 exon 2 and 3 were performed using extracted DNA from 530 sheep samples as described by Heaton et al. (16). Sequencing reactions and electrophoresis were carried out by LGC Genomics GmbH (Berlin, Germany). Sequence chromatograms were analyzed using the software ChromasPro (Technelysium Pty Ltd., South Brisbane, Australia) for DNA and protein sequence variants.

2.7 Statistical analysis

The data were stored and organized in Microsoft Excel 2016 (Microsoft Corporation, Redmond, USA). Associations between potential risk factors and individual and flock-level seropositivity were assessed by using a chi-square or Fisher’s exact test and by calculating the odds ratio (OR) to facilitate comparison with results from other breeds. Due to the small number of positive samples, only a univariable analysis was performed for each independent variable. SAS Studio Version 9.4 program (SAS Institute Inc., Cary, NC, USA) was used for the statistical analyses.

3 Results

3.1 MVV seroprevalence in RPL flocks

Among a total of 849 RPL sheep plasma samples tested with the competitive ELISA, 30 showed a positive result (3.5%). Seropositive animals were detected in six flocks (17.1%), one of them was located in the northwest, and the others in the east of Germany (Table 1; Figure 1). The within-flock prevalence in positive flocks ranged from 5.3 to 37.5%. Four out of 30 seropositive sheep developed typical clinical signs, thus clinical prevalence was 13.3% (Table 2).

Table 2

Table 2. Seroprevalence and clinical prevalence in MVV-positive RPL flocks.

3.2 Risk factors of individual MVV seropositivity

In the group of sheep older than 3 years, significantly more sheep were seropositive (4.8%) than in the group of sheep younger than 3 years (1.5%) (Table 3). Seroprevalence among sheep from the eight different ram lines was not significantly different (p = 0.37). Most seropositive sheep were found in ram line 3 (7.14%). A portion of 13.3% of the tested RPL sheep could not be assigned to any specific ram line, including seven seropositive sheep (6.2%).

Table 3

Table 3. Results of the statistical analysis for individual risk factors associated with MVV seropositivity.

3.3 Risk factors of MVV seropositivity at flock-level

All the participating farmers kept sheep for breeding (100%), with 74.3% also using them for landscape conservation, and 42.9% keeping them as a hobby. The only two management systems of the participating RPL farms were paddock farming and sheep transhumance.

In large positive flocks (>100 sheep), seroprevalence was 40% and therefore higher than in medium-sized (31–100 sheep) (18.9%) and small (1–30 sheep) (10.5%) flocks (Figure 2). Contact with other sheep breeds or the presence of goats in RPL flocks also tended to increase the prevalence (Table 4). However, none of these differences were statistically significant.

In eastern Germany, five out of twelve flocks (41.7%) had a serologically positive flock status (Figure 2). Therefore, region was a significant risk factor at flock-level (p = 0.03). The addition of new RPL sheep in the last year was also significantly associated with a MVV-positive flock status (p = 0.02) (Figure 2). Especially the purchase of female sheep increased the odds of infection by 15.7 times (Table 4).

Figure 2

Figure 2. Numbers of MVV-positive or negative RPL flocks in relation to three different risk factors. Region: north (northwest), east, south (Figure 1); Flock size: small (1–30 sheep), middle (31–100 sheep), large (>100 sheep); Purchase: ♂ (only ram(s)); ♀ (ewe(s) and/or female yearling(s)); no (no purchase).

Table 4

Table 4. Results of the statistical analysis for risk factors associated with MVV seropositivity at flock-level.

3.4 SRLV serotypes and genotypes

Of the 29 plasma samples serotyped using the Eradikit ELISA, 37.9% were positive for one of the defined genotypes A (three samples) or B (eight samples), while 55.2% could not be assigned to any genotype (Table 5). Two sera (6.9%) showed a possible co-infection.

Table 5

Table 5. SRLV serotyping (n = 29) and genotyping (n = 13) results of 30 seropositive RPL sheep and their TMEM154 E35K, T44M, and N70I genotypes.

Thirteen out of 30 seropositive sheep samples were Sanger sequenced. In contrast to the Eradikit ELISA results, virus genotyping by DNA sequencing revealed only the SRLV genotype A, comprising three different subgenotypes, in these sheep: A1 (n = 1), A3 (n = 2), and A21 or A21-like (n = 9). For one sample, the subgenotype was not classified, because its phylogenetic group was not clear. However, also this sample belonged to the SRVL genotype A group (Table 5; Figure 3). Twelve samples were simultaneously serotyped and genotyped. Only one animal showed concordant results (8.33%). In both cases where the Eradikit ELISA indicated a potential co-infection, SRLV genotyping confirmed the presence of genotype A only. Out of the remaining nine discrepant samples, five were serotyped as SRLV serotype B, and four produced indeterminate serotyping results (Table 5).

Figure 3

Figure 3. Phylogenetic tree including SRLV sequences generated in this study (red) and sequences downloaded from GenBank (black), including references for subgenotypes.

3.5 TMEM154 genotypes and results of association analyses

Sequencing of TMEM154 exons 2 and 3 identified variations at codons 35, 44, and 70. All other positions were monomorphic in the investigated 530 sheep. At codon 35 genotype EK occurred at highest frequency (49%), followed by KK (28%) and EE (23%) (Figure 4A). The frequencies of the putative protective allele K and risk allele E were 53 and 47%, respectively. Allele M at codon 44 was present at a frequency of only 18%, whereas allele T was found in 82% of the animals. At codon 70, allele N was observed in 86% of the sheep, while allele I occurred in 14%. The distribution of the different TMEM154 genotypes and the resulting haplotypes among the 530 genotyped RPL sheep is shown in Figure 4.

Figure 4

Figure 4. Percentage distribution (y-axis) and absolute animal counts (within bars) of TMEM154 E35K, T44M, and N70I genotypes (A) and TMEM154 haplotypes (B) among 530 genotyped RPL sheep.

In MVV-tested flocks, 28% of serologically negative sheep (n = 101) carried the putatively protective genotype KK, whereas this genotype was present in 13% of the positive sheep (n = 4) (Table 6). This difference tended to be significant (p = 0.088).

Table 6

Table 6. TMEM154 E35K, T44M, and N70I genotype frequencies of the 395 MVV tested RPL sheep with the statistical results.

Although all 20 individuals homozygous for allele M at codon 44 (MM), and thus homozygous for the linked, putatively protective A4Δ53 frameshift mutation, were tested serologically negative for MVV, this association was not statistically significant (Table 6). However, the exclusive presence of codon 35 allele K and/or codon 44 allele M showed a statistically significant protective effect (p = 0.001). Conversely, codon 70 allele I was significantly associated with susceptibility (p = 0.045).

All other association analyses of TMEM154 genotypes, whether compared individually or grouped (EE vs. EK/KK, TT vs. TM/MM, and II vs. NI/NN), revealed no statistically significant results and were therefore not discussed further in this study.

4 Discussion

This study, the first conducted in Germany, sheds light on seroprevalence of MVV infection in the old, native sheep breed RPL across nine out of sixteen German states, while also investigating potential risk factors and genetically characterizing the circulating SRLV. Therefore, it combines epidemiological, genetic, and molecular insights to provide valuable data for future control strategies. The 3.5% individual and 17.1% flock seroprevalences were considerably lower than the flock prevalence of 51.2% reported in the representative study by Hüttner et al. (40) in sheep flocks in the federal state of Mecklenburg-Western Pomerania. Among other European countries, only Belgium had a similar herd prevalence with 17.2%, but individual seropositivity (9.4%) was higher (51), such as individual seroprevalence in Austria (9.7%), Croatia (10%), Poland (17.3%), and Spain (24.8%) (52–55). The only countries with a lower reported individual seroprevalence were Brazil with 1.07% and Costa Rica with 1.95% (56, 57). According to Straub et al. (15), coarse-wool sheep may be more susceptible to develop clinical MV symptoms than fine-wool sheep. So far, this hypothesis could not be confirmed by the present study, as only four out of 30 positive RPL sheep (13.3%) showed MV typical clinical signs, such as dry cough, cachexia, lameness, or swollen carpal joints (7). Villagra-Blanco et al. (57) detected a higher clinical incidence of 57%, although the individual seroprevalence was lower. However, the actual clinical prevalence could be higher, as it was only a snapshot of the veterinarians at the farm visit or an assessment by the breeders. Additionally, the majority of infected animals were relatively young. In general, the disease occurs in sheep older than 3 years, but in this study, a seropositive yearling showed a dry cough and nasal discharge, which might also be a result of secondary bacterial infection (7).

Nevertheless, when interpreting the obtained seroprevalence in this breed, certain limitations should be taken into account. Besides the possible impact of seroconversion, seroreversion, particularly in sheep older than 3 years, may also lead to false-negative results and consequently to an underestimation of MVV prevalence. Kalogianni et al. (58) observed seroreversion in 8.6% of previously MVV-positive sheep across five farms over 2 years, suggesting a possible suppression of neutralizing antibodies associated with poor general health or individual variations in the humoral immune response, although the exact mechanisms remain unclear. Furthermore, the imperfect sensitivity of the ELISA test used may have contributed to additional misclassification (49). Taken together, due to such factors the actual infection rate in the RPL breed may be higher than the serological data suggest, which should be considered when interpreting results and developing effective control or breeding programs.

4.1 Environmental risk factors for MVV seropositivity at individual and flock-level

In this study, age was the only significant individual risk factor for seropositivity (p = 0.01). Sheep older than 3 years had a threefold higher risk of being serologically MV-positive than younger sheep. The fact that MVV infection rises with animal age, with a peak between 3 and 5 years, has already been established in many other studies (55, 59–61). This is not surprising, as the opportunity for virus exposure, infection with the virus, and antibody production against the virus increases with advancing age.

The purchase of new positive animals in the last year was observed as main risk factor at flock-level for the introduction of MVV into negative RPL flocks (p = 0.02). This result supports similar findings in other studies (55, 62). Interestingly, five of the six positive flocks also purchased female sheep, whereas the integration of foreign rams alone was not associated with MVV-positive flock status. A possible reason for this could be the more intensive MV monitoring of RPL rams, which is required for participation in certain exhibitions. Another explanation might be a predominantly lactogenic transmission of MVV (9), as some of the purchased ewes or their progeny were seropositive. The fact that some of the seropositive sheep were the offspring of MVV-seronegative ewes speaks against an exclusively lactogenic transmission. This assumption is supported by recent studies (10, 63). The finding that the seroprevalence was highest in large (40%) compared to medium (18%) and small-sized flocks (11%) also argues in favor of horizontal transmission, as direct close contact and limited space are known risk factors (64). However, it is important to note the low number of large RPL flocks in this study. Previous studies by Lago et al. (55) and Hüttner et al. (40) also reported that the likelihood of MVV seroprevalence increased with flock size. Given that the largest RPL flocks are located in their original home in an eastern federal state of Germany, it is not surprising that the highest number of positive RPL sheep (5.7%) was identified in this region. Consequently, the three study areas showed a significant difference in the MVV seroprevalence (p = 0.03).

Direct contact with other sheep breeds was identified as a potential risk factor. Although seroprevalence in mixed-breed flocks was fivefold higher than in pure RPL flocks, this association was not significant (p = 0.15). Nevertheless, occasional introduction of MVV into the RPL breed through contact with MVV-positive sheep of other breeds cannot be entirely excluded. Unfortunately, data on the MV status of other sheep breeds were only available from one seropositive flock, in which all non-RPL sheep were tested seronegative. The presence of infected goats can also be considered a potential risk factor for the introduction of lentiviruses into the RPL breed, as several reports have shown that SRLV crossed the species barrier through direct contact (11, 55, 65). However, in our data, keeping of goats in RPL flocks was not significantly associated with their SRLV status (p = 0.64).

Our finding that rams were almost twice as often seropositive as ewes was not statistically significant (p = 0.26) and was likely due to random chance given the small number of seropositive rams. The fact that only the purchase of female sheep is a significant risk factor, along with previous studies (53, 60, 66) reporting higher prevalences in females than in males, supports this hypothesis. However, a possible reason for higher seroprevalence in males could have been the more frequent exchange of rams between flocks for breeding purposes and that they are often kept together with rams of other breeds, or goats.

4.2 SRLV genetics as risk factor for MVV seropositivity

Genetic variability of SRLV is an important tool for epidemiological studies and can provide information on the origin of the virus (67, 68). Moreover, genetic susceptibility of sheep to SLRV infection has been shown at least in some studies also to depend on the genotype of the present SRLVs (25, 69). Serological diagnostics, such as ELISA, serve as a first-line epidemiological tool to characterize SRLV serotypes (70, 71). However, a high variability of circulating strains represents a limitation of this method, as no current serological tests are capable of discriminating among all viral variants with certainty (72, 73). This limitation was confirmed in the present study, where the Eradikit ELISA yielded a high percentage (55.2%) of indeterminate results. In contrast, the more expensive and time-consuming genetic analysis by PCR and Sanger sequencing revealed no indeterminate results, and determined SRLV genotype A21 in four samples with indeterminate ELISA results. In two cases, sheep were serologically tested as coinfected with genotypes A/B and A/E, but only genotype A was detected by sequencing. This discrepancy may be due to the fact that SRLV strains show the ability to infect different organs or tissues in the same host. This phenomenon, called compartmentalization, has been described before (74–77). One probable explanation for the inability to confirm the co-infection is that only one type of tissue was used for Sanger sequencing. Additionally, a recent study showed that Sanger sequencing may have limitations compared to next-generation sequencing (NGS) when it comes to detecting co-infection when there is a low proviral load (78). In such cases, co-infections can be detected by cloning of PCR products before sequencing, but this was not possible in this study. It has also to be admitted that a limitation of SRLV genotyping by sequencing is the usually lower number of samples which can be genotyped compared to serological tests, as the needed minimal amount of provirus DNA usually cannot be obtained from all serologically positive samples (79, 80). However, co-infections with different SRLV genotypes have been reported previously (81, 82), whereas genotype E has only been detected in Italian goats so far (83). Colitti et al. (71) also identified genotype E in sheep using the Eradikit genotyping ELISA, whereas provirus sequencing determined genotype A. Therefore, in our study, we trust the sequencing result (A3) more than the ELISA result (A/E coinfection). By the way, A3 was also found in a second sheep of the same flock.

Due to the low concordance (8.33%) of the serotyping ELISA and genotyping PCR, these results should be interpreted with caution. Similar findings have been reported by Jiménez et al. (72), Colitti et al. (71), and Olech and Kuźmak (73) with concordance rates of only 25.7%, less than 40, and 42.3%, respectively. In contrast, a concordance of more than 97% between serotyping and genotyping was reported in the study by Nogarol et al. (70). Even though the number of samples compared in the present study was low (n = 12), the results suggest that serological serotyping with the Eradikit genotyping ELISA is not a reliable method for predicting the SRLV genotype in infected flocks, as described recently (73). This is of practical importance for eradication programs that focus on specific genotypes, such as in Switzerland, where only genotype B-infected goats are culled, given the considerable risk of inaccurate results (84). Up to now, this is not planned in any control program for MVV worldwide.

As genotype B was not detected by provirus sequencing and the only previous study on SRLV genotypes in German sheep populations (67) only identified A genotype, it appears unlikely that SRLV B genotypes are present in the analyzed RPL flocks. However, we recently identified for the first time SRLV genotype B in a German sheep flock by sequencing, but in another breed and geographical region (unpublished results). To date, no studies have specifically investigated RPL sheep in Germany, therefore we cannot exclude that our serological detection of genotype B may indicate a previously unrecognized presence of this genotype in the breed. Further testing using Sanger sequencing in RPL sheep from the same federal state and/or other regions would be necessary to prove this.

Regardless of whether the results of serotyping ELISA or of provirus sequencing are considered more reliable, both methods revealed viral heterogeneity in the RPL population. This suggests multiple origins and routes of introduction of the virus into the RPL breed and single flocks. In this regard, the provirus sequencing results even indicated a flock-specific SRLV distribution.

4.3 TMEM154 genotype as risk factor for MVV seropositivity

Beyond the environmental factors, the low MVV infection prevalence in RPL sheep may indicate a low genetic susceptibility of this breed. Genotype KK at codon 35 of TMEM154 gene was observed to confer lower susceptibility to MVV infection in various American, Asian, and European sheep breeds (18, 20, 85, 86). Also in the RPL breed, sheep with this genotype were less frequently seropositive compared to sheep with genotypes EK and EE, but this association was not statistically significant (p = 0.09). This could be explained by the low number of serologically positive sheep present in tested flocks (only 30 out of 395). Whether a higher number of serologically positive sheep would lead to a significant result needs to be tested in a further study, for which additional MVV-positive RPL flocks would have to be found. Increasing the sample size was not possible, as the mandatory reporting of MVV in Germany discouraged many voluntarily participating RPL breeders from testing their flocks for MVV. A missing significance of the KK genotype effect on MVV susceptibility, possibly due to low number of seropositive sheep, was also postulated in other studies involving other breeds (19, 69, 86).

The odds of infection for RPL sheep carrying one or two copies of TMEM154 allele E at codon 35 were 2.5 times higher than those for sheep homozygous for the K allele (p = 0.09, OR = 2.487, CI 95% = 0.847–7.303). However, 13.3% (n = 4) of seropositive RPL sheep still carried the putatively protective genotype KK. Previous studies have already shown that the KK genotype does not provide full protection against MVV infection (16, 63). There are also hints that the protective effect of this variant could be breed-dependent. In a survey of German MVV-affected sheep flocks, a significant (even if not absolute) protective effect of the KK genotype was observed in Texel and some milk sheep breed flocks. By contrast, this was not observed in the Merinoland sheep breed, where the frequency of the KK genotype in seropositive sheep (75%) was even higher than in the seronegative animals (59%) (19). Further support for this breed-specific effect comes from a recent study by Materniak-Kornas et al. (22), in which none of the KK-genotyped Cameroonian sheep tested positive for MVV. This finding is consistent with the hypothesis that the protective effect of the KK genotype may be more pronounced in MVV-susceptible breeds such as Texel, East Friesian, and Cameroonian sheep, while being less effective in more resistant breeds like Merinoland and potentially also RPL sheep. Interestingly, the present study showed that 12 seronegative offspring originated from seropositive ewes. Ten of these offspring were born after 2020 and may still become seropositive due to the long incubation period. The remaining two offspring, both over 3 years of age (born before 2020), had TMEM154 genotype KK, which in turn suggests a protective effect. Leymaster et al. (63) already reported this protective effect of genotype KK for the lactogenic but also for the horizontal transmission.

Haplotype 4, defined by a TMEM154 frameshift mutation (A4Δ53) that represents a naturally occurring knock-out allele, has been in the past suggested to potentially offer complete MVV-resistance (16). However, due to its low frequency (less than 5%) in most sheep populations studied so far, association analyses with MVV susceptibility are rare (22, 23). In the RPL breed, the frequency of A4Δ53, determined by genotyping the linked variation at codon 44 in exon 2, is 17.6%. In 2015, Clawson et al. (87) showed for the first time that animals homozygous for this “knockout” allele could still be infected with MVV. Freking et al. (21) later confirmed this finding, although seropositive 4,4 sheep did not show clinical symptoms and had healthy-appearing lung tissues, suggesting that viral replication may be limited in these animals. In the present study, all 20 sheep homozygous for the A4Δ53 frameshift mutation were seronegative, and only six of the 106 heterozygous animals (5.7%) were tested seropositive. Although these results support the hypothesis of reduced susceptibility, statistical significance could not be established, likely due to the low number of infected animals, as discussed before for the variation E35K.

To the best of our knowledge, no study has analyzed until now the combined effect of the two TMEM154 alleles, K at codon 35, and A4Δ53 (determined by M at codon 44), which were assumed to be (at least partially) protective. Indeed, in the RPL sheep, the frequency of seropositive sheep was significantly lower in carriers of exclusively these two alleles (genotypes 35KK, 44MM, or 35 EK in combination with 44 MT) and the odd of infection was 4.2 times higher for sheep without them (p = 0.001, OR = 4.202, CI 95% = 1.678–10.521). A total of 49% of the genotyped RPL sheep carried one of these three genotypes. This provides a promising basis for resistance-oriented breeding in this breed.

Haplotypes 2 and 3 differ only by a single amino acid substitution at position 70 of TMEM154 (N70I) (16). In the past, both variants were considered functionally equivalent dominant risk alleles, with some studies reporting very high MVV prevalence rates among animals carrying either haplotype (16, 21), suggesting limited functional relevance of TMEM154 N70I mutation. However, this assumption was later challenged by Arcangeli et al. (23), who showed that carriers of codon 70 genotypes NI or II had a significantly higher risk of infection than those with genotype NN. This is in accordance with the present study, in which a significantly higher susceptibility was observed for sheep with codon 70 genotypes NI and II.

In addition to host genetics, many other factors play a central role in MVV infection, such as viral dose, route of transmission, management, or coinfection with other diseases (7, 17, 64, 88). Another factor influencing host susceptibility for MVV infection could be a genetic adaptation of the virus to host resistance (25). Clawson et al. (87) reported that a subgroup of the SRLV A2 strain can infect sheep with the TMEM154 A4Δ53 frameshift mutation, even if this abolishes the protein function and thus prevents infection with other genotypes of the virus. However, the present study could not confirm this finding, as none of the sheep homozygous for A4Δ53 tested positive for MVV. A similar phenomenon was first described by Sider et al. (25) regarding the amino acid exchange at TMEM154 position 35 in relation to different SRLV genotypes prevalent in the USA. In our study, SRLV genotyping in the four seropositive sheep which carried genotype KK at TMEM154 codon 35 delivered only a result in two sheep, serotype B by ELISA, and genotype A21 by sequencing, respectively (Table 5). Therefore, no conclusions can be drawn from these results regarding the influence of SRLV genotypes present in Germany on the susceptibility of sheep with the KK genotype at position 35 of TMEM154.

In several breeds, frequencies of the TMEM154 genotypes at position 35 were consistent with the known breed susceptibility (85). In a German study, the highly MVV-susceptible sheep breeds Texel, East Friesian Milk, and Cameroon showed low frequencies of the protective genotype (0–10%), whereas Merinoland and German Grey Heath sheep, not considered to be susceptible to MVV, exhibited 75% and almost 100% of KK genotype, respectively (89). The RPL breed had a KK genotype frequency of 28% and is therefore somewhere between the highly MVV-susceptible and less susceptible German breeds. With the K allele present in 53% of the genotyped RPL sheep, selective breeding would be a realistic opportunity to reduce MVV prevalence, particularly in affected RPL flocks. As the putatively second protective variant, TMEM154 A4Δ53, is also present in the RPL breed, it should be considered to enlarge the selection also for sheep carrying the linked allele M at codon 44 of TMEM154. This approach has potential advantages and disadvantages. On the one hand, it has to be kept in mind that the biological role of the TMEM154 protein is not fully understood, therefore it is not known if it would be better to keep a full-length protein. On the other hand, a second protective allele with a reasonable allele frequency in the breed (17.6%) would allow to keep more sheep for breeding and thus to better maintain genetic diversity. Moreover, the presence of more than one protective variant may prevent an adaptation of the virus to the host’s defence mechanism.

5 Conclusion

This study highlighted for the first time the presence of MVV in the German sheep breed RPL. Although prevalence was low, the disease should not be underestimated, as it can spread unnoticed in the absence of serological monitoring. Therefore, routine serological testing is the most cost-effective preventive strategy, especially in purchased sheep, as the introduction of new ewes with an unknown MV status was identified as main risk factor. Heterogeneous results of the SRLV serotyping and genotyping indicate that MVV does not have a single origin but was introduced into the RPL breed via several different pathways. Moreover, the association between the variation at position 35 in the coding region of TMEM154 gene and MVV susceptibility tended to be significant and may be influenced by the MVV genotype. Nearly 50% of the genotyped RPL sheep carried protective TMEM154 genotypes (35KK, 44MM, or 35EK in combination with 44MT), providing a promising foundation for the implementation of selective breeding strategies aimed at reducing MVV susceptibility within this breed. Among the protective alleles, TMEM154 allele K appears most suitable for breeding programs, as it was more frequent than allele M, encodes a full-length TMEM154 protein, and has shown comparable protective effects in other breeds. However, further investigation is needed to determine whether selective breeding for lower susceptibility could serve as an additional tool to eradicate the disease, both within affected flocks and across the entire breed.

Data availability statement

The datasets presented in this article are not readily available because they originate from German sheep farms and therefore contain privacy issues. Requests to access the datasets should be directed to the corresponding author with the permission of the sheep farmers.

Ethics statement

The animal studies were approved by Baden-Württemberg 35-9185.82/A-11/23 (8 August 2023), Brandenburg LAVG-V6-2347-19-2023- 23-E (7 August 2023), Hesse V 54 19 c 20 15 h 01 Nr. V 3/2023 (21 August 2023), Lower Saxony 33.19-42502-04-23-00387 (4 August 2023), Mecklenburg-Western Pomerania 7221.3-2-012/23 (5 September 2023), North Rhine-Westphalia 81-02.04.40.2023. VG036 (11 August 2023), Saxony 25-5131/564/17 (9 October 2023), Saxony-Anhalt 203.m-42502-6- 011_TiHo_G (25 September 2023), Schleswig Holstein IX 552-117467/2023 (56-9/23 V) (25 September 2023). The studies were conducted in accordance with the local legislation and institutional requirements. Written informed consent was obtained from the owners for the participation of their animals in this study.

Author contributions

CF: Investigation, Formal analysis, Data curation, Writing – original draft, Validation, Resources, Visualization. MB: Validation, Writing – review & editing, Investigation, Methodology. PG: Investigation, Validation, Writing – review & editing, Methodology. SM: Methodology, Validation, Writing – review & editing, Investigation. GL: Funding acquisition, Supervision, Investigation, Writing – review & editing, Conceptualization, Validation, Project administration, Methodology. MG: Methodology, Investigation, Validation, Funding acquisition, Supervision, Conceptualization, Writing – review & editing, Project administration.

Funding

The author(s) declare that financial support was received for the research and/or publication of this article. This work was supported by the Federal Ministry of Food and Agriculture (BMEL) based on a decision of the parliament of the Federal Republic of Germany via the Federal Office for Agriculture and Food (BLE) under the Federal Programme for Ecological Farming (2821OE013/2822OE140). The Sanger sequencing of the SRLV genotypes was partially funded by RC09/2022 IZSUM grant code D63C22000860001. The funding body played no role in the design, analysis, and reporting of the study.

Acknowledgments

The authors would like to thank Christoph Höller, Chairman of the interest group of the Pomeranian Coarsewool sheep (Interessengemeinschaft Rauhwollige Pommersche Landschafe e. V.), who supported and promoted the project from the very beginning. Many thanks also go to the participating sheep breeders and zoos for providing the samples and data. Furthermore, we acknowledge Petra Röhrig, Carina Crispens, and Stephanie Steitz for their excellent technical assistance in carrying out the serological analyses and TMEM154 genotyping, as well as Frederic Kiene for his support in data analysis. Parts of the manuscript’s content have previously appeared online as a preprint on Research Square (90).

Conflict of interest

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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Abbreviations

SRLV, small ruminant lentivirus; MV, maedi-visna; MVV, maedi-visna virus; CAEV, caprine arthritis and encephalitis virus; RPL, Pomeranian Coarsewool Sheep (German, ´Rauhwolliges Pommersches Landschaf´); TMEM154, transmembrane protein 154 gene; ELISA, enzyme-linked immunosorbent assay; EDTA, ethylenediaminetetraacetic acid; OR, odds ratio; CI, confidence interval; OD, optical density; PI, percent inhibition; PCR, polymerase chain reaction; NGS, next-generation sequencing.

References

1. Clements, JE, and Zink, MC. Molecular biology and pathogenesis of animal lentivirus infections. Clin Microbiol Rev. (1996) 9:100–17. doi: 10.1128/cmr.9.1.100

PubMed Abstract | Crossref Full Text | Google Scholar

2. Gorrell, MDBM, Sheffer, D, Adams, RJ, and Narayan, O. Ovine lentivirus is macrophagetropic and does not replicate productively in T lymphocytes. J Virol. (1992) 66:2679–88. doi: 10.1128/jvi.66.5.2679-2688.1992

Crossref Full Text | Google Scholar

3. Blacklaws, BA. Small ruminant lentiviruses: immunopathogenesis of Visna-Maedi and caprine arthritis and encephalitis virus. Comp Immunol Microbiol Infect Dis. (2012) 35:259–69. doi: 10.1016/j.cimid.2011.12.003

PubMed Abstract | Crossref Full Text | Google Scholar

4. Gendelman, HE, Narayan, O, Kennedy-Stoskopf, S, Kennedy, PG, Ghotbi, Z, Clements, JE, et al. Tropism of sheep lentiviruses for monocytes: susceptibility to infection and virus gene expression increase during maturation of monocytes to macrophages. J Virol. (1986) 58:67–74. doi: 10.1128/JVI.58.1.67-74.1986

PubMed Abstract | Crossref Full Text | Google Scholar

5. Narayan, O, and Cork, LC. Lentiviral diseases of sheep and goats: chronic pneumonia Leukoencephalomyelitis and arthritis. Rev Infect Dis. (1985) 7:89–98. doi: 10.1093/clinids/7.1.89

PubMed Abstract | Crossref Full Text | Google Scholar

6. Narayan, O, and Clements, JE. Biology and pathogenesis of lentiviruses. J Gen Virol. (1989) 70:1617–39. doi: 10.1099/0022-1317-70-7-1617

PubMed Abstract | Crossref Full Text | Google Scholar

7. Christodoulopoulos, G. Maedi-visna: clinical review and short reference on the disease status in Mediterranean countries. Small Rumin Res. (2006) 62:47–53. doi: 10.1016/j.smallrumres.2005.07.046

Crossref Full Text | Google Scholar

8. Reina, R, de Andrés, D, and Amorena, B. Immunization against small ruminant lentiviruses. Viruses. (2013) 5:1948–63.

Google Scholar

9. Preziuso, S, Renzoni, G, Allen, TE, Taccini, E, Rossi, G, DeMartini, JC, et al. Colostral transmission of maedi visna virus: sites of viral entry in lambs born from experimentally infected ewes. Vet Microbiol. (2004) 104:157–64. doi: 10.1016/j.vetmic.2004.09.010

PubMed Abstract | Crossref Full Text | Google Scholar

10. Blacklaws, BA, Berriatua, E, Torsteinsdottir, S, Watt, NJ, de Andres, D, Klein, D, et al. Transmission of small ruminant lentiviruses. Vet Microbiol. (2004) 101:199–208. doi: 10.1016/j.vetmic.2004.04.006

PubMed Abstract | Crossref Full Text | Google Scholar

11. Shah, C, Huder, JB, Boni, J, Schonmann, M, Muhlherr, J, Lutz, H, et al. Direct evidence for natural transmission of small-ruminant lentiviruses of subtype A4 from goats to sheep and vice versa. J Virol. (2004) 78:7518–22. doi: 10.1128/JVI.78.14.7518-7522.2004

PubMed Abstract | Crossref Full Text | Google Scholar

12. Minguijon, E, Reina, R, Perez, M, Polledo, L, Villoria, M, Ramirez, H, et al. Small ruminant lentivirus infections and diseases. Vet Microbiol. (2015) 181:75–89. doi: 10.1016/j.vetmic.2015.08.007

PubMed Abstract | Crossref Full Text | Google Scholar

13. Houwers, D, König, C, De Boer, G, and Schaake, JJ. Maedi-visna control in sheep I. Artificial rearing of colostrum-deprived lambs. Vet Microbiol. (1983) 8:179–85. doi: 10.1016/0378-1135(83)90064-0

PubMed Abstract | Crossref Full Text | Google Scholar

14. White, SN, and Knowles, DP. Expanding possibilities for intervention against small ruminant lentiviruses through genetic marker-assisted selective breeding. Viruses. (2013) 5:1466–99. doi: 10.3390/v5061466

PubMed Abstract | Crossref Full Text | Google Scholar

15. Straub, OC. Maedi-Visna virus infection in sheep. History and present knowledge. Comp Immunol Microbiol Infect Dis. (2004) 27:1–5. doi: 10.1016/S0147-9571(02)00078-4

PubMed Abstract | Crossref Full Text | Google Scholar

16. Heaton, MP, Clawson, ML, Chitko-Mckown, CG, Leymaster, KA, Smith, TP, Harhay, GP, et al. Reduced lentivirus susceptibility in sheep with Tmem154 mutations. PLoS Genet. (2012) 8:e1002467. doi: 10.1371/journal.pgen.1002467

PubMed Abstract | Crossref Full Text | Google Scholar

17. Alshanbari, FA, Mousel, MR, Reynolds, JO, Herrmann-Hoesing, LM, Highland, MA, Lewis, GS, et al. Mutations in Ovis Aries Tmem154 are associated with lower small ruminant lentivirus proviral concentration in one sheep flock. Anim Genet. (2014) 45:565–71. doi: 10.1111/age.12181

PubMed Abstract | Crossref Full Text | Google Scholar

18. Yaman, Y, Keles, M, Aymaz, R, Sevim, S, Sezenler, T, Önaldi, AT, et al. Association of Tmem154 variants with Visna/Maedi virus infection in Turkish sheep. Small Rumin Res. (2019) 177:61–7. doi: 10.1016/j.smallrumres.2019.06.006

Crossref Full Text | Google Scholar

19. Molaee, V, Eltanany, M, and Lühken, G. First survey on association of Tmem154 and Ccr5 variants with serological maedi-visna status of sheep in German flocks. Vet Res. (2018) 49:36. doi: 10.1186/s13567-018-0533-y

PubMed Abstract | Crossref Full Text | Google Scholar

20. Molaee, V, Otarod, V, Abdollahi, D, and Lühken, G. Lentivirus susceptibility in Iranian and German sheep assessed by determination of E35k. Animals-Basel. (2019) 9:685. doi: 10.3390/ani9090685

Crossref Full Text | Google Scholar

21. Freking, BA, Murphy, TW, Chitko-McKown, CG, Workman, AM, and Heaton, MP. Impact of four ovine haplotypes on ewes during multiyear lentivirus exposure. Int J Mol Sci. (2022) 23:14966. doi: 10.3390/ijms232314966

Crossref Full Text | Google Scholar

22. Materniak-Kornas, M, Piórkowska, K, Ropka-Molik, K, Musiał, AD, Kowalik, J, Kycko, A, et al. First report of Snps detection in Tmem154 gene in sheep from Poland and their association with Srlv infection status. Pathogens. (2024) 14:16. doi: 10.3390/pathogens14010016

PubMed Abstract | Crossref Full Text | Google Scholar

23. Arcangeli, C, Lucarelli, D, Torricelli, M, Sebastiani, C, Ciullo, M, Pellegrini, C, et al. First survey of SNPs in Tmem154, Tlr9, Myd88 and Ccr5 genes in sheep reared in Italy and their association with resistance to SRLVs infection. Viruses. (2021) 13:1290. doi: 10.3390/v13071290

PubMed Abstract | Crossref Full Text | Google Scholar

24. Glaria, I, Reina, R, Ramirez, H, de Andres, X, Crespo, H, Jauregui, P, et al. Visna/Maedi virus genetic characterization and serological diagnosis of infection in sheep from a neurological outbreak. Vet Microbiol. (2012) 155:137–46. doi: 10.1016/j.vetmic.2011.08.027

PubMed Abstract | Crossref Full Text | Google Scholar

25. Sider, LH, Heaton, MP, Chitko-McKown, CG, Harhay, GP, Smith, TP, Leymaster, KA, et al. Small ruminant lentivirus genetic subgroups associate with sheep Tmem154 genotypes. Vet Res. (2013) 44:64. doi: 10.1186/1297-9716-44-64

PubMed Abstract | Crossref Full Text | Google Scholar

26. Olech, M, and Kuzmak, J. Genetic diversity of the Ltr region of polish Srlvs and its impact on the transcriptional activity of viral promoters. Viruses. (2023) 15:15. doi: 10.3390/v15020302

PubMed Abstract | Crossref Full Text | Google Scholar

27. Pisoni, G, Quasso, A, and Moroni, P. Phylogenetic analysis of small-ruminant lentivirus subtype B1 in mixed flocks: evidence for natural transmission from goats to sheep. Virology. (2005) 339:147–52. doi: 10.1016/j.virol.2005.06.013

PubMed Abstract | Crossref Full Text | Google Scholar

28. Shah, C, Boni, J, Huder, JB, Vogt, HR, Muhlherr, J, Zanoni, R, et al. Phylogenetic analysis and reclassification of caprine and ovine lentiviruses based on 104 new isolates: evidence for regular sheep-to-goat transmission and worldwide propagation through livestock trade. Virology. (2004) 319:12–26. doi: 10.1016/j.virol.2003.09.047

PubMed Abstract | Crossref Full Text | Google Scholar

29. Gjerset, B, Jonassen, CM, and Rimstad, E. Natural transmission and comparative analysis of small ruminant lentiviruses in the Norwegian sheep and goat populations. Virus Res. (2007) 125:153–61. doi: 10.1016/j.virusres.2006.12.014

PubMed Abstract | Crossref Full Text | Google Scholar

30. Giammarioli, M, Bazzucchi, M, Puggioni, G, Brajon, G, Dei Giudici, S, Taccori, F, et al. Phylogenetic analysis of small ruminant lentivirus (Srlv) in Italian flocks reveals the existence of novel genetic subtypes. Virus Genes. (2011) 43:380–4. doi: 10.1007/s11262-011-0653-1

PubMed Abstract | Crossref Full Text | Google Scholar

31. Grego, E, Bertolotti, L, Quasso, A, Profiti, M, Lacerenza, D, Muz, D, et al. Genetic characterization of small ruminant lentivirus in Italian mixed flocks: evidence for a novel genotype circulating in a local goat population. J Gen Virol. (2007) 88:3423–7. doi: 10.1099/vir.0.83292-0

PubMed Abstract | Crossref Full Text | Google Scholar

32. Bazzucchi, M, Pierini, I, Gobbi, P, Pirani, S, Torresi, C, Iscaro, C, et al. Genomic epidemiology and heterogeneity of Srlv in Italy from 1998 to 2019. Viruses. 13:13. doi: 10.3390/v13122338

PubMed Abstract | Crossref Full Text | Google Scholar

33. Thormar, H. The origin of lentivirus research: Maedi-Visna virus. Curr HIV Res. (2021) 11:2–9. doi: 10.2174/157016213804999212

PubMed Abstract | Crossref Full Text | Google Scholar

34. Sigurdsson, B, Grimsson, H, and Palsson, PA. Maedi, a chronic, progressive infection of sheep's lungs. J Infect Dis. (1952) 90:233–41. doi: 10.1093/infdis/90.3.233

PubMed Abstract | Crossref Full Text | Google Scholar

35. Sigurdsson, B, Palsson, P, and Grimsson, H. Visna, a demyelinating transmissible disease of sheep. J Neuropathol Exp Neurol. (1957) 16:389–403. doi: 10.1097/00005072-195707000-00010

PubMed Abstract | Crossref Full Text | Google Scholar

36. Ganter, M, Henze, P, and Wohlsein, P. Clinic and CNS pathology of natural visna cases. Tierärztliche Praxis Ausgabe G: Großtiere / Nutztiere. (2007) 35:219–24. doi: 10.1055/s-0037-1621546

Crossref Full Text | Google Scholar

37. de Miguel, R, Arrieta, M, Rodríguez-Largo, A, Echeverría, I, Resendiz, R, Pérez, E, et al. Worldwide prevalence of small ruminant lentiviruses in sheep: a systematic review and meta-analysis. Animals. (2021) 11. doi: 10.3390/ani11030784

PubMed Abstract | Crossref Full Text | Google Scholar

38. Peterhans, E, Greenland, T, Badiola, J, Harkiss, G, Bertoni, G, Amorena, B, et al. Routes of transmission and consequences of small ruminant lentiviruses (Srlvs) infection and eradication schemes. Vet Res. (2004) 35:257–74. doi: 10.1051/vetres:2004014

PubMed Abstract | Crossref Full Text | Google Scholar

39. Graber, G, and Ganter, M. Maedi-Visna Und Caprine Arthritis-Encephalitis in Deutschland. Vorkommen, Diagnostik Und Bekämpfungsstrategien. Teil 1: Vorkommen Und Bekämpfung. Tierarztl Umsch. (2005) 60:300–10.

Google Scholar

40. Hüttner, K, Seelmann, M, and Feldhusen, F. Prevalence and risk factors for Maedi-Visna in sheep farms in Mecklenburg-Western-Pomerania. Berl Munch Tierarztl Wochenschr. (2010) 123:463–7. doi: 10.2376/0005-9366-123-10

PubMed Abstract | Crossref Full Text | Google Scholar

41. Fitzinger, LJ. Das Pommersche Schaf. In: Über Die Racen Des Zahmen Schafes, Band 2 Gerold (1860).

Google Scholar

42. Grumbach, S, and Zupp, W. Rauhwolliges Pommersches Landschaf Wieder Im Aufwind. Vienna, Dtsch Schafz. (1992) 22:512–4.

Google Scholar

43. Grumbach, S. Rauhwollige Pommersche Landschafe 1982 Bis 2002 20 Jahre Erfolgreiche Erhaltungszucht. Landesschaf- und Ziegenzuchtverband Mecklenburg-Vorpommern e.V. (2002). 2–17.

Google Scholar

44. IGRPL. (2024) 20 Jahre Erfolgreiche Erhaltungszucht 1982–2002. Rinteln: Christoph Höller. Available online at: https://www.ig-pommernschafe.de/historisches/historie-1982-2002.html.

Google Scholar

45. BLE, TGRDEU. Sheep: Rauhwolliges Pommersches Landschaf (2024). Available online at: https://tgrdeu.genres.de/en/livestock-animals/search-for-animal-species/genetic-resource-details?tx_sttgrdeu_nutztier%5Baction%5D=genetikDetail&tx_sttgrdeu_nutztier%5Bcontroller%5D=Nutztier&tx_sttgrdeu_nutztier%5Bg_id%5D=530&cHash=11308349533043f66aea37d4dc466c18.

Google Scholar

46. Drogemuller, C, de Vries, F, Hamann, H, Leeb, T, and Distl, O. Breeding German sheep for resistance to scrapie. Vet Rec. (2004) 154:257–60. doi: 10.1136/vr.154.9.257

PubMed Abstract | Crossref Full Text | Google Scholar

47. Grumbach, S, and Zupp, W. Online-Broschüre Schwerpunkt Schafe, Ziegen Gebrauchshunde (2008). Available online at: https://wwwg-e-hde/images/stories/rassebeschreib/schaf/Das%20Rauhwollige%20Pommerschepdf.

Google Scholar

48. Ganter, M, Herrmann, J, and Waibl, H. Die Blutentnahme Aus Der Vena Cava Cranialis Bei Schaf Und Ziege Mit Aufgesetzter Kanüle. Tierarztl Prax. (2001) 29:37–40.

Google Scholar

49. Herrmann, LM, Cheevers, WP, Marshall, KL, McGuire, TC, Hutton, MM, Lewis, GS, et al. Detection of serum antibodies to ovine progressive pneumonia virus in sheep by using a caprine arthritis-encephalitis virus competitive-inhibition enzyme-linked immunosorbent assay. Clin Diagn Lab Immunol. (2003) 10:862–5. doi: 10.1128/cdli.10.5.862-865.2003

PubMed Abstract | Crossref Full Text | Google Scholar

50. Hall, TA. Bioedit: A user-friendly biological sequence alignment editor and analysis program for windows 95/98/Nt. Nucleic acids symposium series Oxford (1999).

Google Scholar

51. Michiels, R, Van Mael, E, Quinet, C, Welby, S, Cay, AB, and De Regge, N. Seroprevalence and risk factors related to small ruminant lentivirus infections in Belgian sheep and goats. Prev Vet Med. (2018) 151:13–20. doi: 10.1016/j.prevetmed.2017.12.014

PubMed Abstract | Crossref Full Text | Google Scholar

52. Hönger, D, Leitold, B, and Schuller, W. Serological studies of antibodies against Maedi-visna virus in sheep in Austria. Berl Munch Tierarztl Wochenschr. (1990) 103:39–41.

PubMed Abstract | Google Scholar

53. Pavlak, M, Vlahovic, K, Cvitkovic, D, Mihelc, D, Kilvain, I, Udiljak, Z, et al. Seroprevalence and risk factors associated with Maedi Visna virus in sheep population in southwestern Croatia. Vet Arhiv. (2022) 92:277–89. doi: 10.24099/vet.arhiv.1333

Crossref Full Text | Google Scholar

54. Kuzmak, J, Junkuszew, A, Kozaczynska, B, and Gruszecki, TM. The relations between breed and age associated susceptibility/resistance of sheep infection with Meadi-Visna virus (Mvv). Arch Tierz Dummerstorf. (2006) 49:160–5.

Google Scholar

55. Lago, N, López, C, Panadero, R, Cienfuegos, S, Pato, J, Prieto, A, et al. Seroprevalence and risk factors associated with Visna/Maedi virus in semi-intensive lamb-producing flocks in northwestern Spain. Prev Vet Med. (2012) 103:163–9. doi: 10.1016/j.prevetmed.2011.09.019

PubMed Abstract | Crossref Full Text | Google Scholar

56. Da Costa LSPdL, PP, Callado, AKC, do Nascimento, SA, and de Castro, RS. Lentivírus De Pequenos Ruminantes Em Ovinos Santa Inês: Isolamento, Identificação Pela Pcr E Inquérito Sorológico No Estado De Pernambuco. Arq Inst Biol. (2022) 74:11–6. doi: 10.1590/1808-1657v74p0112007

Crossref Full Text | Google Scholar

57. Villagra-Blanco, R, Dolz, G, Solórzano-Morales, A, Alfaro, A, Montero-Caballero, D, and Romero-Zúñiga, JJ. Presence of maedi-visna in Costa Rican sheep flocks. Small Rumin Res. (2015) 124:132–6. doi: 10.1016/j.smallrumres.2015.01.010

Crossref Full Text | Google Scholar

58. Kalogianni, AI, Bouzalas, I, Bossis, I, and Gelasakis, AI. Seroepidemiology of Maedi-Visna in intensively reared dairy sheep: A two-year prospective study. Animals-Basel. (2023) 13:2273. doi: 10.3390/ani13142273

PubMed Abstract | Crossref Full Text | Google Scholar

59. Keen, I, Hungerford, L, Wittum, T, Kwang, J, and Littledike, E. Rick factors for seroprevalence of ovine lentivirus in breeding ewe flocks in Nebraska, USA. Prev Vet Med. (1997) 30:81–94. doi: 10.1016/S0167-5877(96)01121-X

PubMed Abstract | Crossref Full Text | Google Scholar

60. Preziuso, S, Or, ME, Giammarioli, M, Kayar, A, Feliziani, F, Gönül, R, et al. Maedi-Visna virus in Turkish sheep: A preliminary serological survey using Elisa tests. Turk J Vet Anim Sci. (2010) 34:289–93. doi: 10.3906/vet-0905-35

PubMed Abstract | Crossref Full Text | Google Scholar

61. Norouzi, B, Taghavi Razavizadeh, A, Azizzadeh, M, Mayameei, A, and Najar Nezhad Mashhadi, V. Serological study of small ruminant lentiviruses in sheep population of Khorasan-E-Razavi province in Iran. Vet Res Forum. (2015) 6:245–9.

Google Scholar

62. Heinrichs, R, Wilkins, W, Schroeder, G, and Campbell, J. Prevalence of Maedi-visna in Saskatchewan sheep. Can Vet J. (2017) 58:183–6.

Google Scholar

63. Leymaster, KA, Chitko-McKown, CG, and Heaton, MP. Incidence of infection in 39-month-old ewes with Diplotypes "1 1," "1 3," and "3 3" after natural exposure to ovine progressive pneumonia virus. J Anim Sci. (2015) 93:41–5. doi: 10.2527/jas.2014-8553

Crossref Full Text | Google Scholar

64. Leginagoikoa, I, Minguijon, E, Juste, RA, Barandika, J, Amorena, B, de Andres, D, et al. Effects of housing on the incidence of Visna/Maedi virus infection in sheep flocks. Res Vet Sci. (2010) 88:415–21. doi: 10.1016/j.rvsc.2009.11.006

PubMed Abstract | Crossref Full Text | Google Scholar

65. Alba, A, Allepuz, A, Serrano, E, and Casal, J. Seroprevalence and spatial distribution of Maedi-Visna virus and pestiviruses in Catalonia (Spain). Small Rumin Res. (2008) 78:80–6. doi: 10.1016/j.smallrumres.2008.05.004

Crossref Full Text | Google Scholar

66. Simard, C, and Morley, RS. Seroprevalence of maedi-visna in Canadian sheep. Can J Vet Res. (1991) 55:269–73.

Google Scholar

67. Molaee, V, Bazzucchi, M, De Mia, GM, Otarod, V, Abdollahi, D, Rosati, S, et al. Phylogenetic analysis of small ruminant lentiviruses in Germany and Iran suggests their expansion with domestic sheep. Sci Rep-Uk. (2020) 10:2243. doi: 10.1038/s41598-020-58990-9

PubMed Abstract | Crossref Full Text | Google Scholar

68. Carrozza, ML, Niewiadomska, AM, Mazzei, M, Abi-Said, MR, Hue, S, Hughes, J, et al. Emergence and pandemic spread of small ruminant lentiviruses. Virus Evol. (2023) 9:vead005. doi: 10.1093/ve/vead005

PubMed Abstract | Crossref Full Text | Google Scholar

69. Moretti, R, Sartore, S, Colitti, B, Profiti, M, Chessa, S, Rosati, S, et al. Susceptibility of diferent Tmem154 genotypes in three Italian sheep breeds infected by diferent Srlv genotypes. Vet Res. (2022) 53:60. doi: 10.1186/s13567-022-01079-0

Crossref Full Text | Google Scholar

70. Nogarol, C, Bertolotti, L, Klevar, S, Profiti, M, Gjerset, B, and Rosati, S. Serological characterization of small ruminant lentiviruses: a complete tool for serotyping lentivirus infection in goat. Small Rumin Res. (2019) 176:42–6. doi: 10.1016/j.smallrumres.2019.05.010

Crossref Full Text | Google Scholar

71. Colitti, B, Daif, S, Choukri, I, Scalas, D, Jerre, A, El Berbri, I, et al. Serological and molecular characterization of small ruminant lentiviruses in Morocco. Animals. (2024) 14:14. doi: 10.3390/ani14040550

PubMed Abstract | Crossref Full Text | Google Scholar

72. Acevedo Jimenez, GE, Tortora Perez, JL, Rodriguez Murillo, C, Arellano Reynoso, B, and Ramirez Alvarez, H. Serotyping versus genotyping in infected sheep and goats with small ruminant lentiviruses. Vet Microbiol. (2021) 252:108931. doi: 10.1016/j.vetmic.2020.108931

PubMed Abstract | Crossref Full Text | Google Scholar

73. Olech, M, and Kuzmak, J. Comparison of serological and molecular methods for differentiation between genotype a and genotype B strains of small ruminant lentiviruses. J Vet Res. (2024) 68:181–8. doi: 10.2478/jvetres-2024-0025

PubMed Abstract | Crossref Full Text | Google Scholar

74. Pisoni, G, Moroni, P, Turin, L, and Bertoni, G. Compartmentalization of small ruminant lentivirus between blood and colostrum in infected goats. Virology. (2007) 369:119–30. doi: 10.1016/j.virol.2007.06.021

PubMed Abstract | Crossref Full Text | Google Scholar

75. Deubelbeiss, M, Blatti-Cardinaux, L, Zahno, ML, Zanoni, R, Vogt, HR, Posthaus, H, et al. Characterization of small ruminant lentivirus A4 subtype isolates and assessment of their pathogenic potential in naturally infected goats. Virol J. (2014) 11:65. doi: 10.1186/1743-422X-11-65

PubMed Abstract | Crossref Full Text | Google Scholar

76. Blatti-Cardinaux, L, Pisoni, G, Stoffel, MH, Zanoni, R, Zahno, ML, and Bertoni, G. Generation of a molecular clone of an attenuated lentivirus, a first step in understanding cytopathogenicity and virulence. Virology. (2016) 487:50–8. doi: 10.1016/j.virol.2015.09.027

PubMed Abstract | Crossref Full Text | Google Scholar

77. Olech, M, and Kuzmak, J. Compartmentalization of subtype A17 of small ruminant lentiviruses between blood and colostrum in infected goats is not exclusively associated to the env gene. Viruses. (2019) 11. doi: 10.3390/v11030270

PubMed Abstract | Crossref Full Text | Google Scholar

78. Bouzalas, I, Apostolidi, ED, Scalas, D, Davidopoulou, E, Chassalevris, T, Rosati, S, et al. A combined approach for the characterization of small ruminant lentivirus strains circulating in the islands and mainland of Greece. Animals. (2024) 14:270. doi: 10.3390/ani14071119

PubMed Abstract | Crossref Full Text | Google Scholar

79. Barquero, N, Gomez-Lucia, E, Arjona, A, Toural, C, Heras, A, Fernandez-Garayzabal, JF, et al. Evolution of specific antibodies and proviral DNA in Milk of small ruminants infected by small ruminant lentivirus. Viruses. (2013) 5:2614–23. doi: 10.3390/v5102614

PubMed Abstract | Crossref Full Text | Google Scholar

80. Olech, M. The genetic variability of small-ruminant lentiviruses and its impact on tropism, the development of diagnostic tests and vaccines and the effectiveness of control programmes. J Vet Res. (2023) 67:479–502. doi: 10.2478/jvetres-2023-0064

PubMed Abstract | Crossref Full Text | Google Scholar

81. Santry, LA, de Jong, J, Gold, AC, Walsh, SR, Menzies, PI, and Wootton, SK. Genetic characterization of small ruminant lentiviruses circulating in naturally infected sheep and goats in Ontario, Canada. Virus Res. (2013) 175:30–44. doi: 10.1016/j.virusres.2013.03.019

PubMed Abstract | Crossref Full Text | Google Scholar

82. Olech, M, and Kuzmak, J. Molecular characterization of small ruminant lentiviruses in polish mixed flocks supports evidence of cross species transmission, dual infection, a recombination event, and reveals the existence of new subtypes within group A. Viruses. (2021) 13. doi: 10.3390/v13122529

PubMed Abstract | Crossref Full Text | Google Scholar

83. Grego, E, Lacerenza, D, Arias, RR, Profiti, M, and Rosati, S. Serological characterization of the new genotype E of small ruminant lentivirus in Roccaverano goat flocks. Vet Res Commun. (2009) 33:137–40. doi: 10.1007/s11259-009-9266-8

PubMed Abstract | Crossref Full Text | Google Scholar

84. Nardelli, S, Bettini, A, Capello, K, Bertoni, G, and Tavella, A. Eradication of caprine arthritis encephalitis virus in the goat population of South Tyrol, Italy: analysis of the tailing phenomenon during the 2016-2017 campaign. J Vet Diagn Invest. (2020) 32:589–93. doi: 10.1177/1040638720934055

PubMed Abstract | Crossref Full Text | Google Scholar

85. Heaton, MP, Kalbfleisch, TS, Petrik, DT, Simpson, B, Kijas, JW, Clawson, ML, et al. Genetic testing for Tmem154 mutations associated with lentivirus susceptibility in sheep. PLoS One. (2013) 8:e55490. doi: 10.1371/journal.pone.0055490

PubMed Abstract | Crossref Full Text | Google Scholar

86. Ramírez, H, Echeverría, I, Benito, AA, Glaria, I, Benavides, J, Pérez, V, et al. Accurate diagnosis of small ruminant lentivirus infection is needed for selection of resistant sheep through Tmem154 E35k genotyping. Pathogens. (2021) 10:83. doi: 10.3390/pathogens10010083

PubMed Abstract | Crossref Full Text | Google Scholar

87. Clawson, ML, Redden, R, Schuller, G, Heaton, MP, Workman, A, Chitko-McKown, CG, et al. Genetic subgroup of small ruminant lentiviruses that infects sheep homozygous for frameshift deletion mutation A4δ53. Vet Res. (2015) 46:22. doi: 10.1186/s13567-015-0162-7

PubMed Abstract | Crossref Full Text | Google Scholar

88. Leymaster, KA, Chitko-McKown, CG, Clawson, ML, Harhay, GP, and Heaton, MP. Effects of Tmem154 haplotypes 1 and 3 on susceptibility to ovine progressive pneumonia virus following natural exposure in sheep. J Anim Sci. (2013) 91:5114–21. doi: 10.2527/jas.2013-6663

PubMed Abstract | Crossref Full Text | Google Scholar

89. Lühken, G, and Simon, R. Maedi-Visna-Empfänglichkeit Beim Schaf. Schafzucht. (2022):20–1.

Google Scholar

90. Frölich, C, Lühken, G, and Ganter, M. Maedi-visna in Pomeranian Coarsewool sheep in Germany: seroprevalence, environmental and genetic risk factors. Research Square (PREPRINT). (2024) 17. doi: 10.21203/rs.3.rs-4678425/v1

Crossref Full Text | Google Scholar

91. Reina, R, Berriatua, E, Lujan, L, Juste, R, Sanchez, A, de Andres, D, et al. Prevention Strategies against Small Ruminant Lentiviruses: An Update. Vet J. (2009) 182:31–7. doi: 10.1016/j.tvjl.2008.05.008

Crossref Full Text | Google Scholar