Skip to main content

Experimental infection of Korean native goats (Capra aegagrus hircus) with bovine viral diarrhea virus 1b



Bovine viral diarrhea virus (BVDV) infects various ungulates and causes reproductive failure in infected goats. BVDV has been detected among goats in the Republic of Korea, but the route of transmission remains unclear. Here, we aimed to investigate whether BVDV-1b circulating among Korean cattle can be transmitted to Korean native goats (Capra aegagrus hircus) and characterize the outcomes of BVDV infection in these goats.


Four goats were inoculated intranasally with the Korean noncytopathic (ncp) BVDV-1b strain. Two goats exhibited clinical signs of illness, including coughing and nasal discharge. Nasal swabs and blood were collected to screen for viral RNA and BVDV antibodies. Using the 5′-untranslated region (UTR), viral RNA was detected in the nasal swabs of two goats (Goat 1 and 3) on 12 day post-inoculation (dpi) and in the blood sample of one goat (Goat 1) on 7 and 19 dpi. Using the N-terminal protease (Npro) region, viral RNA was detected in the blood sample of Goat 1 on 7 and 12 dpi. Antibodies to BVDV were detected in Goats 1 and 3 on 16–21 dpi using enzyme-linked immunosorbent assay. Sequence analysis of the virus from nasal swabs and blood samples, which was detected via RT-PCR, using the 5′-UTR and Npro regions led to the identification of the strain as ncp BVDV-1b and revealed changes in the nucleotide sequence of these goats.


Our results indicate that changes in the nucleotide sequence are associated with the establishment of BVDV infection in Korean native goats; these changes may be owing to a process required for the establishment of infection in a new host reservoir. Broadly, these findings highlight the importance of BVDV surveillance in ungulates other than cattle.


The genus Pestivirus within the family Flaviviridae comprises four species: border disease virus (BDV), bovine viral diarrhea virus (BVDV)-1, BVDV-2, and classical swine fever virus (CSFV). Other putative pestivirus species include Giraffe virus [1]; Pronghorn virus, isolated from a blind pronghorn antelope in the United States [2]; Bungowannah virus, detected in swine affected by reproductive failures including stillbirth and neonatal death in Australia [3]; and HoBi-like virus, also known as BVDV-3 or atypical pestivirus, and recognized as a bovine pathogen having a clinical presentation similar to that of BVDV-1 or BVDV-2 [4,5,6]. Unlike CSFV, BDV and BVDV are not host-specific and can infect a wide range of artiodactyls [7]. Transplacental infection of the fetus with pestivirus before the onset of immunological maturity leads to the birth of persistently infected animals, which are the main source of viral transmission [8,9,10].

BVDV can infect small ruminants, and clinical signs in these animals are similar to those in cattle [11]. Field cases of BVDV in goats are typically characterized by reproductive failure, including abortion and poor viability of neonates [12,13,14]. In a recent study conducted by our group, natural infection with BVDV was detected in Saanen goats [15]. None of the goats infected with BVDV exhibited clinical signs. However, the BVDV transmission route in these goats has not yet been identified. Because most goats raised in the Republic of Korea (ROK) share their habitats with cattle, it is possible that BVDV transmission frequency is higher among these animals.

The population of Korean native goats (Capra aegagrus hircus) has been increasing gradually in the ROK, and their main products are meat and milk. Few studies have reported BVDV infection in goats in the ROK [15, 16], and little is known about BVDV circulation and transmission in goat farms. Recently, we found that BVDV-1b is the predominant subtype circulating among Korean indigenous cattle [17]. Therefore, here, we aimed to investigate whether BVDV-1b infection can be established in Korean native goats via intranasal (IN) inoculation, evaluate the circulation of BVDV between cattle and Korean native goats, and characterize the outcome of BVDV infection in Korean native goats.


Of the four Korean native goats infected with BVDV via IN inoculation, two goats were symptomatic, exhibiting coughing and nasal discharge; one goat displayed respiratory signs with coughing on 2–5 days post-infection (dpi) and nasal discharge on 12–19 dpi. The respiratory signs were marked and persistent in this goat. On the other hand, the other goat exhibited only nasal discharge (Table 1); however, other clinical signs, including loss of appetite, diarrhea, and pyrexia, were not observed. Additionally, the remaining two goats showed no clinical symptoms until the end of this experiment. No clinical signs were observed in mock-infected animals. In this study, no other diagnostic assay for respiratory signs was performed, and the nasal discharge of the two goats was tested only for BVDV.

Table 1 Clinical signs of the Korean native goats after intranasal inoculation

Prior to inoculation, the goats were negative for BVDV antigen and antibody. Viral RNA was not detected in mock-infected animals. As shown in Table 2, using the 5′-untranslated region (UTR) gene, viral RNA was detected in the nasal swabs of two BVDV-inoculated goats on 12 dpi (Goats 1 and 3) and in the blood sample of one goat (Goat 1) on 7 and 19 dpi. Viral RNA was not detected in nasal swabs but was detected in the blood of Goat 1 on 7 and 12 dpi when primers against the N-terminal protease (Npro) region were used (Table 2). The mock-infected goats remained seronegative throughout the experiment. Two BVDV-inoculated goats (Goat 1 and 3) produced viral antibodies, whereas the other two goats remained seronegative during the experiment (Table 2).

Table 2 Viral RNA detection by RT-PCR of nasal swabs and blood, and the presence of anti-BVDV antibodies by ELISA in four Korean native goats following intranasal infection

Nucleotide sequence analyses were conducted using both the 5′-UTR and Npro regions to confirm BVDV infection in goats and compare the genomic sequences of isolates obtained from BVDV-inoculated goats and 11Q472 (parent strain) with those obtained from BVDV cattle isolates (Additional file 1: Table S1 and Additional file 2: Table S2). Phylogenetic analysis conducted based on 5′-UTR revealed that sequences obtained from the nasal swabs of two goats (Goats 1 and 3) and from the blood of Goat 1 on 7 and 19 dpi were those corresponding to ncp BVDV-1b. These three sequences (identified in the nasal swabs of Goats 1 and 3 and in the blood of Goat 1 on 7 dpi) belonged to the same clade, whereas the sequence obtained from blood on 19 dpi belonged to a separate clade (Fig. 1a). The phylogenetic tree of the Npro region showed that the sequences amplified from the blood of Goat 1 on 7 and 12 dpi were those of ncp BVDV-1b, and these sequences were divergent from those of cattle isolates (Fig. 1b). As shown in Fig. 2 and Fig. 3, there were significant differences in the nucleotide sequences before (11Q472) and after inoculation. These differences in the nucleotide sequences were more frequent in the Npro region than in the 5′-UTR (Fig. 3).

Fig. 1

Phylogenetic tree based on 5′-UTR (288 bp) (a) and Npro region (423 bp) (b) nucleotide sequences. The isolates sequenced in this study and reference BVDV strains/isolates were used to construct the phylogenetic tree by employing the neighbor-joining method using MEGA6 software. Bootstrap values are shown at branch nodes

Fig. 2

Nucleotide alignment of the 5′-UTR of BVDV. Nucleotide sequences from the nasal swabs (NS) of Goats 1 and 3 on 12 dpi and blood of Goat 1 on 7 and 19 dpi were determined and compared with the sequences of the parent strain (11Q472) and other cattle isolates (JN715035 and GU395535). Dots represent identical nucleotides

Fig. 3

Nucleotide alignment of the Npro region of BVDV. Nucleotide sequences from the blood of Goat 1 on 7 and 12 dpi were determined and compared with the sequences of the parent strain (11Q472) and other cattle isolates (EF101530 and U03912). Dots represent identical nucleotides


BVDV infection in small ruminants is similar to those in cattle. Postnatal infections cause mild clinical symptoms, including pyrexia and leukopenia, and infections in pregnant small ruminants may cause reproductive failures [11, 18,19,20,21,22]. Our findings revealed that experimental BVDV infection in Korean native goats via IN route showed a normal appearance with only mild respiratory signs such as coughing and nasal discharge and did not cause pyrexia, anorexia, and diarrhea. In a previous study, we reported BVDV infection in Saanen goats, which exhibited no clinical symptoms [15]. Several studies have demonstrated that clinical manifestation of BVDV infection in sheep and goats may differ depending upon inoculation route, age of the hosts, and the BVDV strain used [18,19,20,21,22]. Taken together, BVDV infection in Korean native goats induced via IN inoculation is accompanied only by respiratory symptoms. Further studies should focus toward investigating the clinical signs of BVDV infection in goats using various inoculation routes and viral strains.

In this study, viral RNA and serologic conversion were detected in only two of the inoculated goats, which exhibited coughing and nasal discharge. Two primers were used to detect BVDV infection in these goats, but PCR results were slightly different between the 5′-UTR and Npro regions (Table 2). Although we could not conclude which gene is better for BVDV diagnosis, we suggest that both genes can be used for a more accurate diagnosis. Moreover, the serological response in goats was relatively different from that in cattle. Seroprevalence rates of 3–35% were detected in small ruminants, and they were relatively lower in goats than in sheep [20]. It is speculated that the antibody production varies between goats and sheep. In our previous study on calves, seroconversion was evident for less than a week after inoculation [23], whereas in the present study, BVDV-specific antibodies were produced by goats for approximately 2 weeks after inoculation (Table 2), indicating a host-specific difference in seroconversion. These results suggested that BVDV requires time to induce infection in a new host.

The present results revealed that the nucleotide sequences of both the 5′-UTR and Npro regions observed in Korean native goats after inoculation were substantially different from those observed in cattle isolates (11Q472, JN715035, and GU395535 for 5′-UTR; EF101530 and U03912 for Npro). The changes in nucleotide sequences observed in BVDV-infected goats might be explained as a strategy for viral transmission to infect a new host. Consequently, these differences in the sequence may be essential to the process of establishing a BVDV infection in a new host. It has been reported that RN`11A viruses have a tendency to change hosts more frequently than other pathogens [24,25,26]. The virus may be pre-adapted to a new host, and these nucleotide changes may enhance the ability of the virus to establish infection in a new host. Bachofen et al. evaluated interspecific transmission from cattle to goats but did not identify any nucleotide changes in 5′-UTR [18]. However, changes were observed in the E2-coding region, which were quite different from our results. Although we did not evaluate nucleotide differences in the E2 region in this study, we found genetic changes in Korean native goats after BVDV infection in both the 5′-UTR and Npro regions; however, genetic changes were highly significant in the Npro region. Nucleotide changes may be necessary for the transmission of BVDV infection from cattle to goats. These results suggest that Korean native goats may possibly act as a reservoir host of BVDV.


Our results show that BVDV can establish an infection in Korean native goats and exhibits minimal clinical symptoms. The observed genetic changes may be explained as a result of the establishment of BVDV infection in a new host, and these consequences were found to be host-dependent. The shedding of BVDV from infected animals may facilitate the maintenance and transmission of BVDV to other hosts. Based on our findings, it is likely that Korean native goats play a role in BVDV maintenance in the ROK. Therefore, understanding interspecific transmission may be important for BVDV eradication programs.


Ethics statement

Korean native goats used in this study were acquired from a private farm located in Mungyeong, ROK. All goats were sacrificed by a veterinarian in accordance with the guidelines of the Animal Ethics Committee of Kyungpook National University (approval number: 2016–00040).

Virus preparation

The noncytopathic (ncp) BVDV-1b strain 11Q472 was isolated from a 2-year-old Korean indigenous cow presenting with emaciation, loss of appetite, and diarrhea in the ROK [23]. The virus strain was obtained after culturing for 5 days with one freeze-thaw cycle and centrifuging at 1900×g for 10 min to remove large cellular debris. The supernatant was aliquoted and then frozen at − 80 °C until inoculation. Viruses in the 5th passage of Madin-Darby Bovine Kidney (MDBK) cells were selected for inoculation; they were titrated from the MDBK cell cultures, and the median tissue culture-infective dose (TCID50) was calculated. MDBK cells were confirmed to be free of mycoplasma contamination using PCR (Takara Bio Inc., Japan). The MDBK cells were maintained in minimum essential medium (MEM; Life Technologies Corp., Carlsbad, CA, USA) supplemented with 5% horse serum (Thermo Fisher Scientific Inc., Waltham, MA, USA).

Experimental animals and virus infection

Korean native goats were maintained under pathogen-free conditions and handled in strictly in accordance with the guidelines and protocols approved by the Kyungpook National University Animal Care and Use Committee (approval number: 2016–00040). Goats used for this study were 3-month-old healthy females. The goats were negative for the BVDV antibody and antigen, as determined by the enzyme-linked immunosorbent assay and RT-PCR. Four goats were infected with 2.5 × 106 TCID50 of ncp BVDV-1b via IN route, and two goats served as uninfected controls (mock-infected) and received MEM (Life Technologies). The goats were housed individually. All experiments were conducted twice.

Sample collection, RT-PCR, and sequence analysis

Goats were monitored daily during the experiment. The body temperature was measured and recorded every other day at approximately the same time. Nasal swabs and blood samples were collected on 0, 2, 5, 7, 9, 12, 14, 16, 19, and 21 dpi, and the animals were euthanized on 21 dpi. Blood samples were obtained in Vacutainer tubes containing ethylenediaminetetraacetic acid (Becton Dickinson, Franklin Lakes, NJ, USA). RNA was extracted from the nasal swabs and blood samples using the PureLink® Total RNA Blood Kit (Invitrogen, Carlsbad, CA, USA). RT-PCR was performed using primers designed to amplify the 5′-UTR and Npro region (Table 3). The PCR products were purified using the AccuPrep® PCR Purification Kit (Bioneer, Daejeon, Korea), and the amplicons were used for direct sequencing (Bioneer). The sequence data were analyzed using Chromas software (version 2.33,, and alignments were obtained using CLUSTAL X (version 1.8). The amplified sequences were compared with those of the virus used for experimental inoculations. A phylogenetic tree was constructed based on nucleotide alignments using the neighbor-joining method [27]. A bootstrap analysis was performed with 1000 replicates using MEGA software version 6 [28].

Table 3 Primers used to detect BVDV in this study

Enzyme-linked immunosorbent assay

Antibody detection was performed using serum samples from the ncp BVDV-1b-infected and mock-infected goats with SVANOVIR® BVDV p80-Ab (SVANOVA, Uppsala, Sweden) according to the manufacturer’s instructions. Serum samples were added to a 96-well plate which was pre-coated with BVDV antigen, and subsequently incubated them for 1 h at 37 °C. Plates were washed, supplemented with horseradish peroxidase-labeled conjugate, and incubated for 1 h at 37 °C. After washing, the substrate solution was added and incubated for 10 min, followed by the addition of stop solution. The optical density was measured at 450 nm using a microplate reader (Tecan, Mannedorf, Switzerland). Samples were classified as positive when inhibition percentage was ≥45%.

Availability of data and materials

The datasets used and analyzed during the current study are available from the corresponding author on reasonable request.



5′- untranslated region


Border disease virus


Bovine viral diarrhea virus


Classical swine fever virus


Days post-infection





Npro :

N-terminal protease


Republic of Korea


Reverse transcription-polymerase chain reaction

TCID50 :

Median tissue culture infective dose


  1. 1.

    Avalos-Ramirez R, Orlich M, Thiel HJ, Becher P. Evidence for the presence of two novel pestivirus species. Virology. 2001;286:456–65.

    CAS  Article  Google Scholar 

  2. 2.

    Vilcek S, Ridpath JF, Van Campen H, Cavender JL, Warg J. Characterization of a novel pestivirus originating from a pronghorn antelope. Virus Res. 2005;108:187–93.

    CAS  Article  Google Scholar 

  3. 3.

    Kirkland PD, Frost MJ, Finlaison DS, King KR, Ridpath JF, Gu X. Identification of a novel virus in pigs--Bungowannah virus: a possible new species of pestivirus. Virus Res. 2007;129:26–34.

    CAS  Article  Google Scholar 

  4. 4.

    Bauermann FV, Ridpath JF, Weiblen R, Flores EF. HoBi-like viruses: an emerging group of pestiviruses. J Vet Diagn Invest. 2013;25:6–15.

    Article  Google Scholar 

  5. 5.

    Liu L, Xia H, Wahlberg N, Belak S, Baule C. Phylogeny, classification and evolutionary insights into pestiviruses. Virology. 2009;385:351–7.

    CAS  Article  Google Scholar 

  6. 6.

    Schirrmeier H, Strebelow G, Depner K, Hoffmann B, Beer M. Genetic and antigenic characterization of an atypical pestivirus isolate, a putative member of a novel pestivirus species. J Gen Virol. 2004;85:3647–52.

    CAS  Article  Google Scholar 

  7. 7.

    Krametter-Froetscher R, Duenser M, Preyler B, Theiner A, Benetka V, Moestl K, Baumgartner W. Pestivirus infection in sheep and goats in West Austria. Vet J. 2010;186:342–6.

    CAS  Article  Google Scholar 

  8. 8.

    Krametter-Froetscher R, Loitsch A, Kohler H, Schleiner A, Schiefer P, Moestl K, Golja F, Baumgartner W. Prevalence of antibodies to pestiviruses in goats in Austria. J Vet Med B Infect Dis Vet Public Health. 2006;53:48–50.

    CAS  Article  Google Scholar 

  9. 9.

    Toplak I, Sandvik T, Barlic-Maganja D, Grom J, Paton DJ. Genetic typing of bovine viral diarrhoea virus: most Slovenian isolates are of genotypes 1d and 1f. Vet Microbiol. 2004;99:175–85.

    CAS  Article  Google Scholar 

  10. 10.

    Bolin SR, Ridpath JF. Differences in virulence between two noncytopathic bovine viral diarrhea viruses in calves. Am J Vet Res. 1992;53:2157–63.

    CAS  PubMed  Google Scholar 

  11. 11.

    Passler T, Riddell KP, Edmondson MA, Chamorro MF, Neill JD, Brodersen BW, Walz HL, Galik PK, Zhang Y, Walz PH. Experimental infection of pregnant goats with bovine viral diarrhea virus (BVDV) 1 or 2. Vet Res. 2014;45:38.

    Article  Google Scholar 

  12. 12.

    Broaddus CC, Lamm CG, Kapil S, Dawson L, Holyoak GR. Bovine viral diarrhea virus abortion in goats housed with persistently infected cattle. Vet Pathol. 2009;46:45–53.

    CAS  Article  Google Scholar 

  13. 13.

    Lamm CG, Broaddus CC, Holyoak GR. Distribution of bovine viral diarrhea virus antigen in aborted fetal and neonatal goats by immunohistochemistry. Vet Pathol. 2009;46:54–8.

    CAS  Article  Google Scholar 

  14. 14.

    Pratelli A, Martella V, Cirone F, Buonavoglia D, Elia G, Tempesta M, Buonavoglia C. Genomic characterization of pestiviruses isolated from lambs and kids in southern Italy. J Virol Methods. 2001;94:81–5.

    CAS  Article  Google Scholar 

  15. 15.

    Han YJ, Chae JB, Chae JS, Yu DH, Park J, Park BK, Kim HC, Yoo JG, Choi KS. Identification of bovine viral diarrhea virus infection in Saanen goats in the Republic of Korea. Trop Anim Health Prod. 2016;48:1079–82.

    Article  Google Scholar 

  16. 16.

    Kim IJ, Hyun BH, Shin JH, Lee KK, Lee KW, Cho KO, Kang MI. Identification of bovine viral diarrhea virus type 2 in Korean native goat (Capra hircus). Virus Res. 2006;121:103–6.

    CAS  Article  Google Scholar 

  17. 17.

    Han DG, Ryu JH, Park J, Choi KS. Identification of a new bovine viral diarrhea virus subtype in the Republic of Korea. BMC Vet Res. 2018;14:233.

    Article  Google Scholar 

  18. 18.

    Bachofen C, Vogt HR, Stalder H, Mathys T, Zanoni R, Hilbe M, Schweizer M, Peterhans E. Persistent infections after natural transmission of bovine viral diarrhoea virus from cattle to goats and among goats. Vet Res. 2013;44:32.

    Article  Google Scholar 

  19. 19.

    Loken T, Bjerkas I, Larsen HJ. Experimental pestivirus infections in newborn goat kids. J Comp Pathol. 1990;103:277–88.

    CAS  Article  Google Scholar 

  20. 20.

    Passler T, Walz PH. Bovine viral diarrhea virus infections in heterologous species. Anim Health Res Rev. 2010;11:191–205.

    Article  Google Scholar 

  21. 21.

    Taylor WP, Okeke AN, Shidali NN. Experimental infection of Nigerian sheep and goats with bovine virus diarrhoea virus. Trop Anim Health Prod. 1977;9:249–51.

    CAS  Article  Google Scholar 

  22. 22.

    Wolff PL, Schroeder C, McAdoo C, Cox M, Nelson DD, Evermann JF, Ridpath JF. Evidence of bovine viral diarrhea virus infection in three species of sympatric wild ungulates in Nevada: life history strategies may maintain endemic infections in wild populations. Front Microbiol. 2016;7:292.

    Article  Google Scholar 

  23. 23.

    Seong G, Oem JK, Choi KS. Pathogenetic differences after experimental infection of calves with Korean non-cytopathic BVDV-1 and BVDV-2 isolates. Vet Immunol Immunopathol. 2013;156:147–52.

    CAS  Article  Google Scholar 

  24. 24.

    Bourret V, Lyall J, Frost SDW, Teillaud A, Smith CA, Leclaire S, Fu J, Gandon S, Guerin JL, Tiley LS. Adaptation of avian influenza virus to a swine host. Virus Evol. 2017;3:vex007.

    Article  Google Scholar 

  25. 25.

    Longdon B, Brockhurst MA, Russell CA, Welch JJ, Jiggins FM. The evolution and genetics of virus host shifts. PLoS Pathog. 2014;10:e1004395.

    Article  Google Scholar 

  26. 26.

    Woolhouse ME, Haydon DT, Antia R. Emerging pathogens: the epidemiology and evolution of species jumps. Trends Ecol Evol. 2005;20:238–44.

    Article  Google Scholar 

  27. 27.

    Saitou N, Nei M. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Mol Biol Evol. 1987;4:406–25.

    CAS  PubMed  Google Scholar 

  28. 28.

    Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics analysis version 6.0. Mol Biol Evol. 2013;30:2725–9.

    CAS  Article  Google Scholar 

Download references


We thank Young-Je Kwon and Hyung-Tae Kim at the Department of Animal Biotechnology, Kyungpook National University; Young-Jae Kwon at the Department of Animal Science, Kyungpook National University for animal care and technical support; and Yoon-Joo Lee at the Department of Animal Science and Biotechnology, Kyungpook National University for the modification of figures.


This work was supported by the National Research Foundation of Korea (NRF), funded by the Korea government (MSIP) (No. 2018R1D1A1B07048271). The funders had no role in the study design, data collection and analysis, interpretation of results, writing of the report, decision to submit the paper for publication.

Author information




KSC designed the experiments and wrote the manuscript. JKO and DGH performed the experiments and conducted data analysis. JKO contributed to writing of the manuscript. All authors have read and approved the final manuscript.

Corresponding author

Correspondence to Kyoung-Seong Choi.

Ethics declarations

Ethics approval

All procedures involving animals in this study were performed in accordance with the ethical standards of the Kyungpook National University Animal Care and Use Committee. We obtained written informed consent from the owner of the animals for using them in our study.

Consent for publication

Not applicable

Competing interests

The authors declare that they have no competing interests.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Additional files

Additional file 1:

Table S1. BVDV isolates used in the phylogenetic tree based on the 5′-UTR. (DOCX 17 kb)

Additional file 2:

Table S2. BVDV isolates used in the phylogenetic tree based on the Npro region. (DOCX 18 kb)

Rights and permissions

Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (, which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver ( applies to the data made available in this article, unless otherwise stated.

Reprints and Permissions

About this article

Verify currency and authenticity via CrossMark

Cite this article

Oem, J., Han, D. & Choi, K. Experimental infection of Korean native goats (Capra aegagrus hircus) with bovine viral diarrhea virus 1b. BMC Vet Res 15, 202 (2019).

Download citation


  • Bovine viral diarrhea virus
  • Noncytopathic
  • Korean native goat
  • 5′-untranslated region
  • N-terminal protease region