Skip directly to site content Skip directly to page options Skip directly to A-Z link Skip directly to A-Z link Skip directly to A-Z link
Volume 10, Number 6—June 2004

Bovine Spongiform Encephalopathy Infectivity in Greater Kudu (Tragelaphus strepsiceros)

Article Metrics
citations of this article
EID Journal Metrics on Scopus
Andrew A. Cunningham*Comments to Author , James K. Kirkwood*1, Michael Dawson†2, Yvonne I. Spencer†, Robert B. Green†, and Gerald A.H. Wells†
Author affiliations: *Institute of Zoology, Regent’s Park, London, UK; †Veterinary Laboratories Agency, Addlestone, Surrey, UK; Bovine Spongiform Encephalopathy in Greater Kudu

Cite This Article


Of all the species exposed naturally to the bovine spongiform encephalopathy (BSE) agent, the greater kudu (Tragelaphus strepsiceros), a nondomesticated bovine from Africa, appears to be the most susceptible to the disease. We present the results of mouse bioassay studies to show that, contrary to findings in cattle with BSE in which the tissue distribution of infectivity is the most limited recorded for any of the transmissible spongiform encephalopathies (TSE), infectivity in greater kudu with BSE is distributed in as wide a range of tissues as occurs in any TSE. Agent was also detected in skin, conjunctiva, and salivary gland, tissues in which infectivity has not previously been reported in any naturally occurring TSE. The distribution of infectivity in greater kudu with BSE suggests possible routes for transmission of the disease and highlights the need for further research into the distribution of TSE infectious agents in other host species.

To date, 13 species of zoo animal have been confirmed as having died with a novel scrapie-like spongiform encephalopathy (SE) concurrent with the bovine spongiform encephalopathy (BSE) epidemic (Table 1). The disease is thought, in some, if not all, of these species, to be caused by infection with the BSE agent. In addition, natural infection with BSE has been reported in five species of primate in French zoos (11), but these results are considered equivocal for the confirmation of a spongiform encephalopathy (12, G.A.H. Wells, unpub. data). BSE was diagnosed in six of eight greater kudu (Tragelaphus strepsiceros), a member of the family Bovidae, subfamily Bovinae, that died at the London Zoo from 1989 through 1992 (2,13,14). The epidemiology of this disease in greater kudu is consistent with either a particularly high susceptibility to infection, the occurrence of direct animal-to-animal transmission of the disease, or with a combination of these factors (2,13,15). To investigate further the biology of BSE in greater kudu, the distribution of the infectious agent in greater kudu with BSE was determined by using the mouse bioassay method.

Materials and Methods


Tissues from four greater kudu that died with spongiform encephalopathy were tested for infectivity by bioassay in C57Bl-J6 mice (Table 2). The epidemiologic, clinical, and pathologic findings of the disease in the kudu have been described in detail previously (2,1416), and a summary of the relevant details is given in Table 3. Tissues for bioassay were collected principally from kudu A1212; each sample was collected in a sterile container by using new disposable instruments and gloves to prevent cross contamination between tissues. As TSE infectivity has been demonstrated previously by bioassay in tissues preserved in formalin and paraffin wax (13,17,18), tissue samples obtained opportunistically after routine postmortem examinations of three additional kudu (A664, A666, and A1221) were also tested for infectivity by using mouse bioassay. Samples collected from kudu A666, A1212, and A1221 were stored in separate, sterile containers and were either frozen at –70°C or fixed in neutral buffered 10% formalin. Non-neural tissues from kudu A664 were fixed in neutral buffered 10% formalin in a common container. The brain from this animal was the last organ removed at necropsy and was fixed in 10% formol saline in a separate container.

Previously, infectivity had been detected in formalin-fixed brain tissue from kudu A664 by bioassay using five inbred mouse strains, with similar incubation periods and lesion profiles to those demonstrated for the BSE agent from domestic cattle with BSE (13). Bioassay of brain from kudu A664 was not repeated in the current study, but a positive control sample of fresh brain tissue from clinically affected kudu A1212 was tested.


The tissues injected into mice for BSE-bioassay are listed in Table 2. Most of the tissue homogenates were prepared from thawed samples of fresh tissues frozen at –20°C. Tissue homogenates prepared from formalin-fixed tissues were rinsed overnight in running water to leach out the fixative, while formalin-fixed, paraffin-embedded tissues were dewaxed in chloroform (two changes) and washed in several changes of absolute alcohol before being rehydrated by immersion in a series of aqueous solutions of descending concentrations of alcohol, through to 100% water. Material for each tissue homogenate was dissected from the center of each tissue sample by using single-use disposable instruments and rigorous sterile procedures. Each sample was homogenized in 10% physiologic saline to make a 10% wt/vol suspension, which was then passed through a gauze filter. To tissue homogenates containing distal ileum or feces, ampicillin was added at the rate of 1.25 mg/mL.

For each tissue homogenate, 20 C57Bl-J6 mice (4–7 weeks old) were each injected by the intracranial route (0.02 mL) and by the intraperitoneal route (0.10 mL). Single tissue or pooled tissue samples were prepared and injected into C57Bl-J6 mice for a standard qualitative assay of infectivity (13,19).

Mice injected with different tissue or tissue-pool homogenates were housed in separate cages. Injected mice were coded, and detailed clinical monitoring of the mice was carried out by using a standard protocol. The clinical endpoint was determined when mice either showed clear signs of neurologic disease (20) or other deterioration of health. Surviving mice were killed 950 days after injection. Postmortem confirmation of disease in mice was routinely carried out by histopathologic examination of the brain for morphologic changes of spongiform encephalopathy.

After the histopathologic assessment of mice, immunohistochemical examination (IHC) for evidence of spongiform encephalopathy disease-specific PrP (PrPSc) was performed on the brains of all mice in selected tissue groups. The groups were mice in which either a low number were positive, testing was inconclusive on histopathologic assessment, or the results indicated a novel or anomalous distribution of the agent in kudu compared to that in other TSE. Additional groups of interest (skeletal muscle, endometrium, and mammary gland), which were negative on the histopathologic examination of mouse brains, were also examined by IHC. Immunohistochemical detection of PrPSc was introduced to the standard protocol to improve specificity and sensitivity of detecting BSE transmission to mice (21,22) and interpret inconclusive histopathologic results (23). For control purposes, the brains from mice that had been injected with pathologically affected cranial thoracic spinal cord from kudu A1212 were also immunostained. Brains from normal mice that were not injected with infected tissues were similarly examined to provide negative controls.

The immunohistochemical method used was essentially that applied previously to cattle central nervous system tissues (24) and adapted for use in mouse brain tissue. Anti-bovine PrPSc serum (971) was used at 1/8,000 and 1/16,000 dilutions in an avidin-biotin-peroxidase (ABC) complex technique. Transmission was defined histopathologic evidence of spongiform encephalopathy, or, where applied, immunohistochemical presence of disease-specific PrP (PrPSc) in the brains of the mice.


The results of the bioassay of tissues are given in Table 2. Based on the histopathologic examination of mouse brains, 15 of the 32 tissue homogenates were positive. The nine histopathologically positive groups examined immunohistochemically were confirmed positive by this method with marginal improved sensitivity of detection (2–3 more mice positive) in only three groups. For nine of the positive tissue homogenates, prepared from fresh central nervous, lymphoreticular, or distal ileum tissue, the proportion of positive mice (>40%) indicated moderate or high levels of spongiform encephalopathy infectivity. The remaining six positive groups (Table 2) had low proportions of positive mice (6%–27%), indicative of relatively low titres or only traces of infectivity. Low numbers (1–2) of histopathologically inconclusive mice in five tissue homogenate groups, which included two groups (A1212, popliteal and submandibular lymph nodes) that contained no histopathologically positive mice, were resolved almost exclusively as negative when examined using immunohistochemistry. The one exception was an inconclusive mouse in A1212 retropharyngeal lymph node group, which proved immunohistochemically positive.


Fifteen of the 32 kudu tissue homogenates transmitted BSE to mice. The positive result for brain tissue from kudu A1221 confirms the diagnosis of subclinical spongiform encephalopathy in this animal (15) and is the first to demonstrate transmission from a subclinical natural case of spongiform encephalopathy in a bovine species. Also, this report is the first of infectivity in the ileum from a field case of spongiform encephalopathy other than scrapie in sheep.

Apparently low titers or only traces of infectivity were detected in spleen, lung, submandibular salivary gland, conjunctiva, and skin. In bioassays of TSE infectivity, the possibility that trace levels of infectivity in tissues may represent postmortem or laboratory contamination of uninfected tissues with infected material has to be considered. Such an explanation is unlikely in the present study for the following reasons. Many of the tissues which contained only traces of infectivity were taken from kudu A1212, an animal from which tissues were collected by using rigorous sterile procedures to prevent cross-contamination, and no pattern between the order of tissue sampling and the bioassay results from this animal suggests a sequence of tissue contamination. A low incidence of disease in the mice or failure to detect infectivity from tissues previously fixed or processed to paraffin wax may be attributable to a reduced titer of infectivity, which can occur as a result of such treatments. The wide range of survival periods for positive mice in these assays (Table 2) is similar to those seen when brain tissue from confirmed cases of BSE in domestic cattle was injected into C57Bl-J6 mice of the same source as used in the current study (G.A.H. Wells and M. Dawson, unpub. data). These results contrast with those from a previous study (13) in which 20 of 21 C57Bl mice were positive for spongiform encephalopathy after injection of formalin-fixed brain from the index case of TSE in greater kudu (kudu A664), with a mean incubation period of 465±14 days (M. Bruce, pers. comm.). We conclude that the low incidence of positive mice in certain tissue groups is due to a lower titer of infectious agent within these tissues when compared with the CNS or ileum.

The distribution of BSE infectivity in tissues of greater kudu contrasts with that in tissues of BSE-infected cattle (24,25) but is more like the distribution found in genetically susceptible sheep infected with scrapie or experimental BSE (2628). In field cases of cattle with BSE, infectivity has been found only in the CNS (25), but in cattle experimentally challenged orally with the agent of BSE, ileum and bone marrow have also been shown to contain infectivity (21,24,29). In classical studies of scrapie in sheep and goats, infectivity was detected in the nervous and lymphoreticular systems, placenta, adrenal gland, nasal mucosa, lung, pancreas, liver, bone marrow, thymus, and alimentary tract, although most of the non-neural, non-lymphoreticular peripheral tissues contained only low titers of agent (30,31).

The finding of infectivity in kudu skin, conjunctiva, and submandibular salivary gland was unexpected as these tissues have not been previously shown to be infective in scrapie, BSE, or any naturally occurring TSE. Nonetheless, infectivity has been found previously in salivary glands of mice after injection of infected tissues with a high titer of scrapie agent (32) and of mink injected with the transmissible mink encephalopathy agent (33). In experimentally-induced transmissible mink encephalopathy agent, low concentrations of agent occurred in liver, kidney, intestine, and salivary gland only after replication in the CNS and in some lymphoreticular tissues (24). The inconsistent observation of low levels or traces of infection in certain non-neural and non-lymphoreticular tissues is in general a feature of both natural and experimental TSE. Given the relative paucity of data on the tissue distribution of infectivity in TSEs, the finding of infection in any given tissue is unprecedented. The infectivity in certain tissues subsequent to CNS involvement may be a rare event incidental to the pathogenesis of the disease.

We have previously indicated that the epidemiology of BSE in the small kudu herd at London Zoo was consistent with either a particularly high susceptibility to infection, the occurrence of direct animal-to-animal transmission of the disease, or with a combination of these factors (46). The presence of infectivity in tissues, such as the skin and salivary gland, suggests possible routes by which direct transmission could occur. Eklund et al. (32), for example, suggested infection of the salivary gland as an explanation for contact infection of scrapie between mice.

Given the extended survival period range with BSE in the C57Bl-J6 mice used in the current study compared to the incubation periods in C57Bl mice used previously (13) and the relative insensitivity of the mouse model (24), these results may be an underestimate of the extent of infectivity in the kudu tissues assayed. A recently reported rapid immunoassay shown to be capable of detecting PrPBSE in the brainstems of cattle with a sensitivity similar to that of the infectivity levels determined by end-point titration in Tg(BoPrP) mice (34) possibly offers prospects for more sensitive detection of disease-related PrP as a proxy for infectivity bioassay. An important area for further research, therefore, is to investigate whether our results represent true qualitative differences in the biology of BSE in the greater kudu and the domestic cow or possibly indicate similarities, unapparent only because of the variables inherent in the sensitivities of current bioassay methods.

Dr. Cunningham, a veterinary pathologist, is a senior research fellow and head of Wildlife Epidemiology at the Institute of Zoology, Zoological Society of London. His research interests include emerging infectious diseases of wildlife and disease threats to biodiversity conservation.



We thank the staff of the Veterinary Laboratories Agency–Weybridge for technical support.

The study was funded by the former Ministry of Agriculture, Fisheries and Food.



  1. Jeffery  M, Wells  GAH. Spongiform encephalopathy in a nyala (Tragelaphus angasi). Vet Pathol. 1988;25:3989. DOIPubMedGoogle Scholar
  2. Kirkwood  JK, Cunningham  AA, Wells  GAH, Wilesmith  JW, Barnett  JEF. Spongiform encephalopathy in a herd of greater kudu Tragelaphus strepsiceros: epidemiological observations. Vet Rec. 1993;133:3604. DOIPubMedGoogle Scholar
  3. Kirkwood  JK, Wells  GAH, Wilesmith  JW, Cunningham  AA, Jackson  SI. Spongiform encephalopathy in an Arabian oryx (Oryx leucoryx) and a greater kudu (Tragelaphus strepsiceros). Vet Rec. 1990;127:41820.PubMedGoogle Scholar
  4. Department for Environment. Food and Rural Affairs. BSE information. 2002; Available from:
  5. Kirkwood  JK, Cunningham  AA. Spongiform encephalopathies in captive wild animals in Europe, 1986–99. In: Proceedings of the 26th World Veterinary Congress, Lyon; 1999 Sept 23–26. World Veterinary Association; 1999.
  6. Baron  T, Belli  P, Madec  JY, Moutou  F, Vitaud  C, Savey  M. Spongiform encephalopathy in an imported cheetah in France. Vet Rec. 1997;141:2701. DOIPubMedGoogle Scholar
  7. Vitaud  C, Flach  EJ, Thornton  SM, Capello  R. Clinical observations in four cases of feline spongiform encephalopathy in cheetahs (Acionyx jubatus). European Association of Zoo and Widlife Veterinarians (EAZWV) Second scientific meeting May 21–24, 1998. Chester, UK. p. 133–8.
  8. Lezmi  S, Bencsik  A, Monks  E, Petit  T, Baron  T. First case of feline spongiform encephalopathy in a captive cheetah born in France: PrPsc analysis in various tissues revealed unexpected targeting of kidney and adrenal gland. Histochem Cell Biol. 2003;119:41522.PubMedGoogle Scholar
  9. Willoughby  K, Kelly  DF, Lyon  DG, Wells  GAH. Spongiform encephalopathy in a captive puma (Felis concolor). Vet Rec. 1992;131:4314. DOIPubMedGoogle Scholar
  10. Young  S, Slocombe  RF. Prion-associated spongiform encephalopathy in an imported Asiatic golden cat (Catopuma temmincki). Aust Vet J. 2003;81:2956. DOIPubMedGoogle Scholar
  11. Bons  N, Mestre-Francés  N, Belli  P, Cathala  F, Gajdusek  DC, Brown  P. Natural and experimental oral infection of nonhuman primates by bovine spongiform encephalopathy agents. Proc Natl Acad Sci U S A. 1999;96:404651. DOIPubMedGoogle Scholar
  12. Baker  HF, Ridley  RM, Wells  GAH, Ironside  JW. Spontaneous spongiform encephalopathy in a monkey. Lancet. 1996;348:9556. DOIPubMedGoogle Scholar
  13. Bruce  M, Chree  A, McConnell  I, Foster  J, Pearson  G, Fraser  H. Transmission of bovine spongiform encephalopathy and scrapie to mice: strain variation and the species barrier. Philos Trans R Soc Lond B Biol Sci. 1994;343:40511. DOIPubMedGoogle Scholar
  14. Kirkwood  JK, Cunningham  AA. Epidemiological observations on spongiform encephalopathies in captive wild animals in the British Isles. Vet Rec. 1994;135:296303. DOIPubMedGoogle Scholar
  15. Cunningham  AA, Wells  GAH, Scott  AC, Kirkwood  JK, Barnett  JEF. Transmissible spongiform encephalopathy in greater kudu (Tragelaphus strepsiceros). Vet Rec. 1993;132:68. DOIPubMedGoogle Scholar
  16. Kirkwood  JK, Cunningham  AA, Austin  AR, Wells  GAH, Sainsbury  AW. Spongiform encephalopathy in a greater kudu (Tragelaphus strepsiceros) introduced into an affected group. Vet Rec. 1994;134:1678. DOIPubMedGoogle Scholar
  17. Brown  P, Rohwer  RG, Green  EM, Gajdusek  DC. Effect of chemicals, heat and histopathological processing on high-infectivity hamster-adapted scrapie virus. J Infect Dis. 1982;145:6837.PubMedGoogle Scholar
  18. Fraser  H, Bruce  ME, Chree  A, McConnell  I, Wells  GAH. Transmission of bovine spongiform encephalopathy and scrapie to mice. J Gen Virol. 1992;73:18917. DOIPubMedGoogle Scholar
  19. Wells  GAH, Dawson  M, Hawkins  SAC, Austin  AR, Green  RB, Dexter  I, Preliminary observations on the pathogenesis of experimental bovine spongiform encephalopathy. In: Gibbs CJ, editor. Bovine spongiform encephalopathy: the BSE dilemma. New York: Springer-Verlag; 1996. p. 28–44.
  20. Dickinson  AG, Meikle  VMH, Fraser  H. Identification of a gene which controls the incubation period of some strains of scrapie agent in mice. J Comp Pathol. 1968;78:2939. DOIPubMedGoogle Scholar
  21. Wells  GAH, Hawkins  SAC, Green  RB, Spencer  YI, Dexter  I, Dawson  M. Limited detection of sternal bone marrow infectivity in the clinical phase of experimental bovine spongiform encephalopathy (BSE). Vet Rec. 1999;144:2924. DOIPubMedGoogle Scholar
  22. Wells  GAH, Hawkins  SAC, Austin  AR, Ryder  SJ, Done  SH, Green  RB, Studies of the transmissibility of the agent of bovine spongiform encephalopathy to pigs. J Gen Virol. 2003;84:102131. DOIPubMedGoogle Scholar
  23. Fraser  H, McBride  PA. Parallels and contrasts between scrapie and dementia of the Alzheimer type and ageing: strategies and problems for experiments involving lifespan studies. In: Traber J, Gispen WH, editors. Senile dementia of the Alzheimer type. Berlin: Springer-Verlag; 1985. p. 250–68.
  24. Wells  GAH, Hawkins  SAC, Green  RB, Austin  AR, Dexter  I, Spencer  YI, Preliminary observations on the pathogenesis of experimental bovine spongiform encephalopathy (BSE): an update. Vet Rec. 1998;142:1036. DOIPubMedGoogle Scholar
  25. Fraser  H, Foster  J. Transmission to mice, sheep and goats and bioassay of bovine tissues. In: Transmissible spongiform encephalopathies. A consultation on BSE with the Scientific Veterinary Committee of the Commission of the European Communities held in Brussels, September 14–15, 1993. Bradley R, Marchant B, editors. Document VI/4131/94-EN. Brussels, European Commission Agriculture. 1994. p. 145–159.
  26. European Commission. Update of the opinion on TSE infectivity distribution in ruminant tissues (Initially adopted by the Scientific Steering Committee at its meeting of January 10–11, 2002 and amended at its meeting of November 7–8, 2002) following the submission of (1) a risk assessment by the German Federal Ministry of Consumer Protection, food and Agriculture and (2) new scientific evidence regarding BSE infectivity distribution in tonsils; 2002. Available from:
  27. Foster  JD, Parnham  DW, Hunter  N, Bruce  M. Distribution of the prion protein in sheep terminally affected with BSE following experimental oral transmission. J Gen Virol. 2001;82:231926.PubMedGoogle Scholar
  28. Jeffrey  M, Ryder  S, Martin  S, Hawkins  SAC, Terry  L, BerthelinBaker C, et al. Oral inoculation of sheep with the agent of bovine spongiform encephalopathy (BSE). 1. Onset and distribution of disease-specific PrP accumulation in brain and viscera. J Comp Pathol. 2001;124:2809. DOIPubMedGoogle Scholar
  29. Wells  GAH, Dawson  M, Hawkins  SAC, Green  RB, Dexter  I, Francis  ME, Infectivity in the ileum of cattle challenged orally with bovine spongiform encephalopathy. Vet Rec. 1994;135:401. DOIPubMedGoogle Scholar
  30. Hadlow  WJ, Kennedy  RC, Race  RE. Natural infection of Suffolk sheep with scrapie virus. J Infect Dis. 1982;146:65764.PubMedGoogle Scholar
  31. Hadlow  WJ, Kennedy  RC, Race  RE, Eklund  CM. Virologic and neurohistologic findings in dairy goats affected with natural scrapie. Vet Pathol. 1980;17:18799.PubMedGoogle Scholar
  32. Eklund  CM, Kennedy  RC, Hadlow  WJ. Pathogenesis of scrapie virus infection in the mouse. J Infect Dis. 1967;117:1522.PubMedGoogle Scholar
  33. Hadlow  WJ, Race  RE, Kennedy  RC. Temporal distribution of transmissible mink encephalopathy virus in mink inoculated subcutaneously. J Virol. 1987;61:323540.PubMedGoogle Scholar
  34. Safar  JG, Scott  M, Monaghan  J, Deering  C, Didorenko  S, Vergara  J, Measuring prions causing bovine spongiform encephalopathy or chronic wasting disease by immunoassays and transgenic mice. Nat Biotechnol. 2002;20:114750. DOIPubMedGoogle Scholar




Cite This Article

DOI: 10.3201/eid1006.030615

1Present affiliation is Universities Federation for Animal Welfare, Wheathampstead, Hertfordshire, UK.

2Present affiliation is National Scrapie Plan Administration Centre, Worcester, UK.

Table of Contents – Volume 10, Number 6—June 2004

EID Search Options
presentation_01 Advanced Article Search – Search articles by author and/or keyword.
presentation_01 Articles by Country Search – Search articles by the topic country.
presentation_01 Article Type Search – Search articles by article type and issue.



Please use the form below to submit correspondence to the authors or contact them at the following address:

Institute of Zoology, Regent’s Park, London NW1 4RY, UK; fax: +44-0-20-7586-1457

Send To

10000 character(s) remaining.


Page created: February 22, 2011
Page updated: February 22, 2011
Page reviewed: February 22, 2011
The conclusions, findings, and opinions expressed by authors contributing to this journal do not necessarily reflect the official position of the U.S. Department of Health and Human Services, the Public Health Service, the Centers for Disease Control and Prevention, or the authors' affiliated institutions. Use of trade names is for identification only and does not imply endorsement by any of the groups named above.