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Chronic wasting disease of cervids
Small Ruminant Research 128 (2015) 72–78
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Chronic wasting disease of cervids夽 D.C. Bourne ∗ Wildlife Information Network, Twycross Zoo – East Midland Zoological Society, Atherstone, Warwickshire, CV9 3PX, UK
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Article history: Available online 18 March 2015 Keywords: Chronic wasting disease (CWD) Transmissible spongiform encephalopathy Cervids
a b s t r a c t Chronic wasting disease (CWD) of cervids is the only known transmissible spongiform encephalopathy (TSE) found in non-domestic, free-ranging animals. To date, it is found in wild cervids only in North America, and natural infection has been detected in only four species (Cervus elaphus nelsoni, Odocoileus hemionus, Odocoileus virginianus and recently Alces alces) although there are concerns that it could spread to other species, particularly Rangifer tarandus. The infectious PrPCWD spreads throughout the body, particularly in lymphoid tissue, although lesions (typical of TSEs) are found only in the brain and are associated with development of clinical disease (e.g. wasting, behavioural changes). Transmission is lateral and probably oral. Infectious prions are shed in faeces, urine and saliva, and are present in various body tissues, all of which may contribute to ante- or post-mortem environmental contamination, increasing transmission. Infectious prions persist in soils. There is presently no evidence of spread to other sympatric wildlife, domestic livestock or humans. Intracerebral inoculation enables transmission to various species of ungulates, rodents, carnivores and even primates, but oral transmission to non-cervids has mainly been unsuccessful. © 2015 Elsevier B.V. All rights reserved.
1. Introduction
2. Emergence, aetiology and possible origins
Chronic wasting disease (CWD) is one of the group of diseases generally known as the transmissible spongiform encephalopathies (TSE) or prion diseases, and the only known prion disease of free-living non-domestic animals. As with other prion diseases, CWD is associated with a misfolded isoform (PrPCWD or PrPres ) of a normal cellular protein (PrPC ), which accumulates in the CNS, with resultant neurodegenerative changes; this abnormal PrP is transmissible (Bourne, 2004).
CWD was first detected as a clinical wasting syndrome in mule deer (Odocoileus hemionus hemionus) at a research facility in Fort Collins, CO, USA, in 1967; it was recognised as a spongiform encephalopathy (SE) in 1978 (Williams and Young, 1980) and was shown to be transmissible by intracerebral (IC) inoculation into ferrets and mule deer fawns (Williams et al., 1982). The first diagnosis in captive Rocky Mountain elk (Cervus elaphus nelsoni) occurred in 1978 (Williams and Young, 1982); it was first diagnosed in wild cervids in 1981 (Spraker et al., 1997) and in farmed cervids in Saskatchewan, Canada (Sullivan, 1996). The origin of CWD is uncertain; the favoured theory is that it derived from scrapie (Williams, 2005); scrapie inoculation IC into elk produced typical CWD lesions (Hamir et al., 2004). It is also possible that CWD arose de novo in deer and became transmissible (Bourne, 2004; Williams et al., 2002). Alternatively, it originated in another species,
夽 This paper is part of a set of articles on the theme of ‘Diseases of Cervids’, guest-edited by C. Billinis, who gratefully acknowledges the contribution of the authors and the editorial staff. ∗ Current address: MA Healthcare Ltd., St Jude’s Church, Dulwich Road, London, SE24 0PB, UK. E-mail addresses: [email protected], [email protected] http://dx.doi.org/10.1016/j.smallrumres.2015.03.008 0921-4488/© 2015 Elsevier B.V. All rights reserved.
D.C. Bourne / Small Ruminant Research 128 (2015) 72–78
as yet unknown (Williams, 2005). Based on strain typing by inoculation into genotypically characterised mice, the CWD agent is different from BSE agent, TME agent and tested strains of CJD agent and scrapie agent (Bruce et al., 1997, 2000; Laplanche et al., 1999). It is not yet certain whether there is a single or multiple strains of CWD. Epidemiological data and the marked similarity of the lesions strongly suggest that the same CWD agent is responsible for the disease in captive and free-ranging deer and elk (Williams and Miller, 2002). Studies in transgenic mice expressing cervid PrP indicated the same prion strain in analysed elk and mule deer (Browning et al., 2004), while ferret-passaged isolates from two sources showed differences in clinical presentation, survival period, lesion distribution, glycoform profile and resistance to proteolysis, suggesting different strains (Perrott et al., 2012). It is difficult to draw conclusions from such experiments because interspecies transmission can alter the characteristics of TSE agents (Sigurdson, 2008). 3. Geographical distribution CWD was first found in research facilities in Colorado and Wyoming (Williams and Young, 1992; Williams and Miller, 2002). It has been confirmed in free-living cervids in 17 states in the USA (Colorado, Wyoming, Utah, Kansas, Nebraska, New Mexico, Texas, North Dakota, South Dakota, Wisconsin, Illinois, Maryland, Missouri, New York, Minnesota, Virginia, West Virginia) and in Canada (Alberta and Saskatchewan). In farmed cervids, it has been detected in Colorado, Nebraska, Montana, South Dakota, Kansas, Oklahoma, Minnesota, Iowa, Wisconsin, Missouri, Michigan, Pennsylvania and New York, and in Canada in Alberta and Saskatchewan (APHIS, 2012; NWHC, 2012). Occurrence and prevalence of CWD varies between states and between regions within a state; even within affected regions there are clusters of affected animals (Joly et al., 2006). There have been very few cases in cervids in North American zoos, with no evidence of the disease remaining in zoos in recent years (Bourne, 2004; Dubé et al., 2006; Williams and Young, 1992). Outside North America, CWD has been detected only in farmed cervids in Korea, associated with elk imported from Canada (Kim et al., 2005; ProMED-Mail, 2011; Sohn et al., 2002). 4. Host range The known natural hosts of chronic wasting disease are mule deer, white-tailed deer, Rocky Mountain elk and moose (Alces alces shirasi) (Baeten et al., 2007; Williams et al., 2002). European red deer (Cervus elaphus elaphus) are susceptible on oral inoculation (Balachandran et al., 2010). Natural infection has not been reported in Rangifer tarandus (reindeer or caribou) (Lapointe et al., 2002; Sigurdarson, 2004). Oral transmission was successful in some reindeer; it is possible that some reindeer PRNP polymorphisms are protective against CWD (Mitchell et al., 2012). In Korea, CWD has been confirmed in Sika deer (Cervus nippon) and in red deer × sika deer (ProMED-Mail, 2011). Fallow deer (Dama dama) have been infected by IC inoculation from elk or white-tailed deer, with development of SE
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lesions (Hamir et al., 2010), but not by exposure to infected mule deer and a CWD-contaminated environment, suggesting lower susceptibility or delayed progression in this species (Rhyan et al., 2011). In Europe, surveillance of roe deer (Capreolus capreolus), red deer, fallow deer, muntjac (Muntiacus reevesi) and reindeer has not yet detected any cases of TSE (A.M. Barlow, pers. comm., 2004; Schettler et al., 2004; Schwaiger et al., 2004; Sigurdarson, 2004). It has been possible to infect domestic cattle, goats and sheep by IC inoculation, although not in all individuals and with several-year incubation periods in cattle and goats (Hamir et al., 2006b, 2011; Williams and Young, 1992). Cattle failed to develop SE although PrPres was detected in the CNS (Hamir et al., 2011). In sheep, only two of eight became infected: one (ARQ/VRQ at codons 136, 154, and 171) developing clinical signs, SE lesions and was PrPres positive at 35 months post inoculation (mpi) and one (ARQ/ARQ) SE and PrPres positive but no clinical signs at 72 months pi; two ARQ/ARQ and four ARQ/ARR were negative for PrPres at 36–72 mpi (Hamir et al., 2006b). Oral transmission into cattle failed (to nine years post infection) (Hamir et al., 2011). In carnivores, initial IC inoculation of mule deer CWD material into ferrets gave an incubation period of over a year to clinical signs (Williams et al., 1982). Serial passage reduced the incubation period to as low as 5 months (Bartz et al., 1998). Inoculation into American mink (Mustela vison) has also been successful, but IC inoculation of Common raccoon (Procyon lotor) kits with CWD failed (Hamir et al., 2007). In Wisconsin, brains of 812 mammalian scavengers were tested; the IDEXX HerdChek ELISA gave positive results in 1/11 Mustela vison, 1/259 Procyon lotor and 2/202 Didelphis virginiana but none were confirmed by Western blotting. The rest were all negative by ELISA (Jennelle et al., 2009). In rodents, IC inoculation into laboratory mice produced infection in only “a very few mice”, after >500 days incubation; serial passage in mice produced clinical disease (Bruce et al., 2000). Golden (Syrian) hamsters (Mesocricetus auratus) were not infected by IC inoculation of material from CWD mule deer (Williams and Young, 1992; Williams et al., 1992), but were infected following passage in ferrets, indicating that the host range of CWD can be altered by passage through other species (Bartz et al., 1998). To date there are no known cases of human prion disease attributable to CWD transmitted to humans. Epidemiological investigations have failed to show links between prion disease in hunters or young people in North America and CWD and there is no evidence of increased CJD in Colorado or Wyoming (Belay et al., 2001, 2004; CDC, 2003). In vitro, CWD-associated PrPres was shown to convert human PrPsen only at very low efficiency (Raymond et al., 2000). IC inoculation of CWD-infected cervid material into transgenic mice expressing human PrP failed (Kong et al., 2005; Sandberg et al., 2010). Squirrel monkeys (Saimiri sciureus) inoculated IC with one CWD strain became ill at 31–34 mpi and were PrPres positive (Marsh et al., 2005). Further experiments produced clinical disease in 33–53 months with 7/8 isolates IC and infection of 3/15 monkeys orally. In contrast, 15 cynomolgus macaques inoculated IC and orally remained well at 70 mpi and no PrPres was detected by IHC or Western blot in the
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brain, spinal cord, spleen and lymph nodes of one individual euthanased at 48 months (Race et al., 2009). 5. CWD in cervids The natural incubation period for CWD in naturally exposed cervids is not known and probably varies depending on frequency of exposure and the dose received (O’Rourke et al., 2004; Spraker, 2003; Spraker et al., 2004) In early studies, clinical signs were first noted in deer at 18 months (Williams and Young, 1992); the youngest recorded affected free-ranging elk was 21 months old (Spraker et al., 1997). Cervids less than 12 months old have been confirmed CWD-positive by IHC (Kahn et al., 2004). In orally inoculated mule deer fawns, PrPCWD was first detected in lymphoid tissues draining the oral and intestinal mucosa: the retropharyngeal lymph nodes (by as early as 6 weeks), tonsils, Peyer’s patches and ileocaecal lymph nodes (Sigurdson et al., 1999). By 3 months, it is widely spread in lymphoid tissues and can first be detected in the brain. By 9 months, it can be detected in peripheral neural tissue associated with the gastrointestinal tract (myenteric and submucosal plexi) and the vagus nerve; by 16 months, throughout the brain and spinal cord (Fox et al., 2006). In both Odocoileus hemionus and Cervus elaphus nelsoni, the parasympathetic region of the vagus nerve in the medulla appears to be the earliest site of PrPCWD accumulation in the brain (Miller and Williams, 2002; Peters et al., 2000; Sigurdson et al., 2001; Spraker et al., 2002b, 2004; Williams and Miller, 2000). Progression of infection in Cervus elaphus nelsoni differs somewhat from that in Odocoileus spp. deer, with involvement of lymphoid tissue, even tonsil and retropharyngeal lymph node, being more variable (Spraker et al., 2004; Williams et al., 2002). Clinical signs typically last a few weeks to months and may be shorter in free-ranging cervids (which must forage, find water and avoid predators) (Kahn et al., 2004; Miller, 1994; Miller and Williams, 2004; Wild et al., 2002; Williams et al., 2002). The main clinical signs in Odocoileus spp. deer and in Cervus elaphus nelsoni are loss of body condition and changes in behaviour, more subtle in the elk (Williams and Miller, 2002). In captive animals, behavioural signs may include increased or decreased interactions with handlers, repetitive behaviours (e.g. walking in a set pattern), lowered head and ear carriage, and periods of somnolence or depression (rousable). They have reduced appetite and loss of body condition. Later, polyuria and polydipsia (and associated reduced urine specific gravity), increased salivation and associated drooling of saliva, and neurological signs, e.g. incoordination, posterior ataxia, wide-based stance and fine head tremors, may occur. Occasionally, dilatation of the oesophagus, hyperexcitability and syncope are noted. Aspiration pneumonia is probably due to difficulties in swallowing; occasionally there is apparent sudden death after handling (Bourne, 2004; Williams, 2005; Williams and Young, 1992). There are no gross lesions of the central nervous system (Williams and Young, 1992). Cervids dying late in the course of the disease may be emaciated. Sometimes aspiration pneumonia is noted. Abnormal ruminal contents (watery or gassy, sometimes with excessive sand
or gravel) are more common in Odocoileus spp. deer than in elk (Williams and Young, 1980, 1992; Williams et al., 1992; Spraker et al., 1997; Williams and Miller, 2002). In a few mule deer, the oesophagus is dilated and fluidfilled (Williams and Young, 1992). Marked adrenal gland enlargement has been seen in some mule deer (Spraker et al., 1997). Histological lesions are found within the CNS, with the distribution of lesions being similar in elk, mule deer and white-tailed deer (Williams and Miller, 2002). Lesions are characterised by bilaterally symmetrical spongiform transformation (microcavitation) of the grey matter with formation of neuronal intracytoplasmic vacuoles; loss and degeneration of neurons, and hypertrophy and hyperplasia of astrocytes are present but not prominent, and there is no inflammatory response (Spraker et al., 1997, 2002c; Williams, 2005; Williams and Young, 1980, 1982, 1993). The first region of the brain to develop lesions of spongiform encephalopathy is the dorsal motor nucleus of the vagus (DMNV) in the dorsal portion of the medulla oblongata at the obex (Spraker et al., 2002b). All cervids with clinical CWD have lesions in the olfactory tubercle and cortex, hypothalamus and DMNV; variable spongiform change may be present in other areas, particularly the thalamus and cerebellum. Lesions in the cerebral cortex, hippocampus and basal ganglia are generally mild (Williams and Miller, 2002; Williams and Young, 1993). Lesions in some thalamic nuclei are more severe in elk than in deer (Williams and Young, 1993). White matter lesions are rare in deer; mild lesions may be present in elk, usually rarefaction in the cerebrum and cerebellum, particularly in cerebellar subcortical regions (Williams and Young, 1993). Amyloid plaques can be seen with haematoxylin and eosin staining as slightly pale eosinophilic fibrillar areas of neuropil, and may be surrounded by vacuoles to form florid plaques; they are most common in white-tailed deer, less in mule deer and least common in elk. Silver or Congo red stains show the plaques more vividly, as does IHC (Williams, 2005; Williams and Young, 1993). Ultrastructural findings include extensive membrane-bound vacuolation in neurons and neuronal processes. Dystrophic neurites may contain electron-dense bodies as well as degenerating mitochondria and pleomorphic membrane-bound structures (Guiroy et al., 1993a, 1994; Williams, 2005). IHC staining for PrP using any of a range of polyclonal and monoclonal antibodies clearly reveals PrPCWD plaques in the brains of affected animals; it may reveal granularity and amorphous clumps on neuronal membranes, perivascular aggregates of PrPCWD and large apparently extracellular accumulations of PrPCWD (Williams and Miller, 2002). 6. Diagnosis Clinical signs (loss of body condition and behavioural changes) are not pathognomonic; a wide variety of diseases causing emaciation and/or nervous signs must be included as differential diagnoses (Williams and Miller, 2002; Williams et al., 2001). Clinical signs can vary, and affected individuals may not be detected (Miller and Williams, 2004).
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Histopathological CNS lesions (particularly in the DMNV) are diagnostic, but are only present in individuals with clinical signs (lesions and clinical signs develop at the same time); additionally, autolysis may make histopathological examination problematic (Kahn et al., 2004; Spraker et al., 2002a; Williams, 2005). IHC using monoclonal or polyclonal antibodies can detect PrPCWD in lymphoid and CNS tissues (including in the absence of SE lesions); it is highly sensitive and specific, also allows precise anatomical considerations of PrPCWD deposition, can be used on tissue which is somewhat autolysed, and is considered the “gold standard” for diagnosis (Bourne, 2004; Hall, 2004; Kahn et al., 2004; Peters et al., 2000; Sigurdson et al., 1999; Spraker et al., 2002a; Williams, 2005). IHC of the DMNV at the obex is sensitive and specific for CWD diagnosis in Odocoileus spp. deer and in elk. In Odocoilus spp. deer, testing of the retropharyngeal lymph nodes is recommended, due to early deposition of PrPCWD at this site. In elk, 10–15% may have detectable PrPCWD in the brain but not in lymphoid tissues; this situation occurs only very rarely (<1%) in Odocoilus spp. deer (Williams, 2005). Tonsils and retropharyngeal lymph nodes are easily collected from the heads of hunted deer (Miller and Williams, 2004; Williams et al., 2002) and tonsillar biopsies can be collected in live, anaesthetised Odocoileus spp. deer; the requirement for anaesthesia makes this unsuitable for large-scale surveillance (Wild et al., 2002; Wolfe et al., 2002). Sampling of conjunctival lymphoid tissue is not useful because this tissue is sparse in cervids (Kahn et al., 2004; Williams and Miller, 2002). Rapid tests for TSEs (ELISA and Western blot) are being used increasingly as screening tests. They use unfixed tissues, which reduces the time from sampling to results. As with IHC the sensitivity of such tests depends on the tissue(s) being tested. In general, these tests have a high sensitivity but a lower specificity, leading to some falsepositive results (Bollinger et al., 2004). Standardisation of both tissue collection and tissue trimming techniques are essential for accuracy of high-throughput tests (Hibler et al., 2003; O’Rourke et al., 2003). In a few cases, IHC may be more sensitive due to detection in specimens with only occasional single-cell staining (O’Rourke et al., 2003). Where rapid tests are used for surveillance, IHC is recommended to confirm positives and reject false positives (Bollinger et al., 2004). Several ELISA kits from different companies have been approved by USDA Center for Veterinary Biologicals for use as CWD screening tests; one strip test has also been approved (Williams, 2005). Western blot assays may be used for detection of PrPCWD in the brains of both clinically affected and (presumably) preclinical cervids (Laplanche et al., 1999; Spraker et al., 2004). A dot–blot assay using monoclonal antibody F99/97.6.1 on tonsillar tissue had high sensitivity and allowed quantification (O’Rourke et al., 2003). A conformation-dependent immunoassay (CDI) has been developed (Safar et al., 2002). 7. Epidemiology and transmission It is thought that CWD is spread mainly by lateral transmission, including between cervid species; it is possible that maternal transmission also occurs (Kahn et al., 2004;
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Miller et al., 2000; Williams and Miller, 2002; Williams and Young, 1992). Transmission probably occurs by both direct and indirect routes (Williams and Miller, 2002) and oral transmission is considered likely to be the main route of natural transmission (Sigurdson et al., 1999; Williams, 2005). In cervid PrP transgenic mice, minor oral abrasions facilitated oral CWD transmission; such lesions are common in cervids (Denkers et al., 2011). Mice exposed to aerosolised scrapie prions efficiently developed scrapie, indicating a previously unappreciated risk route for prion exposure (Haybaeck et al., 2011). It appears likely that shedding of the CWD agent is progressive during the course of the disease (Miller et al., 2000; Williams and Miller, 2002) and infectious prions have been confirmed in faeces (Tamgüney et al., 2009), urine (Haley et al., 2009), saliva (Mathiason et al., 2006) and velvet (Angers et al., 2009). After death, environmental contamination with PrPCWD could occur by scavenging (including by invertebrates) and decomposition (Williams and Miller, 2003). PrPCWD has been detected in the brain, spinal cord, eyes, peripheral nerves, various lymphoid tissues including the spleen, thymus, pituitary, adrenals and pancreas (Sigurdson et al., 2001; Spraker et al., 2002c), cardiac muscle myocytes (Jewell et al., 2006), skeletal muscles (Angers et al., 2006; Daus et al., 2011), antler velvet (Angers et al., 2009), blood, saliva (Mathiason et al., 2006), salivary glands, urinary bladder, nasal epithelium and distal intestine (Haley et al., 2011). Pastures contaminated by infected faeces or carcasses were proven as sources of infection for cervids (Miller et al., 2004). Carcasses remaining naturally on native ranges will be available to act as a source of infection for free-ranging animals (Miller et al., 2004). PrPres attaches to and remains infectious in soils for long periods (years), which may increase the persistence and transmission of CWD (Smith et al., 2011). Transmission probably occurs between sympatric populations of the different cervid species (Williams et al., 2002). Spread of CWD between geographical areas is probably due to natural movement of live animals and translocation of both captive and free-ranging cervids (Bollinger et al., 2004; Gross and Miller, 2001; Miller et al., 2000; Williams et al., 2002). The social behaviour of deer may concentrate populations and increase the chance of direct or indirect lateral transmission; this may be enhanced artificially when deer congregate at feeding or baiting sites (Bartelt et al., 2003; Miller and Williams, 2003; Williams et al., 2002). Deer feeding on frozen piles of food use heat from their nose and mouth to thaw the food, and leave saliva and nasal droppings behind (Bartelt et al., 2003). Transmission may occur faster in high-density white-tailed deer populations in the eastern USA (Joly et al., 2003). 8. Factors affecting susceptibility of cervids Infection has been confirmed in deer and elk of all ages from under a year to elderly animals (O’Rourke et al., 2004; Spraker et al., 1997) and in males, females and castrates (Williams et al., 1990). Higher rates of infection sometimes noted in adult males may be associated with behavioural factors (e.g. larger home ranges, close contact in bachelor
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groups or males contacting large numbers of females for mating) (Langenberg, 2004; T.R. Kreeger, pers. comm.; Wisconsin Department of Natural Resources, 2004). PRNP gene polymorphisms in elk, mule deer and whitetailed deer may affect the rate of infection or progression of CWD (Robinson et al., 2012). Rocky Mountain elk have a polymorphism (Met/Leu) at cervid codon 132 of the PRNP gene (O’Rourke et al., 1999). Based on oral inoculation, elk homozygous for M (132MM) have the shortest incubation time while 132LL individuals appear to have reduced susceptibility or prolonged incubation time (Hamir et al., 2006a), Work with transgenic mice indicates that propagation of PrPres is severely restricted by PrP L132, but latent infection may develop (Green et al., 2008). In mule deer, the incubation time is shorter for deer homozygous for serine at codon 225 (225SS) compared to individuals heterozygous (225 SF) (F = phenylalanine) at this codon, with marked delay in accumulation of CWD, and development of SE lesions, in the CNS in 225 SF, although accumulation of PrPres in the lymphoid tissues was almost identical (Fox et al., 2006; Robinson et al., 2012). In Colorado and Wyomine, 225SS deer were much more likely to be positive than 225SF deer, while in Canada, allele 225F was not present but deer with 20G were more than twice as likely to be infected than those with 20D (Robinson et al., 2012). In white-tailed deer, at least three polymorphisms may affect development of CWD. 96S deer have a longer incubation period than 96G deer, and 96 GS deer have lower PrPres accumulation than 96GG deer; in both free-ranging and captive deer, prevalence is lower in 96S (serine) than in 96G (glycine) individuals. Additionally (based on more limited data), 95H (histadine) deer have a longer incubation period than 95Q (glutamine) deer and reduced rates of infection, while in one Nebraska herd 116A (alanine) deer were more than twice as likely to be CWD-positive than 116G (glycine) deer (Robinson et al., 2012). 9. Concluding remarks While much has been learned about CWD in the past 25 years, it is clear that there is still much to be learned about this unique TSE. Conflict of interest statement None declared. References APHIS, 2012. http://www.aphis.usda.gov/animal health/animal diseases/ cwd/downloads/distribution cwd.pdf (accessed 10.12.2012). Angers, R.C., Browning, S.R., Seward, T.S., Sigurdson, C.J., Miller, M.W., Hoover, E.A., Telling, G.C., 2006. Prions in skeletal muscles of deer with chronic wasting disease. Science 311, 1117. Angers, R.C., Seward, T.S., Napier, D., Green, M., Hoover, E., Spraker, T., O’Rourke, K., Balachandran, A., Telling, G.C., 2009. Chronic wasting disease prions in elk antler velvet. Emerg. Infect. Dis. 15, 696–703. Baeten, L.A., Powers, B.E., Jewell, J.E., Spraker, T.R., Miller, M.W., 2007. A natural case of chronic wasting disease in a free-ranging moose (Alces alces shirasi). J. Wildl. Dis. 43, 309–334. Balachandran, A., Harrington, N.P., Algire, J., Soutyrine, A., Spraker, T.R., Jeffrey, M., González, L., O’Rourke, K.I., 2010. Experimental oral transmission of chronic wasting disease to red deer (Cervus elaphus elaphus): early detection and late stage distribution of proteaseresistant prion protein. Can. Vet. J. 51, 169–178.
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Tamgüney, G., Miller, M.W., Wolfe, L.L., Sirochman, T.M., Glidden, D.V., Palmer, C., Lemus, A., DeArmond, S.J., Prusiner, S.B., 2009. Asymptomatic deer excrete infectious prions in faeces. Nature 461, 529–532. Wild, M.A., Spraker, T.R., Sigurdson, C.J., O’Rourke, K.I., Miller, M.W., 2002. Preclinical diagnosis of chronic wasting disease in captive mule deer (Odocoileus hemionus) and white-tailed deer (Odocoileus virginianus) using tonsillar biopsy. J. Gen. Virol. 83, 2629– 2634. Williams, E.S., 2005. Chronic wasting disease. Vet. Pathol. 42, 530–549. Williams, E.S., Kirkwood, J.K., Miller, M.W., 2001. Transmissible spongiform encephalopathies. In: Williams, E.S., Barker, I.K. (Eds.), Infectious Diseases of Wild Mammals. , third ed. Iowa State University Press, Ames, pp. 292–310. Williams, E.S., Miller, M.W., 2000. Pathogenesis of chronic wasting disease in orally exposed mule deer (Odocoileus hemionus): preliminary results [Abstract]. In: 49th Annual Wildlife Disease Association Conference, Programme and Abstracts of Papers Presented. 4–8 June 2000, Grand Teton National Park, Wyoming, USA, p. 29. Williams, E.S., Miller, M.W., 2002. Chronic wasting disease in deer and elk in North America. Rev. Sci. Tech. 21, 305–316. Williams, E.S., Miller, M.W., 2003. Transmissible spongiform encephalopathies in non-domestic animals: origin, transmission and risk factors. Revue Sci. Tech. 22, 145–156. Williams, E.S., Miller, M.W., Kreeger, T.J., Kahn, R.H., Thorne, E.T., 2002. Chronic wasting disease of deer and elk: a review with recommendations for management. J. Wildl. Man. 66, 551–563. Williams, E.S., Young, S., 1980. Chronic wasting disease of captive mule deer: a spongiform encephalopathy. J. Wildl. Dis. 16, 89–98. Williams, E.S., Young, S., 1982. Spongiform encephalopathy of Rocky Mountain elk. J. Wildl. Dis. 18, 465–471. Williams, E.S., Young, S., 1992. Spongiform encephalopathies in Cervidae. Rev. Sci. Tech. 11, 551–567. Williams, E.S., Young, S., 1993. Neuropathology of chronic wasting disease of mule deer (Odocoileus hemionus) and elk (Cervus elaphus nelsoni). Vet. Pathol. 30, 36–45. Williams, E.S. Young, S., Marsh, R.F., 1982. Preliminary evidence of transmissibility of chronic wasting disease of mule deer [Abstract]. 31st Wildlife Disease Association Conference. Programme and Abstracts of Papers Presented. August 16-20, 1982, Madison, Wisconsin USA. 31, 22. Williams, E.S., Miller, M.W., Young, S., Thorne, E.T., 1992. Chronic wasting disease: a spongiform encephalopathy of mule deer (Odocoileus hemionus) and Rocky Mountain elk (Cervus elaphus nelsoni) in Colorado and Wyoming, USA. New and Emerging Infectious Diseases: 12th International Symposium Proceedings, World Association of Veterinary Microbiologists, Immunologists and Specialists in Infectious Diseases (WAVMI), September 8-1, 1992. 207-212. Williams, E.S., Thorne, E.T., Miller, M. Neil, P., 1990. Epizootiology of cervid spongiform encephalopathy (chronic wasting disease), 6th International Conference of Wildlife Diseases, Berlin, Germany. Wisconsin Department of Natural Resources, 2004. CWD in Wisconsin, http://www.dnr.state.wi.us/org/land/wildlife/Whealth/issues/Cwd/ CWD Display 2003.pdf (accessed 06.01.2004). Wolfe, L.L., Conner, M.M., Baker, T.H., Dreitz, V.J., Burham, K.P., Williams, E.S., Miller, M.W., 2002. Evaluation of antemortem sampling to estimate chronic wasting disease prevalence in free-ranging mule deer. J. Wildl. Man. 66, 564–573.
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Chronic wasting disease of cervids夽 D.C. Bourne ∗ Wildlife Information Network, Twycross Zoo – East Midland Zoological Society, Atherstone, Warwickshire, CV9 3PX, UK
a r t i c l e
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Article history: Available online 18 March 2015 Keywords: Chronic wasting disease (CWD) Transmissible spongiform encephalopathy Cervids
a b s t r a c t Chronic wasting disease (CWD) of cervids is the only known transmissible spongiform encephalopathy (TSE) found in non-domestic, free-ranging animals. To date, it is found in wild cervids only in North America, and natural infection has been detected in only four species (Cervus elaphus nelsoni, Odocoileus hemionus, Odocoileus virginianus and recently Alces alces) although there are concerns that it could spread to other species, particularly Rangifer tarandus. The infectious PrPCWD spreads throughout the body, particularly in lymphoid tissue, although lesions (typical of TSEs) are found only in the brain and are associated with development of clinical disease (e.g. wasting, behavioural changes). Transmission is lateral and probably oral. Infectious prions are shed in faeces, urine and saliva, and are present in various body tissues, all of which may contribute to ante- or post-mortem environmental contamination, increasing transmission. Infectious prions persist in soils. There is presently no evidence of spread to other sympatric wildlife, domestic livestock or humans. Intracerebral inoculation enables transmission to various species of ungulates, rodents, carnivores and even primates, but oral transmission to non-cervids has mainly been unsuccessful. © 2015 Elsevier B.V. All rights reserved.
1. Introduction
2. Emergence, aetiology and possible origins
Chronic wasting disease (CWD) is one of the group of diseases generally known as the transmissible spongiform encephalopathies (TSE) or prion diseases, and the only known prion disease of free-living non-domestic animals. As with other prion diseases, CWD is associated with a misfolded isoform (PrPCWD or PrPres ) of a normal cellular protein (PrPC ), which accumulates in the CNS, with resultant neurodegenerative changes; this abnormal PrP is transmissible (Bourne, 2004).
CWD was first detected as a clinical wasting syndrome in mule deer (Odocoileus hemionus hemionus) at a research facility in Fort Collins, CO, USA, in 1967; it was recognised as a spongiform encephalopathy (SE) in 1978 (Williams and Young, 1980) and was shown to be transmissible by intracerebral (IC) inoculation into ferrets and mule deer fawns (Williams et al., 1982). The first diagnosis in captive Rocky Mountain elk (Cervus elaphus nelsoni) occurred in 1978 (Williams and Young, 1982); it was first diagnosed in wild cervids in 1981 (Spraker et al., 1997) and in farmed cervids in Saskatchewan, Canada (Sullivan, 1996). The origin of CWD is uncertain; the favoured theory is that it derived from scrapie (Williams, 2005); scrapie inoculation IC into elk produced typical CWD lesions (Hamir et al., 2004). It is also possible that CWD arose de novo in deer and became transmissible (Bourne, 2004; Williams et al., 2002). Alternatively, it originated in another species,
夽 This paper is part of a set of articles on the theme of ‘Diseases of Cervids’, guest-edited by C. Billinis, who gratefully acknowledges the contribution of the authors and the editorial staff. ∗ Current address: MA Healthcare Ltd., St Jude’s Church, Dulwich Road, London, SE24 0PB, UK. E-mail addresses: [email protected], [email protected] http://dx.doi.org/10.1016/j.smallrumres.2015.03.008 0921-4488/© 2015 Elsevier B.V. All rights reserved.
D.C. Bourne / Small Ruminant Research 128 (2015) 72–78
as yet unknown (Williams, 2005). Based on strain typing by inoculation into genotypically characterised mice, the CWD agent is different from BSE agent, TME agent and tested strains of CJD agent and scrapie agent (Bruce et al., 1997, 2000; Laplanche et al., 1999). It is not yet certain whether there is a single or multiple strains of CWD. Epidemiological data and the marked similarity of the lesions strongly suggest that the same CWD agent is responsible for the disease in captive and free-ranging deer and elk (Williams and Miller, 2002). Studies in transgenic mice expressing cervid PrP indicated the same prion strain in analysed elk and mule deer (Browning et al., 2004), while ferret-passaged isolates from two sources showed differences in clinical presentation, survival period, lesion distribution, glycoform profile and resistance to proteolysis, suggesting different strains (Perrott et al., 2012). It is difficult to draw conclusions from such experiments because interspecies transmission can alter the characteristics of TSE agents (Sigurdson, 2008). 3. Geographical distribution CWD was first found in research facilities in Colorado and Wyoming (Williams and Young, 1992; Williams and Miller, 2002). It has been confirmed in free-living cervids in 17 states in the USA (Colorado, Wyoming, Utah, Kansas, Nebraska, New Mexico, Texas, North Dakota, South Dakota, Wisconsin, Illinois, Maryland, Missouri, New York, Minnesota, Virginia, West Virginia) and in Canada (Alberta and Saskatchewan). In farmed cervids, it has been detected in Colorado, Nebraska, Montana, South Dakota, Kansas, Oklahoma, Minnesota, Iowa, Wisconsin, Missouri, Michigan, Pennsylvania and New York, and in Canada in Alberta and Saskatchewan (APHIS, 2012; NWHC, 2012). Occurrence and prevalence of CWD varies between states and between regions within a state; even within affected regions there are clusters of affected animals (Joly et al., 2006). There have been very few cases in cervids in North American zoos, with no evidence of the disease remaining in zoos in recent years (Bourne, 2004; Dubé et al., 2006; Williams and Young, 1992). Outside North America, CWD has been detected only in farmed cervids in Korea, associated with elk imported from Canada (Kim et al., 2005; ProMED-Mail, 2011; Sohn et al., 2002). 4. Host range The known natural hosts of chronic wasting disease are mule deer, white-tailed deer, Rocky Mountain elk and moose (Alces alces shirasi) (Baeten et al., 2007; Williams et al., 2002). European red deer (Cervus elaphus elaphus) are susceptible on oral inoculation (Balachandran et al., 2010). Natural infection has not been reported in Rangifer tarandus (reindeer or caribou) (Lapointe et al., 2002; Sigurdarson, 2004). Oral transmission was successful in some reindeer; it is possible that some reindeer PRNP polymorphisms are protective against CWD (Mitchell et al., 2012). In Korea, CWD has been confirmed in Sika deer (Cervus nippon) and in red deer × sika deer (ProMED-Mail, 2011). Fallow deer (Dama dama) have been infected by IC inoculation from elk or white-tailed deer, with development of SE
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lesions (Hamir et al., 2010), but not by exposure to infected mule deer and a CWD-contaminated environment, suggesting lower susceptibility or delayed progression in this species (Rhyan et al., 2011). In Europe, surveillance of roe deer (Capreolus capreolus), red deer, fallow deer, muntjac (Muntiacus reevesi) and reindeer has not yet detected any cases of TSE (A.M. Barlow, pers. comm., 2004; Schettler et al., 2004; Schwaiger et al., 2004; Sigurdarson, 2004). It has been possible to infect domestic cattle, goats and sheep by IC inoculation, although not in all individuals and with several-year incubation periods in cattle and goats (Hamir et al., 2006b, 2011; Williams and Young, 1992). Cattle failed to develop SE although PrPres was detected in the CNS (Hamir et al., 2011). In sheep, only two of eight became infected: one (ARQ/VRQ at codons 136, 154, and 171) developing clinical signs, SE lesions and was PrPres positive at 35 months post inoculation (mpi) and one (ARQ/ARQ) SE and PrPres positive but no clinical signs at 72 months pi; two ARQ/ARQ and four ARQ/ARR were negative for PrPres at 36–72 mpi (Hamir et al., 2006b). Oral transmission into cattle failed (to nine years post infection) (Hamir et al., 2011). In carnivores, initial IC inoculation of mule deer CWD material into ferrets gave an incubation period of over a year to clinical signs (Williams et al., 1982). Serial passage reduced the incubation period to as low as 5 months (Bartz et al., 1998). Inoculation into American mink (Mustela vison) has also been successful, but IC inoculation of Common raccoon (Procyon lotor) kits with CWD failed (Hamir et al., 2007). In Wisconsin, brains of 812 mammalian scavengers were tested; the IDEXX HerdChek ELISA gave positive results in 1/11 Mustela vison, 1/259 Procyon lotor and 2/202 Didelphis virginiana but none were confirmed by Western blotting. The rest were all negative by ELISA (Jennelle et al., 2009). In rodents, IC inoculation into laboratory mice produced infection in only “a very few mice”, after >500 days incubation; serial passage in mice produced clinical disease (Bruce et al., 2000). Golden (Syrian) hamsters (Mesocricetus auratus) were not infected by IC inoculation of material from CWD mule deer (Williams and Young, 1992; Williams et al., 1992), but were infected following passage in ferrets, indicating that the host range of CWD can be altered by passage through other species (Bartz et al., 1998). To date there are no known cases of human prion disease attributable to CWD transmitted to humans. Epidemiological investigations have failed to show links between prion disease in hunters or young people in North America and CWD and there is no evidence of increased CJD in Colorado or Wyoming (Belay et al., 2001, 2004; CDC, 2003). In vitro, CWD-associated PrPres was shown to convert human PrPsen only at very low efficiency (Raymond et al., 2000). IC inoculation of CWD-infected cervid material into transgenic mice expressing human PrP failed (Kong et al., 2005; Sandberg et al., 2010). Squirrel monkeys (Saimiri sciureus) inoculated IC with one CWD strain became ill at 31–34 mpi and were PrPres positive (Marsh et al., 2005). Further experiments produced clinical disease in 33–53 months with 7/8 isolates IC and infection of 3/15 monkeys orally. In contrast, 15 cynomolgus macaques inoculated IC and orally remained well at 70 mpi and no PrPres was detected by IHC or Western blot in the
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brain, spinal cord, spleen and lymph nodes of one individual euthanased at 48 months (Race et al., 2009). 5. CWD in cervids The natural incubation period for CWD in naturally exposed cervids is not known and probably varies depending on frequency of exposure and the dose received (O’Rourke et al., 2004; Spraker, 2003; Spraker et al., 2004) In early studies, clinical signs were first noted in deer at 18 months (Williams and Young, 1992); the youngest recorded affected free-ranging elk was 21 months old (Spraker et al., 1997). Cervids less than 12 months old have been confirmed CWD-positive by IHC (Kahn et al., 2004). In orally inoculated mule deer fawns, PrPCWD was first detected in lymphoid tissues draining the oral and intestinal mucosa: the retropharyngeal lymph nodes (by as early as 6 weeks), tonsils, Peyer’s patches and ileocaecal lymph nodes (Sigurdson et al., 1999). By 3 months, it is widely spread in lymphoid tissues and can first be detected in the brain. By 9 months, it can be detected in peripheral neural tissue associated with the gastrointestinal tract (myenteric and submucosal plexi) and the vagus nerve; by 16 months, throughout the brain and spinal cord (Fox et al., 2006). In both Odocoileus hemionus and Cervus elaphus nelsoni, the parasympathetic region of the vagus nerve in the medulla appears to be the earliest site of PrPCWD accumulation in the brain (Miller and Williams, 2002; Peters et al., 2000; Sigurdson et al., 2001; Spraker et al., 2002b, 2004; Williams and Miller, 2000). Progression of infection in Cervus elaphus nelsoni differs somewhat from that in Odocoileus spp. deer, with involvement of lymphoid tissue, even tonsil and retropharyngeal lymph node, being more variable (Spraker et al., 2004; Williams et al., 2002). Clinical signs typically last a few weeks to months and may be shorter in free-ranging cervids (which must forage, find water and avoid predators) (Kahn et al., 2004; Miller, 1994; Miller and Williams, 2004; Wild et al., 2002; Williams et al., 2002). The main clinical signs in Odocoileus spp. deer and in Cervus elaphus nelsoni are loss of body condition and changes in behaviour, more subtle in the elk (Williams and Miller, 2002). In captive animals, behavioural signs may include increased or decreased interactions with handlers, repetitive behaviours (e.g. walking in a set pattern), lowered head and ear carriage, and periods of somnolence or depression (rousable). They have reduced appetite and loss of body condition. Later, polyuria and polydipsia (and associated reduced urine specific gravity), increased salivation and associated drooling of saliva, and neurological signs, e.g. incoordination, posterior ataxia, wide-based stance and fine head tremors, may occur. Occasionally, dilatation of the oesophagus, hyperexcitability and syncope are noted. Aspiration pneumonia is probably due to difficulties in swallowing; occasionally there is apparent sudden death after handling (Bourne, 2004; Williams, 2005; Williams and Young, 1992). There are no gross lesions of the central nervous system (Williams and Young, 1992). Cervids dying late in the course of the disease may be emaciated. Sometimes aspiration pneumonia is noted. Abnormal ruminal contents (watery or gassy, sometimes with excessive sand
or gravel) are more common in Odocoileus spp. deer than in elk (Williams and Young, 1980, 1992; Williams et al., 1992; Spraker et al., 1997; Williams and Miller, 2002). In a few mule deer, the oesophagus is dilated and fluidfilled (Williams and Young, 1992). Marked adrenal gland enlargement has been seen in some mule deer (Spraker et al., 1997). Histological lesions are found within the CNS, with the distribution of lesions being similar in elk, mule deer and white-tailed deer (Williams and Miller, 2002). Lesions are characterised by bilaterally symmetrical spongiform transformation (microcavitation) of the grey matter with formation of neuronal intracytoplasmic vacuoles; loss and degeneration of neurons, and hypertrophy and hyperplasia of astrocytes are present but not prominent, and there is no inflammatory response (Spraker et al., 1997, 2002c; Williams, 2005; Williams and Young, 1980, 1982, 1993). The first region of the brain to develop lesions of spongiform encephalopathy is the dorsal motor nucleus of the vagus (DMNV) in the dorsal portion of the medulla oblongata at the obex (Spraker et al., 2002b). All cervids with clinical CWD have lesions in the olfactory tubercle and cortex, hypothalamus and DMNV; variable spongiform change may be present in other areas, particularly the thalamus and cerebellum. Lesions in the cerebral cortex, hippocampus and basal ganglia are generally mild (Williams and Miller, 2002; Williams and Young, 1993). Lesions in some thalamic nuclei are more severe in elk than in deer (Williams and Young, 1993). White matter lesions are rare in deer; mild lesions may be present in elk, usually rarefaction in the cerebrum and cerebellum, particularly in cerebellar subcortical regions (Williams and Young, 1993). Amyloid plaques can be seen with haematoxylin and eosin staining as slightly pale eosinophilic fibrillar areas of neuropil, and may be surrounded by vacuoles to form florid plaques; they are most common in white-tailed deer, less in mule deer and least common in elk. Silver or Congo red stains show the plaques more vividly, as does IHC (Williams, 2005; Williams and Young, 1993). Ultrastructural findings include extensive membrane-bound vacuolation in neurons and neuronal processes. Dystrophic neurites may contain electron-dense bodies as well as degenerating mitochondria and pleomorphic membrane-bound structures (Guiroy et al., 1993a, 1994; Williams, 2005). IHC staining for PrP using any of a range of polyclonal and monoclonal antibodies clearly reveals PrPCWD plaques in the brains of affected animals; it may reveal granularity and amorphous clumps on neuronal membranes, perivascular aggregates of PrPCWD and large apparently extracellular accumulations of PrPCWD (Williams and Miller, 2002). 6. Diagnosis Clinical signs (loss of body condition and behavioural changes) are not pathognomonic; a wide variety of diseases causing emaciation and/or nervous signs must be included as differential diagnoses (Williams and Miller, 2002; Williams et al., 2001). Clinical signs can vary, and affected individuals may not be detected (Miller and Williams, 2004).
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Histopathological CNS lesions (particularly in the DMNV) are diagnostic, but are only present in individuals with clinical signs (lesions and clinical signs develop at the same time); additionally, autolysis may make histopathological examination problematic (Kahn et al., 2004; Spraker et al., 2002a; Williams, 2005). IHC using monoclonal or polyclonal antibodies can detect PrPCWD in lymphoid and CNS tissues (including in the absence of SE lesions); it is highly sensitive and specific, also allows precise anatomical considerations of PrPCWD deposition, can be used on tissue which is somewhat autolysed, and is considered the “gold standard” for diagnosis (Bourne, 2004; Hall, 2004; Kahn et al., 2004; Peters et al., 2000; Sigurdson et al., 1999; Spraker et al., 2002a; Williams, 2005). IHC of the DMNV at the obex is sensitive and specific for CWD diagnosis in Odocoileus spp. deer and in elk. In Odocoilus spp. deer, testing of the retropharyngeal lymph nodes is recommended, due to early deposition of PrPCWD at this site. In elk, 10–15% may have detectable PrPCWD in the brain but not in lymphoid tissues; this situation occurs only very rarely (<1%) in Odocoilus spp. deer (Williams, 2005). Tonsils and retropharyngeal lymph nodes are easily collected from the heads of hunted deer (Miller and Williams, 2004; Williams et al., 2002) and tonsillar biopsies can be collected in live, anaesthetised Odocoileus spp. deer; the requirement for anaesthesia makes this unsuitable for large-scale surveillance (Wild et al., 2002; Wolfe et al., 2002). Sampling of conjunctival lymphoid tissue is not useful because this tissue is sparse in cervids (Kahn et al., 2004; Williams and Miller, 2002). Rapid tests for TSEs (ELISA and Western blot) are being used increasingly as screening tests. They use unfixed tissues, which reduces the time from sampling to results. As with IHC the sensitivity of such tests depends on the tissue(s) being tested. In general, these tests have a high sensitivity but a lower specificity, leading to some falsepositive results (Bollinger et al., 2004). Standardisation of both tissue collection and tissue trimming techniques are essential for accuracy of high-throughput tests (Hibler et al., 2003; O’Rourke et al., 2003). In a few cases, IHC may be more sensitive due to detection in specimens with only occasional single-cell staining (O’Rourke et al., 2003). Where rapid tests are used for surveillance, IHC is recommended to confirm positives and reject false positives (Bollinger et al., 2004). Several ELISA kits from different companies have been approved by USDA Center for Veterinary Biologicals for use as CWD screening tests; one strip test has also been approved (Williams, 2005). Western blot assays may be used for detection of PrPCWD in the brains of both clinically affected and (presumably) preclinical cervids (Laplanche et al., 1999; Spraker et al., 2004). A dot–blot assay using monoclonal antibody F99/97.6.1 on tonsillar tissue had high sensitivity and allowed quantification (O’Rourke et al., 2003). A conformation-dependent immunoassay (CDI) has been developed (Safar et al., 2002). 7. Epidemiology and transmission It is thought that CWD is spread mainly by lateral transmission, including between cervid species; it is possible that maternal transmission also occurs (Kahn et al., 2004;
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Miller et al., 2000; Williams and Miller, 2002; Williams and Young, 1992). Transmission probably occurs by both direct and indirect routes (Williams and Miller, 2002) and oral transmission is considered likely to be the main route of natural transmission (Sigurdson et al., 1999; Williams, 2005). In cervid PrP transgenic mice, minor oral abrasions facilitated oral CWD transmission; such lesions are common in cervids (Denkers et al., 2011). Mice exposed to aerosolised scrapie prions efficiently developed scrapie, indicating a previously unappreciated risk route for prion exposure (Haybaeck et al., 2011). It appears likely that shedding of the CWD agent is progressive during the course of the disease (Miller et al., 2000; Williams and Miller, 2002) and infectious prions have been confirmed in faeces (Tamgüney et al., 2009), urine (Haley et al., 2009), saliva (Mathiason et al., 2006) and velvet (Angers et al., 2009). After death, environmental contamination with PrPCWD could occur by scavenging (including by invertebrates) and decomposition (Williams and Miller, 2003). PrPCWD has been detected in the brain, spinal cord, eyes, peripheral nerves, various lymphoid tissues including the spleen, thymus, pituitary, adrenals and pancreas (Sigurdson et al., 2001; Spraker et al., 2002c), cardiac muscle myocytes (Jewell et al., 2006), skeletal muscles (Angers et al., 2006; Daus et al., 2011), antler velvet (Angers et al., 2009), blood, saliva (Mathiason et al., 2006), salivary glands, urinary bladder, nasal epithelium and distal intestine (Haley et al., 2011). Pastures contaminated by infected faeces or carcasses were proven as sources of infection for cervids (Miller et al., 2004). Carcasses remaining naturally on native ranges will be available to act as a source of infection for free-ranging animals (Miller et al., 2004). PrPres attaches to and remains infectious in soils for long periods (years), which may increase the persistence and transmission of CWD (Smith et al., 2011). Transmission probably occurs between sympatric populations of the different cervid species (Williams et al., 2002). Spread of CWD between geographical areas is probably due to natural movement of live animals and translocation of both captive and free-ranging cervids (Bollinger et al., 2004; Gross and Miller, 2001; Miller et al., 2000; Williams et al., 2002). The social behaviour of deer may concentrate populations and increase the chance of direct or indirect lateral transmission; this may be enhanced artificially when deer congregate at feeding or baiting sites (Bartelt et al., 2003; Miller and Williams, 2003; Williams et al., 2002). Deer feeding on frozen piles of food use heat from their nose and mouth to thaw the food, and leave saliva and nasal droppings behind (Bartelt et al., 2003). Transmission may occur faster in high-density white-tailed deer populations in the eastern USA (Joly et al., 2003). 8. Factors affecting susceptibility of cervids Infection has been confirmed in deer and elk of all ages from under a year to elderly animals (O’Rourke et al., 2004; Spraker et al., 1997) and in males, females and castrates (Williams et al., 1990). Higher rates of infection sometimes noted in adult males may be associated with behavioural factors (e.g. larger home ranges, close contact in bachelor
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groups or males contacting large numbers of females for mating) (Langenberg, 2004; T.R. Kreeger, pers. comm.; Wisconsin Department of Natural Resources, 2004). PRNP gene polymorphisms in elk, mule deer and whitetailed deer may affect the rate of infection or progression of CWD (Robinson et al., 2012). Rocky Mountain elk have a polymorphism (Met/Leu) at cervid codon 132 of the PRNP gene (O’Rourke et al., 1999). Based on oral inoculation, elk homozygous for M (132MM) have the shortest incubation time while 132LL individuals appear to have reduced susceptibility or prolonged incubation time (Hamir et al., 2006a), Work with transgenic mice indicates that propagation of PrPres is severely restricted by PrP L132, but latent infection may develop (Green et al., 2008). In mule deer, the incubation time is shorter for deer homozygous for serine at codon 225 (225SS) compared to individuals heterozygous (225 SF) (F = phenylalanine) at this codon, with marked delay in accumulation of CWD, and development of SE lesions, in the CNS in 225 SF, although accumulation of PrPres in the lymphoid tissues was almost identical (Fox et al., 2006; Robinson et al., 2012). In Colorado and Wyomine, 225SS deer were much more likely to be positive than 225SF deer, while in Canada, allele 225F was not present but deer with 20G were more than twice as likely to be infected than those with 20D (Robinson et al., 2012). In white-tailed deer, at least three polymorphisms may affect development of CWD. 96S deer have a longer incubation period than 96G deer, and 96 GS deer have lower PrPres accumulation than 96GG deer; in both free-ranging and captive deer, prevalence is lower in 96S (serine) than in 96G (glycine) individuals. Additionally (based on more limited data), 95H (histadine) deer have a longer incubation period than 95Q (glutamine) deer and reduced rates of infection, while in one Nebraska herd 116A (alanine) deer were more than twice as likely to be CWD-positive than 116G (glycine) deer (Robinson et al., 2012). 9. Concluding remarks While much has been learned about CWD in the past 25 years, it is clear that there is still much to be learned about this unique TSE. Conflict of interest statement None declared. References APHIS, 2012. http://www.aphis.usda.gov/animal health/animal diseases/ cwd/downloads/distribution cwd.pdf (accessed 10.12.2012). Angers, R.C., Browning, S.R., Seward, T.S., Sigurdson, C.J., Miller, M.W., Hoover, E.A., Telling, G.C., 2006. Prions in skeletal muscles of deer with chronic wasting disease. Science 311, 1117. Angers, R.C., Seward, T.S., Napier, D., Green, M., Hoover, E., Spraker, T., O’Rourke, K., Balachandran, A., Telling, G.C., 2009. Chronic wasting disease prions in elk antler velvet. Emerg. Infect. Dis. 15, 696–703. Baeten, L.A., Powers, B.E., Jewell, J.E., Spraker, T.R., Miller, M.W., 2007. A natural case of chronic wasting disease in a free-ranging moose (Alces alces shirasi). J. Wildl. Dis. 43, 309–334. Balachandran, A., Harrington, N.P., Algire, J., Soutyrine, A., Spraker, T.R., Jeffrey, M., González, L., O’Rourke, K.I., 2010. Experimental oral transmission of chronic wasting disease to red deer (Cervus elaphus elaphus): early detection and late stage distribution of proteaseresistant prion protein. Can. Vet. J. 51, 169–178.
Bartelt, G., Pardee, J., Thiede, K., 2003. Environmental impact statement on rules to eradicate chronic wasting disease from Wisconsin’s free-ranging white-tailed deer herd. In: Wisconsin Department of Natural Resources, http://wildpro.twycrosszoo.org/S/00Ref/ MiscellaneousContents/D109 EIS CWD Wisc/Appendix J.htm (accessed 15.12.2012). Bartz, J.C., Marsh, R.F., McKenzie, D.I., Aiken, J.M., 1998. The host range of chronic wasting disease is altered on passage in ferrets. Virology 251, 297–301. Belay, E.D., Gambetti, O., Schonberger, L.B., Parchi, P., Lyon, D.R., Capellari, S., McQuiston, J.H., Bradley, K., Dowdle, G., Crutcher, J.M., Nichols, C.R., 2001. Creutzfeldt-Jakob disease in unusually young patients who consumed venison. Arch. Neurol. 58, 1673–1678. Belay, E.D., Maddox, R.A., Williams, E.S., Miller, M.W., Gambetti, P., Schonberger, L.B., 2004. Chronic wasting disease and potential transmission to humans. Emerg. Infect. Dis. 10, 977–984. 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Guiroy, D.C., Williams, E.S., Liberski, P.P., Wakayama, I., Gajdusek, D.C., 1993a. Ultrastructural neuropathology of chronic wasting disease in captive mule deer. Acta Neuropathol. 85, 437–444. Guiroy, D.C., Liberski, P.P., Williams, E.S., Gajdusek, D.C., 1994. Electron microscopic findings in brain of Rocky Mountain elk with chronic wasting disease. Folia Neuropathol. 32, 171–173. Haley, N.J., Mathiason, C.K., Carver, S., Zabel, M., Telling, G.C., Hoover, E.A., 2011. Detection of chronic wasting disease prions in salivary, urinary, and intestinal tissues of deer: potential mechanisms of prion shedding and transmission. J. Virol. 85, 6309–6318. Haley, N.J., Seelig, D.M., Zabel, M.D., Telling, G.C., Hoover, E.A., 2009. Detection of CWD prions in urine and saliva of deer by transgenic mouse bioassay. PLoS One 4, e4848. Hall, S.M., 2004. Transmissible spongiform encephalopathy diagnostics [Abstract]. Animal Prion Diseases and the Americas. Ames, Iowa, USA 14-16 October. 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D.C. Bourne / Small Ruminant Research 128 (2015) 72–78 of genetic susceptibility of elk (Cervus elaphus nelsoni) to chronic wasting disease by experimental oral inoculation. J. Vet. Diagn. Invest. 18, 110–114. Hamir, A.N., Greenlee, J.J., Nicholson, E.M., Kunkle, R.A., Richt, J.A., Miller, J.M., Hall, M., 2010. Experimental transmission of chronic wasting disease (CWD) from elk and white-tailed deer to fallow deer by intracerebral route: final report. Can. J. Vet. Res. 75, 152–156. Hamir, A.N., Miller, J.M., Cutlip, R.C., Kunkle, R.A., Jenny, A.L., Stack, M.J., Chaplin, M.J., Richt, J.A., 2004. Transmission of sheep scrapie to elk (Cervus elaphus nelsoni) by intracerebral inoculation: final outcome of the experiment. J. Vet. Diagn. Invest. 16, 316–321. Hamir, A.N., Kehrli Jr., M.E., Kunkle, R.A., Greenlee, J.J., Nicholson, E.M., Richt, J.A., Miller, J.M., Cutlip, R.C., 2011. 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