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Review

Transmission, Strain Diversity, and Zoonotic Potential of Chronic Wasting Disease

by
Sandra Pritzkow
Mitchell Center for Alzheimer’s Disease and Related Brain Disorders, Department of Neurology, University of Texas McGovern Medical School, Houston, TX 77030, USA
Viruses 2022, 14(7), 1390; https://doi.org/10.3390/v14071390
Submission received: 7 May 2022 / Revised: 11 June 2022 / Accepted: 12 June 2022 / Published: 25 June 2022
(This article belongs to the Special Issue The Future of the Chronic Wasting Disease Epizootic)
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Abstract

:
Chronic wasting disease (CWD) is a prion disease affecting several species of captive and free-ranging cervids. In the past few decades, CWD has been spreading uncontrollably, mostly in North America, resulting in a high increase of CWD incidence but also a substantially higher number of geographical regions affected. The massive increase in CWD poses risks at several levels, including contamination of the environment, transmission to animals cohabiting with cervids, and more importantly, a putative transmission to humans. In this review, I will describe the mechanisms and routes responsible for the efficient transmission of CWD, the strain diversity of natural CWD, its spillover and zoonotic potential and strategies to minimize the CWD threat.

1. Background

Prion diseases, also known as Transmissible spongiform encephalopathies (TSEs), are a group of fatal, infectious, neurodegenerative disorders of the brain affecting humans and several other species of mammals, including sheep, goats, cattle, cervids, mink, felines, and camelids [1]. In humans, TSEs are exceedingly rare. Even the most common human TSE is still very infrequent, as is the case of Creutzfeldt–Jakob disease (CJD) which appears at an average rate of 1 new case per million people per year [1,2]. In animals, the most common TSEs are sheep scrapie, bovine spongiform encephalopathy (BSE) affecting cattle, and chronic wasting disease (CWD) affecting captive and wild cervids [2].
The infectious agent in prion disease, usually referred as prion, is likely composed exclusively by a misfolded form of the prion protein (PrPSc), which has the uncanny ability to convert the natively folded prion protein (PrPC) naturally present in many cell types, mostly neurons [3,4]. Molecularly, PrPSc is an aggregate of variable size adopting an intermolecular β-sheet-rich structure [4]. The aggregate acts as a seed to capture the mostly α-helical PrPC, which becomes incorporated into the aggregate adopting the same misfolded structure as the parent PrPSc [5]. Strikingly, this protein-based infectious agent satisfies the Koch postulates used to define a causative relationship between a microbe and a disease [5,6]. Indeed, prions can faithfully multiply in an appropriate host, be transmitted among individuals by various routes including foodborne and bloodborne, can be titrated by infectivity bioassays and display strain diversity, transmission controlled by species barrier, and the ability to cross biological membranes [5].
Among TSEs, CWD is perhaps the most prevalent and worrisome member of the group. It is so far the only prion disease in wildlife, affecting various species of cervid, including white-tailed deer (Odocoileus virginianus), mule deer (Odocoileus hemionus), reindeer (Rangifer tarandus), red deer (Cervus elaphus), elk (Cervus canadensis), moose (Alces alces), sika (Cervus nippon), and muntjac (Muntiacus muntjak) [7]. CWD is very contagious and its origin, prevalence, and mechanisms of transmission are not entirely understood [7]. Symptoms of late-stage CWD infection include emaciation, ataxia, excessive salivation, depression, muscle wasting, and weakness [7,8,9]. It has been shown that CWD produces a significant decline of the wild deer population in areas with high incidence to CWD [10]. CWD-affected deer were 4.5 times more likely to die per year than noninfected deer [10].
CWD was first reported in 1967 in a captive deer facility in Colorado [8]. The origin of CWD is unknown [11], but several possibilities have been suggested (Figure 1). It has been proposed that CWD emerged due to infection with scrapie from sheep that commonly cohabit with cervids [12,13]. Supporting this conclusion, experimental infection of cervids with scrapie-contaminated brain homogenate showed that scrapie prions from sheep can infect elk and white-tailed deer [13]. Moreover, studies by Greenlee and collaborators showed that experimental infection of sheep with CWD produce a disease very similar to conventional scrapie [14], providing further evidence for a possible origin of CWD from scrapie. Other alternative for CWD origin is the appearance of mutations in the prion protein gene (Prnp) which resulted in a protein more prone to convert to PrPSc (Figure 1) [7]. In humans, up to 15% of the TSE cases arise from Prnp mutations [1]. In animals, a single mutation led to the emergence of one of the BSE cases in USA [15]. Another alternative is a spontaneous switch on PrPC conformation (Figure 1), perhaps triggered by changes on the cellular milieu or a defect on the cellular response to protein misfolding [7].
In the past several decades, the disease has been rapidly and progressively spreading geographically and now affects 30 states in the USA, four Canadian provinces, South Korea, and has been recently reported in various countries of Northern Europe, including Norway, Sweden, and Finland [16,17,18,19]. By far the country most affected by CWD is the USA. CWD prevalence is highly variable in North America, but can reach 30% for wild populations in endemic areas, and in exceptional cases can reach 80–90% in captive populations [7,10,20]. Over 200 captive deer facilities have detected CWD on their premises [20].

2. Mechanisms and Routes Responsible for the Efficient Natural Spreading of CWD

The mechanisms and factors explaining the highly efficient transmission of CWD in nature are not completely understood. It has been shown that CWD prion infection can be transmitted vertically or horizontally. Vertical transmission from infected mothers to newborns has been reported in deer as well as in other animal TSEs, including prion diseases in cattle and sheep [21,22]. In the case of cervids, it was shown that CWD-positive muntjac dams produced a progeny in which 80% of fetuses were infected with CWD, providing strong evidence for in utero prion infection [21]. Maternal CWD infection also appears to result in a lower percentage of live-birth offspring [21]. Natural CWD transmission from cows to calves has also been demonstrated to contribute to the CWD epidemic in free-ranging elk in Colorado [23]. Furthermore, several studies have reported the presence of PrPSc in fetal, gestational, and reproductive tissues of CWD-infected deer [24,25,26].
Horizontal transmission is possibly the most efficient route of natural spreading and as such the biggest contributor to the rapid spread of CWD [7,27,28]. Horizontal transmission can occur by direct animal-to-animal contact or indirectly through environmental fomites. Epidemiological studies of natural CWD spreading in captive mule deer found that in cohabiting animals, horizontal transmission is highly efficient, with an estimated incidence of 89% in herds in which vertical transmission was excluded [28]. Direct horizontal transmission of the disease from a CWD-infected animal to a healthy cervid likely occurs during mating or fighting through contact with saliva, blood, or mucosal tissues where CWD prions have been detected [29,30,31]. The efficiency of this type of transmission might be exacerbated by lesions on oral mucous membranes, which are frequent in cervids. Hoover et al. showed that minor lingual abrasions substantially facilitate CWD transmission in transgenic mouse models of CWD transmission [32]. Although in general prions are not considered airborne pathogens, some studies have shown that CWD can be transmitted by aerosol exposure [33,34], raising the possibility that simple cohabitation of uninfected and infected animals may result in prion transmission.
Indirect horizontal transmission occurs when uninfected animals become in contact with environmental materials that were previously contaminated by prions released by infected animals. CWD is a disease with a high peripheral distribution of infectious prions and it has been shown that infected animals can excrete a relatively large quantity of prions into the environment through urine, feces, or saliva [2,29,30,35]. Even though the prion-infectivity titer in CWD excreta is low, the repetitive excretion and accumulation of these materials lead to massive release of infectious prions into the environment. Estimations of the amount of prions excreted through feces and urine conducted by infectivity bioassays and high-sensitivity amplification techniques have shown that over the course of the incubation period and clinical disease, an infected deer sheds a higher amount of prions through excreta compared to the level of infectious agent present in the brain at terminal stage of the disease [29,36,37,38,39]. These calculations indicate that per year an infected animal may release >10 mg of infectious prions through feces and urine. Taking into account the population of CWD-infected cervids in the USA (likely to be in the order of hundreds of thousands of animals), the environment in the country possibly receive Kg quantities of CWD prions each year from urine and feces alone. The putative role of excreta in CWD transmission is supported by an experiment in which 11% of mule deer exposed to excreta-contaminated paddocks became infected [40]. CWD prions have been also found in deer saliva during the course of the disease, even in the asymptomatic incubation period [41,42,43]. Cervids produce a large amount of saliva, which can contaminate foliage and soil. Nevertheless, the largest amount of infectious material deposited at one point into the environment occurs when sick animals die and decomposing carcases remain in soil or are taken up by plants, insects, or scavenger animals. This source was proven in an experiment in which healthy mule deer were cohoused with CWD decomposed carcases. The result of this study showed a 25% infection rate in these animals [40].
Compelling evidence indicates that once prions are released into the environment, they can accumulate in association with various natural elements of the environment and remain infectious for several years. Soil is likely the environmental element most likely to interact with prions released via feces, urine, saliva, or animal carcasses. Prions can tightly bind to soil and remain infectious [44,45,46,47,48,49,50,51]. Interestingly, it has been reported that certain soil components (e.g., montmorillonite) may even increase the prion infectivity titer by over two orders of magnitude [52]. It is also possible that rainwater may elute and spread prions from contaminated soil, potentially carrying them to other areas, including groundwater [53]. Interestingly, CWD prions were detected in water in a CWD endemic area [54].
The documented presence of CWD prions in soil suggest that organisms living in contact with soil (e.g., plants, earthworms, and insects), may also be exposed and participate in prion spreading. In this sense, we showed that grass plants effectively bind prions from CWD-infected brain and excreta [55]. Prions can persist for prolonged periods of time bound to living plants [55], and importantly, prions bound to plants are infectious to experimental animals upon oral inoculation [55]. Surprisingly, we found that plants can uptake prions from the soil and transport them to the stem and leaves [55]. We also recently showed that earthworms living in prion-contaminated soil may contribute to spread prions across the environment. An experiment mimicking the interaction of earthworms with soil previously exposed to infectious prions found that worms can bind, uptake, accumulate, and scatter infectious prions in the soil [56]. Earthworms carrying prions maintain infectious properties [56]. Finally, we and others have shown that prions can bind to many natural and manmade environmental surfaces, including rocks, wood, metals, glass, plastic, concrete, etc. [57,58,59]. These surfaces commonly found in areas endemic for CWD can efficiently bind prions, which remain highly infectious. Strikingly, casual indirect contact with the prion contaminated surface (e.g., licking, sniffing, or brushing alongside the surface) is enough to transfer prions from the surface to the animal leading to disease [57]. Altogether, these findings suggest that prion contamination and progressive accumulation in the environment play a major role in CWD natural spreading. In addition, it is possible that prions attached to elements of the environment may also contribute to modify some prion properties, including infectivity titer, strain features, and possibly the potential to infect other animal species or even humans.

3. CWD Strain Diversity

Akin to conventional micro-organisms, prions can adopt multiple strains in a single species of animals [60,61]. However, in contrast from other microbes where strain diversity depends on changes on the nucleic acids, in the case of prions, strains are thought to arise from different conformations of PrPSc [60]. Prion strains can produce diseases with distinct clinical phenotypes, neurodegeneration profiles, incubation periods, areas of the brain affected by vacuolation and accumulation of PrPSc, and biochemical/biophysical properties of PrPSc. In humans affected by sCJD, five different prion strains have been recognized [62]. In animals, diverse prion strains have been reported in cattle and sheep, as well as in experimental rodents [63]. For CWD, we still do not know how many natural strains are present in nature [16]. Earlier studies from Telling et al. using transgenic mice models showed the existence of at least two different prion strains in CWD infected animals [64]. It was shown that host factors can modulate CWD prion diversity [65]. In addition, prion protein polymorphisms (Table 1), as well as small differences on the amino-acid sequence between different cervid species, may lead to the emergence of new prion strains when animals are infected with PrPSc carrying a mismatch sequence from host PrPC [66,67,68,69].
It is well-established that the presence of certain polymorphic variants in cervids (Table 1) may confer resistance to CWD infection [18,70,71]. For example, infection of white-tailed deer harboring rarer polymorphisms at position 95 and 96 of PrPC led to a prolonged incubation period compared to animals with more prevalent polymorphisms [72,73]. Supporting this conclusion, it was shown that wild white-tailed deer harboring the S96 polymorphism were less common in naturally CWD-infected animals [74]. Similarly, captive white-tailed deer containing uncommon polymorphisms (e.g., H95, S96, A116, or K226) have been shown to have a lower rate of CWD infection than those animals containing the more common polymorphisms [75]. Similar findings were observed in elk and mule deer, where the L132 and F225 polymorphisms, respectively, are underrepresented in CWD-infected animals [76,77]. Given the relatively large number of CWD-infected cervids in some areas of North America, it is possible that the differential susceptibility to CWD infection in certain polymorphic variants may alter the distribution of Prnp polymorphisms in cervids [78]. However, it is important to highlight that up to now, no polymorphisms confer absolute protection to CWD. Furthermore, it is likely that new emergent CWD strains may have a distinct preferred host selection than the commonly existing strains.
The recent emergence of CWD in Northern Europe has provided additional evidence for strain diversity in cervids. European CWD appears to have a different origin from North American CWD, since no cervids have been imported into Scandinavia from North America in the past several decades [18]. The first cases of CWD in Europe were reported in 2016–2019 by Benestad et al. in reindeer, moose, and red deer native to Norway [79,80,81]. In subsequent years, CWD was reported in moose living in Finland and Sweden [18]. Experiments by infectivity bioassay in bank voles provided strong evidence for European CWD corresponding to different strains from their North American counterparts [82]. This conclusion was further supported by infectivity studies in transgenic and gene-targeted mice expressing deer Prnp, showing convincing prion strain differences between European and North American CWD isolates [83]. Interestingly, repetitive passage in mice of some of the Norwegian CWD infectious material led to the adaptation and maturation of the prion strain to reach a similar stage as the current North America CWD strains [83]. This intriguing result suggests that European CWD is probably at a more primitive state of maturation than North American CWD. It has been shown that prions can evolve, mutate, adapt, and mature upon successive passages, leading to strain changes and resulting in the emergence of prions with different virulence and ability to cross species barriers [66]. Indeed, we previously showed that maturation of the CWD agent by many cycles of prion replication leads to increase ability to infect human models [84]. The strain diversity observed in CWD raises the possibility that distinct strains may have differential abilities to generate infection into other mammal species. Our studies using in vitro prion replication showed that Norwegian CWD has a higher potential to convert PrPC from various animal species than North America CWD, but a lower potential to convert human PrPC [85].

4. Species Barrier and CWD Zoonotic Potential

Another typical characteristic of prions in common with conventional micro-organisms is their capacity to infect only some animal species, a phenomenon usually referred as “species barrier” [86,87]. The ability of prions from one species to infect animals from a different species depends on the sequence homology between the prion protein from donor and acceptor as well as the specific prion strain properties of the infectious material [86,87]. Cervids cohabit in nature with several animal species susceptible to prion infection, such as livestock, rodents, scavengers, carnivores, pets, and humans (Figure 2) [7]. Particularly at risk are predators and scavenger animals, which may consume CWD-infected carcasses, but also herbivores through exposure to contaminated environment, including plants and soil. Several studies of experimental infection with CWD prions showed that CWD can transmit disease to various animal species, including cattle, sheep, goats, ferrets, and raccoons [88,89,90,91,92,93,94]. Interspecies transmission typically results in a lower attack rate (proportion of animals developing the disease out of the total number of animals exposed) and longer incubation periods compared to CWD transmission to cervids. Importantly, CWD transmission to other animal species may increase the virulence and zoonotic potential of newly generated prion strains in the host animals [2]. This appears to be the case of scrapie passaged into cattle. While traditional scrapie in sheep is considered not infectious to humans, cattle infected by scrapie can infect humans producing variant CJD (vCJD) [95].
In reference to biomedical relevance, the most important species barrier is the cervid-to-human barrier; in other words, the zoonotic potential of CWD prions. Many studies have been conducted using different technologies and model systems to assess the possibility that CWD may be able to infect humans under certain conditions. Currently, the evidence for transmission of CWD to humans is contentious. Studies using transgenic mice expressing human PrPC injected with CWD prions have usually produced negative results, suggesting the species barriers between human and cervids is very large [96,97,98,99]. On the contrary, infectivity experiments in squirrel monkeys showed that CWD is highly infectious in this nonhuman primate model [100,101]. However, experiments in cynomolgus macaques, which are considered evolutionarily closer than squirrel monkeys to humans, have produced contradictory results. Two published studies from Chesebro’s group reported no clinical, pathological, or biochemical evidence of CWD transmission to macaques [101,102]. Studies were conducted in seven macaques inoculated with CWD-and sacrificed 11 to 13 years after CWD infection. Conversely, a yet-unpublished study by Czub, Schaetzl et al. found that upon infection of groups of macaques with the CWD agent by various routes, including oral inoculation of muscle tissue from infected cervids, many of the animals developed clinical, neuropathological, and biochemical alterations consistent with prion disease [103]. Analysis of the cervid-to-human species barrier by in vitro prion replication using the protein-misfolding cyclic-amplification assay (PMCA) or real-time quacking-induced conversion (RT-QuIC) indicated that under certain conditions, PrPSc from CWD-infected cervids is able to convert the human PrPC into the pathogenic form [84,85,104,105,106]. Interestingly, the efficiency of crossing the cervid/human species barrier in vitro depended on the Prnp polymorphisms of the agent and the host [104] as well as on the degree of strain stabilization [84]. It has been shown that repetitive amplification of CWD prions as expenses of cervid PrPC resulted in a PrPSc form more capable to convert human PrPC [84]. These findings suggest that the longer animals are propagating CWD prions, the easier it might be to infect humans. Nevertheless, up to now there has not been any evidence that CWD has been naturally transmitted to humans. Epidemiological studies did not find a correlation between incidence of human TSE and CWD prevalence or consumption of deer meat [107,108].
Up to now, the only animal prion disease that has been transmitted to humans is BSE in cattle [95]. When compared to BSE, CWD appears to have a wider peripheral distribution of prions and much more efficient horizontal transmission. CWD prions have been readily detected in many peripheral tissues and biological fluids, including muscle, skin, lymph nodes, salivary glands, urinary bladder, pancreas, kidney, intestine, blood, urine, feces, and saliva [19]. This makes CWD prions more accessible to exposure to other animals and humans than BSE prions.

5. Strategies for Minimizing CWD Spreading: Is Eradication of CWD a Possibility?

Given the large number of CWD-infected animals in North America, the likely massive contamination of the environment, and the efficient horizontal spreading of the disease, it is unlikely that CWD can be eradicated from the United States. Nevertheless, it is possible to implement strategies to minimize and control CWD transmission. Following is an outline of six different strategies that may contribute to achieve this goal (Figure 3).

5.1. Surveillance

Routine active surveillance for CWD-infected cervids and for putative cases of CWD transmission to other animals or humans may contribute to limit the further spreading of the disease. CWD surveillance includes monitoring the appearance of new CWD cases both in wild and captive animals, the geographical location of these cases, possible origins, and animal species affected. This strategy might be particularly useful to prevent spreading of CWD to geographical areas not currently affected by the disease. An efficient surveillance program would require a tight interaction among state and federal authorities, owners of deer farms, hunters, wildlife agencies, and scientists experienced in CWD biology.

5.2. Selective Breeding

As described above, several reports have confirmed that some Prnp polymorphisms confer relative resistance to CWD infection [18,70,71,72,73,74,75,76,77]. Thus, one strategy to minimize CWD spreading might be to perform selective breeding to produce animals with several polymorphic variants known to decrease infection. A recent study has shown that selective breeding might contribute to reduce CWD spreading [109]. A similar strategy was previously employed to attempt reducing prevalence of scrapie in sheep [110]. However, the emergence of atypical strains of scrapie, which transmitted efficiently to animals with Prnp polymorphisms resistant to classical scrapie, complicated this approach [110]. Perhaps a safer and more effective strategy would be to produce knockout cervids for Prnp. It has been demonstrated that elimination of the Prnp gene in mice confers complete resistance to all forms of prions [111]. Interestingly, a Prnp null cow was previously generated, which showed no detectable abnormality and was completely resistant to BSE prion infection [112].

5.3. Implementation of High-Sensitivity Tests for CWD Prion Detection

Availability of a noninvasive test to detect live CWD-infected animals before the onset of the clinical disease would certainly help to minimize CWD spreading by identifying infected deer before they can excrete too many prions into the environment or transmit the disease horizontally or vertically. Currently, postmortem analysis of the brain or lymphoid tissues for vacuolation and PrPSc deposition by histology or by Western blot or ELISA techniques are the only ways to definitively diagnose CWD [18]. These assays have a rather low sensitivity and cannot be performed in live animals. In recent years, in vitro prion-replication assays, including PMCA and RT-QuIC, have been extensively used to detect prions in biological fluids of different animal species, including CWD [113,114,115,116,117,118]. These assays rely on mimicking in vitro the mechanism of prion replication to amplify minute amounts of PrPSc at expenses of large quantities of PrPC using a cyclic-amplification procedure involving cycles of incubation and fragmentation. Both PMCA and RT-QuIC have shown high-sensitivity detection of CWD prions in a variety of biological samples, including blood, urine, saliva, and feces [31,37,39,41,43,119,120,121,122]. These assays have been also shown to be useful to detect prions in diverse environmental materials, including soil, plants, water, diverse natural and man-made surfaces, and in organisms living in the environment [47,54,55,56,57]. However, since they are not regularly performed, it is unclear how they will operate for routine prion detection.

5.4. Selective Animal Culling

Early removal of CWD-infected animals or cervids at a high risk for infection might be a possibility to decrease CWD prevalence [123,124]. Selective animal culling coupled with massive live animal testing with highly efficient assays for detection of CWD prions may represent a feasible strategy. For example, a study involving selective culling of CWD-infected mule deer (determined by immunohistochemistry of tonsil biopsies) in a limited area of Colorado showed a reduction of CWD prevalence in males [125]. However, other studies have found no evidence of CWD reduction after animal culling [126]. A massive nonselective culling of the entire population of deer in the Nordfjella part of Norway (>2000 animals) was carried out in 2017 after the emergence of CWD in this region [127]. This approach may well work to prevent new cases of the disease, especially at the beginning of the CWD epidemic, but it has serious ethical, economic, and political consequences.

5.5. Prion Decontamination

Although prions are resistant to many of the decontamination procedures utilized for conventional micro-organisms, there are very well-established chemical procedures for prion elimination [128]. The best-established procedures include treatment with solutions of concentrated sodium hypochlorite, sodium hydroxide, or guanidine. Unfortunately, some of these procedures are corrosive or not compatible with some surfaces. There are several reports of other prion-decontamination procedures (e.g., proteases, detergents, hydrogen peroxide) that can be less damaging to materials [129,130,131,132,133,134]. Coupling an efficient and noninvasive prion-detection methodology with an effective decontamination procedure could be very important to decrease further indirect horizontal spreading of the disease [2]. It is important to highlight that effort should be made to test the efficiency of decontamination in “real-life” treatments, and not assume the procedures work based on experimental studies. Fortunately, with the emergence of ultrasensitive prion-detection techniques (e.g., PMCA or RT-QuIC) that work in a variety of surfaces [57,135,136,137,138], it is now feasible to quality control the effectiveness of prion decontamination.

5.6. Treatment for CWD

Unfortunately, at this time there is no treatment available for CWD or any other prion disease [139,140]. Various different approaches have been proposed and tested in vitro on cells or animal models, including small-molecule drugs, passive and active immunization, aptamers, peptides, and RNA-interference techniques [139,140,141]. Although some of these strategies delay the onset of prion disease, none of them completely prevent or reverse the disease. In the case of CWD, a prophylactic treatment that can make animals immune to prion infection, such as a vaccine, could be ideal for massive use in preventing CWD spreading. Various studies investigating the use of a vaccine for CWD [141,142] have been reported, but more studies are needed to assess the efficacy of such approach in the field. Surprisingly, treatment of elk with a vaccine targeting a YYR disease-specific epitope resulted in an acceleration of disease onset compared to controls [143].

6. Conclusions and Perspectives

CWD is currently the most dangerous prion disease, because it affects wild animals, spreads efficiently, and has a high rate of peripheral excretion of infectious agent and a long incubation period. The mechanisms implicated in the facile transmission of CWD are not completely understood, but likely involve substantial contamination of the environment [2,144]. Also unknown are the number and properties of prion strains implicated in natural CWD and their contribution to disease spreading. Finally, the zoonotic potential of CWD remains a dangerous enigma.
Some of the most relevant pending questions in relation to CWD are: Is CWD infectious to humans under certain conditions? What is the exact contribution of the environment to CWD spreading? How many natural CWD strains exist and what are they properties? Has CWD spread to other species of animals? What is the atomic resolution structure of PrPSc responsible for CWD? How CWD prions produce brain damage and disease? What are the most efficient strategies to minimize and control CWD transmission? Is it possible to cure and eradicate CWD?
Much more research at all levels needs to be conducted to properly combat this insidious disease and to avoid the emergence of new diseases.

Funding

This study was supported in part by a grant from the National Institute of Health P01AI077774.

Acknowledgments

I would like to thank Claudio Soto and members of his team for continuous support and stimulating discussions.

Conflicts of Interest

S.P. has a conflict of interest related to the PMCA technology, which is under development for commercialization by Amprion Inc. The University of Texas System has licensed intellectual property to Amprion. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

References

  1. Collinge, J. Prion diseases of humans and animals: Their causes and molecular basis. Annu. Rev. Neurosci. 2001, 24, 519–550. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  2. Pritzkow, S.; Gorski, D.; Ramirez, F.; Soto, C. Prion Dissemination through the Environment and Medical Practices: Facts and Risks for Human Health. Clin. Microbiol. Rev. 2021, 34, e0005919. [Google Scholar] [CrossRef] [PubMed]
  3. Prusiner, S.B. Prions. Proc. Natl. Acad. Sci. USA 1998, 95, 13363–13383. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Soto, C. Prion Hypothesis: The end of the Controversy? Trends Biochem. Sci. 2011, 36, 151–158. [Google Scholar] [CrossRef] [Green Version]
  5. Soto, C. Transmissible Proteins: Expanding the Prion Heresy. Cell 2012, 149, 968–977. [Google Scholar] [CrossRef] [Green Version]
  6. Walker, L.; LeVine, H.; Jucker, M. Koch’s postulates and infectious proteins. Acta Neuropathol. 2006, 112, 1–4. [Google Scholar] [CrossRef]
  7. Escobar, L.E.; Pritzkow, S.; Winter, S.N.; Grear, D.A.; Kirchgessner, M.S.; Dominguez-Villegas, E.; Machado, G.; Townsend Peterson, A.; Soto, C. The ecology of chronic wasting disease in wildlife. Biol. Rev. Camb. Philos. Soc. 2020, 95, 393–408. [Google Scholar] [CrossRef]
  8. Williams, E.S.; Young, S. Chronic wasting disease of captive mule deer: A spongiform encephalopathy. J. Wildl. Dis. 1980, 16, 89–98. [Google Scholar] [CrossRef] [Green Version]
  9. Spraker, T.R.; Miller, M.W.; Williams, E.S.; Getzy, D.M.; Adrian, W.J.; Schoonveld, G.G.; Spowart, R.A.; O’Rourke, K.I.; Miller, J.M.; Merz, P.A. Spongiform encephalopathy in free-ranging mule deer (Odocoileus hemionus), white-tailed deer (Odocoileus virginianus) and Rocky Mountain elk (Cervus elaphus nelsoni) in northcentral Colorado. J. Wildl. Dis. 1997, 33, 1–6. [Google Scholar] [CrossRef]
  10. Edmunds, D.R.; Kauffman, M.J.; Schumaker, B.A.; Lindzey, F.G.; Cook, W.E.; Kreeger, T.J.; Grogan, R.G.; Cornish, T.E. Chronic Wasting Disease Drives Population Decline of White-Tailed Deer. PLoS ONE 2016, 11, e0161127. [Google Scholar] [CrossRef]
  11. Williams, E.S.; Miller, M.W. Transmissible spongiform encephalopathies in non-domestic animals: Origin, transmission and risk factors. Rev. Sci. Tech. 2003, 22, 145–156. [Google Scholar] [CrossRef]
  12. Miller, M.W.; Williams, E.S. Chronic wasting disease of cervids. Curr. Top. Microbiol. Immunol. 2004, 284, 193–214. [Google Scholar]
  13. Greenlee, J.J.; Smith, J.D.; Kunkle, R.A. White-tailed deer are susceptible to the agent of sheep scrapie by intracerebral inoculation. Vet. Res. 2011, 42, 107. [Google Scholar] [CrossRef] [Green Version]
  14. Cassmann, E.D.; Frese, R.D.; Greenlee, J.J. Second passage of chronic wasting disease of mule deer to sheep by intracranial inoculation compared to classical scrapie. J. Vet. Diagn. Investig. 2021, 33, 711–720. [Google Scholar] [CrossRef]
  15. Richt, J.A.; Hall, S.M. BSE case associated with prion protein gene mutation. PLoS Pathog. 2008, 4, e1000156. [Google Scholar] [CrossRef] [Green Version]
  16. Benestad, S.L.; Telling, G.C. Chronic wasting disease: An evolving prion disease of cervids. Handb. Clin. Neurol. 2018, 153, 135–151. [Google Scholar]
  17. Hazards, E.P.O.B.; Ricci, A.; Allende, A.; Bolton, D.; Chemaly, M.; Davies, R.; Fernandez Escamez, P.S.; Girones, R.; Herman, L.; Koutsoumanis, K.; et al. Chronic wasting disease (CWD) in cervids. EFSA J. 2017, 15, e04667. [Google Scholar]
  18. Silva, C.J. Chronic Wasting Disease (CWD) in Cervids and the Consequences of a Mutable Protein Conformation. ACS Omega 2022, 7, 12474–12492. [Google Scholar] [CrossRef]
  19. Otero, A.; Velasquez, C.D.; Aiken, J.; McKenzie, D. Chronic wasting disease: A cervid prion infection looming to spillover. Vet. Res. 2021, 52, 115. [Google Scholar] [CrossRef]
  20. Rivera, N.A.; Brandt, A.L.; Novakofski, J.E.; Mateus-Pinilla, N.E. Chronic Wasting Disease In Cervids: Prevalence, Impact And Management Strategies. Vet. Med. 2019, 10, 123–139. [Google Scholar] [CrossRef] [Green Version]
  21. Nalls, A.V.; McNulty, E.; Powers, J.; Seelig, D.M.; Hoover, C.; Haley, N.J.; Hayes-Klug, J.; Anderson, K.; Stewart, P.; Goldmann, W.; et al. Mother to offspring transmission of chronic wasting disease in reeves’ muntjac deer. PLoS ONE 2013, 8, e71844. [Google Scholar] [CrossRef] [Green Version]
  22. Foster, J.D.; Goldmann, W.; Hunter, N. Evidence in sheep for pre-natal transmission of scrapie to lambs from infected mothers. PLoS ONE 2013, 8, e79433. [Google Scholar] [CrossRef] [Green Version]
  23. Selariu, A.; Powers, J.G.; Nalls, A.; Brandhuber, M.; Mayfield, A.; Fullaway, S.; Wyckoff, C.A.; Goldmann, W.; Zabel, M.M.; Wild, M.A.; et al. In utero transmission and tissue distribution of chronic wasting disease-associated prions in free-ranging Rocky Mountain elk. J. Gen. Virol. 2015, 96, 3444–3455. [Google Scholar] [CrossRef]
  24. Kramm, C.; Gomez-Gutierrez, R.; Soto, C.; Telling, G.; Nichols, T.; Morales, R. In Vitro detection of Chronic Wasting Disease (CWD) prions in semen and reproductive tissues of white tailed deer bucks (Odocoileus virginianus). PLoS ONE 2019, 14, e0226560. [Google Scholar] [CrossRef] [Green Version]
  25. Nalls, A.V.; McNulty, E.; Hoover, C.E.; Pulscher, L.A.; Hoover, E.A.; Mathiason, C.K. Infectious Prions in the Pregnancy Microenvironment of Chronic Wasting Disease-Infected Reeves’ Muntjac Deer. J. Virol. 2017, 91, e00501-17. [Google Scholar] [CrossRef] [Green Version]
  26. Nalls, A.V.; McNulty, E.E.; Mayfield, A.; Crum, J.M.; Keel, M.K.; Hoover, E.A.; Ruder, M.G.; Mathiason, C.K. Detection of Chronic Wasting Disease Prions in Fetal Tissues of Free-Ranging White-Tailed Deer. Viruses 2021, 13, 2430. [Google Scholar] [CrossRef]
  27. Moore, S.J.; Kunkle, R.; Greenlee, M.H.; Nicholson, E.; Richt, J.; Hamir, A.; Waters, W.R.; Greenlee, J. Horizontal Transmission of Chronic Wasting Disease in Reindeer. Emerg. Infect. Dis. 2016, 22, 2142–2145. [Google Scholar] [CrossRef] [Green Version]
  28. Miller, M.W.; Williams, E.S. Prion disease: Horizontal prion transmission in mule deer. Nature 2003, 425, 35–36. [Google Scholar] [CrossRef] [PubMed]
  29. Haley, N.J.; Seelig, D.M.; Zabel, M.D.; Telling, G.C.; Hoover, E.A. Detection of CWD prions in urine and saliva of deer by transgenic mouse bioassay. PLoS ONE 2009, 4, e4848. [Google Scholar] [CrossRef] [PubMed]
  30. Mathiason, C.K.; Powers, J.G.; Dahmes, S.J.; Osborn, D.A.; Miller, K.V.; Warren, R.J.; Mason, G.L.; Hays, S.A.; Hayes-Klug, J.; Seelig, D.M.; et al. Infectious prions in the saliva and blood of deer with chronic wasting disease. Science 2006, 314, 133–136. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  31. Kramm, C.; Pritzkow, S.; Lyon, A.; Nichols, T.; Morales, R.; Soto, C. Detection of Prions in Blood of Cervids at the Asymptomatic Stage of Chronic Wasting Disease. Sci. Rep. 2017, 7, 17241. [Google Scholar] [CrossRef] [Green Version]
  32. Denkers, N.D.; Telling, G.C.; Hoover, E.A. Minor oral lesions facilitate transmission of chronic wasting disease. J. Virol. 2011, 85, 1396–1399. [Google Scholar] [CrossRef] [Green Version]
  33. Denkers, N.D.; Hayes-Klug, J.; Anderson, K.R.; Seelig, D.M.; Haley, N.J.; Dahmes, S.J.; Osborn, D.A.; Miller, K.V.; Warren, R.J.; Mathiason, C.K.; et al. Aerosol transmission of chronic wasting disease in white-tailed deer. J. Virol. 2013, 87, 1890–1892. [Google Scholar] [CrossRef] [Green Version]
  34. Denkers, N.D.; Seelig, D.M.; Telling, G.C.; Hoover, E.A., Jr. Aerosol and Nasal Transmission of Chronic Wasting Disease in Cervidized Mice. J. Gen. Virol. 2010, 91, 1651–1658. [Google Scholar] [CrossRef]
  35. Safar, J.G.; Lessard, P.; Tamguney, G.; Freyman, Y.; Deering, C.; Letessier, F.; DeArmond, S.J.; Prusiner, S.B. Transmission and detection of prions in feces. J. Infect. Dis 2008, 198, 81–89. [Google Scholar] [CrossRef]
  36. Tamguney, G.; Miller, M.W.; Wolfe, L.L.; Sirochman, T.M.; Glidden, D.V.; Palmer, C.; Lemus, A.; DeArmond, S.J.; Prusiner, S.B. Asymptomatic deer excrete infectious prions in faeces. Nature 2009, 461, 529–532. [Google Scholar] [CrossRef] [Green Version]
  37. Pulford, B.; Spraker, T.R.; Wyckoff, A.C.; Meyerett, C.; Bender, H.; Ferguson, A.; Wyatt, B.; Lockwood, K.; Powers, J.; Telling, G.C.; et al. Detection of PrPCWD in feces from naturally exposed Rocky Mountain elk (Cervus elaphus nelsoni) using protein misfolding cyclic amplification. J. Wildl. Dis. 2012, 48, 425–434. [Google Scholar] [CrossRef]
  38. Henderson, D.M.; Tennant, J.M.; Haley, N.J.; Denkers, N.D.; Mathiason, C.K.; Hoover, E.A. Detection of chronic wasting disease prion seeding activity in deer and elk feces by real-time quaking-induced conversion. J. Gen. Virol. 2017, 98, 1953–1962. [Google Scholar] [CrossRef] [Green Version]
  39. Haley, N.J.; Mathiason, C.K.; Carver, S.; Zabel, M.; Telling, G.C.; Hoover, E.A. Detection of chronic wasting disease prions in salivary, urinary, and intestinal tissues of deer: Potential mechanisms of prion shedding and transmission. J. Virol. 2011, 85, 6309–6318. [Google Scholar] [CrossRef] [Green Version]
  40. Miller, M.W.; Williams, E.S.; Hobbs, N.T.; Wolfe, L.L. Environmental sources of prion transmission in mule deer. Emerg. Infect. Dis. 2004, 10, 1003–1006. [Google Scholar] [CrossRef]
  41. Davenport, K.A.; Hoover, C.E.; Denkers, N.D.; Mathiason, C.K.; Hoover, E.A. Modified Protein Misfolding Cyclic Amplification Overcomes Real-Time Quaking-Induced Conversion Assay Inhibitors in Deer Saliva To Detect Chronic Wasting Disease Prions. J. Clin. Microbiol. 2018, 56, e00947-18. [Google Scholar] [CrossRef] [Green Version]
  42. Davenport, K.A.; Mosher, B.A.; Brost, B.M.; Henderson, D.M.; Denkers, N.D.; Nalls, A.V.; McNulty, E.; Mathiason, C.K.; Hoover, E.A. Assessment of Chronic Wasting Disease Prion Shedding in Deer Saliva with Occupancy Modeling. J. Clin. Microbiol. 2018, 56, e01243-17. [Google Scholar] [CrossRef] [Green Version]
  43. Henderson, D.M.; Manca, M.; Haley, N.J.; Denkers, N.D.; Nalls, A.V.; Mathiason, C.K.; Caughey, B.; Hoover, E.A. Rapid antemortem detection of CWD prions in deer saliva. PLoS ONE 2013, 8, e74377. [Google Scholar] [CrossRef] [Green Version]
  44. Wyckoff, A.C.; Kane, S.; Lockwood, K.; Seligman, J.; Michel, B.; Hill, D.; Ortega, A.; Mangalea, M.R.; Telling, G.C.; Miller, M.W.; et al. Clay Components in Soil Dictate Environmental Stability and Bioavailability of Cervid Prions in Mice. Front. Microbiol. 2016, 7, 1885. [Google Scholar] [CrossRef] [Green Version]
  45. Giachin, G.; Narkiewicz, J.; Scaini, D.; Ngoc, A.T.; Margon, A.; Sequi, P.; Leita, L.; Legname, G. Prion protein interaction with soil humic substances: Environmental implications. PLoS ONE 2014, 9, e100016. [Google Scholar] [CrossRef] [Green Version]
  46. Smith, C.B.; Booth, C.J.; Pedersen, J.A. Fate of prions in soil: A review. J. Environ. Qual. 2011, 40, 449–461. [Google Scholar] [CrossRef]
  47. Nagaoka, K.; Yoshioka, M.; Shimozaki, N.; Yamamura, T.; Murayama, Y.; Yokoyama, T.; Mohri, S. Sensitive detection of scrapie prion protein in soil. Biochem. Biophys. Res. Commun. 2010, 397, 626–630. [Google Scholar] [CrossRef]
  48. Jacobson, K.H.; Lee, S.; Somerville, R.A.; McKenzie, D.; Benson, C.H.; Pedersen, J.A. Transport of the pathogenic prion protein through soils. J. Environ. Qual. 2010, 39, 1145–1152. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  49. Seidel, B.; Thomzig, A.; Buschmann, A.; Groschup, M.H.; Peters, R.; Beekes, M.; Terytze, K. Scrapie Agent (Strain 263K) can transmit disease via the oral route after persistence in soil over years. PLoS ONE 2007, 2, e435. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  50. Johnson, C.J.; Phillips, K.E.; Schramm, P.T.; McKenzie, D.; Aiken, J.M.; Pedersen, J.A. Prions Adhere to Soil Minerals and Remain Infectious. PLoS Pathog. 2006, 2, e32. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  51. Brown, P.; Gajdusek, D.C. Survival of scrapie virus after 3 years’ interment. Lancet 1991, 337, 269–270. [Google Scholar] [CrossRef]
  52. Johnson, C.J.; Pedersen, J.A.; Chappell, R.J.; McKenzie, D.; Aiken, J.M. Oral Transmissibility of Prion Disease Is Enhanced by Binding to Soil Particles. PLoS Pathog. 2007, 3, e93. [Google Scholar] [CrossRef]
  53. Somerville, R.A.; Fernie, K.; Smith, A.; Bishop, K.; Maddison, B.C.; Gough, K.C.; Hunter, N. BSE infectivity survives burial for five years with only limited spread. Arch. Virol. 2019, 164, 1135–1145. [Google Scholar] [CrossRef] [Green Version]
  54. Nichols, T.A.; Pulford, B.; Wyckoff, A.C.; Meyerett, C.; Michel, B.; Gertig, K.; Hoover, E.A.; Jewell, J.E.; Telling, G.C.; Zabel, M.D. Detection of protease-resistant cervid prion protein in water from a CWD-endemic area. Prion 2009, 3, 171–183. [Google Scholar] [CrossRef] [Green Version]
  55. Pritzkow, S.; Morales, R.; Moda, F.; Khan, U.; Telling, G.C.; Hoover, E.; Soto, C. Grass plants bind, retain, uptake, and transport infectious prions. Cell Rep. 2015, 11, 1168–1175. [Google Scholar] [CrossRef] [Green Version]
  56. Pritzkow, S.; Morales, R.; Camacho, M.; Soto, C. Uptake, Retention, and Excretion of Infectious Prions by Experimentally Exposed Earthworms. Emerg. Infect. Dis. 2021, 27, 3151–3154. [Google Scholar] [CrossRef]
  57. Pritzkow, S.; Morales, R.; Lyon, A.; Concha-Marambio, L.; Urayama, A.; Soto, C. Efficient prion disease transmission through common environmental materials. J. Biol. Chem. 2018, 293, 3363–3373. [Google Scholar] [CrossRef] [Green Version]
  58. Flechsig, E.; Hegyi, I.; Enari, M.; Schwarz, P.; Collinge, J.; Weissmann, C. Transmission of scrapie by steel-surface-bound prions. Mol. Med. 2001, 7, 679–684. [Google Scholar] [CrossRef]
  59. Maddison, B.C.; Baker, C.A.; Terry, L.A.; Bellworthy, S.J.; Thorne, L.; Rees, H.C.; Gough, K.C. Environmental sources of scrapie prions. J. Virol. 2010, 84, 11560–11562. [Google Scholar] [CrossRef] [Green Version]
  60. Carta, M.; Aguzzi, A. Molecular foundations of prion strain diversity. Curr. Opin. Neurobiol. 2021, 72, 22–31. [Google Scholar] [CrossRef]
  61. Bartz, J.C. Prion Strain Diversity. Cold Spring Harb. Perspect. Med. 2016, 6, a024349. [Google Scholar] [CrossRef] [Green Version]
  62. Parchi, P.; Castellani, R.; Capellari, S.; Ghetti, B.; Young, K.; Chen, S.G.; Farlow, M.; Dickson, D.W.; Sima, A.A.; Trojanowski, J.Q.; et al. Molecular basis of phenotypic variability in sporadic Creutzfeldt-Jakob disease. Ann. Neurol. 1996, 39, 767–778. [Google Scholar] [CrossRef]
  63. Morales, R.; Abid, K.; Soto, C. The prion strain phenomenon: Molecular basis and unprecedented features. Biochim. Biophys. Acta 2007, 1772, 681–691. [Google Scholar] [CrossRef] [Green Version]
  64. Angers, R.C.; Kang, H.E.; Napier, D.; Browning, S.; Seward, T.; Mathiason, C.; Balachandran, A.; McKenzie, D.; Castilla, J.; Soto, C.; et al. Prion strain mutation determined by prion protein conformational compatibility and primary structure. Science 2010, 328, 1154–1158. [Google Scholar] [CrossRef] [Green Version]
  65. Crowell, J.; Hughson, A.; Caughey, B.; Bessen, R.A. Host Determinants of Prion Strain Diversity Independent of Prion Protein Genotype. J. Virol. 2015, 89, 10427–10441. [Google Scholar] [CrossRef] [Green Version]
  66. Duque Velasquez, C.; Kim, C.; Haldiman, T.; Kim, C.; Herbst, A.; Aiken, J.; Safar, J.G.; McKenzie, D. Chronic wasting disease (CWD) prion strains evolve via adaptive diversification of conformers in hosts expressing prion protein polymorphisms. J. Biol. Chem. 2020, 295, 4985–5001. [Google Scholar] [CrossRef] [Green Version]
  67. Moore, J.; Tatum, T.; Hwang, S.; Vrentas, C.; West Greenlee, M.H.; Kong, Q.; Nicholson, E.; Greenlee, J. Novel Strain of the Chronic Wasting Disease Agent Isolated From Experimentally Inoculated Elk With LL132 Prion Protein. Sci. Rep. 2020, 10, 3148. [Google Scholar] [CrossRef] [Green Version]
  68. Bian, J.; Christiansen, J.R.; Moreno, J.A.; Kane, S.J.; Khaychuk, V.; Gallegos, J.; Kim, S.; Telling, G.C. Primary structural differences at residue 226 of deer and elk PrP dictate selection of distinct CWD prion strains in gene-targeted mice. Proc. Natl. Acad. Sci. USA 2019, 116, 12478–12487. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  69. Hannaoui, S.; Triscott, E.; Duque Velasquez, C.; Chang, S.C.; Arifin, M.I.; Zemlyankina, I.; Tang, X.; Bollinger, T.; Wille, H.; McKenzie, D.; et al. New and distinct chronic wasting disease strains associated with cervid polymorphism at codon 116 of the Prnp gene. PLoS Pathog. 2021, 17, e1009795. [Google Scholar] [CrossRef] [PubMed]
  70. Arifin, M.I.; Hannaoui, S.; Chang, S.C.; Thapa, S.; Schatzl, H.M.; Gilch, S. Cervid Prion Protein Polymorphisms: Role in Chronic Wasting Disease Pathogenesis. Int. J. Mol. Sci. 2021, 22, 2271. [Google Scholar] [CrossRef] [PubMed]
  71. Moazami-Goudarzi, K.; Andreoletti, O.; Vilotte, J.L.; Beringue, V. Review on PRNP genetics and susceptibility to chronic wasting disease of Cervidae. Vet. Res. 2021, 52, 128. [Google Scholar] [CrossRef]
  72. Miller, M.W.; Wolfe, L.L.; Sirochman, T.M.; Sirochman, M.A.; Jewell, J.E.; Williams, E.S. Survival patterns in white-tailed and mule deer after oral inoculation with a standardized, conspecific prion dose. J. Wildl. Dis. 2012, 48, 526–529. [Google Scholar] [CrossRef]
  73. Johnson, C.J.; Herbst, A.; Duque-Velasquez, C.; Vanderloo, J.P.; Bochsler, P.; Chappell, R.; McKenzie, D. Prion protein polymorphisms affect chronic wasting disease progression. PLoS ONE 2011, 6, e17450. [Google Scholar] [CrossRef]
  74. Johnson, C.; Johnson, J.; Vanderloo, J.P.; Keane, D.; Aiken, J.M.; McKenzie, D. Prion protein polymorphisms in white-tailed deer influence susceptibility to chronic wasting disease. J. Gen. Virol. 2006, 87 Pt 7, 2109–2114. [Google Scholar] [CrossRef]
  75. Haley, N.J.; Merrett, K.; Buros Stein, A.; Simpson, D.; Carlson, A.; Mitchell, G.; Staskevicius, A.; Nichols, T.; Lehmkuhl, A.D.; Thomsen, B.V. Estimating relative CWD susceptibility and disease progression in farmed white-tailed deer with rare PRNP alleles. PLoS ONE 2019, 14, e0224342. [Google Scholar] [CrossRef] [Green Version]
  76. Jewell, J.E.; Conner, M.M.; Wolfe, L.L.; Miller, M.W.; Williams, E.S. Low frequency of PrP genotype 225SF among free-ranging mule deer (Odocoileus hemionus) with chronic wasting disease. J. Gen. Virol. 2005, 86 Pt 8, 2127–2134. [Google Scholar] [CrossRef]
  77. O’Rourke, K.I.; Besser, T.E.; Miller, M.W.; Cline, T.F.; Spraker, T.R.; Jenny, A.L.; Wild, M.A.; Zebarth, G.L.; Williams, E.S. PrP genotypes of captive and free-ranging Rocky Mountain elk (Cervus elaphus nelsoni) with chronic wasting disease. J. Gen. Virol. 1999, 80 Pt 10, 2765–2769. [Google Scholar] [CrossRef]
  78. Robinson, S.J.; Samuel, M.D.; Johnson, C.J.; Adams, M.; McKenzie, D.I. Emerging prion disease drives host selection in a wildlife population. Ecol. Appl. 2012, 22, 1050–1059. [Google Scholar] [CrossRef] [Green Version]
  79. Pirisinu, L.; Tran, L.; Chiappini, B.; Vanni, I.; Di Bari, M.A.; Vaccari, G.; Vikoren, T.; Madslien, K.I.; Vage, J.; Spraker, T.; et al. Novel Type of Chronic Wasting Disease Detected in Moose (Alces alces), Norway. Emerg. Infect. Dis. 2018, 24, 2210–2218. [Google Scholar] [CrossRef] [Green Version]
  80. Benestad, S.L.; Mitchell, G.; Simmons, M.; Ytrehus, B.; Vikoren, T. First case of chronic wasting disease in Europe in a Norwegian free-ranging reindeer. Vet. Res. 2016, 47, 88. [Google Scholar] [CrossRef]
  81. Vikoren, T.; Vage, J.; Madslien, K.I.; Roed, K.H.; Rolandsen, C.M.; Tran, L.; Hopp, P.; Veiberg, V.; Heum, M.; Moldal, T.; et al. First Detection of Chronic Wasting Disease in a Wild Red Deer (Cervus elaphus) in Europe. J. Wildl. Dis. 2019, 55, 970–972. [Google Scholar] [CrossRef]
  82. Nonno, R.; Di Bari, M.A.; Pirisinu, L.; D’Agostino, C.; Vanni, I.; Chiappini, B.; Marcon, S.; Riccardi, G.; Tran, L.; Vikoren, T.; et al. Studies in bank voles reveal strain differences between chronic wasting disease prions from Norway and North America. Proc. Natl. Acad. Sci. USA 2020, 117, 31417–31426. [Google Scholar] [CrossRef]
  83. Bian, J.; Kim, S.; Kane, S.J.; Crowell, J.; Sun, J.L.; Christiansen, J.; Saijo, E.; Moreno, J.A.; DiLisio, J.; Burnett, E.; et al. Adaptive selection of a prion strain conformer corresponding to established North American CWD during propagation of novel emergent Norwegian strains in mice expressing elk or deer prion protein. PLoS Pathog. 2021, 17, e1009748. [Google Scholar] [CrossRef]
  84. Barria, M.A.; Telling, G.C.; Gambetti, P.; Mastrianni, J.A.; Soto, C. Generation of a New Form of Human PrPSc in Vitro by Interspecies Transmission from Cervid Prions. J. Biol. Chem. 2011, 286, 7490–7495. [Google Scholar] [CrossRef] [Green Version]
  85. Pritzkow, S.; Gorski, D.; Ramirez, F.; Telling, G.C.; Benestad, S.L.; Soto, C. North American and Norwegian Chronic Wasting Disease prions exhibit different potential for interspecies transmission and zoonotic risk. J. Infect. Dis. 2021, 225, 542–551. [Google Scholar] [CrossRef]
  86. Moore, R.A.; Vorberg, I.; Priola, S.A. Species barriers in prion diseases-brief review. In Infectious Diseases from Nature: Mechanisms of Viral Emergence and Persistence; Springer: Berlin/Heidelberg, Germany, 2005; pp. 187–202. [Google Scholar]
  87. Kurt, T.D.; Sigurdson, C.J. Cross-species transmission of CWD prions. Prion 2016, 10, 83–91. [Google Scholar] [CrossRef] [Green Version]
  88. Moore, S.J.; Smith, J.D.; Richt, J.A.; Greenlee, J.J. Raccoons accumulate PrP(Sc) after intracranial inoculation of the agents of chronic wasting disease or transmissible mink encephalopathy but not atypical scrapie. J. Vet. Diagn. Investig. 2019, 31, 200–209. [Google Scholar] [CrossRef] [Green Version]
  89. Moore, S.J.; West Greenlee, M.H.; Kondru, N.; Manne, S.; Smith, J.D.; Kunkle, R.A.; Kanthasamy, A.; Greenlee, J.J. Experimental Transmission of the Chronic Wasting Disease Agent to Swine after Oral or Intracranial Inoculation. J. Virol. 2017, 91, e00926-17. [Google Scholar] [CrossRef] [Green Version]
  90. Greenlee, J.J.; Nicholson, E.M.; Smith, J.D.; Kunkle, R.A.; Hamir, A.N. Susceptibility of cattle to the agent of chronic wasting disease from elk after intracranial inoculation. J. Vet. Diagn. Investig. 2012, 24, 1087–1093. [Google Scholar] [CrossRef] [Green Version]
  91. Hamir, A.N.; Kunkle, R.A.; Cutlip, R.C.; Miller, J.M.; O’Rourke, K.I.; Williams, E.S.; Miller, M.W.; Stack, M.J.; Chaplin, M.J.; Richt, J.A. Experimental transmission of chronic wasting disease agent from mule deer to cattle by the intracerebral route. J. Vet. Diagn. Investig. 2005, 17, 276–281. [Google Scholar] [CrossRef] [Green Version]
  92. Hamir, A.N.; Kunkle, R.A.; Cutlip, R.C.; Miller, J.M.; Williams, E.S.; Richt, J.A. Transmission of chronic wasting disease of mule deer to Suffolk sheep following intracerebral inoculation. J. Vet. Diagn. Investig. 2006, 18, 558–565. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Sigurdson, C.J.; Mathiason, C.K.; Perrott, M.R.; Eliason, G.A.; Spraker, T.R.; Glatzel, M.; Manco, G.; Bartz, J.C.; Miller, M.W.; Hoover, E.A. Experimental chronic wasting disease (CWD) in the ferret. J. Comp. Pathol. 2008, 138, 189–196. [Google Scholar] [CrossRef] [PubMed]
  94. Williams, E.S.; Young, S. Spongiform encephalopathies in Cervidae. Rev. Sci. Tech. 1992, 11, 551–567. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Collinge, J. Variant Creutzfeldt-Jakob disease. Lancet 1999, 354, 317–323. [Google Scholar] [CrossRef]
  96. Kong, Q.; Huang, S.; Zou, W.; Vanegas, D.; Wang, M.; Wu, D.; Yuan, J.; Zheng, M.; Bai, H.; Deng, H.; et al. Chronic wasting disease of elk: Transmissibility to humans examined by transgenic mouse models. J. Neurosci. 2005, 25, 7944–7949. [Google Scholar] [CrossRef]
  97. Tamguney, G.; Giles, K.; Bouzamondo-Bernstein, E.; Bosque, P.J.; Miller, M.W.; Safar, J.; DeArmond, S.J.; Prusiner, S.B. Transmission of elk and deer prions to transgenic mice. J. Virol. 2006, 80, 9104–9114. [Google Scholar] [CrossRef] [Green Version]
  98. Sandberg, M.K.; Al-Doujaily, H.; Sigurdson, C.J.; Glatzel, M.; O’Malley, C.; Powell, C.; Asante, E.A.; Linehan, J.M.; Brandner, S.; Wadsworth, J.D.; et al. Chronic wasting disease prions are not transmissible to transgenic mice overexpressing human prion protein. J. Gen. Virol. 2010, 91 Pt 10, 2651–2657. [Google Scholar] [CrossRef]
  99. Wadsworth, J.D.F.; Joiner, S.; Linehan, J.M.; Jack, K.; Al-Doujaily, H.; Costa, H.; Ingold, T.; Taema, M.; Zhang, F.; Sandberg, M.K.; et al. Humanised transgenic mice are resistant to chronic wasting disease prions from Norwegian reindeer and moose. J. Infect. Dis. 2022; (In press). [Google Scholar] [CrossRef]
  100. Marsh, R.F.; Kincaid, A.E.; Bessen, R.A.; Bartz, J.C. Interspecies Transmission of Chronic Wasting Disease Prions to Squirrel Monkeys (Saimiri sciureus). J. Virol. 2005, 79, 13794–13796. [Google Scholar] [CrossRef] [Green Version]
  101. Race, B.; Meade-White, K.D.; Miller, M.W.; Barbian, K.D.; Rubenstein, R.; LaFauci, G.; Cervenakova, L.; Favara, C.; Gardner, D.; Long, D.; et al. Susceptibilities of nonhuman primates to chronic wasting disease. Emerg. Infect. Dis. 2009, 15, 1366–1376. [Google Scholar] [CrossRef]
  102. Race, B.; Williams, K.; Orru, C.D.; Hughson, A.G.; Lubke, L.; Chesebro, B. Lack of Transmission of Chronic Wasting Disease to Cynomolgus Macaques. J. Virol. 2018, 92, e00550-18. [Google Scholar] [CrossRef] [Green Version]
  103. Czub, S.; Schulz-Schaeffer, W.; Stahl-Hennig, C.; Beekes, M.; Schaetzl, H.; Motzkus, D. First evidence of intracranial and peroral transmission or chronic wasting disease (CWD) into Cynomolgus macaques: A work in progress. In Prion 2017 Deciphering Neurodegenerative Disorders; Taylor & Francis: Edingurgh, UK, 2017. [Google Scholar]
  104. Barria, M.A.; Libori, A.; Mitchell, G.; Head, M.W. Susceptibility of Human Prion Protein to Conversion by Chronic Wasting Disease Prions. Emerg. Infect. Dis. 2018, 24, 1482–1489. [Google Scholar] [CrossRef]
  105. Barria, M.A.; Balachandran, A.; Morita, M.; Kitamoto, T.; Barron, R.; Manson, J.; Knight, R.; Ironside, J.W.; Head, M.W. Molecular barriers to zoonotic transmission of prions. Emerg. Infect. Dis. 2014, 20, 88–97. [Google Scholar] [CrossRef] [Green Version]
  106. Davenport, K.A.; Henderson, D.M.; Bian, J.; Telling, G.C.; Mathiason, C.K.; Hoover, E.A. Insights into Chronic Wasting Disease and Bovine Spongiform Encephalopathy Species Barriers by Use of Real-Time Conversion. J. Virol. 2015, 89, 9524–9531. [Google Scholar] [CrossRef] [Green Version]
  107. Mawhinney, S.; Pape, W.J.; Forster, J.E.; Anderson, C.A.; Bosque, P.; Miller, M.W. Human prion disease and relative risk associated with chronic wasting disease. Emerg. Infect. Dis. 2006, 12, 1527. [Google Scholar] [CrossRef]
  108. Abrams, J.Y.; Maddox, R.A.; Harvey, A.R.; Schonberger, L.B.; Belay, E.D. Travel history, hunting, and venison consumption related to prion disease exposure, 2006–2007 FoodNet Population Survey. J. Am. Diet. Assoc. 2011, 111, 858–863. [Google Scholar] [CrossRef]
  109. Haley, N.; Donner, R.; Merrett, K.; Miller, M.; Senior, K. Selective Breeding for Disease-Resistant PRNP Variants to Manage Chronic Wasting Disease in Farmed Whitetail Deer. Genes 2021, 12, 1396. [Google Scholar] [CrossRef]
  110. Buschmann, A.; Groschup, M.H. TSE eradication in small ruminants--quo vadis? Berl. Munch. Tierarztl. Wochenschr. 2005, 118, 365–371. [Google Scholar]
  111. Bueler, H.; Aguzzi, A.; Sailer, A.; Greiner, R.A.; Autenried, P.; Aguet, M.; Weissmann, C. Mice devoid of PrP are resistant to scrapie. Cell 1993, 73, 1339–1347. [Google Scholar] [CrossRef]
  112. Richt, J.A.; Kasinathan, P.; Hamir, A.N.; Castilla, J.; Sathiyaseelan, T.; Vargas, F.; Sathiyaseelan, J.; Wu, H.; Matsushita, H.; Koster, J.; et al. Production of cattle lacking prion protein. Nat. Biotechnol. 2007, 25, 132–138. [Google Scholar] [CrossRef]
  113. Haley, N. Amplification Techniques for the Detection of Misfolded Prion Proteins in Experimental and Clinical Samples. Curr. Protoc. Mol. Biol. 2020, 130, e118. [Google Scholar] [CrossRef] [Green Version]
  114. Erana, H.; Charco, J.M.; Gonzalez-Miranda, E.; Garcia-Martinez, S.; Lopez-Moreno, R.; Perez-Castro, M.A.; Diaz-Dominguez, C.M.; Garcia-Salvador, A.; Castilla, J. Detection of Pathognomonic Biomarker PrP(Sc) and the Contribution of Cell Free-Amplification Techniques to the Diagnosis of Prion Diseases. Biomolecules 2020, 10, 469. [Google Scholar] [CrossRef] [Green Version]
  115. McNulty, E.; Nalls, A.V.; Mellentine, S.; Hughes, E.; Pulscher, L.; Hoover, E.A.; Mathiason, C.K. Comparison of conventional, amplification and bio-assay detection methods for a chronic wasting disease inoculum pool. PLoS ONE 2019, 14, e0216621. [Google Scholar] [CrossRef] [Green Version]
  116. Brandel, J.P.; Culeux, A.; Grznarova, K.; Levavasseur, E.; Lamy, P.; Privat, N.; Welaratne, A.; Denouel, A.; Laplanche, J.L.; Haik, S. Amplification techniques and diagnosis of prion diseases. Rev. Neurol. 2019, 175, 458–463. [Google Scholar] [CrossRef]
  117. Haley, N.J.; Richt, J.A. Evolution of Diagnostic Tests for Chronic Wasting Disease, a Naturally Occurring Prion Disease of Cervids. Pathogens 2017, 6, 35. [Google Scholar] [CrossRef] [Green Version]
  118. Morales, R.; Duran-Aniotz, C.; Diaz-Espinoza, R.; Camacho, M.V.; Soto, C. Protein misfolding cyclic amplification of infectious prions. Nat. Protoc. 2012, 7, 1397–1409. [Google Scholar] [CrossRef]
  119. Kramm, C.; Soto, P.; Nichols, T.A.; Morales, R. Chronic wasting disease (CWD) prion detection in blood from pre-symptomatic white-tailed deer harboring PRNP polymorphic variants. Sci. Rep. 2020, 10, 19763. [Google Scholar] [CrossRef]
  120. McNulty, E.E.; Nalls, A.V.; Xun, R.; Denkers, N.D.; Hoover, E.A.; Mathiason, C.K. In vitro detection of haematogenous prions in white-tailed deer orally dosed with low concentrations of chronic wasting disease. J. Gen. Virol. 2020, 101, 347–361. [Google Scholar] [CrossRef]
  121. Cheng, Y.C.; Hannaoui, S.; John, T.R.; Dudas, S.; Czub, S.; Gilch, S. Real-time Quaking-induced Conversion Assay for Detection of CWD Prions in Fecal Material. J. Vis. Exp. 2017, 29, 56373. [Google Scholar] [CrossRef]
  122. Elder, A.M.; Henderson, D.M.; Nalls, A.V.; Wilham, J.M.; Caughey, B.W.; Hoover, E.A.; Kincaid, A.E.; Bartz, J.C.; Mathiason, C.K. In vitro detection of prionemia in TSE-infected cervids and hamsters. PLoS ONE 2013, 8, e80203. [Google Scholar] [CrossRef] [Green Version]
  123. Uehlinger, F.D.; Johnston, A.C.; Bollinger, T.K.; Waldner, C.L. Systematic review of management strategies to control chronic wasting disease in wild deer populations in North America. BMC Vet. Res. 2016, 12, 173. [Google Scholar] [CrossRef] [PubMed]
  124. Manjerovic, M.B.; Green, M.L.; Mateus-Pinilla, N.; Novakofski, J. The importance of localized culling in stabilizing chronic wasting disease prevalence in white-tailed deer populations. Prev. Vet. Med. 2014, 113, 139–145. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Wolfe, L.L.; Watry, M.K.; Sirochman, M.A.; Sirochman, T.M.; Miller, M.W. Evaluation of a Test and Cull Strategy for Reducing Prevalence of Chronic Wasting Disease in Mule Deer (Odocoileus Hemionus). J. Wildl. Dis. 2018, 54, 511–519. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  126. Haley, N.J.; Henderson, D.M.; Donner, R.; Wyckoff, S.; Merrett, K.; Tennant, J.; Hoover, E.A.; Love, D.; Kline, E.; Lehmkuhl, A.D.; et al. Management of chronic wasting disease in ranched elk: Conclusions from a longitudinal three-year study. Prion 2020, 14, 76–87. [Google Scholar] [CrossRef] [Green Version]
  127. Mysterud, A.; Rolandsen, C.M. A reindeer cull to prevent chronic wasting disease in Europe. Nat. Ecol. Evol. 2018, 2, 1343–1345. [Google Scholar] [CrossRef]
  128. Taylor, D.M. Inactivation of transmissible degenerative encephalopathy agents: A review. Vet. J. 2000, 159, 10–17. [Google Scholar] [CrossRef]
  129. Secker, T.J.; Leighton, T.G.; Offin, D.G.; Birkin, P.R.; Herve, R.C.; Keevil, C.W. A cold water, ultrasonically activated stream efficiently removes proteins and prion-associated amyloid from surgical stainless steel. J. Hosp. Infect. 2020, 106, 649–656. [Google Scholar] [CrossRef]
  130. McDonnell, G.; Dehen, C.; Perrin, A.; Thomas, V.; Igel-Egalon, A.; Burke, P.A.; Deslys, J.P.; Comoy, E. Cleaning, disinfection and sterilization of surface prion contamination. J. Hosp. Infect. 2013, 85, 268–273. [Google Scholar] [CrossRef]
  131. Hirata, Y.; Ito, H.; Furuta, T.; Ikuta, K.; Sakudo, A. Degradation and destabilization of abnormal prion protein using alkaline detergents and proteases. Int. J. Mol. Med. 2010, 25, 267–270. [Google Scholar]
  132. Suyama, K.; Yoshioka, M.; Akagawa, M.; Murayama, Y.; Horii, H.; Takata, M.; Yokoyama, T.; Mohri, S. Assessment of prion inactivation by fenton reaction using protein misfolding cyclic amplification and bioassay. Biosci. Biotechnol. Biochem. 2007, 71, 2069–2071. [Google Scholar] [CrossRef]
  133. Suyama, K.; Yoshioka, M.; Akagawa, M.; Murayama, Y.; Horii, H.; Takata, M.; Yokoyama, T.; Mohri, S. Prion inactivation by the Maillard reaction. Biochem. Biophys. Res. Commun. 2007, 356, 245–248. [Google Scholar] [CrossRef]
  134. Yoshioka, M.; Miwa, T.; Horii, H.; Takata, M.; Yokoyama, T.; Nishizawa, K.; Watanabe, M.; Shinagawa, M.; Murayama, Y. Characterization of a proteolytic enzyme derived from a Bacillus strain that effectively degrades prion protein. J. Appl. Microbiol. 2007, 102, 509–515. [Google Scholar] [CrossRef]
  135. Belondrade, M.; Jas-Duval, C.; Nicot, S.; Bruyere-Ostells, L.; Mayran, C.; Herzog, L.; Reine, F.; Torres, J.M.; Fournier-Wirth, C.; Beringue, V.; et al. Correlation between Bioassay and Protein Misfolding Cyclic Amplification for Variant Creutzfeldt-Jakob Disease Decontamination Studies. mSphere 2020, 5, e00649-19. [Google Scholar] [CrossRef] [Green Version]
  136. Belondrade, M.; Nicot, S.; Beringue, V.; Coste, J.; Lehmann, S.; Bougard, D. Rapid and Highly Sensitive Detection of Variant Creutzfeldt-Jakob Disease Abnormal Prion Protein on Steel Surfaces by Protein Misfolding Cyclic Amplification: Application to Prion Decontamination Studies. PLoS ONE 2016, 11, e0146833. [Google Scholar] [CrossRef] [Green Version]
  137. Moudjou, M.; Castille, J.; Passet, B.; Herzog, L.; Reine, F.; Vilotte, J.L.; Rezaei, H.; Beringue, V.; Igel-Egalon, A. Improving the Predictive Value of Prion Inactivation Validation Methods to Minimize the Risks of Iatrogenic Transmission With Medical Instruments. Front. Bioeng. Biotechnol. 2020, 8, 591024. [Google Scholar] [CrossRef]
  138. Pritzkow, S.; Wagenfuhr, K.; Daus, M.L.; Boerner, S.; Lemmer, K.; Thomzig, A.; Mielke, M.; Beekes, M. Quantitative detection and biological propagation of scrapie seeding activity in vitro facilitate use of prions as model pathogens for disinfection. PLoS ONE 2011, 6, e20384. [Google Scholar] [CrossRef]
  139. Zattoni, M.; Legname, G. Tackling prion diseases: A review of the patent landscape. Expert Opin. Ther. Pat. 2021, 31, 1097–1115. [Google Scholar] [CrossRef]
  140. Sim, V.L.; Caughey, B. Recent advances in prion chemotherapeutics. Infect. Disord. Drug Targets 2009, 9, 81–91. [Google Scholar] [CrossRef] [Green Version]
  141. Sassoon, J.; Sadowski, M.; Wisniewski, T.; Brown, D.R. Therapeutics and prion disease: Can immunisation or drugs be effective? Mini. Rev. Med. Chem. 2005, 5, 361–366. [Google Scholar] [CrossRef]
  142. Goni, F.; Mathiason, C.K.; Yim, L.; Wong, K.; Hayes-Klug, J.; Nalls, A.; Peyser, D.; Estevez, V.; Denkers, N.; Xu, J.; et al. Mucosal immunization with an attenuated Salmonella vaccine partially protects white-tailed deer from chronic wasting disease. Vaccine 2015, 33, 726–733. [Google Scholar] [CrossRef] [Green Version]
  143. Wood, M.E.; Griebel, P.; Huizenga, M.L.; Lockwood, S.; Hansen, C.; Potter, A.; Cashman, N.; Mapletoft, J.W.; Napper, S. Accelerated onset of chronic wasting disease in elk (Cervus canadensis) vaccinated with a PrP(Sc)-specific vaccine and housed in a prion contaminated environment. Vaccine 2018, 36, 7737–7743. [Google Scholar] [CrossRef]
  144. Saunders, S.E.; Bartelt-Hunt, S.L.; Bartz, J.C. Prions in the environment: Occurrence, fate and mitigation. Prion 2008, 2, 162–169. [Google Scholar] [CrossRef] [Green Version]
Viruses 14 01390 g001 550
Figure 1. Schematic representation of the different hypotheses to explain the CWD origin. Although the origin of CWD is largely unknown, various hypotheses have been proposed, including transmission from a scrapie-infected sheep, a mutation in the Prnp gene (illustrated as a star in the left panel) and a spontaneous misfolding of PrPC into PrPSc.
Figure 1. Schematic representation of the different hypotheses to explain the CWD origin. Although the origin of CWD is largely unknown, various hypotheses have been proposed, including transmission from a scrapie-infected sheep, a mutation in the Prnp gene (illustrated as a star in the left panel) and a spontaneous misfolding of PrPC into PrPSc.
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Figure 2. Animal species susceptible or at risk to be infected with CWD. Various animals cohabit with cervid, including livestock, rodents, scavengers, carnivores, domestic animals, and humans. Several studies of experimental infection of animals by CWD have shown that sheep, goats, cattle, ferrets, raccoons, and rodents can be infected with CWD [88,89,90,91,92,93,94].
Figure 2. Animal species susceptible or at risk to be infected with CWD. Various animals cohabit with cervid, including livestock, rodents, scavengers, carnivores, domestic animals, and humans. Several studies of experimental infection of animals by CWD have shown that sheep, goats, cattle, ferrets, raccoons, and rodents can be infected with CWD [88,89,90,91,92,93,94].
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Figure 3. An overview of various strategies to minimize CWD spreading. Various alternative approaches have been proposed to decrease the risk of further CWD spreading, including detailed surveillance, selective breeding of animals harboring rare polymorphisms partially resistant to CWD, development of a highly efficient test for CWD detection, selective culling of infected animals, prion decontamination, and development of a treatment for CWD.
Figure 3. An overview of various strategies to minimize CWD spreading. Various alternative approaches have been proposed to decrease the risk of further CWD spreading, including detailed surveillance, selective breeding of animals harboring rare polymorphisms partially resistant to CWD, development of a highly efficient test for CWD detection, selective culling of infected animals, prion decontamination, and development of a treatment for CWD.
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Table
Table 1. Natural Prnp polymorphisms found in various species of cervids.
Table 1. Natural Prnp polymorphisms found in various species of cervids.
Cervid SpeciesPolymorphisms
White-tailed deer (Odocoileus virginianus)G37V, G96S, G96R, A123T, Q230L
Mule deer (Odocoileus hemionus)D20G, S225F, Q226K
Elk (Cervus canadensis)M132L, E226
Reindeer (Rangifer tarandus)V2M, Del 84-91, G129S, S138N, Y153F, V169M, N176D, S225Y, P242L
Red deer (Cervus elaphus)G59S, T98A, P168S, M208I, Q226E, I247L
Moose (Alces alces)T36N, S100R, K109Q, M209I
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Pritzkow, S. Transmission, Strain Diversity, and Zoonotic Potential of Chronic Wasting Disease. Viruses 2022, 14, 1390. https://doi.org/10.3390/v14071390

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Pritzkow, Sandra. 2022. "Transmission, Strain Diversity, and Zoonotic Potential of Chronic Wasting Disease" Viruses 14, no. 7: 1390. https://doi.org/10.3390/v14071390

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Pritzkow, S. (2022). Transmission, Strain Diversity, and Zoonotic Potential of Chronic Wasting Disease. Viruses, 14(7), 1390. https://doi.org/10.3390/v14071390

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