Detection of chronic wasting disease prions in soil at an illegal white-tailed deer carcass disposal site

ABSTRACT

Chronic wasting disease (CWD) is a contagious prion disorder affecting cervids such as deer, elk, caribou, and moose, causing progressive and severe neurological degeneration followed by eventual death. As CWD prions (PrP^Sc) accumulate in the body, they are shed through excreta and secreta, as well as through decomposing carcasses. Prions can persist in the environment for years, posing significant concerns for ongoing transmission to susceptible cervids and pose an unknown risk to sympatric species. We used a validated protocol for real-time quaking-induced conversion (RT-QuIC) in vitro prion amplification assay to detect prions in soil collected within and around an illegal white-tailed deer (Odocoileus virginianus, WTD) carcass disposal site and associated captive WTD farm in Beltrami County, Minnesota. We detected PrP^Sc in 26 of 201 soil samples across 15 locations within the illegal disposal site and one on the farm that housed the cervids. Importantly, a subset of RT-QuIC positive soil samples was collected from soils where carcasses were recovered, providing direct evidence that environmental contamination resulted from this illegal activity. These findings reveal that improper cervid carcass disposal practices may have important implications for ongoing CWD transmission through the environment.

Introduction

Chronic wasting disease (CWD) is a transmissible prion disease affecting cervids such as deer, elk, caribou, and moose, causing progressive and severe neurological degeneration followed by eventual death [1–3]. Cervids infected with CWD prions experience abnormal folding of the host encoded prion protein (PrP^C), leading to the accumulation of infectious prions (PrP^Sc) in the central nervous system and elsewhere in the body [4–7]. Specifically, when PrP^Sc come into contact with healthy PrP^C they convert PrP^C into the lethal and contagious PrP^Sc isoform [6,7]. As CWD prions accumulate in the body, they are externally shed in excreta and secreta such as saliva, blood, urine, and faeces, as well as during carcass decomposition [8–11].

While CWD has primarily been studied in the context of animal health and direct disease transmission [4,7,12], numerous studies over the past twenty years have raised concerns about the possible indirect transmission of CWD prions through exposure to prion-contaminated environments [13–16]. One area of environmental concern is the contamination of soils with PrP^Sc, as soils may serve as a reservoir for persistent prion transmission in cervids [14,17–19]. It is well documented that soils can bind and adsorb infectious prions, as well as retain infectivity, raising questions about the influence of soil composition, infectious dose, persistence, and long-term bioavailability [20–24].

Indeed, PrP^Sc can bind to soils, and some studies show it remains infectious when adsorbed to specific soil types and environmental surfaces [21,25–27]. However, the precise impact of soil binding on prions remains unclear, as studies report differing impacts on both infectivity and bioavailability of mineral-bound prions [16,24,28–30]. Thus, understanding the dynamics of PrP^Sc contamination in the environment, including soils and waters, is crucial for developing effective management strategies and mitigating further spread of CWD [23,31,32].

PrP^Sc likely persists in small concentrations in environmental matrices, such as soils and waters, necessitating the use of protein amplification assays for detection. To address this, researchers have employed methods to amplify exceedingly small PrP^Sc concentrations, such as real-time quaking-induced conversion (RT-QuIC) in vitro prion amplification assay [33,34] and protein misfolding conversion assay [35–37]. Previous studies [14,38–40] have demonstrated the utility of prion amplification assays for prion detection in a variety of environmental sample types, such as soils. However, assay performance can vary considerably and requires predefining sample behaviour to understand that variation [40]. In particular, a myriad of soil characteristics, including minerals, textures, as well as clay and organic matter contents and types, can interact with prion detection methods in complex ways [21,22,39]. These interactions may interfere with detection methods, making assay interpretation challenging [18,24,39].

Here, we describe the use of RT-QuIC in vitro prion amplification assay for the detection of prion contamination of native Minnesota soils. We employed a method of RT-QuIC [41] optimized to reduce false positive reactions, providing a more rigorous analysis of soil samples for the presence of prions. We applied the optimized method to screen for PrP^Sc contamination within an illegal WTD carcass disposal site in Beltrami County, Minnesota, where we previously recovered remains from PrP^Sc-positive WTD carcasses on this publicly-owned property [10]. Given the history of PrP^Sc-positive carcass decomposition at this site, we aimed to characterize the extent of contamination in the associated soils.

Results

Field site investigation

Following full quarantine and depopulation of a captive WTD facility in Beltrami County, Minnesota, state and federal officials determined that the owner had disposed WTD carcasses from the facility approximately 1.5 km away at a site on publicly-owned, tax-forfeited property [10], herein the study site (Figure 1). Upon discovery, the Minnesota Department of Natural Resources secured the approximately 6-hectare study site area with 914.4 metres of 3.048 metre-high woven wire fencing [10]. Approximately 90% of the disposed carcass material had been recovered from 2.4 hectares of the site.

Figure 1.

Geographic location where CWD-positive white-tailed deer (Odocoileus virginianus) carcasses were disposed, along with the captive facility of origin, in Beltrami County, Minnesota. The inset map shows the study site location within Minnesota. Map lines delineate study areas and do not necessarily depict accepted national boundaries.

In 2021, our team conducted four site visits in May, June, July, and September to collect a total of 201 soil samples. Within the study site, WTD carcass remains were spread across 44 locations consisting of nine adult, three yearling, 19 fawn, and 13 of unknown age [10] Figure 1). When gathering animal remains for forensics investigation in May and June of 2021, we collected an initial set of 28 soil samples ~2 cm deep directly beneath animal carcass remains and marked these locations with metal and wooden posts, flagging tape, and GPS coordinates [10].

In July and September 2021, we established 11 soil sampling transects at locations with WTD remains with previously detected PrP^Sc within the study site [10]. Transects were designed to consist of a central sample location surrounded by sample locations at one- and two-metre distances in all cardinal directions, totalling nine sampling points for 2 cm-deep surface soils (Figure 2). However, there were some instances where all nine samples within a transect could not be collected. Three additional transects were created within the study site that were not directly associated with PrP^Sc-positive carcass material. One of these transects was directly associated with remains that were previously tested and PrP^Sc was not detected [10]. However, those remains had been genetically determined to have originated from the same animal as other remains recovered from the study site that had tested PrP^Sc-positive. Another transect was located at the collection site of an equine carcass, which had previously tested negative for PrP^Sc. We established the final transect > 26 metres away from where carcass materials had been previously recovered. Thus, we collected 96 soil samples at all accessible transect locations within the study site (n = 14), in addition to the 28 samples collected earlier in the summer at point locations below recovered carcass materials.

Figure 2.

The transect configuration for collecting soils associated with recovered carcass materials at a white-tailed deer carcass disposal site. Transects consisted of a central sample location surrounded by sample locations at 1 and 2 metre distances in all cardinal directions (i.e. Center, N1, N2, S1, S2, E1, E2, W1, and W2; n = 9). Figure diagram created by Roxanne Larsen (modified from [42].

Outside of the study site, we collected 15 soil samples from five point locations immediately surrounding the study site perimeter, 33 soil samples from 11 locations within the WTD captive facility, as well as 29 sediment and soil samples from nine locations spanning a waterway that ran directly through the WTD captive facility. For all sampling conducted outside of the study site, we did not establish long-term sampling transects, but instead collected three individual 2 cm-deep soil samples from each point location. We handled 11 blank quartz sand samples as a field control during perimeter and off-site soil sampling.

Approximately 1.5–2 years after field collection, we thawed, dried, and extracted each study site soil sample following an optimized RT-QuIC protocol for PrP^Sc detection in soils [41]. Using RT-QuIC, we determined that seven of the 11 sites (63.6%) with previously documented PrP^Sc-positive WTD remains, also had PrP^Sc detections in at least one soil sample from its corresponding soil transect (Table 1, Figure 3). Across all 201 soil samples for PrP^Sc analysis, 26 contained PrP^Sc as indicated by statistically significant prion seeding activity (Table 1). Of those PrP^Sc detections, 25 were located within the study site, and one was not. Seventeen of the 25 PrP^Sc detections in study site soil samples, or 68%, were located across seven transects that had previously demonstrated statistically significant prion seeding activity in biological samples from WTD remains. The other eight study site soil samples positive for PrP^Sc were among 21 soil samples (38.09%) collected across three soil transects and five point source sites where PrP^Sc had not been previously detected in WTD remains. We also detected PrP^Sc in one soil sample collected outside of the study site. This sample (identified as MPF 2) was collected near a deer feeder at the centre of a central deer pen within the captive WTD facility.

Table 1.

RT-QuIC results of soil samples collected from a site where CWD-positive white-tailed deer (Odocoileus virginianus, WTD) had been disposed and the associated captive WTD facility in Beltrami County, Minnesota [10]. Sampling occurred through either 1–3 samples at a single point location or transect collection design (with nine samples per transect) at each sampling location, as indicated. Statistical significance is represented by one to three stars indicative of p-value strength, where we represented p-values <0.05 with one star (*), p-values <0.01 with 2 stars (**), and p-values <0.001 with three stars (***). Results of associated WTD biological remains were reported as identified in [10]. We collected all samples testing positive from the site of carcass disposal, with the exception of one (MPF 2) that we collected from the captive facility. Shading of coloured rows indicates associated remains were genetically-unique WTD individuals (n = 4); uncoloured rows depict sites where genetic testing data were unavailable. ^aCWD and genetic testing of carcass remains described in [10].

Figure 3.

Geographical distribution of soil sampling for PrP^Sc detection at a site where CWD-positive white-tailed deer (Odocoileus virginianus, WTD) carcasses were disposed of in Beltrami County, Minnesota. Sample ID labels are provided for each soil sampling location. The map also depicts the locations of WTD remains previously collected and tested as described in [10]. Map lines delineate study areas and do not necessarily depict accepted national boundaries.

Discussion

In this study, we document how disposal of CWD-infected cervids can establish an environmental prion reservoir in soils. Our results show that disposal of PrP^Sc-positive WTD [10] contributed to dispersed environmental PrP^Sc contamination across the study site. The initial discovery of CWD-infected carcasses on the landscape [10] resulted in an opportunity to investigate if this above-ground carcass disposal practice could lead to the establishment of environmental CWD prions following infections in a captive herd. We documented PrP^Sc contamination of soils in the study area by utilizing an optimized [41] RT-QuIC assay protocol for the detection of prions in the soil types found at the location. Specifically, we detected PrP^Sc in 26 soil samples across 15 sampling locations within the study site and one sampling location from the CWD-positive WTD captive facility. While some of the CWD-positive sampling locations were directly associated with CWD-positive carcasses, others were not (Table 1). We suspect that the detections unassociated with CWD-positive carcass material may be due to local dissemination by scavengers [43–45], soil erosion, surface water transport [46], or the limitations of RT-QuIC to detect PrP^Sc in all sample types [10]. Importantly, we did not detect PrP^Sc in soil samples collected immediately outside of the study site fence enclosure (i.e. perimeter samples from locations PER001-PER005; Figure 3), areas characterized by the same soil orders and land cover as within our study site. This observation increases the rigour of the findings of RT-QuIC testing of soil samples from within the study site and farm enclosure and suggests that prions, at this time post environmental exposure, have not spread to the local surrounding area. Although others have shown RT-QuIC’s utility in detecting PrP^Sc in other environmental sample types or in combination with PMCA for soils [38,40,44,47], this is the first report applying an optimized soil extraction protocol for prion detection by RT-QuIC with novelty in its application to understand point source contamination associated with decomposing WTD carcass material.

In analysing point source contamination, it is worth noting that we detected prions in soils more frequently at the study site where carcasses had been disposed than at the WTD captive facility where the infected animals had originated. This finding was surprising given infected carcasses had also been discovered at and removed from the captive cervid facility at the time of depopulation [10]. In contrast, we did not know the locations at the facility where the carcasses had been found, thus could not directly sample and test soil from those locations and instead targeted areas for soil sampling where animals were known to congregate at the facility. Thus, the differences in prion detection frequency between these two locations provides some insight about the relative contribution of excreta/secreta versus carcasses for establishment of environmental prions. This may be important for new geographical areas of CWD invasion, where removal of infected carcasses may have a greater impact on mitigating the establishment of environmental prions compared to areas that have a long history of endemic CWD. The role of soil composition and erosion in environmental prion persistence and detection also warrants consideration. For instance, the animal enclosures at the captive facility likely experienced higher levels of animal activity and reduced vegetative cover compared to the study area, leading to increased soil erosion. This erosion may have contributed to heightened prion transport via movement of excreta, secreta, and soil particles into a water channel that traversed the enclosures. Consequently, prion distribution and transport in the environment may have differed significantly between these locations due to variations in soil erosion and water-mediated transport.

Distribution of detections across and within transects

Within the established perimeter of the study site, we observed epidemiologically-linked patterns of PrP^Sc detection. For example, five soil transects exhibited multiple PrP^Sc detections. These were concentrated within an area of only 3000 m² in the west-central area of the study site (Figure 3). This area not only contained much of the WTD carcass material previously recovered from the site [10], but also contained more localized areas of PrP^Sc detections in both carcass materials and soils (Figure 3). In particular, three transects (PL13, PL18, and GR09) within approximately 630 m² of this area contained 44% of the positive soil detections. Limited DNA recovery and genetic analysis in a previous study demonstrated that these particular sites contained biological materials from distinct individuals [10]. Regardless, the numerous PrP^Sc detections across multiple soil transects in this area suggest a region of high prion contamination, as might be expected with point source contamination.

In addition to the localization of CWD-positive carcass materials, landscape slope may also be associated with the distribution of positive soil detections (Figure 3). For instance, the elevation gradually decreased from northwest to southeast across the area where carcasses were deposited within the study site. Across all positive transects, detections in the southern arm of transects were most frequent (n = 7 versus n = 2–4 in the other sampling directions or centre). Looking more closely at a transect located on a more extreme slope, we detected PrP^Sc in soil sample 013-S1, where positive biological remains were recovered at the transect centre [10]. The location of soil sample 013-S1 was unique due to a notable 0.3 m drop in elevation over a 1 m distance from the transect centre towards the southwest, a relatively steep slope for this area. We observed a similar pattern with soil transect 028, where PrP^Sc was detected in soil samples 028-S2 and 028-N2, two sample locations along the transect that are approximately 0.3 m lower in elevation from the transect centre, where positive carcass material was previously recovered. While we did not design our study with hypotheses about the distribution of prion contamination and slope of the landscape, these patterns warrant further investigation. Because scavengers had also distributed carcass materials widely across the site prior to investigation [10] and they can also disseminate prions further in their faeces [43–45], it is difficult to draw clear conclusions about the role of biotic and abiotic processes on prion distribution from our work.

Also notable among our findings was the detection of PrP^Sc in soil samples from three locations (GR06, PL22, PL21) where PrP^Sc had not been detected previously in associated carcass materials. This could point to the limitations of RT-QuIC related either to sensitivity of PrP^Sc detection in substantially decomposed and aged carcass materials [10] or specificity associated with false seeding by these soil types. While we applied stringent criteria to the analysis of detection, we cannot exclude the latter possibility. However, further examination of these sites provided some epidemiological and environmental insights that may explain these findings. For instance, the carcass material recovered from one of these locations (GR06), while negative for PrP^Sc by RT-QuIC, was genetically linked to PrP^Sc-positive animal remains recovered from a different nearby location (GR05A). Soil samples from the transect directly associated with the PrP^Sc-positive carcass material (GR05A; located 4 m from GR06), however, contained no PrP^Sc detections. No other genetic links had been made between carcass materials recovered across the sites examined in this study; carcass materials originally associated with four PrP^Sc-positive soil sites were genetically unique (Table 1 [10]. Additionally, PL22 was a transect site located approximately 26 m from the nearest site of recovered carcass material and soil samples with PrP^Sc detections, and PL21 was a site where equine carcass material had been recovered. Both sites were at lower elevations from the concentrated area of carcass deposition and PrP^Sc detections, and also along surface water flow pathways, observed to be areas of concentrated overland flow, to intermittent stream channels and wetlands. Hence, one possible mechanism is physical and water-mediated movement downhill. Unfortunately, we did not know the locations where CWD-positive animals were originally deposited, and these detection patterns could be influenced by multiple factors, such as assay performance, movement of PrP^Sc or PrP^Sc-containing carcass materials by scavengers, and/or physical downslope prion transport via soils, waters, and sediments.

Study limitations

There was a paucity of literature related to the interaction or testing of prions in the soil types found in our study area, yet previous work suggested that the clay (5.5–13%) and organic matter (3–35%) content found in these soil types [41] might not only play a role in prion binding and infectivity [16,24,28–30], but also affect detection in complex ways [18,21,22,39]. Thus, we made substantial effort to understand how these native soil types would interfere with RT-QuIC in the presence and absence of prion and to optimize the analytical pipeline for detection [41]. However, experimental controls revealed some challenges remained. For instance, even after optimization of both the soil extraction protocol and data analysis, we still observed some degree of false seeding in negative soil controls. This necessitated more stringent criteria for the determination of prion detection in a sample using the RT-QuIC output. While that approach increased our confidence in determinations of prion detection in samples, we also recognized there may be subsequent trade-off in detection sensitivity. Thus, it is possible that soil samples with low concentrations of prions (e.g., below 10⁻³ [41], produced inconsistent seeding activity in RT-QuIC, which then did not meet our analytical criteria for detection. However, among those samples determined positive for PrP^Sc by our criteria, we observed similar patterns of amyloid formation in RT-QuIC as we observed with positive tissue controls and previously prion-inoculated soils (Figure S1), giving confidence to positive detections. Nevertheless, given the inherent challenges in prion detection associated with these soil types, there is a need for further research into the specific soil properties that incite false seeding in RT-QuIC, assay optimization to mitigate their occurrence, and validation of alternative methodologies that provide additional evidence of prion detection and infectivity.

During field sampling, we included handling of field blanks containing quartz sand, one of which had been handled at the southern perimeter of the study site and also displayed statistically significant seeding activity in RT-QuIC. Given that this sample type consisted of only pure quartz sand instead of compositions of clays and organic matter, these samples shouldn’t have been at risk for false seeding like the native soils. The presence of seeding activity in the pure quartz sand sample was unexpected, and suggested contamination at the field site or within the laboratory. Given we did not see evidence of contamination across other experimental controls (field and laboratory), we do not believe that artificial contamination can account for the numerous detections observed across soil samples from the site. While it is unclear at which stage of the sampling and testing process cross-contamination occurred, the event demonstrates the need for careful attention to sample handling practices, particularly given the ‘sticky’ nature of prions and their affinity for binding to dust and surfaces [48,49].

Future considerations

We did not design our study to test hypotheses about the influence of environmental or biological processes on local prion transport and detection patterns, though our study has raised several questions regarding these topics. Given the dissemination of carcass materials around the study site, scavenger activity was evident [10], and they could play a role in further dissemination of prions across the site or locally [43–45]. We did not collect scat from scavengers or other local fauna (e.g., invertebrates [50], that might interact with the local environment in various ways. However, this is an area of research that is needed to further understand the biological processes that may impact prion persistence and/or transport over time, particularly in places where environmental prion contamination has occurred.

The topography of our site was also characterized by changes in elevation, where carcass materials were concentrated on an upland point. Our site experienced seasonal surface water flow upon snow melt and seasonally large rain events, thus it is also possible that prion transport via surface water and/or hillslope erosion may have occurred [51,52]. Prions were detected in sediments from a small intermittent stream channel exiting the study site a year after carcass removal [46], further supporting the idea that water may play a role in the storage and transport of prions. Given the persistence of prions over weeks to years in aquatic environments [26,32,53], the hydrological transport of prions may contribute to further dispersal of environmentally-contaminating prions in ways previously unrecognized. Thus, more research is needed to understand environmental dispersal and the hydrological fate of prions over time.

Conclusions

Using RT-QuIC, we detected PrP^Sc in 26 of 201 soil samples collected across 16 locations on public land where WTD carcasses had been disposed and the captive facility from where they originated. Within the disposal site, 25 out of 124 soil samples (20%) tested positive for PrP^Sc. Among those positive detections, 17, or 68%, were collected from locations where CWD-positive WTD remains had been previously recovered. This environmental investigation demonstrates how improper cervid carcass disposal practices can result in persistent environmental contamination, posing a potential risk to wildlife health. Given that disposal of livestock on the landscape is a common practice among producers [54–56], these findings underscore the need for improved disposal practices and further investigation of environmental impacts. Expanding on this area of environmental research is crucial as the geographic range of CWD continues to expand [57]. The use of RT-QuIC for prion detection in environmental samples offers an exciting advancement to environmental surveillance for prions, though as we demonstrate here and in Grunklee et al. [41], assay optimization and validation for use with different environmental samples, including new soil types, is still necessary. Further enhancements to RT-QuIC and other methodologies for prion detection will facilitate more opportunities to explore the persistence, degradation, transport, and remediation of environmental prions.

Materials and methods

Study site

The study site was located approximately 1.5 km from a depopulated captive WTD facility in Beltrami County, Minnesota. The landscape within and surrounding the study site comprised a matrix of relatively level and gently undulating topography dominated by deciduous shrub and hardwood forests, with intermittent wetlands. The soil orders present are characteristic of hardwood forests (Alfisols) and wetlands (Histosols [58]. Testing of these soils by Grunklee et al. [41] revealed levels of clay (5.5–13%) and organic matter (3–35%) known to enhance prion binding [15,21,59,60] and/or cause RT-QuIC interference [24,61]. The area downstream of the study site and upstream of the captive facility is a definitional wetland, characterized by typical wetland hydrological features, soils, and vegetation.

Soil sampling

We assigned unique IDs to each sample collection location, and marked these locations with metal and wooden posts, flagging tape, and GPS coordinates. Prior to soil sampling, we removed and collected forest litter and duff layers. We then used 10-gram Terra Core® plastic disposable soil core samplers (QED Environmental Systems, Dexter, MI, USA) placed into 18 oz Whirl-Pak® bags (Nasco, Fort Atkinson, WI, USA) to collect 1–3 surface level soil samples ~2 cm deep directly beneath animal carcass remains or along transects associated with PrP^Sc-positive carcass material. After field collection, all soil samples were stored at −20°C

Sample processing

Approximately 1.5–2 years after field collection, we thawed, dried, and extracted ~2 g of each study site soil sample [41]. In doing so, we placed all thawed, wet 2-g soil aliquots into individually labelled kraft paper stock drying envelopes (‘coin envelopes’, Innovative, Minneapolis, MN, USA), sealed them with autoclave or bench tape, and placed the samples into a Thermo Scientific™ Precision™ incubator (ThermoFisher Scientific, Rochester, MN, USA) to dry.

After drying, we measured ~200 mg dried soil for extraction and followed the extraction protocol detailed in Grunklee et al. [41]. Briefly, we added 10 μL of Proteinase K (PK; Sigma-Aldrich, St. Louis, MO, USA) and 1000 μL of myristyl sulfobetaine buffer (MSB buffer; Sigma-Aldrich T7763, St. Louis, MO, USA) to each soil aliquot and rotated for 60 minutes. We centrifuged the aliquots briefly at 8000 × g. We removed 750 μL of supernatant and transferred it to clean 1.5 mL tubes with care not to disturb the soil pellet. To the soil pellet, we added 250 μL MSB buffer and 2.5 μL PK, rotated the mixtures for another 60 minutes, and centrifuged the mixture briefly at 8000 × g. We separated 250 μL of supernatant from the pellets and added it to the originally retained supernatant. We then added 20 μL aminoethyl benzenesulfonyl fluoride hydrochloride (AEBSF) protease inhibitor (Sigma-Aldrich A8456, St. Louis, MO, USA) to the consolidated supernatant, vortexed to mix, and centrifuged the mixture at 8000 × g briefly. We removed ~950 μL of the supernatant while avoiding the pellet and transferred it to a clean tube. To the supernatant, we added 90 μL of 4% sodium phosphotungstate powder (NaPTA; Sigma-Aldrich 496,626, Burlington, MA, USA) mixed with 170 mm magnesium chloride (MgCl₂) and incubated at room temperature for 1 hour. Following incubation, we centrifuged the tubes at 16,000 × g for 30 minutes. We carefully removed the supernatant to leave the pellet undisturbed. We added 200 μL of dH₂O to each pellet and performed the last centrifuge step at 16,000 × g for 30 minutes. We carefully removed the supernatants without disturbing the pellet a final time. We froze the resultant extraction pellets at −80°C.

Real-time quaking-induced conversion assay (RT-QuIC)

Prior to performing RT-QuIC, we removed soil extracts from −80°C storage and allowed at least 15 minutes to thaw at room temperature. Once samples thawed, we resuspended soil pellets in 50 μL of dilution buffer (0.1% Scientific™ sodium dodecyl sulphate (SDS) solution (10% stock solution, Fisher 15,553,027), 1 × phosphate-buffered saline (PBS; Fisher BP3991; ThermoFisher Scientific, Rochester, MN, USA), N-2 Supplement (Life Technologies Corporation, Carlsbad, CA, USA) and vigorously vortexed. In addition, we sonicated all sample extracts that were in −80°C storage for more than one week for 30 seconds on a 36 amplitude level with no pulsing using a Qsonica Sonicator Q700 (Qsonica, ThermoFisher Scientific, Rochester, MN, USA). Immediately before conducting the RT-QuIC reaction, we diluted each extract to 10⁻¹ in dilution buffer to reduce false seeding activity in RT-QuIC as described in Grunklee et al. [41].

We performed RT-QuIC on diluted soil extracts as described in [62] and [10]. Briefly, we filtered rPrP hamster substrate at 3000 × g and created a reaction mixture using filtered reagents: 5X phosphate-buffered saline (PBS; Fisher BP3991), 100 mm Ethylenediaminetetraacetic acid (EDTA; 0.5 M EDTA, 15–575–020; ThermoFisher Scientific, Rochester, MN, USA), 100 mm Thioflavin T (ThT; 102598–926 VWR), 2 M Sodium Chloride (NaCl; Neat SIAL-S5150-1 L), and 0.1 mg/mL rPrP. We pipetted 98 μL of the reaction mixture into a 96-well format optical-bottom black microplate (ThermoFisher Scientific, Rochester, MN, USA). On the microplate, we seeded eight wells with 2 μL of each soil sample extract. Each plate contained wells of blank dilution buffer, 2 μL positive tissue plate control, and 2 μL negative tissue plate control for quality assurance. We also included octuplicate wells of 2 μL of Alfisol and Histosol negative control soils each to ensure appropriate soil order comparisons within each reaction. These Alfisol and Histosol control soil samples have been described in Grunklee et al. [41] and originate 0.5 km southwest of the study site and 2.3 km southwest from the associated captive cervid facility in an area with no history of CWD prion contamination (Figure 1). Finally, we sealed the microplates with clear plate tape and performed all experiments using BMG FLUOStar® Omega plate readers (BMG Labtech, Ortenberg, Germany) at 42°C, 1600 gain, and recorded readings every 45 minutes with one-minute cycles of double-orbital shaking and resting for 48 hours.

Statistical analysis

We used RStudio in the R Software v4.2.3 [63] to manage, curate, and visualize RT-QuIC data. We followed the statistical analysis procedures for maxpoint ratio (MPR) as described by Grunklee et al. [41], for optimal sensitivity and specificity for prion detection in soils. Briefly, we calculated the MPR of each RT-QuIC reaction well by dividing the maximum relative fluorescent units (RFU) value obtained within 48 hours of RT-QuIC by the background RFU value (i.e. MPR = maxRFU/backgroundRFU [64]. Following Grunklee et al.’s (criterion 3 approach [41]; approach using statistical testing to obtain a binomial test result from multiple RT-QuIC replicates, we conducted an uncorrected Fisher’s LSD test following ANOVA comparing test samples to negative soil controls (Alfisol or Histosol; n = 8). However, because some false seeding activity among negative controls (associated with these soil orders) was still observed even after extraction optimization, the MPRs of some soil samples were statistically different from negative controls even though there was very little seeding activity among those sample replicates. Thus, we applied a more stringent approach to determination of prion detection by conducting statistical testing on only those samples where at least 50% of the test sample replicate wells (i.e. x ≥ 4 of 8 replicate wells) had an MPR ≥ the threshold T_MPR = 2.504 (combining criteria 1 and 3 approaches [41]. Statistical significance was set at p < 0.05.

Funding Statement

This work was supported by the Minnesota Environment and Natural Resource Trust Fund as recommended by the Legislative-Citizen Commission on Minnesota Resources [2020–087 and 2022–217] and the College of Food, Agricultural and Natural Resource Sciences at the University of Minnesota Twin Cities.

Disclosure statement

Marc D. Schwabenlander and Peter A. Larsen hold financial and business interests in Priogen Corp., which has a licence from the University of Minnesota to commercialize prion diagnostics technology. These interests have been reviewed and managed by the University of Minnesota in accordance with its Conflict of Interest policies.

Data availability statement

Data from this study have been made publicly available through the Data Repository of the University of Minnesota (https://doi.org/10.13020/P2P3-1509).

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/19336896.2025.2514947