CWD prions are present in the blood of healthy-appearing, naturally infected free-ranging white-tailed deer

Abstract

Chronic wasting disease (CWD) is an emergent, fatal prion disease in deer, elk, and moose that is recognized in 37 U.S. states, impacting $447 billion in annual wildlife activities and cervid farming industries. The continued detection of CWD in new geographic areas supports the need for the development of sensitive antemortem diagnostics capable of detecting CWD during the early stages of disease. Early detection will identify infected animals in the initial stages of prion shedding, helping to curtail further spread through improved surveillance and cull programs. Current CWD diagnostics, while providing valuable insights to our understanding of the magnitude of the disease, fall short in their ability to detect low concentrations of prions associated with nascent infections. Here we analyzed blood buffy coat cells collected from free-ranging white-tailed deer (WTD) (Odocoileus virginianus) in CWD endemic regions by a modified amyloid amplification assay to probe for the presence of blood-borne prions in naturally exposed deer. Our findings provide evidence that prions can be detected in the blood of healthy-appearing, naturally exposed and infected free-ranging WTD. These findings provide a tool for improved surveillance, herd management, and future studies to reveal therapeutic targets for this and related protein misfolding disorders.

Introduction

Chronic wasting disease (CWD) is the only prion disease identified in free-ranging wildlife. The disease is known for its characteristic long incubation period (≥ 2 years) and facile transmission among deer, elk, and moose (cervids)¹. The geographical expansion of CWD continues across North America, Asia, and Scandinavia, with prevalence rates as high as 80% in some United States (U.S.) cervid populations². Transmission of the disease among cervids includes direct animal-to-animal contact, indirect contact with prions shed to the environment, and gestational transmission from mother to offspring^3–5. CWD-infected cervids appear healthy for most of the infection, yet studies have demonstrated that they actively shed the infectious prion agent within bodily fluids, excretions, and tissues (urine, saliva, feces, blood, birthing-related, carcass) during this time^6–11. Diagnosis of CWD using USDA official testing is primarily based on analysis of (postmortem) retropharyngeal lymph nodes or brain stem obex tissue by immunohistochemistry (IHC) for the accumulation of disease-associated prion protein (PrP^CWD)¹². Antemortem IHC testing of tonsil and RAMALT biopsies has revealed CWD positivity in naturally infected farmed and free-ranging cervids^13–15. Here we provide evidence for the presence of prions in the blood of healthy-appearing free-ranging CWD infected white-tailed deer.

We and others have demonstrated the presence of the infectious prion agent within the blood of infected animals (bovine spongiform encephalopathy, CWD, and scrapie), including humans (variant Creutzfeldt-Jakob disease [vCJD]), capable of initiating disease in susceptible hosts^4,16–18. Due to the presumed low concentration of circulating prions in blood and the presence of blood related inhibitors that interfere with in vitro assays, detection of hematogenous prions has proven to be difficult^19–23. Despite these challenges, further development of reliable, sensitive blood testing will enhance our ability to surveil the continued geographical expansion and increased prevalence of CWD in captive and free-ranging cervids²⁴, and will enhance the drive for effective intervention therapeutics to mitigate the progression of Alzheimer’s disease and related dementias (ADRDs) that affect more than 50 million people globally²⁴. The amyloid amplification methods, protein misfolding cyclic amplification (PMCA) and real-time quaking-induced conversion (RT-QuIC), have successfully revealed the presence of prions in the blood of asymptomatic CWD-infected captive cervids, BSE-infected sheep, and human vCJD patients^26–30 as well as in platelets from wild cervids³⁰. We previously reported on the longitudinal profiling of hematogenous CWD prions in experimentally-infected cervids confirming their presence in both symptomatic and asymptomatic deer³¹. What remains to be addressed is if our modified RT-QuIC assay is able to detect prions in blood cells harvested from free-ranging naturally infected cervids.

Here we demonstrate the presence of prions in blood buffy coat cells collected from healthy-appearing CWD-infected free-ranging white-tailed deer (WTD) in populations native to southeastern U.S. states. Our findings demonstrate: (I) the relationship between RT-QuIC detectable amyloid seeding activity in blood (i.e. prionemia) and tissue positivity, providing insight into disease progression, and (II) the ability to detect blood-borne prions in WTD with polymorphisms associated with delayed disease kinetics and detection,^33,34 extending our knowledge of RT-QuIC’s diagnostic capabilities. These findings offer promising potential for early CWD antemortem detection, and by extension a means for early detection of other protein misfolding diseases.

Results

Prion detection in blood buffy coat cells collected from healthy-appearing free-ranging WTD

To determine if hematogenous CWD prions can be detected in healthy-appearing and diseased, free-ranging WTD, we assessed blood buffy coats from n = 37 (n = 33 from 3 CWD endemic regions: n = 4 from a CWD non-endemic area) WTD by lipase iron-oxide bead real-time quaking-induced conversion (LIQ)³¹ for the presence of amyloid seeding activity (i.e. prions). Our findings revealed blood-borne prions in 19/33 (57.6%) blood samples collected from WTD in CWD endemic areas of southeastern U.S. (Arkansas (4/11, 36.4%), Tennessee (11/14, 78.6%), and West Virginia (4/8, 50%)) (Fig. 1; Table 1). Blood harvested from deer in the CWD non-endemic area were negative (Georgia (0/4, 0%)) by LIQ. These findings demonstrate the ability to detect prionemia in free-ranging WTD populations across multiple U.S states.

Fig. 1.

Lipase iron-oxide bead RT-QuIC (LIQ) analysis of buffy coat cells from free-ranging white-tailed deer, showing CWD prion detection in 19 of 33 blood samples tested. Samples were grouped based on the state of collection. Pooled results from Georgia blood samples served as negative controls, while samples from experimentally inoculated and known CWD-positive WTD served as positive controls. P values are reported as follows < 0.0001****, < 0.001***, < 0.01**, and < 0.05*.

Table 1.

Summary of white-tailed deer (WTD) data including sex, age, genotype based on Prnp codon 96, and chronic wasting disease (CWD) status of buffy coat, lymphoid tissues, and obex. RT-QuIC and IHC results are shown for retropharyngeal lymph node (RPLN), tonsil, rectoanal mucosal-associated lymphoid tissue (RAMALT) and obex, and RT-QuIC results are shown for buffy coat cells. Data are grouped by state (Georgia, Arkansas, Tennessee, and West Virginia) and assay (RT-QuIC or immunohistochemistry), with “+” and “-” denoting CWD-positive and negative detection. Statistical significance was determined for buffy coat samples by comparison to the samples collected from Georgia, in an area with no CWD detections, P values reported as follows <0.0001****, <0.001***, <0.01**, and <0.05*, and NS was non-significant. NA (not available), no tissue available for testing.

Prionemia demonstrated in free-ranging WTD prior to and after lymphoid and central nervous system (CNS) involvement

Previous experimental studies in WTD provided evidence that prionemia is most prominent two to three months post infection, near the time of lymphoid biopsy positivity, and later in infection when CNS involvement is presumed^31,34. As the timing of CWD infection is unknown for field collected samples, terminal harvests provide an opportunity to assess the presence of blood-borne prions when specific tissue CWD status is known. Lymphoid and brain tissues collected from this cohort of deer were analyzed by RT-QuIC and immunohistochemistry (IHC) to determine each deer’s CWD status. We identified RT-QuIC prion seeding activity in 23/33 (69.7%) and IHC PrP^CWD deposition in 15/33 (45%) WTD (Table 1). Blood, lymphoid and brain tissues harvested from deer in a CWD non-endemic region remained negative by both RT-QuIC and IHC (0/4, 0%).

Stage dependent patterns of prionemia revealed in free-ranging WTD

To assess how prionemia correlated with lymphoid and CNS involvement we determined each animal’s tissue CWD status (RT-QuIC and IHC) and compared the results with those from blood LIQ assessment. This analysis allowed us to classify prionemia into four distinct stages: (I) prionemia without lymphoid or CNS involvement, (II) prionemia with single lymphoid tissue involvement (retropharyngeal lymph node, RPLN; tonsil; or rectoanal mucosa-associated lymphoid tissue, RAMALT), (III) prionemia with multiple lymphoid tissues involved (RPLN, tonsil, and/or RAMALT), and (IV) prionemia with both lymphoid and CNS (obex) involvement. This categorization showed a distinct pattern of increased prevalence of prionemia with more extensive tissue involvement (Fig. 2). Overall we detected prionemia in 19/33 (58%) free-ranging healthy appearing WTD from CWD endemic areas. Specifically, we revealed hematogenous prions in 3/10 (30%) free-ranging WTD prior to the presence of prion deposition in lymphoid or brain tissue. Among deer with single lymphoid involvement, prionemia were demonstrated in 2/5 (40%) WTD. Detection of prionemia increased in deer with multiple lymphoid involvement to 6/10 (60%), and when deer had both lymphoid and CNS involvement prionemia was detected in all deer 8/8 (100%) (Table 1; Fig. 2). LIQ was unable to detect prionemia in 7/23 (30.4%) WTD with demonstrated tissue positivity. Blood and tissue samples collected from n = 4 WTD harvested from the CWD negative area remained negative by RT-QuIC and IHC (Table 1; Fig. 1) Collectively these findings demonstrate that prionemia can be detected in blood buffy coat cells from free-ranging deer throughout disease course, and importantly, prior to CNS involvement.

Fig. 2.

Congruency between prion detection in buffy coat and in lymphoid and CNS tissues. The CWD status of each WTD was determined by RT-QuIC and IHC of lymphoid and obex tissues. Prionemia was detected by LIQ and classified into four categories: (I) prionemia without lymphoid or CNS involvement, (II) prionemia with single lymphoid tissue (RPLN, tonsil, or RAMALT) involvement, (III) prionemia with multiple lymphoid tissues involved (RPLN, tonsil, and/or RAMALT), and (IV) prionemia with both lymphoid and CNS (obex) involvement. Ratios represent the number of deer with a CWD positive buffy coat sample, out of total deer in each category (I-IV). Congruency between tissue RT-QuIC/IHC and buffy coat LIQ is reported below, and showed LIQ was able to detect prionemia in three deer (3/10) prior to deposition in lymphoid or CNS tissue. LIQ was also able to detect prionemia in 40% of WTD with deposition in a single lymphoid tissue, 60% of WTD with deposition in two or more lymphoid tissues, and 100% of WTD with prior deposition in both lymphoid and CNS tissue. Figure created using Canva (2026) and adapted from original artwork commissioned by C. Mathiason and drawn by L. O’Keefe.

Prionemia demonstrated in free-ranging WTD with Prnp codon 96 GG and GS polymorphisms

As Prnp codon 96 polymorphisms, glycine (G) and serine (S), have been identified to impact a WTD’s susceptibility to CWD infection³⁵, we correlated prionemia with each deer’s genotype. In our study cohort from endemic regions (n = 33), 22/33 (66.7%) were wildtype 96GG, 10/33 (30.3%) were 96GS, and 1/33 (3.03%) was 96SS. Among WTD from a non-endemic region (n = 4), 2/4 (50%) were wildtype 96GG, 1/4 (25%) was 96GS, and 1/4 (25%) was 96SS. The distribution of codon 96 polymorphisms in our study cohort was similar to that previously reported in native populations³⁶. We identified prionemia in 13/22 (59.1%) 96GG, 6/10 (60%) 96GS, and 0/1 (0%) 96SS expressing deer (Table 2). We found CWD prionemia in 96GG deer in all three CWD endemic states and CWD prionemia in 96GS deer in 2 of the endemic states (Tennessee and West Virginia) (Table 1). These results provide evidence that CWD prionemia can be detected in free-ranging WTD with polymorphisms at codon 96.

Table 2.

Prnp codon 96 genotype distribution and buffy coat prion detection in white-tailed deer (WTD). Summary of the frequency of each of the three Prnp codon 96 genotypes (GG, GS, and SS) observed across a cohort of 37 WTD, compared to buffy coat lipase iron- oxide bead RT-QuIC (LIQ) positivity. Percentages represent the proportion of buffy coat, CWD positive WTD from each genotype.

WTD p Prnp codon 96 Polymorphisms and CWD Prionemia
Genotype (Codon 96)	Total deer	Buffy coat positive deer	Percentage of positives
GG	24	13	54.2
GS	11	6	54.5
SS	2	0	0

Discussion

The continued geographical expansion and increased prevalence of CWD exhibits the need for rapid and reliable tests to surveil the disease in captive and free-ranging cervid populations²⁴. Our findings support the continued development of prion amplification assays for use in antemortem testing and demonstrate the ability to detect prionemia in healthy-appearing, free-ranging WTD from CWD endemic regions within the United States.

We further demonstrate the ability to detect prionemia in free-ranging WTD prior to lymphoid and CNS involvement, supporting experimental studies revealing early and persistent prionemia in WTD and sheep^27,34,37,39. Surprisingly we noted that RT-QuIC amyloid conversion rates in blood cell components of free-ranging WTD were generally higher than those demonstrated in our experimental studies in deer³¹, suggesting higher prion load within this cohort of free-ranging CWD-infected deer, or enhanced conversion competency of the prions present in the blood of these animals.

Using RT-QuIC and IHC to determine tissue CWD status, we classified prionemia into four main categories: (I) prionemia without lymphoid or CNS involvement, (II) prionemia with single lymphoid tissue (RPLN, tonsil, or RAMALT) involvement, (III) prionemia with multiple lymphoid tissues involved (RPLN, tonsil, and/or RAMALT), and (IV) prionemia with both lymphoid and CNS (obex) involvement. This classification revealed a higher prevalence of prionemia in animals with more expansive lymphoid tissue involvement, with amyloid seeding rates highest in animals with concurrent CNS involvement. In WTD positive by RPLN, tonsil, RAMALT, or obex based assays, LIQ did not always detect prionemia, suggesting that blood-based and other tissue-based assays may capture different aspects of disease detection. Importantly, the ability to detect prionemia in animals lacking lymphoid or CNS involvement mirrors findings from experimentally inoculated WTD cohorts, in which RT-QuIC detected circulating prions as early as one to three months post exposure, prior to detection in lymphoid or CNS tissues^31,34. Of additional interest, we were not able to detect hematogenous prions in some deer with confirmed lymphoid or CNS positivity, supporting experimental findings that prionemia may be intermittent throughout disease progression³¹. A noted limitation of this work is the small number of contemporary blood samples collected from deer free-ranging in areas with no CWD detection that were used as negative control comparison. The current findings will be enhanced by extended studies to increase sample size and sampling across a more broad geopgraphic range of cervid populations.

Compared to the inherently invasive procedures required to obtain lymphoid or brain tissue biopsies, blood represents a substantially less invasive sample type for CWD prion detection. Notably, our modified RT-QuIC assay requires only minimal amounts of anticoagulated whole blood to achieve adequate sensitivity. This small volume of blood makes the assay feasible for farmed and field conditions, expanding its applicability in situations where larger blood volumes are impractical. This provides a powerful tool for CWD surveillance and management.

The demonstration of prionemia in free-ranging WTD with Prnp polymorphisms known to modulate disease initiation and progression (codon G96S)⁷ provides evidence that blood testing may be suitable to surveil WTD populations for early CWD detection. Additional studies incorporating larger cohorts of WTD representing this and other Prnp polymorphisms will be needed to assess the robustness of these findings. Furthermore, chances of detecting a true positive may be increased if multiple antemortem sample types are collected (e.g., blood, tonsil biopsy, RAMALT biopsy).

The implications of sensitive hematogenous prion detection extend beyond wildlife disease surveillance. Previous studies have shown mechanistic similarities between prion diseases and other protein misfolding disorders^40,41, providing a potential link between the detection of CWD prionemia and the detection of peripheralized misfolded proteins associated with other amyloid diseases. The detection of CWD in blood prior to its deposition in lymphoid or nervous tissue enhances the importance to further investigations in unraveling the mystery of amyloid disease peripheralization. As globally more than 55 million people are affected by Alzheimer’s Disease and Related Dementias (ADRDs)²⁴, methodologies developed within the prion field, including the prion amyloid amplification assays continue to fill gaps in our understanding of proteopathy trafficking and peripheralization^42,43. The ability to detect and monitor early ADRD is important to drive effective intervention therapeutics to mitigate the progression of these diseases. Here we describe an amyloid detection method using a less-invasive, readily available tissue (blood), that may have applicability in human proteopathy detection.

This study highlights the ability of a contemporary prion amplification detection assay (RT-QuIC) to identify CWD prions in the blood of healthy appearing free-ranging WTD in southeastern U.S. states. Further assessments, including access to longitudinal blood collections from a larger cervid population, would be beneficial in addressing the role blood and blood components play in CWD disease pathogenesis, as well as being vital for surveillance, management, and therapeutic development for CWD and other proteopathies.

Methods

Study area, white-tailed deer, tissue and blood collection

Healthy-appearing, free-ranging WTD from CWD endemic areas (Newton County, Arkansas; Fayette County, Tennessee; and Hampshire County, West Virginia) and from a CWD nonendemic site (Clarke County, Georgia) were collected by the Southeastern Cooperative Wildlife Disease Study (SCWDS; University of Georgia) or by state wildlife management agency personnel, between January 2020 and April 2021. Field sampling was performed in accordance with relevant guidelines and regulations provided by the University of Georgia’s IACUC # A2018 02–010. All methods are reported in accordance with ARRIVE guidelines. All WTD in this study were collected as part of a study on vertical transmission of CWD, and data from all the female deer was previously published⁵, with data collected on male deer presented here. Personnel from SCWDS performed euthanasia via sharp shooter, followed by necropsy and sample collection on each individual animal, using separate, single use instruments and personal protective equipment for each animal. Blood samples were collected directly from the heart of each WTD using a needle and syringe, within five minutes of death. Whole blood (22 to 27.5 mL) was transferred to 50 mL tubes containing citrate phosphate dextrose adenine (CPDA) anticoagulant (14–20%, MWI Animal Health) and stored at 4 C until shipment within 4 days.

Following blood collection, WTD carcasses were transported to a controlled site for necropsy and further sample collection was carried out as previously described⁵. Care was taken to minimize cross contamination within and between WTD when collecting RPLN, palatine tonsil, RAMALT and obex samples, as these tissues were used to determine CWD status for each animal. Each tissue was divided equally, with ½ placed in 10% neutral buffered formalin, and the other ½ frozen at -20 °C. Whole blood, as well as fixed and fresh-frozen tissues were shipped to Colorado State University (CSU) where they were processed and/or stored at -80 °C until analysis.

Blood processing and buffy coat isolation

To isolate buffy coat cells for analysis, whole blood CPDA samples were centrifuged at 300 g for 15 min at 21 °C before removing the plasma layer. Following centrifugation, the buffy coat layer was removed and fully processed as below or was aliquoted into cryovials containing CryoStor CS10 (Biolife Solutions) in a 1:1 ratio and stored at -80˚C until analyzed. Forty milliliters (40 mL) of ammonium-chloride-potassium (ACK) lysing buffer [(97.3 µM ethylenediaminetetraacetic acid (EDTA),10 mM potassium bicarbonate (KHCO3), 154.4 mM ammonium chloride (NH4Cl)] was added to the fresh or cryopreserved buffy coat cells, followed by a second centrifugation as described above. The cell pellet was retained after the supernatant was discarded. The ACK lysing step and centrifugation was repeated until no visible red blood cell contamination remained in the cell pellet. The cells were then washed twice with 1xPBS and centrifuged at 300 g for 10 min at 21 °C and resuspended in 1xPBS. The buffy coat cells were counted using a hemacytometer, aliquoted into tubes at approximately 10⁷ total cells per milliliter and centrifuged at 300 g for 10 min prior to removing the supernatant. Isolated buffy coat cell pellets were stored at -80 °C until homogenization and LIQ analysis.

Lipase-iron oxide bead treatment (Lipase- IOB)

To liberate membrane bound prions present in blood buffy coat cells and to remove lipid inhibitors the frozen buffy coat cell pellets were thawed at room temperature and subjected to Lipase Iron-Oxide Bead Treatment (Lipase-IOB) as previously described³¹. Blood buffy coat cell pellets were resuspended in 1xPBS at a concentration of 10⁷ cells per milliliter and homogenized using an Omni Bead Ruptor 24. Following two 30 s bead Ruptor cycles (power 5) the homogenates were frozen, thawed, and subjected to an additional 30 s bead Ruptor cycle.

Fifty microliters (50 µL) of homogenized blood buffy coat cells, roughly equivalent to 5 × 10⁵ buffy coat cells, were added to 950 mL of 1xPBS containing 4.5 units of lipase B [(Lipase B Candida antarctica, recombinant from Aspergillus oryzae; Sigma-Aldrich), and 0.4 units lipase C (Phospholipase C from Clostridium perfringens (welchii)]. The samples were then placed on a 37 °C thermomixer at 1400 r.p.m for 1 h, before the addition of 2 µL of iron oxide beads (49 mg/ml, ~ 9 μm; Bangs Laboratories, Indiana BioMag superparamagnetic iron-oxide). Samples were incubated with the beads on an end-over-end rocker for 1 h before being placed on a magnetic particle separator (Pure Biotech, NJ, USA) to remove all supernatant. The remaining iron-oxide beads were resuspended in 10 µL of 0.1% Sodium dodecyl sulfate (SDS) (Sigma-Aldrich).

Tissue homogenate preparation for CWD status determination

Upon arrival at CSU the fresh-frozen obex, RAMALT, RPLN, and tonsil were homogenized using an Omni Bead Ruptor 24, at 10% weight to volume (w/v) tissue solution in 1x phosphate buffered saline (PBS) (20mM sodium phosphate (NaPO₄), 150 mM sodium chloride (NaCl), pH 7.4), and stored at -80 °C until analyzed by RT-QuIC. (see RT-QuIC methods below).

Real-time quaking induced conversion (RT-QuIC)

To determine the CWD status for each WTD and assess blood for the presence of prions using contemporary prion amplification methods, RPLN, tonsil, RAMALT, obex, and Lipase-IOB treated blood buffy coat cells were analyzed for CWD amyloid formation using modified RT-QuIC methods previously described¹¹. In brief, 0.1 mg/mL of truncated Syrian hamster recombinant protein (amino acids 90–231) was added to a master mix, containing 20 mM NaPO₄, 320 mM NaCl, 1.0 mM EDTA, and 10 µM thioflavin T (Sigma)⁴³. Protein master mix (98 µL) was added to each well of a 96 well black optical bottom plate (Greiner Bio-One). Tissue homogenates were diluted in 1x PBS (RPLN, tonsil, and RAMALT final concentration 10^− 4; obex final concentration of 10^− 5) and 2 µL of each tissue was added to the 96 well plate in quadruplicate. Lipase-IOB treated buffy coat cells were analyzed by loading 3 µL of the iron oxide bead sample mixture, suspended in 0.1% SDS, to each well of the 96 well plate, again plating each sample in quadruplicate. Each 96-well plate was covered with clear polyolefin acrylate sealing tape (Thermo Scientific), placed in a BMG FLUOstar Omega plate reader and subjected to alternating cycles of quaking and rest for 62.5 h at 55 °C, with data analyzed and collected every 15 min. Each sample in quadruplicate was plated on one to four independent plates, resulting in a total of 4–16 technical replicates per sample.

RT-QuIC controls and statistical analysis

Tissue and blood samples from free-ranging WTD in Georgia, collected prior to CWD detection in the state, served as CWD negative tissue controls. Tissue and blood buffy coat cells collected from experimental WTD of known CWD status were used as negative or positive blood and tissue controls. Additionally, six replicates of CWD negative brain and six replicates of CWD positive brain, from deer with confirmed CWD status, were placed on each plate to serve as RT-QuIC plate controls. Brain control samples were used to confirm RT-QuIC assay sensitivity, monitor inter-plate consistency, and ensure proper amplification of amyloid seeding activity between runs. All experimental sample types (lymphoid tissues, obex, and buffy coat cells) were analyzed alongside appropriate sample-type matched controls to enable direct comparison within each assay. Replicates were assigned rates of amyloid formation equal to 1/time to threshold, with the threshold defined as ten times the standard deviation of the baseline fluorescence for buffy coat cell samples, and five times the standard deviation of the baseline fluorescence for RPLN, tonsil, obex, and RAMALT samples⁴⁴. Replicates for each sample were combined across plates and compared to combined negative buffy coat controls using the Mann-Whitney U non-paired, non-parametric test in GraphPad Prism v9. Samples were considered to be positive for CWD amyloid formation if the median rate of amyloid formation was significantly greater (p-value < 0.05) than that of the combined negative control.

White-tailed deer immunohistochemistry (IHC) CWD status

Immunohistochemistry is the gold standard and USDA-approved test to determine CWD status in cervid species, and was used to assess PrP^CWD deposition in WTD RPLN, tonsil, RAMALT, or obex formalin fixed and paraffin-embedded tissues. Tissue sections.  (5 μm) were cut from blocks and affixed to positive-charged slides. Slides were further processed by deparaffinization in xylene (100%) and rehydration in graded alcohols (100% − 70%) before treatment with 88% formic acid (30 min at room temperature) as previously described⁴⁵. After washing with water, slides were placed in a 2100-Retriever (Prestige Medical) in sodium citrate buffer (0.01 M sodium citrate, 0.05% Tween 20, pH 6.0) for heat-induced epitope antigen retrieval. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide and slides were blocked using TNB Blocking Buffer (0.1 M TRIS-HCl, 0.15 M NaCl, 0.5% blocking reagent) for 60 min before overnight incubation in anti-prion antibody BAR224 (1 mg/mL; Caymen Chemical) diluted 1:750. Envision+ anti-mouse HRP-labeled polymer secondary antibody (Dako) was followed by 3-amino-9-ethylcarbazole (AEC) (Dako) for visualization. Negative control tissues were run concurrently with test samples.

Prnp genotyping

Genotypes for all 33 WTD from CWD endemic areas, as well as 4 WTD from non-endemic Georgia, were determined for codon 96 of the Prnp open reading frame. Female deer genotypes were previously published⁵, male deer genotypes are reported here. DNA was extracted from WTD spleen or RPLN tissue as previously published⁴⁶. Briefly, DNeasy Blood and Tissue kit (QIAGEN) were used to extract DNA from the tissues as per kit instructions. DNA concentration was determined for each sample using a NanoDrop − 1000 UV/Vis Spectrophotometer. The purified DNA samples were amplified by polymerase chain reaction (PCR), using primers 213d AGGTCAACTTTGTCCTTGGAGGAG and 139u TAAGCGCCAAGGGTATTAGCAT, and sequencing primer 86d CAGTCATTCATTATGCTGCAGACT. DNA segments underwent Sanger sequencing by Quintara Biosciences. Results were analyzed at CSU using Chromas sequencing software (Technelysium Pty Ltd v2.6.6).

Author contributions

DJT: data acquisition, analysis, interpretation, and manuscript drafting, AVN: data acquisition, analysis, interpretation and manuscript drafting, EEM: data acquisition, analysis, interpretation and manuscript review, NDD: data acquisition, analysis, interpretation and manuscript review, SKS: data acquisition and interpretation AMS: data acquisition and interpretation, ZO: data acquisition, EB: animal collection, data acquisition and manuscript review JB: data acquisition and manuscript review DMG: design, facilitated field collections, manuscript review JSD: animal collection, data acquisition, manuscript review NS: design, data acquisition, manuscript review CAC: design, data acquisition, manuscript review JMC: state agency coordination, animal collection, financial grant collaborator, manuscript review MGR: study conception, financial support, data acquisition, analysis, interpretation, manuscript drafting, CKM: study conception, design, data analysis and interpretation, manuscript drafting, and financial support.

Data availability

The datasets generated and/or analyzed during the current study are available in the NCBI GenBank database under accession numbers PZ387726–PZ387731.

Declarations

Competing interests

The authors declare no competing interests.