JOURNAL OF VIROLOGY, May 2010, p. 5097–5107
Copyright © 2010, American Society for Microbiology. All Rights Reserved.
Vol. 84, No. 10
B Cells and Platelets Harbor Prion Infectivity in the Blood of Deer
Infected with Chronic Wasting Disease?†
Candace K. Mathiason,1Jeanette Hayes-Klug,1Sheila A. Hays,1Jenny Powers,2David A. Osborn,3
Sallie J. Dahmes,4Karl V. Miller,3Robert J. Warren,3Gary L. Mason,1Glenn C. Telling,5
Alan J. Young,6and Edward A. Hoover1*
Department of Microbiology, Immunology, and Pathology, Colorado State University, Fort Collins, Colorado1; National Park Service,
Fort Collins, Colorado2; Warnell School of Forestry and Natural Resources, University of Georgia, Athens,
Georgia3; WASCO Inc., Monroe, Georgia4; University of Kentucky Medical Center, Lexington,
Kentucky5; and South Dakota State University, Brookings, South Dakota6
Received 13 October 2009/Accepted 23 February 2010
Substantial evidence for prion transmission via blood transfusion exists for many transmissible spongiform
encephalopathy (TSE) diseases. Determining which cell phenotype(s) is responsible for trafficking infectivity
has important implications for our understanding of the dissemination of prions, as well as their detection and
elimination from blood products. We used bioassay studies of native white-tailed deer and transgenic cer-
vidized mice to determine (i) if chronic wasting disease (CWD) blood infectivity is associated with the cellular
versus the cell-free/plasma fraction of blood and (ii) in particular if B-cell (MAb 2-104?), platelet (CD41/61?),
or CD14?monocyte blood cell phenotypes harbor infectious prions. All four deer transfused with the blood
mononuclear cell fraction from CWD?donor deer became PrPCWDpositive by 19 months postinoculation,
whereas none of the four deer inoculated with cell-free plasma from the same source developed prion infection.
All four of the deer injected with B cells and three of four deer receiving platelets from CWD?donor deer
became PrPCWDpositive in as little as 6 months postinoculation, whereas none of the four deer receiving blood
CD14?monocytes developed evidence of CWD infection (immunohistochemistry and Western blot analysis)
after 19 months of observation. Results of the Tg(CerPrP) mouse bioassays mirrored those of the native cervid
host. These results indicate that CWD blood infectivity is cell associated and suggest a significant role for B
cells and platelets in trafficking CWD infectivity in vivo and support earlier tissue-based studies associating
putative follicular B cells with PrPCWD. Localization of CWD infectivity with leukocyte subpopulations may aid
in enhancing the sensitivity of blood-based diagnostic assays for CWD and other TSEs.
Chronic wasting disease (CWD) is an infectious protein-
misfolding disease, or transmissible spongiform encephalopa-
thy (TSE), affecting cervids in North America (59, 76–79) and
one Asian country (41, 68). CWD is unique among prion dis-
eases in affecting free-ranging wildlife populations (deer, elk,
and moose). Early and subsequent observations made by Wil-
liams and Miller (58, 79) related CWD transmission to direct
contact with clinically affected deer, as well as indirect contact
with environments previously populated by infected deer (57).
Bioassay studies of white-tailed deer have demonstrated that
body fluids and excreta (saliva, urine, feces, and blood) contain
infectious prions (53, 54). Both clinical and preclinical CWD-
infected deer harbored sufficient infectious prions to produce
CWD in naïve white-tailed deer following ingestion of saliva or
transfusion of whole blood (53, 54).
The detection of blood-borne infectious prions has impor-
tant implications for our understanding of the spread of prions
among and within individuals, as well as for the elimination of
prions from blood products (13, 15, 33, 45), given the evidence
for Creutzfeldt-Jakob disease (CJD) transmission via blood
transfusion (16, 29, 47, 50, 62, 72, 73). Identifying the cell
phenotype or cell-free protein fractions that harbor prion in-
fectivity would contribute importantly to this understanding
and to the development of blood-based assays to detect prion
infection. We undertook the present studies to address these
MATERIALS AND METHODS
Bioassay studies of white-tailed deer. White-tailed deer fawns were provided
by the Warnell School of Forestry and Natural Resources, University of Georgia,
Athens—a region in which CWD has not been detected. The deer fawns were
hand raised and human and indoor adapted before overnight transport directly
to the Colorado State University (CSU) CWD research indoor isolation facility
without contact with the native Colorado environment. The 4-month-old fawns
were adapted to the facility housing conditions and diet for 2 months before the
Genotyping. All white-tailed deer were genotyped to determine their GG/GS
(codon 96) status by the laboratory of Katherine O’Rourke, USDA-ARS, Pull-
man, WA. Deer were allocated into inoculation cohorts (n ? 4) without knowl-
edge of their G96 genotypes.
Biocontainment protocols. Protocols to preclude extraneous exposure and
cross contamination between cohorts of animals as previously described (53, 54)
incorporated protective shower-in requirements, Tyvek clothing, masks, head
covers, and footwear while maintaining stringent husbandry. Tonsil biopsy and
terminal sample collections were taken with animal-specific biopsy and sample
collection instruments to minimize the possibility of cross contamination. Bed-
ding and liquid waste from each suite were either incinerated or collected in a
dedicated outdoor underground holding tank and denatured by alkaline diges-
* Corresponding author. Mailing address: Colorado State Univer-
sity, Pathology 1619, Fort Collins, CO 80523. Phone: (970) 491-7587.
Fax: (970) 491-0523. E-mail: Edward.Hoover@ColoState.edu.
† Supplemental material for this article may be found at http://jvi
?Published ahead of print on 10 March 2010.
Deer inoculation cohorts. Groups of 6-month-old fawns (usually four per
group) (Table 1) were housed in separate isolation suites throughout the study.
Suite-dedicated protective clothing, utensils, and waste disposal were incorpo-
rated to exclude cross contamination by fomites, bedding, food, excretions, or
contact. Deer cohorts 1 to 6 were inoculated by the intravenous (i.v.) route with
blood components from CWD-infected donor deer housed in the CSU CWD
isolation facility as follows: cohort 1, whole blood (250 ml); cohort 2, blood
mononuclear cell fraction (1 ? 107to 1.24 ? 108white blood cells [WBC] plus
platelets); cohort 3, cell-free plasma fraction (140 to 150 ml) recovered from 250
ml citrated whole blood; cohort 4, B cells (1 ? 106to 5 ? 106) separated by
Dynatec magnetic bead separation (98% purity) using anti-sheep B-cell mono-
clonal antibody (MAb) 2-104, which identifies peripheral blood B cells and may
identify follicular dendritic cells (FDCs) in lymphoid germinal centers (80);
cohort 5, CD41/61?platelets (6 ? 109to 35 ? 109) magnetically separated to
99% purity using MAb CAPPA 2A (VMRD, Pullman, WA); cohort 6, CD14?
cells (4 ? 105) magnetically separated to 98% purity using anti-sheep CD14 MAb
clone VPM65 (Fitzgerald Industries Inc., Concord, MA); cohort 7, naïve white-
tailed deer inoculated with blood from CWD?deer, which served as negative
controls for the study and were housed in a separate suite at the same facility.
Blood donor deer. Six experimentally inoculated CWD?and two CWD?deer
housed at the CSU indoor research facility were recruited from previously
described studies (54) for use as blood donors for these studies (Table 2).
Blood donors for cohorts 1, 2, and 3. Two CWD-infected deer previously
inoculated intracranially (i.c.) with 1 g whole brain homogenate collected from a
naturally infected CWD?deer (TS-989-09147) were the source animals.
Blood donors for cohorts 1, 4, 5, and 6. Four CWD-infected deer served as
blood donors for cohorts 1, 4, 5, and 6. Two donors (designated brain pool) had
been orally inoculated with 10 g (2 g/day for 5 days) brain from naturally infected
field isolates (TS-989-09147 or WDNR). The remaining two donors (designated
blood pool) had received 250 ml of blood via i.v. infusion from an experimentally
inoculated CWD?deer (TS989-CSU112). Two deer from each cohort (1, 4, 5, or
6) were inoculated with whole blood or specific cell phenotypes from the brain
pool donors, while the other two deer from each cohort were inoculated with
similar components from the blood pool donors.
Blood donors for cohort 7. One CWD?deer, housed at the Warnell School of
Forestry and Natural Resources, University of Georgia, Athens (UGA)—a re-
gion where CWD has not been detected—served as the donor for two negative
control donors that each received 250 ml whole blood via i.v. infusion.
Thus, the inocula used reflected both a conscious attempt to assess the uni-
versality of the results obtained given the constraints of a limited number of
recipient animals and limited amounts of inoculum materials (cell fractions etc.)
Blood collections, harvests, and inoculations. (i) Cohorts 1 to 3. One liter
sodium citrate-treated whole blood was collected from each of two CWD?donor
deer (Table 2) for cohort 1 to 3 inoculations (Table 1). The blood was not
pooled. Half (500 ml) of each whole blood collection was immediately adminis-
tered i.v. to two recipient cohort 1 deer (250 ml/deer)/donor deer (four recipi-
ents). Plasma and blood mononuclear cells harvested from 250-ml nonpooled
aliquots of whole blood were administered i.v. to cohorts 2 and 3 (Table 1).
(ii) Cohorts 1, 4, 5, and 6. Similarly, a portion (250 ml) of the whole blood
collected from the blood pool and brain pool donors (Table 2) was administered
i.v. to two recipient cohort 1 deer/donor (Table 1) (four recipients). The remain-
ing blood collected from the blood pool was pooled, as was the remaining blood
from the brain pool, and each pool was further processed to harvest specific cell
phenotypes by magnetic separation (as described below) that were then inocu-
lated by i.v. infusion into cohort 4, 5, or 6 (Table 1).
Deer monitoring and sample collection. All animals were monitored for evi-
dence of CWD infection by serial tonsil biopsies taken at 3, 6, 12, and 15 months
postinoculation (p.i.), and at study termination (19 months p.i.). Tonsil tissue was
divided, and equal portions were either stored at ?70°C or fixed in 10% formalin
for 24 h before processing for immunohistochemistry (IHC) analysis. At the
same sampling intervals, blood, saliva, feces, and urine were collected from each
animal and stored at ?70°C. At necropsy, the palatine tonsils, brainstem (me-
dulla at the obex), and retropharyngeal lymph nodes, as well as other tissues,
were collected for examination by IHC and Western blotting (WB) analyses to
identify the presence of the protease-resistant prion protein associated with
Cervid PrP transgenic mouse bioassay studies. (i) Cervid PrP transgenic
mice. Tg(CerPrP-E226)5037?/?mice (2), which express the elk PrP coding
sequence, were generated in the Telling laboratory at the University of Ken-
tucky. Mice were inoculated and maintained in accord with CSU IACUC guide-
(ii) Genotyping. All mice were screened at weaning for the presence of the
cervid/elk Prnp transgene by both conventional and real-time PCR. All inocu-
lated mice that tested negative for cervid PrPRESat the completion of bioassay
studies were rescreened to confirm the presence of the cervid Prnp transgene.
Biocontainment protocols. The protocols for white-tailed deer described
above also applied to cohorts of mice housed in filter-top isolation cages.
Mouse inoculation cohorts. Groups of five to nine weanling mice (Table 3)
were housed in separate cages throughout the study. Suite-dedicated protective
TABLE 1. White-tailed deer cohorts i.v. inoculated with blood components
CohortNo./cohort Donor statusInoculum source Inoculum
Blood mononuclear cells
250 ml CWD?blood
1 ? 107–1.24 ? 108WBC ? platelets
140–150 ml recovered from 250 ml blood
1 ? 106–5 ? 1062-104?cells (98% pure)
6 ? 109–35 ? 109CD41/61?cells (99% pure)
4 ? 105CD14?cells (98% pure)
250 ml CWD?blood
TABLE 2. CWD?/CWD?blood cell component donor history
Donor statusNo. of donorsDonor source history
1, 2, 3CWD?
2i.c. inoculated with brain homogenate from naturally
infected CWD?deer TS-989-09147
i.v. transfused with 250 ml whole blood from
inoculated CWD?deer TS989-CSU112
p.o. inoculated (2 g/day for 5 days) with CWD?
brain from naturally infected CWD?deer
TS-989-09147 or WDNR (1 each)
i.v. infused with 250 ml CWD?whole blood from
deer residing at Warnell School of Forestry and
Natural Resources, Athens, GA
14 months p.i.a/terminal
24 months p.i./late-stage
23 months p.i./late-stage
1, 4, 5, 6CWD?
2 blood pool
1, 4, 5, 6CWD?
2 brain pool
1 G/G, 1 G/S
5098MATHIASON ET AL.J. VIROL.
clothing, utensils, and waste disposal were incorporated to exclude cross con-
tamination by fomites, bedding, food, excretions, or contact. Cohorts 8 and 9
consisted of naïve Tg(CerPrP-E226)5037?/?mice that served as i.c. brain inoc-
ulate positive or negative controls each receiving 30 ?l of a 1% brain homoge-
nate prepared in phosphate-buffered saline (PBS) of either CWD?deer D10 or
deer UGA. Cohorts 10 to 25 consisted of naïve Tg(CerPrP-
E226)5037?/?mice that were inoculated by the i.c., i.v., intraperitoneal (i.p.), or
per os (p.o.) route using the same CWD?blood components described for
white-tailed deer inoculations above. Cohorts 10 to 13 received whole blood.
Cohorts 14 to 17 received the blood mononuclear cell fraction (106WBC plus
platelets). Cohorts 18 to 21 received the cell-free plasma fraction. Cohorts 22 and
23 received B cells (106) harvested from the retropharyngeal lymph node or
spleen. Cohort 24 received CD41/61?platelets (109). Cohort 25 received CD14?
cells (105). Cohorts 26 to 41, consisting of five to nine naïve Tg(CerPrP-
E226)5037?/?mice, served as the negative controls for this study and were
inoculated i.c., i.v., i.p., or p.o. with blood components from the same negative
control white-tailed deer donors (Table 2) as used for the negative control
white-tailed deer inoculations (Table 1).
Mouse monitoring and sample collection. All mice were monitored daily for
evidence of CWD clinical disease. Upon detection of clinical disease, mice were
euthanized and necropsied. Brain tissue was collected, divided into equal por-
tions, and either stored at ?70°C for WB or fixed in 10% formalin for 24 h before
processing for IHC analysis to identify the presence of PrPCWD.
Blood cell and plasma harvests [white-tailed deer and Tg(CerPrP-
E226)5037?/?mouse inocula]. Total blood cell populations were collected from
250 ml sodium citrate-treated whole blood by centrifugation at 1,200 rpm for 15
min at 4°C. The plasma fraction was collected and set aside on ice. The cell
fraction from this initial centrifugation was diluted 1:1 in 1? PBS (Gibco, Inc.)
and layered over Histopaque 1088 (Sigma) at a 1:1 ratio. These Histopaque
gradients were centrifuged without a brake at 2,500 rpm for 30 min at room
temperature. The discrete bands of WBC were collected, diluted in an equal
volume of 1? PBS, and further centrifuged for 10 min at 2,500 rpm at 4°C
(washed). The cell pellets were washed in wash buffer (1? PBS, 0.2% fetal
bovine serum [FBS], 2 mM EDTA) three times. Platelets were collected from the
plasma fraction by centrifugation at 3,000 rpm for 15 min. Cells and plasma
recovered from Histopaque 1088 gradient separations and plasma centrifuga-
tions were either directly inoculated into deer and mouse bioassay studies or
further processed to separate cell phenotypes.
Retropharyngeal lymph node and spleen cell harvests [Tg(CerPrP-E226)
5037?/?mouse inocula]. Retropharyngeal lymph node and spleen tissues were
pressed through a fine wire mesh (0.45 ?m), and WBC were collected as de-
scribed above for the Histopaque gradient protocol.
Cell phenotype labeling and flow cytometry. Cell phenotype MAbs were used
to recover and determine the purity of 2-104?B cells, CD14?monocytes, and
CD41/61?platelets from the blood donor sources described above. Leukocyte
and platelet blood cells were collected by centrifugation and Histopaque 1088
separation as described above and were then labeled with one of three antibodies
(Table 4), i.e., anti-sheep pan-B-cell MAb 2-104 (equivalent to MAb 2-8 de-
scribed by Young et al. ) (cell supernatant used undiluted), anti-sheep CD14
MAb clone VPM65 (cell supernatant used undiluted; Fitzgerald Industries Inc.,
Concord, MA), or anti-sheep CD41/61 MAb CAPPA 2A (1:100 dilution of a
1-mg/ml stock; VMRD, Pullman, WA). Cell aliquots were incubated with pri-
mary antibody for 20 min on ice and then washed three times in wash buffer. The
secondary antibody, goat anti-mouse IgG or IgM fluorescein isothiocyanate
(FITC), was diluted 1:100 in 1? PBS, 0.2% FBS and placed on the cells for 20
min on ice. The cells were again washed three times in wash buffer. The cells
were then labeled with anti-FITC beads at 10 ?l beads/107cells, again incubated
on ice for 20 min, and passed over LS or LD Dynatec magnetic bead separation
columns (in accordance with the manufacturer’s instructions). Cell populations
of interest were eluted in 1? PBS containing 0.1% FBS. Eluted (FITC-labeled)
cells were analyzed by flow cytometry (Dako-Becton Dickinson). To determine
purity, the cells were gated by forward and side scatter to include primarily
lymphocytes, which were counted, and volumes were adjusted to be equal to or
greater than the total number of each cell-specific phenotype populating 1 ? 107
peripheral blood mononuclear cells (as determined by prior flow cytometric
analysis of specific cell phenotype populations in white-tailed deer; i.e., 2-104?B
cell populations were ?10% and CD14?cells ?2% of the total leukocyte
population). This was done to equate the total number of phenotype-specific
cells (2-104?, CD14?) to that found in the total blood mononuclear cell fraction
TABLE 3. Tg(CerPrP) mouse cohorts inoculated with blood components from CWD?donor deer
Cohort(s)No./cohortDonor status Inoculum Volume/concn/no. of cells Route of inoculation
B cells 2-104?(RLNc)
B cells 2-104?(spleen)
50 ?l/day ? 3 days
50 ?l ? 3 days
2 ? 105
1 group each as for cohorts 10–25
i.c., i.v., i.p., or p.o.
5 or 9
aBMC, blood mononuclear cells.
b106BMC plus platelets.
cRLN, retropharyngeal lymph node.
TABLE 4. MAbs used for cell-specific phenotype harvests
CD72 B cells, lymphoid
germinal center FDCs
Alan Young SDSU,b
VMRD, Pullman, WAc
aMAb 2-104 is equivalent to MAb 2-880; see Table S1 in the supplemental
bSDSU, South Dakota State University.
cVeterinary Diagnostics, MAbs.
VOL. 84, 2010CWD PRIONS IN BLOOD CELL FRACTIONS5099
inoculum (cohort 2) that established infection (107blood mononuclear cells)
(Tables 1 and 5). The cells were either directly inoculated by i.v. inoculation into
deer bioassays or frozen for future i.c. mouse bioassays.
Western blotting. Tissue homogenates were prepared from the obex region of
the medulla oblongata encompassing the dorsal motor vagal nucleus (medulla at
the obex). Ten percent (wt/vol) homogenates were prepared in NP-40 buffer (10
mM Tris-HCl buffer [pH 7.5], 0.5% NP-40, 0.5% sodium deoxycholate) by
Fastprep disruption at a setting of 6.5 for 45 s. Twenty-five microliters of each
homogenate was mixed with 5 ?l of proteinase K (PK; Invitrogen) to a final
concentration of 20 ?g/ml and incubated for 30 min at 37°C with shaking. PK
activity was stopped with 4 ?l 200 mM Pefablock SC, and an equivalent volume
of each sample was mixed with 10 ?l sample buffer (20% 10? reducing agent,
50% 4? LDS sample buffer; Invitrogen) and 5 ?l NP-40 buffer (10 mM Tris-HCl
[pH 7.5], 0.5% deoxycholic acid, 0.5% nonylphenoxylpolyethoxylethanol), heated
to 95°C for 5 min, and separated by 12% Bis-Tris precast polyacrylamide gel
electrophoresis (PAGE) (Invitrogen) at 150 V for 2.5 h in 1? morpholinepro-
panesulfonic acid (MOPS; Invitrogen). Proteins separated by PAGE were trans-
ferred to polyvinylidene fluoride (PVDF) membrane for 1 h at 100 V in transfer
buffer (0.025 M Trizma base, 0.2 M glycine, 20% methanol, pH 8.3). After the
PVDF membranes were blocked overnight at room temperature in Pierce
Blocker, they were probed with PrP-specific antibody BAR224 (kindly supplied
by J. Grassi), followed by horseradish peroxidase-conjugated goat anti-mouse
IgG diluted in Pierce Blocker. Membranes were washed for 1 h after blocking
and between antibodies with wash buffer (0.1 M Tris, 0.15 M NaCl, 0.2% Tween
20, pH 8.0). To visualize PrP bands, the PVDF membranes were developed with
the Amersham ECL detection system and a digital GelDoc (Fuji Intelligent dark
box) using LAS-3000 Lite ImageReader software.
Immunohistochemistry. IHC analysis was performed by employing protocols
described by Spraker et al. (69). Briefly, 3- to 5-mm sections of formalin-fixed,
formic acid-treated tissues were deparaffinized at 60 to 70°C for 1 h, rehydrated
via a series of xylene-ethanol baths, and treated in formic acid a second time (5
min) prior to a 20-min antigen retrieval (10? Dako target retrieval solution)
cycle in a 2100 Retriever (PickCell Laboratories). Slides were further processed
with the aid of a Ventana Discovery autostainer utilizing the Ventana Red Map
stain kit, PrPCWD-specific primary antibody BAR224, and a biotinylated second-
ary goat anti-mouse antibody (Ventana). After autostaining, the slides were
quickly rinsed in a warm water detergent solution, passed through a series of
dehydration baths, and coverslipped.
White-tailed deer bioassays. (i) Cell versus non-cell-associ-
ated inoculates, cohorts 2 and 3. Four of four deer (cohort 2)
inoculated with blood mononuclear cells (leukocytes plus
platelets) from 250 ml CWD?deer blood (1 ? 107to 1.24 ?
108) became tonsil biopsy PrPCWDpositive (by IHC and WB
analyses) between 6 and 19 months p.i. (Table 5). By contrast,
PrPCWDwas not detected in any tissue (tonsil, retropharyngeal
lymph node, or medulla at obex) of any of four deer (cohort 3)
that received the cell-free plasma portion (140 to 150 ml
plasma) from this same 250 ml of CWD?blood.
(ii) Specific blood cell phenotype (2-104?B, CD14?, CD41/
61?) inoculates, cohorts 4 to 6. Four of four deer (cohort 4)
inoculated with 2-104?B cells (1 ? 106to 5 ? 106) and three
of four deer (cohort 5) receiving CD41/61?platelets (6 ? 109
to 35 ? 109) became tonsil biopsy PrPCWDpositive between 12
and 19 months p.i. All four deer (cohort 6) inoculated with
CD14?monocytes (4 ? 105) remained CWD?through the
observation period of 19 months.
(iii) Whole-blood control deer inoculates, cohorts 1 and 7.
PrPCWDwas detected in tonsil biopsies of all eight deer inoc-
ulated with 250 ml whole blood from CWD?deer (cohort 1)
between 6 and 12 months p.i. All deer began to show signs of
TSE disease between 15 and 26 months p.i., including wasting,
hyperphagia, polydipsia, lowered head with wide leg stance,
and lethargy. Both negative control deer (cohort 7) remained
CWD?as determined by IHC and WB analyses.
(iv) White-tailed deer IHC and WB analyses. IHC analysis
(Fig. 1) of terminal lymphoid tissue and medulla obex from
deer inoculated with whole blood, blood mononuclear cells
(leukocytes plus platelets), 2-104?B cells, or CD41/61?plate-
lets demonstrated punctate PrPCWDstaining within lymphoid
follicles and brain tissue typical of that found associated with
CWD-infected cervid controls. The cohort 4 (2-104?B cell)
deer brain IHC analysis in Fig. 1 showed overall less demon-
strable PrPCWDin the medulla than the other CWD?deer
cohorts. Confirmatory WB analysis (Fig. 2) of brain tissue
(medulla at obex) showing the presence of PK-resistant bands
in lanes 6, 8, 12, and 14, corroborates CWD infection. Similar
tissues from deer inoculated with cell-free plasma and CD14?
monocytes did not reveal PrPCWDstaining by either conven-
tional test (IHC or WB analysis) used to verify CWD infection.
Cervidized mouse [Tg(CerPrP-E226)5037?/?] bioassays. (i)
Whole-blood inoculates (i.c., i.v., i.p., or p.o.), cohorts 10 to 13.
Seven of nine mice inoculated i.c. (cohort 10), one of nine mice
inoculated i.v. (cohort 11), five of nine mice inoculated i.p.
(cohort 12), and two of nine mice inoculated p.o. (cohort 13)
with whole blood from a CWD?deer began to show signs of
TSE clinical disease, including weight loss, circling, rigid tail,
hyperactivity, or inactivity, 270 to 490 days p.i. (dpi). Upon
termination (2 to 4 weeks after the initial clinical signs ap-
peared), PrPCWDwas detected in the brains of all of the mice
exhibiting clinical disease by IHC and WB analyses (range, 270
to 490 dpi) (i.c., 275 ? 5 dpi; i.v., 312 dpi; i.p., 340 ? 10 dpi;
p.o., 482 ? 8 dpi).
(ii) Blood mononuclear cell (cell-associated) inoculates
(leukocytes plus platelets) (i.c., i.v., i.p., or p.o.), cohorts 14 to
17. PrPCWDwas detected by IHC and WB analyses of the
TABLE 5. Bioassay results from naı ¨ve deer cohorts inoculated with CWD?blood components
Tonsil PrPCWDresult (no. positive/total) at
postinoculation time of:
at 19 mo p.i.
3 mo6 mo12 mo
Blood mononuclear cells
aRetropharyngeal lymph node, tonsil, and medulla oblongata at obex.
5100MATHIASON ET AL.J. VIROL.
brains of six of nine mice inoculated i.c. (cohort 14), one of
nine mice inoculated i.v. (cohort 15), one of nine mice inocu-
lated i.p. (cohort 16), and zero of nine mice inoculated p.o.
(cohort 17) with blood mononuclear cells (leukocytes plus
platelets) from CWD?deer. Clinical disease included weight
loss and a rough hair coat (range, 275 to 494 dpi) (i.c., 290 ?
15 dpi; i.v., 303 dpi; i.p., 494 dpi; p.o., ?600 dpi). Thus, con-
sistent with results obtained with deer, Tg(cerPrP) mouse bio-
assays also indicated that CWD prion infectivity is associated with
the leukocyte fraction of blood from CWD?cervid donors.
(iii) Non-cell-associated (cell-free) plasma inoculates (i.c.,
i.v., i.p., or p.o.), cohorts 18 to 21. As with bioassay results
obtained with deer, PrPCWDwas not detected in the brains of
mice inoculated with cell-free plasma from CWD?deer which
were monitored for their natural life span (range, 623 to 862
FIG. 1. Terminal lymphoid and brain (medulla at obex) IHC analysis results of naïve deer cohorts inoculated with CWD?blood components.
PrPCWDdemonstrated by IHC analysis in tonsil, brain (medulla oblongata at obex), and retropharyngeal lymph node tissues of deer receiving
whole blood, cell fraction, B cells, or CD41/61?cells from CWD-infected donors. Arrows indicate PrPCWDstaining (red) within the brain and
lymphoid follicles. The arrow with the asterisk indicates a lymphoid follicle negative for PrPCWD.
VOL. 84, 2010CWD PRIONS IN BLOOD CELL FRACTIONS5101
(iv) B-cell inoculates (i.c.), cohorts 22 and 23. In that harvest
of sufficient B cells from blood to permit bioassays in deer and
mice was not possible, 2-104?B cells harvested from retropha-
ryngeal lymph nodes and spleens were analyzed. Five of five
mice inoculated i.c. with retropharyngeal lymph node 2-104?B
cells from a terminal CWD?deer (cohort 22) began to show
clinical TSE disease, including hyperactivity and circling, at
282 ? 7 dpi. One of five mice inoculated i.c. with 2-104?B cells
from the spleen of this same CWD?deer (cohort 23) devel-
oped signs of TSE disease at 180 dpi (weight loss, rough hair
coat, rigid tail). All six mice demonstrating TSE clinical disease
were PrPCWDpositive by IHC and WB analyses of brain tissue
(range, 275 to 289 dpi).
(v) Platelet inoculates (i.c.), cohort 24. Three of five mice
inoculated i.c. with CD41/61?platelets were PrPCWDpositive
by IHC and WB analyses of brain tissue at 305 ? 10 dpi. All
three of these mice exhibited TSE signs of wasting and rough
hair coats (range, 295 to 315 dpi).
(vi) Monocyte blood cell inoculates (i.c.), cohort 25. Neither
TSE clinical disease nor PrPCWDcould be detected in the five
mice inoculated i.c. with CD14?blood cells upon termination
at 600 to 862 dpi (range, 600 to 862 dpi), thus paralleling the
results of deer bioassays.
(vii) Blood component negative control inoculates, cohorts
26 to 41. All mice inoculated with cell components or cell-free
plasma from CWD?white-tailed deer donors remained free of
TSE clinical disease for up to 862 dpi and were PrPCWDneg-
ative upon analysis of brain tissue by IHC and WB.
(viii) Positive and negative control brain inoculates, cohorts
8 and 9. Clinical disease progression (wasting, circling, inability
to right self, hyperactivity, or inactivity) and PrPCWDwere de-
tected by IHC and WB analyses of the brains of all 10 of 10 mice
inoculated i.c. with CWD?brain homogenate at 168 ? 4 dpi,
while 0 of 10 negative control mice were PrPCWDpositive (?600
(ix) Cervidized mouse [Tg(CerPrP-E226)5037?/?] IHC
analysis. IHC analysis (Fig. 3) of sagittal brain tissue sections
FIG. 2. WB analysis of cohorts of naïve deer inoculated with
CWD?blood components. WB demonstration of the typical PK di-
gestion band shift (28 to 35 kDa) associated with prion infection
(medulla at obex) of deer receiving whole blood, mononuclear cell
fraction, B cells, or CD41/61?cells. Deer receiving cell-free plasma or
CD14?cells from CWD-infected donors remained PrPCWDnegative.
Lanes 1 to 4 represent CWD?/CWD?deer controls (10% brain ho-
mogenate) without and with PK digestion at 25 ?g/ml. Lanes 5 to 16,
10% brain homogenate of whole-blood-, mononuclear-cell-, plasma-,
B-cell-, CD41/61?-, or CD14?-inoculated deer without and with PK
digestion at 25 ?g/ml.
FIG. 3. Brain IHC analysis results for Tg(CerPrP-E226)5037?/?mice inoculated with CWD?blood components. PrPCWDdemonstrated by
IHC analysis in sagittal brain sections of Tg(CerPrP-E226)5037?/?mice receiving whole blood, cell fraction, CD41/61?, or B cells from
CWD-infected donors. PrPCWDplaque deposits are typical of those previously described in CWD infection in white-tailed deer (69) and
5102MATHIASON ET AL. J. VIROL.
from terminal mice inoculated with whole blood, blood mono-
nuclear cells (leukocytes plus platelets), 2-104?B cells, or
CD41/61?platelets demonstrated punctate PrPCWDstaining
typically found associated with CWD infection (28). Similar
PrPCWDstaining was not detected in mice inoculated with
cell-free plasma or CD14?monocytes.
(x) Cervidized [Tg(CerPrP-E226)5037?/?] mice as a bioas-
say tool. Multiple routes of inoculation (i.c., i.v., i.p., or p.o.)
were utilized to assess the infectivity of CWD blood fractions
in cervid PrP transgenic (TgCerPrP) mice. Clinical disease and
PrPCWDwere detected in TgCerPrP mice inoculated i.c. with
(i) CWD?brain (100% attack rate), (ii) whole blood (78%),
(iii) blood mononuclear cells (67%), (iv) platelets (60%), (v)
retropharyngeal lymph node 2-104?B cells (100%), or (vi)
splenic 2-104?B cells (20%) (Fig. 4). CWD prion infection was
also detected in mice inoculated with whole blood by the i.p.
(56%) or p.o. (22%) route and mice inoculated i.v. or i.p. with
blood mononuclear cells (11%). All mice inoculated with
CWD?brain (i.c.), monocytes (i.c.), cell-free plasma (i.c., i.v.,
i.p., or p.o.), or blood mononuclear cells (p.o.) were main-
tained for greater than 600 dpi without evidence of clinical
disease or PrPCWDdetection by IHC and WB analyses (Table
6). Thus, the results from Tg(CerPrP-E226)5037?/?bioassay
reinforced the infectious nature of whole blood, blood mono-
nuclear cell, B-cell and platelet blood fractions, and did not
detect prion infectivity in the monocyte or cell-free plasma
blood fractions of blood from CWD-infected deer.
In summary, bioassay results from deer and Tg(CerPrP-
E226)5037?/?mice were concordant. Namely, 79% of the deer
and 35% of the cervidized mice developed CWD prion infec-
tion after inoculation with blood or blood mononuclear cell
fractions [whole blood, 100% of the deer and 42% of the
Tg(CerPrP) mice; blood mononuclear cells, 100% of the deer
and 22% of the mice; 2-104?B cells, 100% of the deer and
60% of the Tg(CerPrP) mice; CD41/61?cells, 75% of the deer
and 60% of the mice], while 0% of those inoculated with either
CD14?cells or cell-free plasma (0% of the deer and 0% of the
mice) became PrPCWDpositive (Table 7). Thus, CWD infec-
tivity segregated with the mononuclear cell, platelet, and
2-104?B-cell-enriched fractions and not with either the
CD14?monocyte or plasma compartments of blood from
Factors influencing the number of cells bioassayed and
routes of inoculation. The number of mononuclear cells, B
cells, monocytes, and platelets bioassayed was influenced by
the total number of cells recoverable from whole blood and
limitations in the volume of inoculum that could safely or
feasibly be administered to mice. The total number of 2-104?
B cells or CD14?monocytes used for inoculation was based on
our previous flow cytometric analyses of white-tailed deer,
which indicated that ?10% of the mononuclear leukocytes
were 2-104?B cells and CD14?cells accounted for ?2% of
the total mononuclear leukocyte population. Because CWD
infection was generated by 107total blood mononuclear cells
(cohort 2), we surmised that a minimum of 1062-104?B cells
(10%) or 2 ? 104CD14?cells (2%) would be sufficient to
determine whether this cell phenotype may carry blood-borne
prion infectivity. Thus, these cell numbers were used for both
deer and mouse cohorts. To mimic blood transfusion dynam-
ics, all deer were inoculated by i.v. infusion. At the time this
study was initiated, very little was known about peripheral
FIG. 4. Attack rates in Tg(CerPrP-E226)5037?/?mice intracranially inoculated with CWD?blood components. Shown are the percentages of
Tg(CerPrP-E226)5037?/?mice that developed prion disease after i.c. inoculation with brain tissue (}), B cells (-), whole blood (Œ), cell fraction
(f), platelets (F), cell-free plasma (?), or monocytes (E) from CWD-infected donor deer or brain tissue from a CWD?donor deer (?).
VOL. 84, 2010CWD PRIONS IN BLOOD CELL FRACTIONS5103
trafficking of CWD in Tg(CerPrP-E226)5037?/?mice. We
therefore used multiple routes of inoculation (i.c., i.v., i.p., or
p.o.) to explore the ability of these mice to support CWD
infection. While attack rates were incomplete, likely due to the
limited volume of blood or blood cells we could administer or
innate differences in sensitivity between deer and Tg(cerPrP)
mice, we did see a similar pattern of CWD infectivity associ-
ated with cell versus cell-free blood components (Fig. 4). These
results suggest that while infection may not be as robust as that
incurred post i.c. inoculation, Tg(CerPrP-E226)5037?/?mice
are capable of establishing and maintaining CWD infection via
peripheral routes of inoculation. Due to logistical reasons and
animal availability, it was not possible in this study to deter-
mine the minimum infectious dose for any of the inocula,
although this information would surely be of interest and could
be approached in subsequent more specific-inoculum-focused
Interval to detection of CWD infection by tonsil biopsy.
Transmission of infectious prions by blood transfusion has now
been established for scrapie, BSE, CJD, and CWD (3, 6, 30, 38,
39, 47, 54, 62). Identifying whether this infectivity is associated
with the cellular, cell-free, or both fractions of blood has been
a bit more challenging. Inherent limitations associated with the
volume of inoculum that can be introduced by i.c. inoculation
in rodent bioassay models impose constraints on assay sensi-
We were able to intravenously inoculate large volumes of
CWD-infected whole blood and equivalent concentrations/
volumes of blood constituents (leukocytes plus platelets,
cell-free plasma, or cell phenotype fractions, i.e., 2-104?B
cells, CD14?monocytes, or CD41/61?platelets) harvested
from the same donor pool into cohorts of naïve white-tailed
deer. In recipient deer, the time to tonsil biopsy positivity
after inoculation of blood, blood mononuclear cells plus
platelets, 2-104?B cells, or CD41/61?platelets was vari-
able—as early as 6 months to as late as 19 months p.i. We
have observed similar PrPCWDdetection kinetics in previous
cervid bioassay studies employing several routes of inocula-
tion and inocula from CWD?deer (53, 54). While we can-
not rule out horizontal transmission from the first positive
animal in each cohort, the time frame for detection in the
remaining deer (6 months) is half that which we have his-
torically observed in deer inoculated with large quantities of
saliva from CWD-infected deer (53, 54), suggesting much
earlier exposure to infectious prions, i.e., more likely to
relate to the experimental inoculum than to animal-to-ani-
TABLE 6. Bioassay of blood components from CWD?deer in Tg(CerPrP) mice
Cohort(s)Donor status Inoculum
Avg time (dpi) of
clinical disease ? SDb
Splenic B cells
RLNgB, splenic B, CD41/61?, or
i.c., i.v., i.p., p.o.
i.c., i.v., i.p., p.o.
i.c., i.v., i.p., p.o.
168 ? 4
270 ? 5
340 ? 10
482 ? 8
290 ? 15
282 ? 7
305 ? 10
bThe natural life spans of Tg(CerPrP-E226)5037?/?mice ranged from 601 to 862 dpi.
cBMC, blood mononuclear cells.
dRLN, retropharyngeal lymph node.
TABLE 7. Summary of white-tailed deer and Tg(CerPrP) mouse
blood component bioassay studies
Blood mononuclear cells
B cells (2-104?)
5104MATHIASON ET AL.J. VIROL.
Infectious prions in cell-associated (leukocyte plus platelet)
blood fraction. Although identification of blood cell-associated
TSE infectivity has been sought with disparate results, in CWD
we detected infectivity in the cellular but not the cell-free plasma
fraction of blood. While some bioassay studies have yielded neg-
ative findings (7, 72), likely due to the restricted sample volume
assayable in rodent models, the transmission of infectious prions
associated with buffy-coat WBC of TSE infected-donors has been
well documented. Kuroda (44), Manuelidis (50), and Brown et al.
(4–6) were the first to demonstrate this association in rodent
models of CJD and Gertsmann-Straussler-Scheinker disease
(GSS). Subsequent to these studies, scrapie prion infectivity in
leukocyte populations has been detected by several researchers
utilizing bioassays in rodent models (3, 10, 36, 50) and in sheep
The replication of prions in the lymphoid tissues precedes
CNS infection in several TSEs (CWD, scrapie, and variant
CJD [vCJD]) (19, 27, 66, 67), raising the potential for hema-
togenous spread via recirculating lymphocytes. Consistent with
this is the fact that leukodepletion reduces blood-borne prion
transmission (24, 63).
Infectious prions in MAb 2-104?B cells. Immunohistolog-
ical detection of PrPRES/Scin lymphoid tissues of scrapie-in-
fected sheep provided some of the first evidence for lymphoid
system involvement in TSE diseases (19, 20, 61, 70). Subse-
quent studies utilizing confocal microscopy confirmed an as-
sociation between PrPRESand immune cells (FDCs, tingible
body macrophages, and B cells) and extended the repertoire of
prion diseases with lymphoid involvement to include CWD and
vCJD (42, 55, 60, 66, 67). B cells have been associated with
PrPREStransport and/or deposition within the lymphoid sys-
tem (9, 10, 18, 22, 23, 32, 43, 56, 64, 66, 81). The present study
supports this contention in demonstrating that MAb 2-104?
primarily B cells harvested from peripheral blood or retropha-
ryngeal lymph nodes contain sufficient infectious prions to
transmit CWD to native or transgenic hosts.
B cells harvested for the deer bioassay studies were collected
from peripheral whole blood. However, as noted earlier, due to
cell loss associated with Ficoll and magnetic bead separation,
we were not able to harvest sufficient 2-104?B cells from
peripheral blood to adequately analyze the infectivity of these
cells in both deer and Tg(CerPrP-E226)5037?/?mice. We
therefore harvested B cells from the spleen and the retropha-
ryngeal lymph nodes from the terminal harvest of one of the
donor deer for bioassays of mice. The B cells harvested from
these tissues (by mechanical disruption/filtration) (for mouse
bioassay) and those collected from whole-blood Ficoll separa-
tions (for deer bioassay) were sorted using MAb 2-104. MAb
2-104 is known to be specific for all peripheral blood B cells in
sheep and has been found to cross-react in a similar fashion in
cervid species (see Fig. S1 in the supplemental material). Mo-
lecular studies suggest that the target antigen is the sheep
homologue of CD72 (80), with more recent findings verifying
that MAb 2-104 may identify FDCs but does not recognize
CD21, T cells, monocytes, or granulocytes (see Fig. S1 in the
supplemental material). It is therefore feasible that MAb 2-104
could label germinal-center FDCs in addition to B cells. It
would be expected that most FDCs would remain within the
connective tissue stroma, be rendered nonviable during me-
chanical disruption, or be removed by the gradient separation
(17). The morphology of the cells harvested for these studies
by these methods supported their identity as B lymphocytes.
We cannot, however, exclude the possibility that CWD infec-
tivity was sheared from dendritic processes of FDCs by me-
chanical disruption and therefore could be a constituent of the
B-cell harvests from retropharyngeal lymph nodes used for the
mouse bioassay study. As FDCs are not found in peripheral
blood, this is not a concern for the B-cell harvests from whole
blood used for deer bioassays.
Differences exist between lymphatic recirculating and non-
recirculating lymphocyte populations found in peripheral
blood and lymphoid tissues. The reassortment of lymphocytes
into these two subsets correlates with lymphocyte lineage and
the expression of maturation and/or adhesion markers. Not all
of the cells in peripheral blood have equal access to the lym-
phatic recirculation pathway (80). Recirculating lymphocytes
constitute 60% of the peripheral blood lymphocytes (PBLs) in
the blood, while nonrecirculating PBLs (40%) are, by defini-
tion, excluded from lymph and confined to blood and the
spleen. CD21 expression correlates with the recirculation com-
petency of these subsets. B cells expressing the CD21 molecule
(CD21?) preferentially migrate across the high endothelial
venules (HEV) and are able to recirculate between the periph-
eral blood and lymphatics, while those not expressing CD21
(CD21?) do not cross the HEV and therefore remain in pe-
ripheral blood (nonrecirculating). Based on previous data, the
spleen would therefore contain representative populations of
both recirculating and nonrecirculating B cells, whereas lymph
nodes would only be expected to contain recirculating B cells
and therefore represent 60% of the total peripheral blood B
cells (25, 80). Given that blood, lymph nodes, and splenic B
cells were able to induce infection in recipient Tg(CerPrP)
mice and deer, it is likely that the recirculating B-cell popula-
tion (i.e., CD21?) was responsible. However, the possibility
cannot be excluded that a unique population of germinal-
center resident B cells was present in samples harvested from
the spleen and lymph nodes, which would not be present in the
peripheral blood. As well, although platelet contamination was
not observed in this cell fraction, it cannot be completely ruled
out as a possible contributor to infectivity.
Infectious prions in CD41/61?platelets. Evidence associat-
ing prion infectivity with platelets has been variable—from no
detection in sporadic-CJD platelets (11) to reported infectivity
in hamster scrapie (36), GSS, and vCJD (10). Here we report
the transmission of infectious CWD via CD41/61?platelets in
both naïve white-tailed deer and cervid transgenic mouse bio-
PrPCis produced endogenously by cells of the platelet lin-
eage, which could implicate platelets in PrPRESpropagation or
trafficking within the body or transmission in contaminated
blood products (11, 34, 35, 37, 48). Forty-three to 53%, 63 to
95%, and 69 to 93% of bovine (3), ovine (4), and white-tailed
deer (unpublished findings) platelets, respectively, express
PrPC, and given the number of circulating platelets versus the
number of leukocytes, the majority of blood-borne PrPCex-
pression is platelet associated (11, 48, 71). This blood compo-
nent could be largely responsible for the transmission of vCJD
by transfusion (45, 46).
Absence of infectious prions in plasma from CWD?deer.
The documentation of TSE infectivity (5, 6, 12, 21, 24, 74), or
VOL. 84, 2010CWD PRIONS IN BLOOD CELL FRACTIONS5105
lack thereof (7, 19, 26, 51, 52), associated with cell-free serum
or plasma is proportionate historically. These discrepancies
may be explained by the inherent limits on the volume of fluid
that can be inoculated in rodent bioassays, the presence/ab-
sence of contaminating cells, or variation in the biology of
prion diseases. Here we report absence of infectious CWD
prions in cell/platelet-free plasma collected from an equivalent
volume of whole blood (250 ml) shown to be capable of in-
fecting naïve white-tailed deer (cohort 1). The virtual elimina-
tion of vCJD transmission by leukoreduction argues strongly
that, as for CWD, prion infectivity is strongly leukocyte and/or
platelet associated in vCJD (14, 15, 45, 46, 75).
Absence of infectious prions in CD14?cells from CWD?
deer. The presence of PrPRESin lymph node tingible body
macrophages of scrapie-infected sheep (1, 31, 40, 56) and
CWD-infected deer (66) led us to investigate the possibility
that circulating CD14?monocytes may contain infectious pri-
ons capable of transmitting disease. Somewhat to our surprise,
the results indicated that in CWD this is not the case. We were
not able to detect PrPCWDin the brain or lymphoid tissues of
white-tailed deer or cervidized mice inoculated with up to 4 ?
105CD14?macrophages (twice the number of cells present in
the blood mononuclear cell inoculum that produced CWD).
In vitro experiments have determined that bone marrow-
derived macrophages can acquire and degrade PrPBSE(65),
which could lead to decreased rates of infection in an in vivo
setting (8). Maignien et al. (49) found that depletion of mac-
rophage numbers at the gut/follicle interface prior to TSE
infection leads to an increase in the infection rate. While mac-
rophages are capable of receptor-mediated uptake of infec-
tious particles, in particular, infectious prions, it appears that
their role may be associated with lysosomal degradation versus
sites of prion amplification or trafficking.
Summary. We have detected infectious prions in the cellular
fraction (mononuclear leukocytes plus platelets) and not in the
cell-free plasma fraction of blood from CWD?deer. B cells
from blood or retropharyngeal lymph nodes and platelets, but
not CD14?monocytes or plasma, contained infectious prions
capable of transmitting CWD. These results (i) support the
identity of a hematogenous route of CWD infection and rein-
force the notion that all tissues are exposed to infection, (ii)
help in understanding the pathogenesis and trafficking of
CWD prions, and (iii) highlight the utility of CWD as a model
in the development of antemortem assays to detect prion in-
We thank A. Avery and M. Zabel for technological conversations
and input into this work, K. O’Rourke and L. Hamburg for performing
the Prnp genotype analysis, J. Grassi and J. Langeveld for their gen-
erous gifts of antibodies, N. Albarado and E. McNulty for sample
collection and tireless dedication to animal welfare, Amy Nalls for
insightful editing, and fellow graduate students D. Seelig, N. Denkers,
and N. Haley for necropsy banter, support, and study critique.
This work was supported by the National Institute of Allergy and
Infectious Diseases, NIH (contract N01-AI-25491).
1. Andre ´oletti, O., P. Berthon, E. Levavasseur, D. Marc, F. Lantier, E. Monks,
J. M. Elsen, and F. Schelcher. 2002. Phenotyping of protein-prion (PrPsc)-
accumulating cells in lymphoid and neural tissues of naturally scrapie-af-
fected sheep by double-labeling immunohistochemistry. J. Histochem. Cy-
2. Angers, R., T. Seward, D. Napier, M. Green, E. Hoover, T. Spraker, K.
O’Rourke, A. Balachandran, and G. Telling. 2009. Chronic wasting disease
prions in elk antler velvet. Emerg. Infect. Dis. 15:696–703.
3. Bons, N., S. Lehmann, N. Mestre-Frances, D. Dormant, and P. Brown. 2002.
Brain and buffy coat transmission of bovine spongiform encephalopathy to
the primate Microcebus murinus. Transfusion 42:513–516.
4. Brown, P. 1995. Can Creutzfeldt-Jakob disease be transmitted by transfu-
sion? Curr. Opin. Hematol. 2:472–477.
5. Brown, P., L. Cervenakova, P. McShane, B. R. Rubenstein, and W. N.
Drohan. 1999. Further studies of blood infectivity in an experimental model
of transmissible spongiform encephalopathy, with an explanation of why
blood components do not transmit Creutzfeldt-Jakob disease in humans.
6. Brown, P., R. G. Rohwer, B. C. Dunstan, C. MacAuley, D. C. Gajdusek, and
W. N. Drohan. 1998. The distribution of infectivity in blood components and
plasma derivatives in experimental models of transmissible spongiform en-
cephalopathy. Transfusion 38:810–816.
7. Bruce, M. E., I. McConnell, R. G. Will, and J. W. Ironside. 2001. Detection
of variant Creutzfeldt-Jakob disease infectivity in extraneural tissues. Lancet
8. Carp, R. I., and S. M. Callahan. 1981. In vitro interaction of scrapie agent
and mouse peritoneal macrophages. Intervirology 16:8–13.
9. Casaccia, P., A. Ladogana, Y. G. Xi, and M. Pocchiari. 1989. Levels of
infectivity in the blood throughout the incubation period of hamsters pe-
ripherally injected with scrapie. Arch. Virol. 108:145–149.
10. Cervenakova, L., O. Yakovleva, C. McKenzie, S. Kolchinsky, L. McShane,
W. N. Drohan, and P. Brown. 2003. Similar levels of infectivity in the blood
of mice infected with human-derived vCJD and GSS strains of transmissible
spongiform encephalopathy. Transfusion 43:1687–1694.
11. Choi, E. M., M. D. Geschwind, C. Deering, K. Pomeroy, A. Kuo, B. L. Miller,
J. G. Safar, and S. B. Prusiner. 2009. Prion proteins in subpopulations of
white blood cells from patients with sporadic Creutzfeldt-Jakob disease. Lab.
12. Clarke, M. C., and D. A. Haig. 1967. Presence of the transmissible agent of
scrapie in the serum of affected mice and rats. Vet. Rec. 80:504.
13. Clarke, P., and A. C. Ghani. 2005. Projections of the future course of the
primary vCJD epidemic in the UK: inclusion of subclinical infection and the
possibility of wider genetic susceptibility. J. R. Soc. Interface 2:19–31.
14. Cleemput, I., M. Leys, D. Ramaekers, and L. Bonneux. 2006. Balancing
evidence and public opinion in health technology assessments: the case of
leukoreduction. Int. J. Technol. Assess. Health Care 22:403–407.
15. Coste, J., C. Prowse, R. Eglin, and C. Fang. 2009. A report on transmissible
spongiform encephalopathies and transfusion safety. Vox Sang. 96:284–291.
16. Deslys, J., C. Lasmezas, and D. Dormont. 1994. Selection of specific strains
in iatrogenic Creutzfeldt-Jakob disease. Lancet 343:848–849.
17. Eaton, S., M.-J. Anderson, S. Hamilton, L. Gonzalez, J. Sales, M. Feffrey,
H. W. Reid, M. S. Rocchi, and F. Chianini. 2009. CD21 B cell populations
are altered following subcutaneous scrapie inoculation in sheep. Vet. Immu-
nol. Immunopathol. 131:105–109.
18. Eklund, C. M., W. Hadlow, and R. C. Kennedy. 1963. Some properties of the
scrapie agent and its behavior in mice. Proc. Soc. Exp. Biol. Med. 112:974–
19. Eklund, C. M., R. C. Kennedy, and W. J. Hadlow. 1967. Pathogenesis of
scrapie virus infection in the mouse. J. Infect. Dis. 117:15–22.
20. Fraser, H., and A. G. Dickinson. 1970. Pathogenesis of scrapie in the mouse:
the role of the spleen. Nature 226:462–463.
21. Gajdusek, D. C., C. J. Gibbs, Jr., and M. Alpers (ed.). 1965. Slow, latent, and
temperate virus infections. NINDB monograph no. 2. U.S. Department of
Health, Education, and Welfare, Washington, DC.
22. Glatzel, M., E. Abela, M. Maissen, and A. Aguzzi. 2003. Extraneural patho-
logic prion protein in sporadic Creutzfeldt-Jakob disease. N. Engl. J. Med.
23. Gordon, W. 1957. Discussion to Palmer, A. C. Studies in scrapie. Vet. Rec.
24. Gregori, L., N. McCombie, D. Palmer, P. Birch, S. O. Sowemimo-Coker, A.
Giulivi, and R. G. Rohwer. 2004. Effectiveness of leucoreduction for removal
of infectivity of transmissible spongiform encephalopathies from blood. Lan-
25. Gupta, V., I. McConnell, R. G. Dalziel, and J. Hopkins. 1998. Two B cell
subpopulations have distinct recirculation characteristics. Eur. J. Immunol.
26. Hadlow, W. J., C. M. Eklund, and R. C. Kennedy. 1974. Course of experi-
mental scrapie virus infection in the goat. J. Infect. Dis. 129:559–567.
27. Hadlow, W. J., R. C. Kennedy, and R. E. Race. 1982. Natural infection of
Suffolk sheep with scrapie virus. J. Infect. Dis. 146:657–664.
28. Haley, N., D. Seelig, M. Zabel, G. Telling, and E. Hoover. 2009. Detection of
CWD prions in urine and saliva of deer by transgenic mouse bioassay. PLoS
29. Health Protection Agency. 18 January 2007, posting date. Fourth case of trans-
fusion-associated vCJD infection in the United Kingdom. http://www.hpa.org
5106MATHIASON ET AL.J. VIROL.
30. Health Protection Agency. 17 February 2009, posting date. vCJD abnor- Download full-text
mal prion protein found in a patient with haemophilia at post mortem.
31. Herrmann, L. M., W. P. Cheevers, W. C. Davis, D. P. Knowles, and K. I.
O’Rourde. 2003. CD21-positive follicular dendritic cells: a possible source of
PrPsc in lymph node macrophages in scrapie infected sheep. Am. J. Pathol.
32. Herzog, C., J. Riviere, N. Lescoutra-Etchegaray, A. Charbonnier, V. Leblanc,
N. Sales, J. P. Deslys, and C. I. Lasmezas. 2005. PrPTSE distribution in a
primate model of variant, sporadic, and iatrogenic Creutzfeldt-Jakob dis-
ease. J. Virol. 79:14339–14345.
33. Hilton, D. A., A. C. Ghani, L. Conyers, P. Edwards, L. McCardle, D. Ritchie,
M. Penney, D. Hegazy, and J. W. Ironside. 2004. Prevalence of lymphore-
ticular prion protein accumulation in UK tissue samples. J. Pathol. 203:733–
34. Holada, K., T. H. Mondoro, J. Juller, and J. G. Vostal. 1998. Increased
expression of phosphatidylinositol-specific phospholipase C resistant prion
proteins on the surface of activated platelets. Br. J. Haematol. 103:276–282.
35. Holada, K., J. Simak, P. Brown, and J. G. Vostal. 2007. Divergent expression
of cellular prion protein on blood cells of human and nonhuman primates.
36. Holada, K., J. G. Vostal, P. W. Theisen, C. MacAuley, L. Gregori, and R. G.
Rohwer. 2002. Scrapie infectivity in hamster blood is not associated with
platelets. J. Virol. 76:4649–4950.
37. Holada, K., and J. G. Vostal. 2000. Different levels of prion protein (PrPc)
expression on hamster, mouse and human blood cells. Br. J. Haematol.
38. Houston, F., J. D. Foster, A. Chong, N. Hunter, and C. J. Bostock. 2000.
Transmission of BSE by blood transfusion in sheep. Lancet 356:999–1000.
39. Hunter, N., J. Foster, A. Chong, S. McCutcheon, D. Parnham, S. Eaton, C.
MacKenzie, and F. Houston. 2002. Transmission of prion diseases by blood
transfusion. J. Gen. Virol. 83:2897–2905.
40. Jeffrey, M., G. McGovern, S. Martin, C. M. Goodsir, and K. L. Brown. 2000.
Cellular and sub-cellular localisation of PrP in the lymphoreticular system of
mice and sheep. Arch. Virol. Suppl. 16:23–38.
41. Kim, T. Y., H. J. Shon, Y. S. Joo, U. K. Mun, K. S. Kang, and Y. S. Lee. 2005.
Additional cases of chronic wasting disease in imported deer in Korea. J.
Vet. Med. Sci. 67:753–759.
42. Kitamoto, T., T. Muramoto, S. Mohri, K. Doh-Ura, and J. Tateishi. 1991.
Abnormal isoform of prion protein accumulates in follicular dendritic cells in
mice with Creutzfeldt-Jakob disease. J. Virol. 65:6292–6295.
43. Klein, M. A., R. Frigg, E. Flechsig, A. J. Raeber, U. Kalinke, H. Bluethmann,
F. Bootz, M. Suter, R. M. Zinkernagel, and A. Aguzzi. 1997. A crucial role for
B cells in neuroinvasive scrapie. Nature 390:687–690.
44. Kuroda, Y., C. J. Gibbs, Jr., L. A. Herbert, and D. C. Gajdusek. 1983.
Creutzfeldt-Jakob disease in mice: persistent viremia and preferential rep-
lication of virus in low-density lymphocytes. Infect. Immun. 41:154–161.
45. Lefre `re, J., and P. Hewit. 2009. From mad cows to sensible blood transfusion:
the risk of prion transmission by labile blood components in the United
Kingdom and in France. Transfusion 49:797–812.
46. Lefre `re, J. J., and B. Danic. 2009. Pictorial representation of transfusion over
the years. Transfusion 49:1007–1017.
47. Llewelyn, C. A., P. E. Hewitt, R. S. Knight, K. Amar, S. Cousens, J. Mack-
enzie, and R. G. Will. 2004. Possible transmission of variant Creutzfeldt-
Jakob disease by blood transfusion. Lancet 363:417–421.
48. MacGregor, I., J. Hope, G. Barnard, L. Kirby, O. Drummond, D. Pepper, V.
Hornsey, R. Barclay, H. Bessos, M. Turner, and C. Prowse. 1999. Applica-
tion of a time-resolved fluoroimmunoassay for the analysis of normal prion
protein in human blood and its components. Vox Sang. 77:88–96.
49. Maignien, T., M. Shakweh, P. Calvo, D. Marce, N. Sales, E. Fattal, J. P.
Deslys, P. Couvreur, and C. I. Lasmezas. 2005. Role of gut macrophages in
mice orally contaminated with scrapie or BSE. Int. J. Pharm. 298:293–304.
50. Manuelidis, E. E., J. H. Kim, J. R. Mericangas, and L. Manuelidis. 1985.
Transmission to animals of Creutzfeldt-Jakob disease from human blood.
51. Marsh, R. F., and R. P. Hanson. 1969. Transmissible mink encephalopathy:
neuroglial response. Am. J. Vet. Res. 30:1637–1642.
52. Marsh, R. F., J. M. Miller, and R. P. Hanson. 1973. Transmissible mink
encephalopathy: studies on the peripheral lymphocyte. Infect. Immun.
53. Mathiason, C. K., S. A. Hays, J. Powers, J. Hayes-Klug, J. Langenberg, S. J.
Dahmes, D. A. Osborn, K. V. Miller, R. J. Warren, G. L. Mason, and E. A.
Hoover. 2009. Infectious prions in pre-clinical deer and transmission of
chronic wasting disease solely by environmental exposure. PLoS One
54. Mathiason, C. K., J. G. Powers, S. J. Dahmes, D. A. Osborn, K. V. Miller,
R. J. Warren, G. L. Mason, S. A. Hays, J. Hayes-Klug, D. M. Seelig, M. A.
Wild, L. L. Wolfe, T. R. Spraker, M. W. Miller, C. J. Sigurdson, G. C. Telling,
and E. A. Hoover. 2006. Infectious prions in the saliva and blood of deer with
chronic wasting disease. Science 314:133–136.
55. McBride, P. A., P. Eikelenboom, G. Kraal, H. Fraser, and M. E. Bruce. 1992.
PrP protein is associated with follicular dendritic cells of spleens and lymph
nodes in uninfected and scrapie-infected mice. J. Pathol. 168:413–418.
56. McGovern, G., and M. Jeffrey. 2007. Scrapie-specific pathology of sheep
lymphoid tissues. PLoS One 2:e1304.
57. Miller, M., E. Williams, N. Hobbs, and L. L. Wolfe. 2004. Environmental
sources of prion transmission in mule deer. Emerg. Infect. Dis. 10:1003–
58. Miller, M. W., and E. S. Williams. 2003. Prion disease: horizontal prion
transmission in mule deer. Nature 425:35–36.
59. Miller, M. W., M. A. Wild, and E. S. Williams. 1998. Epidemiology of chronic
wasting disease in captive Rocky Mountain elk. J. Wildl. Dis. 34:532–538.
60. Muramoto, T., T. Kitamoto, J. Tateishi, and I. Goto. 1992. The sequential
development of abnormal prion protein accumulation in mice with
Creutzfeldt-Jakob disease. Am. J. Pathol. 140:1411–1420.
61. Pattison, I. H., and K. M. Jones. 1968. Detection of the scrapie agent in
tissues of normal mice and in tumours of tumour-bearing but otherwise
normal mice. Nature 218:102–104.
62. Peden, A. H., M. W. Head, D. L. Ritchie, J. E. Bell, and J. W. Ironside. 2004.
Preclinical vCJD after blood transfusion in a PRNP codon 129 heterozygous
patient. Lancet 364:527–529.
63. Prowse, C. V., and A. Bailey. 2000. Validation of prion removal by leukocyte-
depleting filters: a cautionary tale. Vox Sang. 79:248.
64. Raeber, A. J., M. A. Klein, R. Frigg, E. Flechsig, A. Aguzzi, and C. Weiss-
mann. 1999. PrP-dependent association of prions with splenic but not cir-
culating lymphocytes of scrapie-infected mice. EMBO J. 18:2702–2706.
65. Rybner-Barnier, C., C. Jacquemot, C. Cuche, G. Dorre, L. Majlessi, M. M.
Gabellec, A. Moris, O. Schwartz, J. Di Santo, A. Cumano, C. Leclerc, and F.
Lazarini. 2006. Processing of the bovine spongiform encephalopathy-specific
prion protein by dendritic cells. J. Virol. 80:4656–4663.
66. Sigurdson, C. J., C. Barillas-Mury, M. W. Miller, B. Oesch, L. J. van Keulen,
J. P. Langeveld, and E. A. Hoover. 2002. PrP(CWD) lymphoid cell targets in
early and advanced chronic wasting disease of mule deer. J. Gen. Virol.
67. Sigurdson, C. J., E. S. Williams, M. W. Miller, T. R. Spraker, K. I. O’Rourke,
and E. A. Hoover. 1999. Oral transmission and early lymphoid tropism of
chronic wasting disease PrPres in mule deer fawns (Odocoileus hemionus).
J. Gen. Virol. 80(Pt. 10):2757–2764.
68. Sohn, H. J., J. H. Kim, K. S. Choi, J. J. Nah, Y. S. Joo, Y. H. Jean, S. W. Ahn,
O. K. Kim, and A. Balachandran. 2002. A case of chronic wasting disease in
an elk imported to Korea from Canada. J. Vet. Med. Sci. 64:855–858.
69. Spraker, T. R., R. R. Zink, B. A. Cummings, M. A. Wild, M. W. Miller, and
K. I. O’Rourke. 2002. Comparison of histological lesions and immunohisto-
chemical staining of proteinase-resistant prion protein in a naturally occur-
ring spongiform encephalopathy of free-ranging mule deer (Odocoileus
hemionus) with those of chronic wasting disease of captive mule deer. Vet.
70. Stamp, J. T., J. G. Brotherston, I. Zlotnik, J. M. Mackay, and W. Smith.
1959. Further studies on scrapie. J. Comp. Pathol. 69:268–280.
71. Starke, R., P. Harrison, I. Mackie, G. Wang, J. D. Erusalimsky, R. Gale,
J. M. Masse, E. Cramer, A. Pizzey, J. Biggerstaff, and S. Machin. 2005. The
expression of prion protein (PrP(C)) in the megakaryocyte lineage. J.
Thromb. Haemost. 3:1266–1273.
72. Tamai, Y., H. Kojima, R. Kitajima, F. Taguchi, Y. Ohtani, T. Kawaguchi, S.
Miura, M. Sato, and Y. Ishihara. 1992. Demonstration of the transmissible
agent in tissue from a pregnant woman with Creutzfeldt-Jakob disease.
N. Engl. J. Med. 327:649.
73. Tateishi, J., T. Kitamato, and H. Hiratani. 1985. Creutzfeldt-Jakob disease
pathogen in growth hormone preparations is eliminatable. Lancet 2:1299–
74. Taylor, D. M., I. McConnell, and C. E. Ferguson. 2000. Closely similar values
obtained when the ME7 strain of scrapie agent was titrated in parallel by two
individuals in separate laboratories using two sublines of C57BL mice. J. Vi-
rol. Methods 86:35–40.
75. Turner, M. L., and C. A. Ludlam. 2009. An update on the assessment and
management of the risk of transmission of variant Creutzfeldt-Jakob disease
by blood and plasma products. Br. J. Haematol. 144:14–23.
76. U.S. Geological Survey. 23 March 2009, posting date. Map of chronic wasting
disease in North America. http://www.nwhc.usgs.gov/disease_information
77. Williams, E. 2005. Chronic wasting disease. Vet. Pathol. 42:530–549.
78. Williams, E. S., and M. W. Miller. 2002. Chronic wasting disease in deer and
elk in North America. Rev. Sci. Tech. 21:305–316.
79. Williams, E. S., and S. Young. 1992. Spongiform encephalopathies in Cervi-
dae. Rev. Sci. Tech 11:551–567.
80. Young, A. J., W. L. Marston, M. Dessing, L. Dudler, and W. R. Hein. 1997.
Distinct recirculating and non-recirculating B-lymphocyte pools in the pe-
ripheral blood are defined by coordinated expression of CD21 and L-selectin.
81. Zabel, M. D., M. Heikanwalder, M. Prinz, I. Arrighi, P. Schwarz, J. Kranich,
A. von Teichman, K. M. Haas, N. Zeller, T. F. Tedder, J. H. Weis, and A.
Aguzzi. 2007. Stromal complement receptor CD21/35 facilitates lymphoid
prion colonization and pathogenesis. J. Immunol. 179:6144–6152.
VOL. 84, 2010CWD PRIONS IN BLOOD CELL FRACTIONS5107