PrPCWD lymphoid cell targets in early and advanced chronic wasting disease of mule deer

Christina J. Sigurdson1, Carolina Barillas-Mury1, Michael W. Miller2, Bruno Oesch3, Lucien J. M. van Keulen4, Jan P. M. Langeveld4 and Edward A. Hoover1

Department of Microbiology, Immunology and Pathology, College of Veterinary Medicine and Biomedical Sciences, Colorado State University, Fort Collins, CO 80523-1671, USA1
Colorado Division of Wildlife, Wildlife Research Center, 317 West Prospect Road, Fort Collins, CO 80526-2097, USA2
Prionics AG, Wagistrasse 27a, 8952 Schlieren, Switzerland3
Institute for Animal Science and Health (ID-Lelystad), Edelhertweg 15, 8219 PH Lelystad, The Netherlands4

Author for correspondence: Edward Hoover. Fax +1 970 491 0523. e-mail ehoover{at}lamar.colostate.edu


   Abstract
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
Up to 15% of free-ranging mule deer in northeastern Colorado and southeastern Wyoming, USA, are afflicted with a prion disease, or transmissible spongiform encephalopathy (TSE), known as chronic wasting disease (CWD). CWD is similar to a subset of TSEs including scrapie and variant Creutzfeldt–Jakob disease in which the abnormal prion protein isoform, PrPCWD, accumulates in lymphoid tissue. Experimental scrapie studies have indicated that this early lymphoid phase is an important constituent of prion replication interposed between mucosal entry and central nervous system accumulation. To identify the lymphoid target cells associated with PrPCWD, we used triple-label immunofluorescence and high-resolution confocal microscopy on tonsils from naturally infected deer in advanced disease. We detected PrPCWD primarily extracellularly in association with follicular dendritic and B cell membranes as determined by frequent co-localization with antibodies against membrane bound immunoglobulin and CD21. There was minimal co-localization with cytoplasmic labels for follicular dendritic cells (FDC). This finding could indicate FDC capture of PrPCWD, potentially in association with immunoglobulin or complement, or PrPC conversion on FDC. In addition, scattered tingible body macrophages in the germinal centre contained coarse intracytoplasmic aggregates of PrPCWD, reflecting either phagocytosis of PrPCWD on FDC processes, apoptotic FDC or B cells, or actual PrPCWD replication within tingible body macrophages. To compare lymphoid cell targets in early and advanced disease, we also examined: (i) PrPCWD distribution in lymphoid cells of fawns within 3 months of oral CWD exposure and (ii) tonsil biopsies from preclinical deer with naturally acquired CWD. These studies revealed that the early lymphoid cellular distribution of PrPCWD was similar to that in advanced disease, i.e. in a pattern suggesting FDC association. We conclude that in deer, PrPCWD accumulates primarily extracellularly and associated with FDCs and possibly B cells – a finding which raises questions as to the cells responsible for pathological prion production.


   Introduction
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
Chronic wasting disease (CWD) is the only prion disease, or transmissible spongiform encephalopathy (TSE), known to affect free-ranging wildlife (Spraker et al., 1997 ; Williams & Young, 1992 , 1993 ). In endemic areas of Colorado and Wyoming, USA, up to 15% of free-ranging mule deer are infected (Miller et al., 2000 ) and the potential of CWD transmission to livestock or humans is unknown. Even transmission routes among deer remain obscure, although epidemiological evidence suggests lateral transmission (Miller et al., 2000 ). The pathogenesis of CWD is beginning to unfold. Recent studies have revealed the abnormal isoform of the prion protein (PrPres) in lymphoid tissue (Sigurdson et al., 1999 ) in a pattern very similar to that described in natural scrapie of sheep (van Keulen et al., 1996 ) and variant Creutzfeldt–Jakob disease (vCJD) of humans (Hill et al., 1999 ).

Lymphoid tropism differs among the TSEs – these differences possibly reflect variants of prion disease pathogenesis. For example, in bovine spongiform encephalopathy (BSE) no detectable PrPres or infectivity is detectable in spleen (Somerville et al., 1997 ) or lymph nodes (Wells et al., 1998 ), unlike CWD, sheep scrapie and vCJD (Hill et al., 1999 ; Spraker et al., 2002; van Keulen et al., 1996 ). However, experimental inoculation of BSE into sheep does result in detectable lymphoid PrPres (Foster et al., 2001 ; Jeffrey et al., 2001 ). Moreover, lymphotropism appears to be determined not only by host species, but also by prion strain: for example, humans with vCJD have lymphoid PrPres accumulation or infectivity (Bruce et al., 2001 ; Hill et al., 1999 ; Hilton et al., 1998 ) whereas humans with sporadic or iatrogenic CJD do not have lymphoid PrPres accumulation (Hill et al., 1999 ).

Naturally infected deer with advanced CWD have CWD PrPres (PrPCWD) disseminated throughout lymph nodes, spleen, tonsils and Peyer’s patches. In tonsils, PrPCWD accumulation is restricted primarily to germinal centres and is present in >50% of secondary follicles (Spraker et al., 2002). In fawns orally inoculated with CWD brain homogenate, PrPCWD was detected in alimentary-associated lymphoid tissues as early as 6 weeks post-inoculation (p.i.). In these early stages of infection, PrPCWD was limited to <30% of secondary follicles, which were typically clustered, suggesting a common conduit or seeding site into the draining lymph node (Sigurdson et al., 1999 ).

The mechanisms of lymphoid tissue PrPCWD accumulation remain uncertain, although studies in natural and experimental scrapie (Andreoletti et al., 2000 ; Brown et al., 2000 ; Jeffrey et al., 2000 ; Kitamoto et al., 1991 ; McBride et al., 1992 ; Montrasio et al., 2000 ), CJD in mice (Manuelidis et al., 2000 ) and vCJD in humans (Hill et al., 1999 ) provide evidence for PrPres association with follicular dendritic cells (FDC) and/or tingible body (TB) macrophages. With the abundant PrPCWD in lymphoid tissues of deer, it seems possible that PrPCWD-containing lymphoid cells could traffic into the blood. Several studies have established that PrPres strongly correlates with infectivity (Bolton et al., 1991 ; McKinley et al., 1983 ; Race et al., 1998 ). Therefore, with the hope of gaining insight into potential trafficking, conversion or capture sites of PrPCWD, we studied the spatial relationship of the protease-resistant prion protein to lymphoid cell phenotypes in the tonsils and lymph nodes of mule deer naturally or experimentally infected with CWD by triple-immunofluorescent labelling and laser scanning confocal microscopy. We found PrPCWD almost exclusively in association with cell membrane surfaces. In addition, smaller deposits of PrPCWD were detected intracytoplasmically in CD68+ macrophages or dendritic cells within germinal centres and much less commonly within the paracortical zone of lymph nodes. These results are reminiscent of those of Jeffrey et al. (2000) regarding PrPSc and suggest to us that either: (a) PrPCWD conversion occurs at the surface rather than within FDCs or (b) PrPCWD formation occurs at distant sites and is concentrated at FDC surfaces.


   Methods
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
{blacksquare} CWD-infected deer and tissue collection.
Tonsils or retropharyngeal lymph nodes from CWD-positive deer were acquired from three groups of captive mule deer (Odocoileus hemionus) in various stages of infection: (1) tonsils from six deer with naturally occurring, clinical CWD, (2) retropharyngeal lymph nodes from two fawns orally inoculated with a CWD brain homogenate and euthanized at 42 and 78 days p.i., and (3) tonsil biopsies from three naturally infected, asymptomatic deer from a captive herd with endemic CWD. The asymptomatic deer eventually developed clinical signs of CWD and were euthanized (CWD confirmed with brain immunohistochemical staining (IHC) for PrPCWD). Tonsils were fixed in 10% neutral buffered formalin for 1–3 days then immersed in 88% formic acid for 1 h and embedded in paraffin.

The clinically affected CWD-positive deer were diagnosed by: (1) histological lesions of CWD in the medulla oblongata including perikaryonic neuronal vacuoles, spongiform degeneration of the neuropil and astrocytosis, and (2) abundant PrPCWD staining in the medulla oblongata by IHC (methods described in Sigurdson et al., 2001 ). Deer were confirmed as CWD-negative by the absence of histological brain lesions and negative staining for PrPCWD in brain and tonsil.

{blacksquare} Negative control deer and tissues.
Tonsils from CWD-negative mule deer were acquired from two sources: (1) adult deer from the CWD non-endemic area (non-endemic area established by methods in Miller et al., 2000 ) and (2) two mule deer fawns inoculated with CWD-negative brain homogenate from a previous study (Sigurdson et al., 1999 ). Tissues were similarly fixed and processed.

{blacksquare} Phenotype antibodies.
Several antibodies which recognize lymphoid epitopes on deer lymphoid cells were used. These included antibodies which recognize: (1) lambda light chain (DAKO), present in antigen–antibody complexes on FDC membrane surfaces and on B cells, (2) cc21 (CD21 or complement receptor type 2) (antibody generously donated by Dr Chris Howard), a receptor that traps immune complexes on FDC surfaces also expressed by B cells (Zabel & Weis, 2001 ), (3) CD68 (Serotec), an intracytoplasmic, lysosome-associated epitope within macrophages and human DC (Betjes et al., 1991 ), (4) ferritin (DAKO), a large protein surrounding a core of ferric oxide which functions to store and detoxify iron (Morikawa et al., 1995 ) in macrophages (Kindblom et al., 1982 ), (5) heat shock protein 70 (HSP70) (DAKO) in macrophages (Bachelet et al., 1998 ), (6) vimentin (DAKO), an intermediate filament in TB macrophages (Giorno, 1985 ) and FDC (Tsunoda et al., 1990 ), (7) anti-FDC (DAKO), which targets a 120 kDa epitope in FDC of humans (Raymond et al., 1997 ) and has been shown to cross-react with sheep FDC (Lezmi et al., 2001 ), (8) S100 (DAKO), a calcium-binding protein present in FDC and/or TB macrophages, depending on the species (Carbone et al., 1988 ), and (9) CD3 (DAKO), an intracytoplasmic domain of the CD3 epsilon chain of T cells.

{blacksquare} Immunofluorescent staining.
Tissue sections (6 µm) were mounted onto positively charged glass slides, deparaffinized, hydrated, autoclaved in a buffer solution (DAKO Target Antigen Retrieval) for 12 min at 121 °C, and cooled for 5 min. Sections were rinsed in PBS and immersed in 3% H2O2 for 15 min to quench endogenous peroxidase. Sections were then briefly rinsed in PBS and incubated in TNB blocking solution (NEN Sciences) for 30 min followed by exposure to 1–2 lymphoid phenotype antibodies and anti-PrP antibody 6H4 (monoclonal, IgG, 1:200 dilution) or R522 (polyclonal, 1:1500 dilution) for 30 min at room temperature. mAb 6H4 recognizes a conserved sequence of the prion protein, corresponding to the human amino acid sequence 144–152 (Korth et al., 1997 ). R522 recognizes ovine PrP 94–105 (Garssen et al., 2000 ; van Keulen et al., 1995 ). Antibodies were diluted in a protein block containing goat serum (Biogenex).

Since HSP epitopes appear to be destroyed by autoclaving, slides stained for HSP and PrP were initially labelled for HSP, followed by autoclaving and labelling for PrPCWD. In general, phenotype antibodies were labelled with FITC or Alexa 488 (Molecular Probes) and PrP labelled with CY3. In sections labelled for HSP or CD68, PrPCWD was labelled with FITC. Tyramide amplification (NEN Sciences) was used to enhance stain signal on R522, ferritin and HSP labels. Slides were coverslipped using anti-fade mounting media (Molecular Probes). CWD-negative deer tissues were incubated with an anti-PrP antibody and an isotype- and concentration-matched rabbit or mouse antibody to control for the phenotype antibody.

{blacksquare} Confocal microscopy.
To co-localize the cell phenotype marker and PrPCWD, triple immunofluorescently labelled sections were examined using an Olympus FLUOVIEW laser scanning confocal microscope equipped with 12-bit resolution which allows for data acquisition from three fluorescent channels using three lasers, Argon 488 nm, HeNe 543 nm and HeNe 622 nm; these emit in the green, red and far-red spectra, respectively. Secondary follicles were selected from each tonsil section and sequentially scanned using the three lasers.

{blacksquare} Quantification of co-localization of PrPCWD and phenotype marker.
Images from each deer were analysed using Metamorph software (Universal Imaging Corp., West Chester, PA) applying the colour thresholding tool to differentiate the positively stained cells from the unstained cells. Percent co-localization of PrPCWD with the phenotype marker stain was measured using the co-localization tool and recorded on a Microsoft Excel spreadsheet. For each tissue section, two follicles (900x magnification) were analysed for PrPCWD and phenotype marker co-labelling, and the results were averaged. Data were analysed using Student’s t-test. Significance was defined at P<0·05.

{blacksquare} Dual immunocytochemical (ICC) staining.
To determine whether PrPCWD could be associated with individual cells from a CWD-infected lymph node, we collected the retropharyngeal lymph node into cold cell culture medium immediately after euthanasia. Single cell suspensions were prepared by mincing and incubating 2 mm3 sections in serum-enriched medium containing collagenase, dispase and DNase at 37 °C with agitation to digest the stroma and release the cells. The cells were pelleted by centrifugation, washed in PBS, and then cytocentrifuged onto positively charged glass slides. Cells were fixed in 10% buffered formalin for 15 min and pretreated by hydrated autoclaving if necessary immediately prior to immunostaining.

The ICC protocol employed an automated immunostainer (Ventana Medical Systems) and was separated into two stages. First, the cells were labelled with a phenotype marker using the appropriate phenotype antibody, a biotinylated secondary antibody, a horseradish peroxidase–streptavidin conjugate and a diaminobenzadine chromagen. Second, hydrated autoclaving was performed on cell preparations not previously autoclaved and the cells were labelled for PrPCWD using PrP mAb F99/97.6.1 (generously provided by Dr Katherine O’Rourke) (Spraker et al., 2002), a biotinylated secondary antibody, an alkaline phosphatase–streptavidin conjugate, a substrate chromagen (fast red A), and a haematoxylin and bluing counterstain (Ventana Medical Systems). mAb F99/97.6.1 reacts with a conserved epitope (residues QYQRES) on the prion protein of mule deer, Rocky Mountain elk, domestic sheep and cattle (Spraker et al., 2002). An isotype-matched, irrelevant antibody was substituted in the ICC protocol as a negative control for the phenotype marker. The anti-PrP antibody was applied to both CWD-negative and -positive deer cell preparations.

IHC was performed on lymphoid tissue as described for the ICC utilizing anti-PrP mAbs F89/160.1.5 and F99/97.6.1. mAb F89/160.1.5 recognizes a conserved epitope of the prion protein of mule deer, elk, sheep and cattle (residues IHFG) (O’Rourke et al., 1998 ).


   Results
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
Lymphoid cells in the germinal centres include FDC, TB macrophages, T and B lymphocytes, and germinal centre dendritic cells. Germinal centre dendritic cells, a dendritic cell subset in the tonsil that presents antigen to germinal centre B cells, have been described in humans (Grouard et al., 1996 ; Summers et al., 2001 ) but not in ruminants.

Because PrPCWD deposits accumulate within germinal centres of primary and secondary lymphoid follicles, we focused on phenotype marker antibodies which would target FDC, B and T lymphocytes, and TB macrophages. To ensure that the human antigen-derived phenotype antibodies recognized the appropriate target epitope, we compared the cell staining patterns of our phenotype antibodies in human and deer tonsil sections and determined that the antibodies identified lymphoid cells with similar morphology and anatomical distribution.

PrPCWD in lymphoid germinal centres
In tonsils of all CWD-infected deer examined by IHC, PrPCWD was concentrated primarily in lymphoid follicle germinal centres (Fig. 1). Tonsils from deer with clinical CWD or tonsil biopsies from preclinical, CWD-infected deer had a high frequency (~80–100%) of PrPCWD-positive follicles. By contrast, in fawns examined 7 to 11 weeks after oral CWD exposure, <30% of retropharyngeal lymph node follicles contained detectable PrPCWD. Although PrPCWD was found primarily within the germinal centres, it was also detected occasionally in cells within perifollicular areas (Fig. 1).



View larger version (200K):
[in this window]
[in a new window]
 
Fig. 1. Typical abundant follicular PrPCWD deposition in a lymph node follicle from a deer with advanced CWD. Note the network-like lattice of PrPCWD deposition defining the central zone of the follicle (bracket) and isolated PrPCWD-positive cells at the periphery of the follicle (arrows). Anti-PrP antibody used: F99/97.6.1. Bar, 10 µm.

 
PrPCWD accumulates on FDC membranes
To study the association of PrPCWD with germinal centre cells, we co-labelled tonsil sections for PrPCWD, FDC and other lymphoid cell phenotypes. Three cytoplasmic phenotype markers were used to identify FDC: S100, vimentin and anti-FDC (see Methods for details on antibodies). To investigate whether PrPCWD accumulated on the cell membrane or cytoplasmically with respect to FDC, we also triple-labelled tonsil sections with antibodies targeting two membrane-bound epitopes associated with FDC and B cell membranes: lambda light chain and cc21 (CD21 or complement receptor type 2). Using confocal microscopy, we found that co-localization of PrPCWD with the FDC intracellular phenotype markers was rare, though PrPCWD appeared in close association with FDC (Fig. 2).



View larger version (50K):
[in this window]
[in a new window]
 
Fig. 2. PrPCWD is on the cell surface of FDC. Lymphoid follicle within a CWD-positive deer tonsil stained using two triple-labelling protocols. PrPCWD (panels a, e, red, antibody 6H4) co-localizes with cell membrane markers for cc21 and immunoglobulin (Ig) (panels c, g, blue) visible as pink in the merged image (panels d, h). PrPCWD (red) does not co-localize with intracellular FDC labels, S100 and FDC (green), as apparent by the lack of yellow in merged images (panels d, h). Bar, 50 µm.

 
Although the lack of co-localization of PrPCWD with the FDC cytoplasmic markers (S100 and anti-FDC) was visually apparent as assessed by the lack of yellow stain, we quantified and compared the co-localization of PrPCWD with intracytoplasmic and membrane markers using Metamorph software. PrPCWD co-localization with the extracellular markers was approximately four times higher than with intracellular FDC markers (P<0·05 by Student’s t-test; Fig. 5). We concluded that PrPCWD accumulated primarily on membranes associated with FDC and B lymphocytes.



View larger version (12K):
[in this window]
[in a new window]
 
Fig. 5. Percentage PrPCWD co-localization with intracellular markers for FDC (S100, FDC) or cell membrane markers (immunoglobulin, cc21). n=12.

 
To verify our results, we analysed tonsil sections labelled with the intracytoplasmic marker vimentin at a higher magnification (900x with zoom) and created a stack of individual optical sections ~0·3 microns apart through the tissue. At high magnification, PrPCWD co-localized strongly with lambda light chain and poorly with vimentin (Fig. 3a–c); this labelling pattern was consistent throughout the thickness of the tissue section (Fig. 3d–h).



View larger version (65K):
[in this window]
[in a new window]
 
Fig. 3. Lymphoid follicle, CWD-positive deer tonsil. PrPCWD (panel b, red, antibody 6H4) co-localizes strongly with membrane-bound lambda light chain of immunoglobulin (Ig, blue) and poorly with the intracellular marker vimentin (panels a, c). Lower magnification, serial sections from the same field show different planes ~1 µm apart from top to bottom and demonstrate the strong co-localization of PrPCWD and Ig and the poor co-localization of PrPCWD and vimentin through the specimen. Bars, 10 µm (a), 20 µm (b).

 
We then characterized PrPCWD in relation to individual cells at high magnification. Using the FDC intracytoplasmic label (S100) and a nuclear stain, we found that PrPCWD was present on the plasma membrane surface and on the fine processes of the FDC (Fig. 4a). Perpendicular sections through a cell labelled with S100 and cc21 antibodies indicated that even in three-dimensional views PrPCWD co-localized only with the plasma membrane marker and did not appear to be intracytoplasmic (Fig. 4b).



View larger version (92K):
[in this window]
[in a new window]
 
Fig. 4. PrPCWD associates with cell membranes on follicular dendritic and B cells. Tonsillar germinal centre, high magnification. (a) PrPCWD is on membrane surfaces and dendritic processes of FDCs (arrows). (b) PrPCWD co-localizes with cc21 (CD21) on membrane surfaces (arrowheads, pink) and poorly with intracellular S100 (note no yellow) indicating that PrPCWD accumulates on cell surfaces. Nucleus is likely the central round black structure (arrow). FDC cell membrane is designated by the white dotted line. Side panel to the right of main panel is the same cell viewed on a perpendicular plane across the pink line and the lower panel shows a section perpendicular to the yellow line in the main panel (T=top, B=bottom). These views demonstrate that there is no discernible PrPCWD within the cytoplasm of the FDC; PrPCWD is between the cells. Anti-PrP antibody used: 6H4. Bar, 10 µm.

 
To determine whether T cells may harbour PrPCWD, we co-labelled tonsil sections for PrPCWD and T cell receptor CD3. We found that relatively low numbers of T cells were present and no consistent intimate association was evident between these CD3+ cells and PrPCWD (data not shown).

PrPCWD in the cytoplasm of TB macrophages
Perifollicular cells containing PrPCWD were seen in chromagen-based IHC staining of lymph nodes (Fig. 6a, b). To phenotype these cells, we triple-labelled a tonsil section using antibodies against PrPCWD, nuclei and CD68, which labels a lysosomal epitope of macrophages and human dendritic cells, and found that PrPCWD was associated with CD68+ macrophages or dendritic cells (Fig. 6c, d).



View larger version (142K):
[in this window]
[in a new window]
 
Fig. 6. PrPCWD-containing cells (arrows) are peripheral to the lymph node follicle. Cells are labelled by immunohistochemistry (a, b) or by triple-immunofluorescence (c, d). Panels (c) and (d) demonstrate PrPCWD labelling (red, antibody R522) in CD68+ cells (green, macrophages or dendritic cells) using confocal microscopy. Nuclei are labelled blue. Bars, 1 mm (a) or 10 µm (b).

 
We investigated whether TB macrophages were involved in PrPCWD accumulation based on earlier experiments in which PrPCWD staining was visualized in cells morphologically characteristic of TB macrophages (see Fig. 9). To determine whether TB macrophages in germinal centres accumulated PrPCWD, we used three intracellular antibody markers for TB macrophages: CD68, HSP70 and ferritin.



View larger version (65K):
[in this window]
[in a new window]
 
Fig. 9. TB macrophages contain PrPCWD. (a) Tonsil biopsy from a deer with pre-clinical CWD demonstrating PrPCWD in cells morphologically consistent with TB macrophages. Note condensed fragmenting nuclei within the cytoplasm (arrows). (b) S100-positive cell (brown) dually labelled for PrPCWD (red) from a cytospin preparation of an enzymatically dissociated CWD-positive deer lymph node. Anti-PrP antibodies used: F89/160.1.5 (a); F99/97.6.1 (b). Bar, 10 µm.

 
In HSP70-labelled sections examined at high magnification, we found that PrPCWD was closely associated with HSP70 (Fig. 7a, b). Serial optical sections through a single cell consistently demonstrated PrPCWD adjacent to the intracellular TB macrophage marker (HSP70), indicating that PrPCWD was intracellular (Fig. 7c–f). Similar results were seen with the ferritin label (data not shown). At high magnification, PrPCWD occasionally co-localized with the macrophage phenotype marker CD68 (Fig. 8). To conclude, two populations of macrophages contained PrPCWD: (1) TB macrophages within germinal centres and (2) isolated macrophages or possibly dendritic cells in the perifollicular area.



View larger version (89K):
[in this window]
[in a new window]
 
Fig. 7. Tonsillar follicle from a CWD-positive deer. PrPCWD (red, antibody 6H4) is detected within cells positive for HSP (green), presumably macrophages (a, b). (b) Higher magnification of cell from panel (a) (white box) shows close approximation of PrPCWD and HSP likely within a single cell. Serial sections of a single cell (yellow box, panel 1) from top to bottom were obtained ~1 µm apart and demonstrate the presence of PrPCWD and HSP70 within the cytoplasm (c–f). Nuclei are labelled blue. Bars, 15  µm (a), 2·5 µm (b), 5 µm (c–f).

 


View larger version (23K):
[in this window]
[in a new window]
 
Fig. 8. CD68+ cell (green) within the germinal centre contains PrPCWD (red, arrow, antibody R522). Nuclei are labelled blue. Bar, 5 µm.

 
PrPCWD in separated lymphoid cells
In a further attempt to determine whether PrPCWD was membrane associated and affiliated with FDC, we examined lymphoid cells enzymatically digested from a CWD-infected retropharyngeal lymph node. Using cytospin preparations of lymphoid cells stained for S100 and PrPCWD, we found that many PrPCWD-bearing cells labelled for S100, identifying them as FDC (Fig. 9). In addition, however, some PrPCWD-containing cells also stained positively for ferritin, a trait most compatible with macrophages. Occasionally PrPCWD-positive cells also co-labelled for lambda light chain or vimentin, traits compatible with FDC, TB macrophages or B cells. These experiments demonstrated that: (1) PrPCWD was cell associated, and (2) PrPCWD-harbouring cells were positive for either S100, ferritin, vimentin or lambda light chain, confirming that at least FDC and macrophages were accumulating PrPCWD.

PrPCWD lymphoid cell association in preclinical CWD-infected deer
To determine whether the lymphoid cell association of PrPCWD changed through the course of infection, we compared PrPCWD lymphoid target cells from deer in early, asymptomatic stages of infection to deer with clinical signs of advanced CWD. The PrPCWD distribution in tonsil biopsies from asymptomatic, naturally exposed deer was similar to that in the tonsils from clinically affected deer. In contrast, in fawns sacrificed 6–11 weeks post-oral inoculation, PrPCWD was distributed primarily on FDC and B cell membrane surfaces with less involvement of TB macrophages. One fawn (6 weeks p.i.) had no apparent PrPCWD in TB macrophages; PrPCWD was primarily associated with cell membranes. In a second fawn (11 weeks p.i.) PrPCWD was detected in both the cell membrane (FDC/B cells) and intracellular (TB macrophages) patterns. These studies suggested that PrPCWD accumulated first in association with FDC vs macrophages and that no additional cell associations were apparent in early pre-clinical stages of infection.


   Discussion
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
A prominent feature of CWD in mule deer is the abundant PrPCWD accumulation in lymphoid germinal centres, similar to that in variant CJD in humans (Hill et al., 1999 ; Hilton et al., 1998 ) and scrapie in sheep (Andreoletti et al., 2000 ; Heggebø et al., 2000 ; van Keulen et al., 1996 ). PrPSc/PrPCWD or infectivity is initially detectable in alimentary-associated lymphoid tissue within weeks following oral exposure and months before detection in the brain (Andreoletti et al., 2000 ; Hadlow et al., 1982 ; Kimberlin & Walker, 1989 ; Sigurdson et al., 1999 ; van Keulen et al., 2000 ; Williams & Miller, 2000 ). While PrPCWD accumulates in lymph nodes in these early stages of infection, the role of specific immune system cells in prion replication and trafficking to the central nervous system (CNS) remains unclear.

Our observations indicate that PrPCWD accumulates in close association with FDC (Fig. 10). Due to the close contact of FDC processes with numerous B cells (emperiopolesis), it is possible that PrPCWD is also on B cell membranes, or is in the extracellular space between FDC and B cells. This finding is consistent with two recent studies in the mouse TSE models demonstrating FDC membrane-associated PrPSc: Jeffrey et al. (2000) used immunogold labelling to elegantly demonstrate ME7 PrPSc on the plasmalemma of splenic FDC. Secondly, Manuelidis et al. (2000) used confocal microscopy to localize strain FU CJD PrPres on FDC membranes. Interestingly, the localization of infectious agent to FDC is not unique to TSEs. Other infectious agents, especially viruses, have been described on FDC surfaces, including bovine viral diarrhoea (Fray et al., 2000 ) and human immunodeficiency viruses (Fujiwara et al., 1999 ; Joling et al., 1993 ; Schmitz et al., 1994 ).



View larger version (49K):
[in this window]
[in a new window]
 
Fig. 10. Working model for lymphoid cells associated with PrPCWD. PrPCWD (red) accumulates on the cell membrane or extracellularly in association with the FDC and/or B cells and accumulates within the cytoplasm of TB macrophages.

 
Although FDC have been associated with PrPSc (Brown et al., 1999 ; Hill et al., 1999 ; Kitamoto et al., 1991 ; McBride et al., 1992 ; Ritchie et al., 1999 ), whether FDC replicate or merely harbour prions remains controversial. For example, Montrasio et al. (2000) demonstrated that inhibition of FDC development virtually eliminated splenic PrPSc. Mabbott et al. (2000) found similar results if mice had FDC deleted prior to scrapie challenge; however, when FDC were deleted after challenge, mice developed high levels of splenic infectivity. Moreover, in experiments using chimeric mice in which PrPC expression between FDC and other lymphoid cells was mismatched, Brown et al. (1999) found that only those mice expressing PrPC in FDC were susceptible to scrapie, strongly suggesting prion propagation by FDC. In contrast, Manuelidis et al. (2000) concluded that limiting intraperitoneal doses of CJD into FDC-deficient mice resulted in only a slightly prolonged incubation period over wild-type controls, suggesting FDC do not play a key role in this model. In our confocal microscopy study of PrPCWD in deer tonsils, serial images through FDC failed to reveal intracytoplasmic PrPCWD, which might indicate that FDC do not uptake or convert appreciable PrPCWD in the cytoplasmic compartment. This finding suggests that FDC may convert PrPC at the cell membrane or that intracellular conversion may be followed by rapid PrPCWD exocytosis. Another possibility would be that the FDC could act as scaffold for passive capture of PrPCWD on the cell membrane, potentially in association with complement or Fc- {gamma} receptors. The association of PrPCWD on cell membranes is consistent with recent evidence for complement involvement in prion pathogenesis, shown by Klein et al. (2001) and Mabbott et al. (2001) .

Unlike the membrane-associated PrPCWD of FDC, intracytoplasmic large, dense aggregates of PrPCWD were detected in TB macrophages. This finding is reminiscent of studies showing PrPSc deposits associated with CD68+ cells (Andreoletti et al., 2000 ) or cells morphologically consistent with TB macrophages in naturally infected scrapie sheep (van Keulen et al., 1996 ). Moreover, Jeffrey et al. (2000) described PrPSc in lysosomes of TB macrophages, consistent with immunogold electron microscopy studies localizing PrPSc in lysosomes of neurons (Laszlo et al., 1992 ).

There are several potential roles for the TB macrophages in prion pathogenesis. It is possible that CD68+ dendritic cells or macrophages transport PrPCWD into the germinal centre and expose the FDC, T and B cells to PrPCWD. CD68+ cells harbouring PrPCWD or PrPSc (Andreoletti et al., 2000 ) have been localized adjacent to germinal centres. However, TB macrophages are in close contact with FDC and are known to phagocytose immune complex-coated bodies (iccosomes) on FDC membranes (Szakal et al., 1988 ). TB macrophages may phagocytose PrPCWD-retaining FDC cell fragments (Heinen et al., 1993 ) and extracellular PrPCWD amyloid, and may or may not replicate PrPCWD, as suggested by Jeffrey et al. (2000) . In addition, TB macrophages phagocytose apoptotic B cells, which also could serve as a potential source of PrPCWD exposure. Therefore, PrPCWD accumulation in TB macrophages may be a secondary event which follows FDC PrPCWD accumulation.

While CD3+ T cells were present in germinal centres, a consistent association between these cells and PrPCWD deposits was not detected, although this association was difficult to assess due to the low number of T cells. Studies with scrapie in transgenic and immunodeficient mice suggest that T cells do not affect disease susceptibility or splenic infectivity (Klein et al., 1997 , 1998 ). Nevertheless, the involvement of T cells in CWD pathogenesis remains an open question.

Although PrPCWD was primarily localized to germinal centres, PrPCWD was not restricted to follicles in all lymphoid tissue studied. Scattered cells in the paracortical zone and medullary cords of lymph nodes occasionally contained PrPCWD. These cells invariably labelled for CD68, indicating that they were either macrophages or dendritic cells.

Surprisingly few differences in the lymphoid cells associated with PrPCWD were seen in fawns weeks after oral exposure to CWD when compared to naturally infected deer with advanced CWD. One fawn at 6 weeks p.i. had PrPCWD extracellularly with no detectable involvement of TB macrophages. We speculate that the TB macrophages may be phagocytosing extracellular PrPCWD iccosomes and that there is a short lag before TB macrophages contain PrPCWD. This scenario could explain why 1 fawn (6 weeks p.i.) had no apparent PrPCWD in TB macrophages versus a second fawn (11 weeks p.i.). In scrapie-inoculated mice at 70 and 170 days p.i., the cell labelling of PrPSc was similar at both time-points (Jeffrey et al., 2000 ). In contrast, in sheep naturally infected with scrapie, PrPSc was apparent in CD68+ cells prior to detection in FDC (Andreoletti et al., 2000 ).

The close association of PrPCWD with the membrane surfaces of FDC and B cells and the presence of intracytoplasmic PrPCWD in TB macrophages raises questions as to the contribution of each of these cell types to PrPCWD replication and trafficking. Our findings in naturally infected deer add to those in CJD- and scrapie-infected mice, and may lend insight into the lymphoid cell targets in vCJD. Understanding peripheral lymphoid reservoirs may be central to deciphering prion trafficking routes from mucosal surfaces and could be critical to diagnostic and intervention measures during the preclinical stages of prion infections.


   Acknowledgments
 
We gratefully acknowledge Katherine O’Rourke and Chris Howard for their generous donation of antibodies F99/97.6.1 and cc21. We are grateful to Margaret Wild, Kate Larsen and Sam Hendrix for assistance with deer tissue collection and to Robert Zink and Bruce Cummings for histotechnology support. We thank Kevin Keane for guidance in image analysis and Leslie Obert for help with phenotype markers and immunofluorescent staining.

This work was supported by grants from the Colorado Division of Wildlife, the College of Veterinary Medicine and Biomedical Sciences Research Council, Colorado State University, and grant RO1-AI-49171 from NIH, NIAID. C. Sigurdson was supported by USDA fellowship 97-36200-5238 and by grant K08-AI-01802 from NIH, NIAID.


   References
Top
Abstract
Introduction
Methods
Results
Discussion
References
 
Andreoletti, O., Berthon, P., Marc, D., Sarradin, P., Grosclaude, J., van Keulen, L., Schelcher, F., Elsen, J. M. & Lantier, F. (2000). Early accumulation of PrPSc in gut-associated lymphoid and nervous tissues of susceptible sheep from a Romanov flock with natural scrapie. Journal of General Virology 81, 3115-3126.[Abstract/Free Full Text]

Bachelet, M., Adrie, C. & Polla, B. S. (1998). Macrophages and heat shock proteins. Research in Immunology 149, 727-732.[Medline]

Betjes, M. G., Haks, M. C., Tuk, C. W. & Beelen, R. H. (1991). Monoclonal antibody EBM11 (anti-CD68) discriminates between dendritic cells and macrophages after short-term culture. Immunobiology 183, 79-87.[Medline]

Bolton, D. C., Rudelli, R. D., Currie, J. R. & Bendheim, P. E. (1991). Copurification of Sp33–37 and scrapie agent from hamster brain prior to detectable histopathology and clinical disease. Journal of General Virology 72, 2905-2913.[Abstract]

Brown, K. L., Stewart, K., Ritchie, D. L., Mabbott, N. A., Williams, A., Fraser, H., Morrison, W. I. & Bruce, M. E. (1999). Scrapie replication in lymphoid tissues depends on prion protein-expressing follicular dendritic cells. Nature Medicine 5, 1308-1312.[Medline]

Brown, K. L., Stewart, K., Ritchie, D., Fraser, H., Morrison, W. I. & Bruce, M. E. (2000). Follicular dendritic cells in scrapie pathogenesis. Archives of Virology Supplementum 16, 13-21.[Medline]

Bruce, M. E., McConnell, I., Will, R. G. & Ironside, J. W. (2001). Detection of variant Creutzfeldt–Jakob disease infectivity in extraneural tissues. Lancet 358, 208-209.[Medline]

Carbone, A., Poletti, A., Volpe, R. & Manconi, R. (1988). S-100 protein detection in ‘follicular macrophages’ of mouse lymphoid organs by ABC immunoperoxidase method. International Journal of Biological Markers 3, 36-40.[Medline]

Foster, J., Goldmann, W., Parnham, D., Chong, A. & Hunter, N. (2001). Partial dissociation of PrPSc deposition and vacuolation in the brains of scrapie and BSE experimentally affected goats. Journal of General Virology 82, 267-273.[Abstract/Free Full Text]

Fray, M. D., Supple, E. A., Morrison, W. I. & Charleston, B. (2000). Germinal centre localization of bovine viral diarrhoea virus in persistently infected animals. Journal of General Virology 81, 1669-1673.[Abstract/Free Full Text]

Fujiwara, M., Tsunoda, R., Shigeta, S., Yokota, T. & Baba, M. (1999). Human follicular dendritic cells remain uninfected and capture human immunodeficiency virus type 1 through CD54–CD11a interaction. Journal of Virology 73, 3603-3607.[Abstract/Free Full Text]

Garssen, G. J., Van Keulen, L. J., Farquhar, C. F., Smits, M. A., Jacobs, J. G., Bossers, A., Meloen, R. H. & Langeveld, J. P. (2000). Applicability of three anti-PrP peptide sera including staining of tonsils and brainstem of sheep with scrapie. Microscopy Research and Technique 50, 32-39.[Medline]

Giorno, R. (1985). Immunohistochemical analysis of the distribution of vimentin in human peripheral lymphoid tissues. Anatomical Record 211, 43-47.[Medline]

Grouard, G., Durand, I., Filgueira, L., Banchereau, J. & Liu, Y. J. (1996). Dendritic cells capable of stimulating T cells in germinal centres. Nature 384, 364-367.[Medline]

Hadlow, W. J., Kennedy, R. C. & Race, R. E. (1982). Natural infection of Suffolk sheep with scrapie virus. Journal of Infectious Diseases 146, 657-664.[Medline]

Heggebø, R., Press, C. M., Gunnes, G., Lie, K. I., Tranulis, M. A., Ulvund, M., Groschup, M. H. & Landsverk, T. (2000). Distribution of prion protein in the ileal Peyer’s patch of scrapie-free lambs and lambs naturally and experimentally exposed to the scrapie agent. Journal of General Virology 81, 2327-2337.[Abstract/Free Full Text]

Heinen, E., Tsunoda, R., Marcoty, C., Antoine, N., Bosseloir, A., Cormann, N. & Simar, L. (1993). Follicular dendritic cells: isolation procedures, short and long term cultures. Advances in Experimental Medicine and Biology 329, 333-338.[Medline]

Hill, A. F., Butterworth, R. J., Joiner, S., Jackson, G., Rossor, M. N., Thomas, D. J., Frosh, A., Tolley, N., Bell, J. E., Spencer, M., King, A., Al-Sarraj, S., Ironside, J. W., Lantos, P. L. & Collinge, J. (1999). Investigation of variant Creutzfeldt–Jakob disease and other human prion diseases with tonsil biopsy samples. Lancet 353, 183-189.[Medline]

Hilton, D. A., Fathers, E., Edwards, P., Ironside, J. W. & Zajicek, J. (1998). Prion immunoreactivity in appendix before clinical onset of variant Creutzfeldt–Jakob disease [letter]. Lancet 352, 703-704.[Medline]

Jeffrey, M., McGovern, G., Martin, S., Goodsir, C. M. & Brown, K. L. (2000). Cellular and subcellular localization of PrP in the lymphoreticular system of mice and sheep. Archives of Virology Supplementum 16, 23-38.[Medline]

Jeffrey, M., Ryder, S., Martin, S., Hawkins, S. A., Terry, L., Berthelin-Baker, C. & Bellworthy, S. J. (2001). Oral inoculation of sheep with the agent of bovine spongiform encephalopathy (BSE). 1. Onset and distribution of disease-specific PrP accumulation in brain and viscera. Journal of Comparative Pathology 124, 280-289.[Medline]

Joling, P., Bakker, L. J., Van Strijp, J. A., Meerloo, T., de Graaf, L., Dekker, M. E., Goudsmit, J., Verhoef, J. & Schuurman, H. J. (1993). Binding of human immunodeficiency virus type-1 to follicular dendritic cells in vitro is complement dependent. Journal of Immunology 150, 1065-1073.[Abstract/Free Full Text]

Kimberlin, R. H. & Walker, C. A. (1989). Pathogenesis of scrapie in mice after intragastric infection. Virus Research 12, 213-220.[Medline]

Kindblom, L. G., Jacobsen, G. K. & Jacobsen, M. (1982). Immunohistochemical investigations of tumors of supposed fibroblastic–histiocytic origin. Human Pathology 13, 834-840.[Medline]

Kitamoto, T., Muramoto, T., Mohri, S., Doh-Ura, K. & Tateishi, J. (1991). Abnormal isoform of prion protein accumulates in follicular dendritic cells in mice with Creutzfeldt–Jakob disease. Journal of Virology 65, 6292-6295.[Medline]

Klein, M. A., Frigg, R., Flechsig, E., Raeber, A. J., Kalinke, U., Bluethmann, H., Bootz, F., Suter, M., Zinkernagel, R. M. & Aguzzi, A. (1997). A crucial role for B cells in neuroinvasive scrapie [see comments]. Nature 390, 687-690.[Medline]

Klein, M. A., Frigg, R., Raeber, A. J., Flechsig, E., Hegyi, I., Zinkernagel, R. M., Weissmann, C. & Aguzzi, A. (1998). PrP expression in B lymphocytes is not required for prion neuroinvasion. Nature Medicine 4, 1429-1433.[Medline]

Klein, M. A., Kaeser, P. S., Schwarz, P., Weyd, H., Xenarios, I., Zinkernagel, R. M., Carroll, M. C., Verbeek, J. S., Botto, M., Walport, M. J., Molina, H., Kalinke, U., Acha-Orbea, H. & Aguzzi, A. (2001). Complement facilitates early prion pathogenesis. Nature Medicine 7, 488-492.[Medline]

Korth, C., Stierli, B., Streit, P., Moser, M., Schaller, O., Fischer, R., Schulz-Schaeffer, W., Kretzschmar, H., Raeber, A., Braun, U., Ehrensperger, F., Hornemann, S., Glockshuber, R., Riek, R., Billeter, M., Wuthrich, K. & Oesch, B. (1997). Prion (PrPSc)-specific epitope defined by a monoclonal antibody. Nature 390, 74-77.[Medline]

Laszlo, L., Lowe, J., Self, T., Kenward, N., Landon, M., McBride, T., Farquhar, C., McConnell, I., Brown, J., Hope, J. and others (1992). Lysosomes as key organelles in the pathogenesis of prion encephalopathies. Journal of Pathology 166, 333–341.[Medline]

Lezmi, S., Bencsik, A. & Baron, T. (2001). CNA42 monoclonal antibody identifies FDC as PrPsc accumulating cells in the spleen of scrapie affected sheep. Veterinary Immunology and Immunopathology 82, 1-8.[Medline]

Mabbott, N. A., Mackay, F., Minns, F. & Bruce, M. E. (2000). Temporary inactivation of follicular dendritic cells delays neuroinvasion of scrapie. Nature Medicine 6, 719-720.[Medline]

Mabbott, N. A., Bruce, M. E., Botto, M., Walport, M. J. & Pepys, M. B. (2001). Temporary depletion of complement component C3 or genetic deficiency of C1q significantly delays onset of scrapie. Nature Medicine 7, 485-487.[Medline]

McBride, P. A., Eikelenboom, P., Kraal, G., Fraser, H. & Bruce, M. E. (1992). PrP protein is associated with follicular dendritic cells of spleens and lymph nodes in uninfected and scrapie-infected mice. Journal of Pathology 168, 413-418.[Medline]

McKinley, M. P., Bolton, D. C. & Prusiner, S. B. (1983). A protease-resistant protein is a structural component of the scrapie prion. Cell 35, 57-62.[Medline]

Manuelidis, L., Zaitsev, I., Koni, P., Lu, Z. Y., Flavell, R. A. & Fritch, W. (2000). Follicular dendritic cells and dissemination of Creutzfeldt–Jakob disease. Journal of Virology 74, 8614-8622.[Abstract/Free Full Text]

Miller, M. W., Williams, E. S., McCarty, C. W., Spraker, T. R., Kreeger, T. J., Larsen, C. T. & Thorne, E. T. (2000). Epidemiology of chronic wasting disease in free-ranging cervids. Journal of Wildlife Diseases 36, 676-690.[Abstract/Free Full Text]

Montrasio, F., Frigg, R., Glatzel, M., Klein, M. A., Mackay, F., Aguzzi, A. & Weissmann, C. (2000). Impaired prion replication in spleens of mice lacking functional follicular dendritic cells. Science 288, 1257-1259.[Abstract/Free Full Text]

Morikawa, K., Oseko, F. & Morikawa, S. (1995). A role for ferritin in hematopoiesis and the immune system. Leukaemia & Lymphoma 18, 429-433.

O’Rourke, K. I., Baszler, T. V., Miller, J. M., Spraker, T. R., Sadler-Riggleman, I. & Knowles, D. P. (1998). Monoclonal antibody F89/160.1.5 defines a conserved epitope on the ruminant prion protein. Journal of Clinical Microbiology 36, 1750-1755.[Abstract/Free Full Text]

Race, R., Jenny, A. & Sutton, D. (1998). Scrapie infectivity and proteinase K-resistant prion protein in sheep placenta, brain, spleen, and lymph node: implications for transmission and antemortem diagnosis. Journal of Infectious Diseases 178, 949-953.[Medline]

Raymond, I., Al Saati, T., Tkaczuk, J., Chittal, S. & Delsol, G. (1997). CNA.42, a new monoclonal antibody directed against a fixative-resistant antigen of follicular dendritic reticulum cells. American Journal of Pathology 151, 1577-1585.[Abstract]

Ritchie, D. L., Brown, K. L. & Bruce, M. E. (1999). Visualization of PrP protein and follicular dendritic cells in uninfected and scrapie infected spleen. Journal of Cellular Pathology 1, 3-10.

Schmitz, J., van Lunzen, J., Tenner-Racz, K., Grossschupff, G., Racz, P., Schmitz, H., Dietrich, M. & Hufert, F. T. (1994). Follicular dendritic cells retain HIV-1 particles on their plasma membrane, but are not productively infected in asymptomatic patients with follicular hyperplasia. Journal of Immunology 153, 1352-1359.[Abstract/Free Full Text]

Sigurdson, C. J., Williams, E. S., Miller, M. W., Spraker, T. R., O’Rourke, K. I. & Hoover, E. A. (1999). Oral transmission and early lymphoid tropism of chronic wasting disease PrPres in mule deer fawns (Odocoileus hemionus). Journal of General Virology 80, 2757-2764.[Abstract/Free Full Text]

Sigurdson, C. J., Spraker, T. R., Miller, M. W., Oesch, B. & Hoover, E. A. (2001). PrPCWD in the myenteric plexus, vagosympathetic trunk and endocrine glands of deer with chronic wasting disease. Journal of General Virology 82, 2327-2334.[Abstract/Free Full Text]

Somerville, R. A., Birkett, C. R., Farquhar, C. F., Hunter, N., Goldmann, W., Dornan, J., Grover, D., Hennion, R. M., Percy, C., Foster, J. & Jeffrey, M. (1997). Immunodetection of PrPSc in spleens of some scrapie-infected sheep but not BSE-infected cows. Journal of General Virology 78, 2389-2396.[Abstract]

Spraker, T. R., Miller, M. W., Williams, E. S., Getzy, D. M., Adrian, W. J., Schoonveld, G. G., Spowart, R. A., O’Rourke, K. I., Miller, J. M. & Merz, P. A. (1997). Spongiform encephalopathy in free-ranging mule deer (Odocoileus hemionus), white-tailed deer (Odocoileus virginianus) and Rocky Mountain elk (Cervus elaphus nelsoni) in northcentral Colorado. Journal of Wildlife Diseases 33, 1-6.[Abstract]

Spraker, T. R., O’Rourke, K. I., Balachandran, A., Zink, R. R., Cummings, B. A., Miller, M. W. & Powers, B. E. (2002a). Validation of monoclonal antibody F99/97.6.1 for immunohistochemical staining of brain and tonsil in mule deer (Odocoileus hemionus) with chronic wasting disease. Journal of Veterinary Diagnostic Investigation 14, 3-7.[Medline]

Spraker, T. R., Zink, R. R., Cummings, B. A., Wild, M. A., Miller, M. W. & O’Rourke, K. I. (2002b). Comparison of histological lesions and immunohistochemical staining of protease-resistant prion protein in a naturally-occurring spongiform encephalopathy of free-ranging mule deer (Odocoileus hemionus) with those of chronic wasting disease of captive mule deer. Veterinary Pathology 39, 110-119.[Abstract/Free Full Text]

Summers, K. L., Hock, B. D., McKenzie, J. L. & Hart, D. N. (2001). Phenotypic characterization of five dendritic cell subsets in human tonsils. American Journal of Pathology 159, 285-295.[Abstract/Free Full Text]

Szakal, A. K., Kosco, M. H. & Tew, J. G. (1988). A novel in vivo follicular dendritic cell-dependent iccosome-mediated mechanism for delivery of antigen to antigen-processing cells. Journal of Immunology 140, 341-353.[Abstract/Free Full Text]

Tsunoda, R., Nakayama, M., Onozaki, K., Heinen, E., Cormann, N., Kinet-Denoel, C. & Kojima, M. (1990). Isolation and long-term cultivation of human tonsil follicular dendritic cells. Virchows Archiv B Cell Pathology Including Molecular Pathology 59, 95-105.

van Keulen, L. J., Schreuder, B. E., Meloen, R. H., Poelen-van den Berg, M., Mooij-Harkes, G., Vromans, M. E. & Langeveld, J. P. (1995). Immunohistochemical detection and localization of prion protein in brain tissue of sheep with natural scrapie. Veterinary Pathology 32, 299-308.[Abstract]

van Keulen, L. J., Schreuder, B. E., Meloen, R. H., Mooij-Harkes, G., Vromans, M. E. & Langeveld, J. P. (1996). Immunohistochemical detection of prion protein in lymphoid tissues of sheep with natural scrapie. Journal of Clinical Microbiology 34, 1228-1231.[Abstract]

van Keulen, L. J., Schreuder, B. E., Vromans, M. E., Langeveld, J. P. & Smits, M. A. (2000). Pathogenesis of natural scrapie in sheep. Archives of Virology Supplementum 16, 57-71.[Medline]

Wells, G. A., Hawkins, S. A., Green, R. B., Austin, A. R., Dexter, I., Spencer, Y. I., Chaplin, M. J., Stack, M. J. & Dawson, M. (1998). Preliminary observations on the pathogenesis of experimental bovine spongiform encephalopathy (BSE): an update. Veterinary Record 142, 103-106.[Medline]

Williams, E. S. & Young, S. (1992). Spongiform encephalopathies in Cervidae. Revue Scientifique et Technique Office International des Epizooties 11, 551-567.

Williams, E. S. & Young, S. (1993). Neuropathology of chronic wasting disease of mule deer (Odocoileus hemionus) and elk (Cervus elaphus nelsoni). Veterinary Pathology 30, 36-45.[Abstract]

Williams, E. S. & Miller, M. W. (2000). Pathogenesis of chronic wasting disease in orally exposed mule deer (Odocoileus hemionus): preliminary results. In Wildlife Disease Association Conference, pp. 29. Jackson, WY.

Zabel, M. D. & Weis, J. H. (2001). Cell-specific regulation of the CD21 gene. International Immunopharmacology 1, 483-493.[Medline]

Received 16 January 2002; accepted 30 April 2002.