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
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Abstract |
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Introduction |
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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 Peyers 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.
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Methods |
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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.
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.
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 antigenantibody 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.
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 12 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 144152 (Korth et al., 1997 ). R522 recognizes ovine PrP 94105 (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.
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.
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 Students t-test. Significance was defined at P<0·05.
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 peroxidasestreptavidin 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 ORourke) (Spraker et al., 2002), a biotinylated secondary antibody, an alkaline phosphatasestreptavidin 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) (ORourke et al., 1998 ).
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Results |
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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 (
80100%) 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
).
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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
).
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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 611 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.
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Discussion |
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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
).
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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.
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Acknowledgments |
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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.
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Received 16 January 2002;
accepted 30 April 2002.