Neuroinvasion and subsequent destruction of the central nervous system by prions are typically preceded by a colonization phase in lymphoid organs. An important compartment harboring prions in lymphoid tissue is the follicular dendritic cell (FDC), which requires both tumor necrosis factor receptor 1 (TNFR1) and lymphotoxin β receptor (LTβR) signaling for maintenance. However, prions are still detected in TNFR1−/− lymph nodes despite the absence of mature FDCs. Here we show that TNFR1-independent prion accumulation in lymph nodes depends on LTβR signaling. Loss of LTβR signaling, but not of TNFR1, was concurrent with the dedifferentiation of high endothelial venules (HEVs) required for lymphocyte entry into lymph nodes. Using luminescent conjugated polymers for histochemical PrPSc detection, we identified PrPSc deposits associated with HEVs in TNFR1−/− lymph nodes. Hence, prions may enter lymph nodes by HEVs and accumulate or replicate in the absence of mature FDCs.
Prions are unusual infectious agents thought to be comprised solely of an abnormally folded, aggregated isoform (PrPSc) of the endogenous cellular prion protein (PrPC) [1]. Deposition of PrPSc aggregates, vacuolation, and neuronal loss in brain tissue are histopathological hallmarks of a group of neurological disorders collectively known as prion diseases or transmissible spongiform encephalopathies (TSEs), including scrapie in sheep, bovine spongiform encephalopathy (BSE) in bovids, chronic wasting disease (CWD) in cervids, and Creutzfeldt-Jakob disease (CJD) in humans [2]. Although TSEs seem to selectively damage the central nervous system (CNS), peripheral prion exposure leads to the accumulation of prions and PrPSc in secondary lymphoid organs (SLOs) long before neurological symptoms appear [3], [4], [5], [6], [7], [8], and it is largely from these extraneural sites that prions transmigrate to the peripheral nervous system (PNS) and finally the CNS [9], [10], [11]. Extraneural prion accumulation is thought to occur primarily within stromal cells found in germinal centers of lymphoid follicles known as follicular dendritic cells (FDCs) [4], [12], [13], [14], [15], [16], [17]. Maintenance of mature FDC networks depends on signaling through FDC-expressed lymphotoxin β receptor (LTβR) and tumor necrosis factor receptor 1 (TNFR1) by B cell-derived tumor necrosis factor (TNF) and lymphotoxins (LT) α and β [18], [19], [20], [21], [22], [23]. Accordingly, ablation of B cells, and hence loss of LTα/β and TNFα ligands, antagonizes prion deposition in secondary lymphoid organs [24], [25], and intraperitoneal (i. p.) injection of mice with TNFR1 or LTβR blocking antibodies prior to peripheral prion inoculation causes transient de-differentiation of FDCs, reduces splenic prion accumulation, and delays prion neuroinvasion [26], [27], [28], [29]. However, extraneural prion accumulation in SLOs is not strictly dependent on the presence of mature FDCs. Although prion titers remain below detection in spleens of i. p. -inoculated TNFR1−/− and TNFα−/− mice, PrPSc levels and prion infectivity in TNFR1−/− and TNFα−/− lymph nodes are only marginally reduced compared to TNFR1−/−, LTβR−/−, LTα−/−, or LTβ−/− spleens [30], [31]. Furthermore, TNFR1−/− and TNFα−/− mice succumb to terminal disease upon i. p. prion inoculation at a noticeably higher rate than lymphotoxin signaling-deficient mice, indicating that prions are still effectively transmitted to the CNS in the absence of TNFR1 signaling. Since TNFR1−/− lymph nodes are devoid of detectable mature FDCs, this implies that other undefined cell types may also be required for prions to colonize SLOs. However, lymph nodes are either absent or profoundly disrupted in LTβR−/−, LTα−/−, and LTβ−/− mice compared to TNFR1−/− and TNFα−/−, making it difficult to formally conclude that LTβR signaling is specifically required for prion accumulation in lymph nodes while TNFR1 is not. To determine whether continuous LTβR signaling is required for prions to accumulate in TNFR1−/− lymph nodes, we investigated the ability of prions to colonize SLOs of prion-infected TNFR1−/− mice treated with an LTβR-Ig blocking antibody.
We previously showed that mice devoid of TNF signaling accumulate prion infectivity and PrPSc in lymph nodes but not in spleen, in contrast to LT signaling-deficient mice which do not accumulate prions in either spleen or lymph nodes [31]. To determine whether prion accumulation in TNFR1−/− lymph nodes was dependent on continuous LTβR signaling, we administered weekly 100 µg intraperitoneal (i. p.) injections of an LTβR immunoglobulin fusion protein (LTβR-Ig) or control pooled human immunogloblulin (Ig) to wild-type (WT) and TNFR1−/− mice to achieve sustained inhibition of LTβR signaling [32]. One week following the initial LTβR-Ig or control Ig injection, mice were inoculated intraperitoneally (i. p.) with 6 log LD50 RML6 prions. At 60 days post-injection (d. p. i.), spleens and mesenteric lymph nodes (mLNs) from these mice were assessed for accumulation of prion infectivity and PrPSc. To confirm that LTβR-Ig treatment effectively inhibited LTβR signaling, we analyzed follicular dendritic cell marker 1 (FDCM1) immunoreactivity in spleens from WT or TNFR1−/− mice treated with either control Ig or LTβR-Ig. FDCM1 immunoreactivity was absent in spleens and mLNs from TNFR1−/−-Ig, WT-LTβR-Ig, and TNFR1−/−-LTβR-Ig spleens in contrast to WT-Ig, indicating that LTβR-dependent FDCs had de-differentiated in response to LTβR-Ig treatment (Fig. 1A–J). In addition, we analyzed transcriptional targets of the LTβR pathway in spleens from WT or TNFR1−/− mice treated with either control Ig or LTβR-Ig. As expected, levels of both NFκB2 (p100) and CXCL13 mRNA were reduced in TNFR1−/−-Ig, WT-LTβR-Ig, and TNFR1−/−-LTβR-Ig spleens compared to WT-Ig (Fig. 1K & L). Next, to determine the effect of inhibited LTβR signaling on prion accumulation in SLOs, we compared the pattern of PrPSc deposition in spleens and mLNs from prion-infected TNFR1−/−-Ig, WT-LTβR-Ig, and TNFR1−/−-LTβR-Ig mice to WT-Ig mice by histoblotting. As expected, TNFR1−/−-Ig, WT-LTβR-Ig, and TNFR1−/−-LTβR-Ig spleens accumulated less PrPSc than WT-Ig spleens (Fig. 2A–D), though chronic LTβR-Ig administration seemed less effective at preventing PrPSc deposition in WT spleens than genetic ablation of TNFR1 (compare Fig. 2B with 2C). Since the 60 day treatment period approaches the limit of effective LTβR-Ig inhibition, this most likely reflects partial FDC re-maturation in WT SLOs near the end of the experiment (J. Browning; personal communication). Regardless, the ability of LTβR-Ig to block prion replication in WT SLOs is well-established [26], [27]. Consistent with our previous studies, mLNs from WT and TNFR1−/−-Ig-treated mice contained similar numbers of PrPSc deposits (Fig. 2E & F), whereas PK-resistant PrP deposits in WT-LTβR-Ig mLNs were less numerous than in WT-Ig or TNFR1−/−-Ig mLNs (Compare Fig. 2G with Fig. 2E & F). Background PrP immunoreactivity in PK-digested histoblots from non-infected wild-type spleens was negligible (Supp. Fig. S1). Of note, TNFR1−/−-LTβR-Ig mLNs were devoid of PrPSc immunoreactivity (Fig. 2H), demonstrating that prion accumulation in lymph nodes in the absence of TNFR1 is dependent on LTβR signaling. To confirm this result quantitatively, we analyzed prion infectivity in mLNs from Ig-treated versus LTβR-Ig-treated WT and TNFR1−/− mice using the scrapie cell assay (SCA; [33], [34], [35]). Consistent with the corresponding histoblots from these mice, LTβR-Ig treatment decreased prion infectivity in WT and TNFR1−/− mLNs by 2–3 log tissue culture infectious (TCI) units per gram of tissue compared to Ig treatment (Fig. 2I). To determine the effect of LTβR-Ig treatment on PrPC expression in SLOs, which might explain the differential ability of TNFR1−/−-Ig versus TNFR1−/−-LTβR-Ig mLNs to accumulate prions, we measured Prnp mRNA levels in spleens and lymph nodes from all treatment groups by quantitative PCR. Prnp mRNA levels were reduced in WT-LTβR-Ig, TNFR1−/−-Ig, and TNFR1−/−-LTβR-Ig spleens compared to WT-Ig (Fig. 2J), most likely reflecting de-differentiation of FDCs, the primary PrPC-expressing cell type in spleen. In contrast, no differences in Prnp mRNA expression were found in mLNs from mice of different treatment groups. Of particular note, no difference in Prnp mRNA expression was found between TNFR1−/−-Ig and TNFR1−/−-LTβR-Ig mLNs (Fig. 2K). Taken together, our data indicate that TNFR1−/− lymph nodes accumulate prions in the absence of mature FDCs yet in an LTβR-dependent manner. Moreover, inhibited prion deposition in mLNs upon loss of LTβR signaling seems to be unrelated to local Prnp levels. We reasoned that prion accumulation in TNFR1−/− lymph nodes might rely on a putative LTβR signaling-dependent, TNFR1 signaling-independent cell present in lymph nodes but not in spleens. In order to identify such a cell, we screened spleens and mLNs from WT-Ig, WT-LTβR-Ig, TNFR1−/−-Ig, and TNFR1−/−-LTβR-Ig mice for a variety of hematopoietic and stromal cell markers by immunohistochemistry (IHC) whose expression correlated with prion replication ability. As previously noted, the pattern of FDCM1 immunoreactivity was consistent with PrPSc deposition in spleen, but not in mLNs (Fig. 1A–J). Likewise, several other markers for B cells (B220), T cells (CD3), marginal zone macrophages (MOMA-1), monocytes (F4/80), and stromal cells (CD21/35, C4, VCAM1, ICAM1) revealed no staining patterns consistent with an involvement in splenic or lymph nodal prion accumulation (Supp. Figs S2A–H & S3A–H). However, we identified one stromal cell marker that exhibited a staining pattern consistent with prion deposition in both spleen and mLNs: mucosal addressin cell adhesion molecule 1 (MadCam1 [36]). MadCam1 weakly stained FDC networks of both WT-Ig spleen (Fig. 3A & E) and lymph nodes (Fig. 4A), as well as the marginal sinus (MS) of spleens from WT-Ig mice (Fig. 3A). Although FDC and MS-associated MadCam1 immunoreactivity was absent upon loss of TNFR1 or LTβR signaling in both spleen (Fig. 3B–E) and lymph nodes (Fig. 4 B–D), MadCam1 immunoreactivity persisted in TNFR1−/−-Ig mLNs (Fig. 3G & J; 4B). Of note, MadCam1 immunoreactivity was completely absent in mLNs from LTβR-Ig-treated WT and TNFR1−/− mice (Fig. 3I–J & 4C–D). Furthermore, analysis of MadCam1 mRNA expression by Real Time PCR in mLNs quantitatively corroborated the MadCam1 IHC findings: MadCam1 mRNA levels were equally reduced in spleens from WT-LTβR-Ig, TNFR1−/−-Ig, and TNFR1−/−-LTβR-Ig mice, compared to WT-Ig (Fig. 4E). In contrast, MadCam1 mRNA levels were intermediately reduced in TNFR1−/−-Ig mLNs compared to WT-Ig, and a further reduction in MadCam1 mRNA levels was observed in TNFR1−/−-LTβR-Ig mLNs compared to TNFR1−/−-Ig (Fig. 4F). Thus far, our data suggested that the presence of a MadCam1-expressing cell was associated with accumulation of prions in lymph nodes but not in spleen. MadCam1 immunoreactivity in TNFR1−/−-Ig mLNs was restricted to thick vessels (Fig. 3G & 4B) that were absent from TNFR1−/−-LTβR-Ig mLNs (Fig. 3I & Fig. 4D), as well as isotype controls (Supp. Fig. S4B), and were morphologically distinct from MadCam1-positive FDC networks found in WT-Ig spleens (Fig. 3A) and mLNs (Fig. 4A). These MadCam1-positive structures were morphologically consistent with high endothelial venules (HEVs) responsible for controlling lymphocyte entry into LNs, Peyer' s patches (PPs), and other SLOs, with the exception of spleen [37]. Co-localization of Madcam1-positive vessels with peripheral node addressin (PNAd; a. k. a. MECA-79), a specific marker of HEVs [38], confirmed that these structures in TNFR1−/−-Ig mLNs were HEVs (Fig. 5A–F), whereas MadCam1-positive FDC networks in WT-Ig mLNs (Fig. 5A–C) and isotype controls (Supp. Fig. S4A) were PNAd-negative. To gather evidence that HEVs could be potential sites of prion replication, we analyzed PrPC immunoreactivity in HEVs of WT-Ig mLNs. Co-immunofluorescent (co-IF) staining of WT-Ig mLNs with MadCam1 and PrPC antibodies confirmed that PrPC immunoreactivity was present in HEVs (Fig. 5G–I) and absent from isotype controls (Supp. Fig. S4C). Thus, HEVs fulfill at least one requirement of a prion replicating tissue - PrPC expression. To determine whether prions were localized to HEVs in mLNs, we performed co-IF with PrPC and MadCam1 antibodies in prion-infected TNFR1−/−-Ig mLNs. Prion-infected TNFR1−/−-Ig mLNs contained areas of intense PrP-positive deposits which were not detectable in non-infected TNFR1−/−-Ig mLNs (data not shown; [31]). Of note, many of these PrPC-positive areas localized to MadCam1-postive vessels in prion-infected TNFR1−/−-Ig mLNs, indicating that HEVs are probable sites of PrPSc localization in infected lymph nodes (Fig. 6A–C). However, PrP immunostaining of prion-infected tissue cannot reliably distinguish between PrPSc deposits and sites of high PrPC expression, and HEVs expressed relatively high levels of PrPC in uninfected mLNs (Fig. 5G–I). To develop an independent method of distinguishing PrPSc from PrPC in lymphoid organs, we tested the ability of a series of fluorescent amyloid-binding dyes known as luminescent conjugated polymers (LCPs [39], [40], [41], [42]) to stain PrPSc deposits in spleen and lymph node. LCPs were previously shown to recognize PrPSc in brain [43]. One LCP, pentameric formic thiophene acetic acid (p-FTAA; [44]), stained PrP-positive FDC networks of prion-infected spleens from WT mice (Supp. Fig. S5). In contrast, no immunofluorescence could be detected with p-FTAA in spleens and lymph nodes from uninfected mice (Supp. Fig. S6). Using this method, points of PrPSc/MadCam1 co-localization could be reliably identified in prion-infected TNFR1−/− lymph nodes, which again indicated that a proportion of PrPSc localizes to HEVs in TNFR1−/− mLNs (Fig. 6D–F). However, at lower magnification we also noted that a number of PrPSc deposits were located outside of HEVs (Fig. 6G–I). To confirm this observation and to analyze the tissue-wide distribution of PrPSc deposits relative to HEVs in prion-infected TNFR1−/−-Ig mLNs, we performed histoblot co-stains with PNAd antibody, which is highly immunoreactive to HEVs and can be visualized even after proteinase K digestion (Supp. Fig. S7). This analysis confirmed that some PrPSc deposits were localized to HEVs in TNFR1−/−-Ig mLNs. However, PrPSc deposits were also present outside of HEVs (Fig. 6J–K), which is consistent with our previous study showing that strong PrP immunoreactivity in prion-infected TNFR1−/− mLNs can also be found in other cell types (i. e. macrophages [31]). Since HEVs could potentially serve as entry portals for prion-harboring lymphocytes, and it was recently reported that neighboring dendritic cells (DCs) are responsible for HEV differentiation [45], we also performed PrP co-immunofluorescence on prion-infected TNFR1−/−-Ig mLNs using the DC marker, CD11c, to determine whether DCs in the vicinity of HEVs might also contain PrPSc. However, no overlap of PrP and CD11c immunoreactivity in prion-infected TNFR1−/−-Ig mLNs was identified (Supp Fig. S8).
The means by which prions evade the immune system' s numerous defense mechanisms and finally transmigrate to, and selectively damage, neurons of the CNS has been the subject of scientific scrutiny for two decades. A number of studies have implicated FDCs in the germinal centers of SLOs as the primary reservoirs of prions prior to neuroinvasion [4], [12], [13], [14], [15], [16], [17]. Yet the ability of TNFR1−/− mLNs to accumulate prions with a minimal loss of infectivity compared to WT mLNs presents an apparent paradox, since FDC maintenance depends on TNFR1 signaling [21], [22], [23]. Here we have established that lymph nodal prion accumulation in the absence of TNFR1 signaling is LTβR signaling-dependent. Crucially, transient loss of LTβR signaling was sufficient to block TNFR1-independent prion accumulation in lymph nodes, indicating that prion accumulation in lymph nodes specifically requires LTβR signaling and is not simply prevented by general developmental defects or architectural disruptions caused by lack of LTβR signaling in LTβ−/− lymph nodes. We previously showed that intense PrP immunoreactivity was localized to macrophages in prion-infected TNFR1−/− mLNs [31], and others have reported that PK-resistant PrP is localized to macrophages in spleens with PrPC-deficient FDCs [46], indicating that macrophages serve as alternative sites of prion accumulation in the absence of PrPC-expressing FDCs. How this phenomenon was mechanistically linked to LTβR signaling and the pattern of prion accumulation in lymph nodes was initially unclear, since most macrophage populations were preserved in the absence of both TNFR1 and LTβR signaling [31]. Here, we discovered that loss of LTβR signaling in mLNs was also correlated with the dedifferentiation of HEVs – the primary point of entry for lymphocytes into lymph nodes and a likely determinant in the ability of prions to colonize lymph nodes. Consistent with the pattern of prion accumulation in spleens and lymph nodes of mice lacking TNF and/or LT signaling components, HEVs exist in lymph nodes and other SLOs but not spleens [37], and the maintenance of HEV architecture is TNFR1 signaling-independent yet LTβR signaling-dependent [47]. Moreover, we identified sites of PrPSc-HEV overlap in TNFR1−/−-Ig mLNs, indicating that HEVs might replicate prions and/or serve as points of entry for prions or prion-harboring lymphocytes. FDC-deficient mice can succumb to scrapie even in the absence of detectable splenic prion titers [24]. In light of our current results, this phenomenon is most likely explained by HEV-dependent entry and accumulation of prions in lymph nodes and other HEV-containing SLOs, such as Peyer' s patches. HEVs can also form ectopically in certain chronic inflammatory conditions. Prion accumulation at sites of chronic inflammation has previously been observed, but in most cases this could be attributed to local formation of FDC networks [48], [49], [50]. However, we also previously reported LTβR-dependent prion colonization of granulomas in the absence of FDC markers [51]. Since ectopic HEV formation without any evidence of FDC networks has previously been reported in certain inflammatory conditions [52], ectopic HEV formation might be a source of prion replication and/or uptake in granulomas, as well. Of note, LTα/β signaling to HEVs differs from LTα/β signaling to FDCs. Though HEVs express LTβR [53], HEVs persist in the absence of B cells [47], in contrast to FDCs which rely on B cells to provide LTα/β ligand [19], [20]. This indicates that the LTα/β signal to HEVs emanates from another cell type. Until recently the cell type providing LTα/β to HEVs was not known; however a recent publication reports that dendritic cells (DCs) may be the source of LTα/β signaling to HEVs [45]. If HEV differentiation is indeed B cell-independent, this may explain why prion neuroinvasion can occur in the absence of B cells in some cases, since LTβR signaling to HEVs and hence the ability of prions to enter and accumulate in lymph nodes would be preserved [24], [54]. Do prions actually replicate in TNFR1−/− lymph nodes, or do they simply accumulate? Accumulation seems likely, as the only cell type in SLOs known to replicate prions are FDCs, and prion replication in macrophage populations was not observed [46]. In any case, experimental evidence suggests that prion accumulation in SLOs is sufficient for prions to invade the CNS [55], [56], [57]. Moreover, deletion of complement factors dramatically impairs prion accumulation in SLOs and subsequent neuroinvasion without affecting PrPC levels [58], indicating that the ability of SLOs to physically capture prions is indeed critical for the development of downstream pathology. Curiously, we previously showed that PrPC expression is required in either the stromal or the hematopoietic compartment for lymph nodes to accumulate prion infectivity [31], which implies that prion replication can occur in both compartments in lymph nodes. However, an alternative explanation is that a hematopoietic cell is required for delivery of prions to a “trapping” cell within the lymph node, and PrPC expression on the hematopoietic cell mediates efficient uptake of PrPSc in the bloodstream. Hence, HEVs may facilitate the selective uptake and accumulation of prions or prion-containing lymphocytes into lymph nodes, rather than serving as sites of bona fide prion replication. Based on both current and past findings, it is likely that the “trapping” cells in lymph nodes are endothelial cells in HEVs (HEVECs), and the hematopoietic delivery cells are macrophages. Once prions or prion-harboring cells have successfully invaded mLNs through HEVs, they may be transported via conduits to FDCs [59] under normal conditions. However, in the absence of mature FDCs, prions may remain in or be transferred to macrophages.
TNFR1−/− mice carrying a targeted deletion of exons 2,3, and part of exon 4 of the tumor necrosis factor receptor 1 (TNFR1) open reading frame [60] were maintained on a C57BL/6 background in-house. C57BL/6 wild-type control mice were purchased from Harlan Laboratories and bred in-house. Murine LTβR-murine IgG1 (LTβR-Ig) and MOPC21 mouse immunoglobulin control were obtained from Biogen Idec. For analysis of prion-infected tissues (Figs 2A–I, 6 & Supp. Figs S5, S7 & S8), TNFR1−/− or C57BL/6 mice were injected intraperitoneally (i. p.) with 100 µg LTβR-Ig or MOPC21 (n = 3–4/group). One week later, mice were inoculated i. p. with 100 µL 6 log LD50 Rocky Mountain Laboratory mouse-adapted prion strain [61], passage 6 (RML6). Mice were then boostered weekly with 100 µg LTβR-Ig or MOPC21 until 60 days post-inoculation (d. p. i), at which point mice were sacrificed. For analysis of uninfected tissue (Figs 1,2J–K, 3,4, 5 & Supp. Figs S1, S2, S3, S4 & S6), TNFR1−/− or C57BL/6 mice received 2 weekly injections of LTβR-Ig or MOPC21 and were sacrificed 3 weeks following the initial injection (n = 3–4/group). Spleens and lymph nodes from prion-infected and uninfected mice were either flash frozen for scrapie cell assays and gene expression analysis or frozen in 1x Hank' s balanced salt solution for immunohistochemistry. Tissues were stored at −80°C until analysis. All animal experiments were carried out in strict accordance with the rules and regulations for the Protection of Animal Rights (Tierschutzverordnung) of the Swiss Bundesamt für Veterinärwesen and were pre-approved by the Animal Welfare Committee of the Canton of Zürich. Animal permit # 130/2008. CD1 wild-type mice were inoculated i. c. with 30 µL 1% (w/v) RML-5-infected brain homogenate and sacrificed at terminal stage disease. Brains were flash frozen in liquid nitrogen and stored at −80°C. Brains were then homogenized in sterile 0. 32 M sucrose in 1x PBS (20% w/v) using a Ribolyser (Hybaid, Catalys). Subsequent dilutions of inoculum were performed in sterile 5% (w/v) bovine serum albumin (BSA) in 1x PBS. For titer determinations, serial dilutions of RML6 were inoculated i. c. into indicator mice, and LD50 was defined as the dilution that induced a 50% attack rate. Frozen spleens and lymph nodes were homogenized in Qiazol (10% w/v) using a TissueLyser, and total mRNA was isolated using an RNeasy Mini kit, according to the manufacturer' s instructions (Qiagen). 1 µg mRNA from each sample was used for first-strand cDNA synthesis using random hexamers from a Superscript II kit (Invitrogen). ∼100 ng cDNA, 500 nM primers, and 1x Faststart SYBR Green reaction mixture (Roche) in 25 µL reaction volumes were used for Real Time PCR amplification of target sequences from spleen and lymph nodes normalized to GAPDH using an ABI 7900HT (Applied Biosystems). (See Table 1 for list of primer sequences.) Reactions from each tissue were performed in triplicate and averaged. Samples with a triplicate variability exceeding 10% were eliminated from further analysis. Amplification data was analyzed using the relative quantification method (RQ) with WT-Ig samples serving as the calibrator values. RQ values were calculated and averaged for each sample. Average values depicted in the graphs represent the mean value of single spleens or lymph nodes from each individual mouse from each treatment group. N = 2–4 mice per group, depending on the specific tissue and treatment group. 10 µM frozen sections were transferred to nitrocellulose pre-soaked in 1x Tris-buffered saline with Tween 20 (TBST; 50 mM Tris-HCl, pH = 7. 8,150 mM NaCl, 0. 1% Tween 20) and air-dried. Membranes were washed in 1x TBST for 1 hr. and then digested with proteinase K (Roche, 20 µg/mL) diluted in digestion buffer (10 mM Tris-HCl, pH = 7. 8,100 mM NaCl, 0. 1% (v/v) Brij 35) at 37°C for 4 hrs. Membranes were then washed in TBST, incubated in denaturing solution (10 mM Tris-HCl, pH = 7. 8,3 M guanidine thiocyanate) for 10 min, washed in 1x TBST, blocked in 5% dried milk in TBST for 1 hr, and then probed with 0. 1 µg/mL POM1 [62] diluted in 1% milk in 1x TBST overnight at 4°C. Membranes were then washed in 1x TBST, blocked in 1% milk in 1x TBST, and then incubated with 1 µg/mL alkaline phosphatase (AP) -conjugated goat anti-mouse secondary antibody (Dako) for 1 hr. Membranes were then washed in 1x TBST, 10 min. in B3 (100 mM Tris, 100 mM NaCl, 100 mM MgCl2, pH = 9. 5), and then developed for 40 min. with BCIP/NBT (Roche). Histoblot sections were then washed in distilled water, dried, and imaged using an Olympus SZX12 stereomicroscope. Uninfected spleens from wild-type mice served as negative controls for background PrP immunoreactivity (Supp. Fig. S1). Scrapie-susceptible neuroblastoma cells (subclone N2aPK1, [33]) were incubated with uninfected brain homogenate, defined titers of RML6-infected brain homogenate, or 10−3 to 10−6 dilutions of mesenteric lymph node homogenate from WT-Ig, WT-LTβR-Ig, TNFR1−/−-Ig, or TNFR1−/−-LTβR-Ig for 3 days. Infected N2aPK1 cells were passaged 1∶3 three times every 2 days, and then 1∶10 four times every 3 days. After reaching confluence, 2×104 cells from each well were filtered onto the membrane of an ELISPOT plate (Millipore; MultiScreenHTS filter plates with Immobilon-P PVDF membrane) and denatured with 0. 5 µg/mL proteinase K (PK). Individual prion-infected cells were immunodetected with POM1. Wells were scored positive if the spot number exceeded mean background values, determined as three times the standard deviation of the uninfected control. In this experiment, an ELISPOT membrane with ≥3 PrPSc+ colonies was regarded as infected. From the proportion of negative to total wells, the number of tissue culture infectious units per mL was calculated with the Poisson equation. In two independent experiments, a 10−8 dilution of a standard inoculum (brain homogenate from a terminally scrapie-sick mouse) yielded 11/24 or 12/24 positive wells, corresponding to a titer of ∼8. 3 log tissue culture infectivity (TCI) units/g of brain tissue for the initial inoculum. The sensitivity threshold was calculated to be 2. 8 log TCI units/g of brain tissue. 7 µM frozen sections on glass coverslips (Thermofisher) were dried for several hours at room temp, fixed in 4% formalin for 2 min, 50% acetone for 2 min, 100% acetone for 2 min, and 50% acetone for 2 min. Sections were then washed in 1x PBS, then 1x PBS+ 0. 05–0. 1% Tween 20, and blocked for 1 hr. in SuperBlock (Pierce). Sections were then incubated overnight at 4°C in primary antibody (see Table 2 for antibodies and dilutions) diluted in 1∶10 SuperBlock. For vessel stains, isotype controls (rat IgM for PNAd, Pharmingen # 553941; rat IgG2a for MadCam1, eBioscience # 14-4321; and mouse IgG1 for PrP, Sigma # 15381) were performed using equivalent concentrations to the corresponding primary antibodies (Supp. Fig. S4). Sections were then washed in 1x PBS, followed by 1x PBST, then incubated with 0. 2–0. 4 µg/mL Alexa Fluor secondary antibody (Invitrogen) diluted in 1x PBST (for immunofluorescence [IF]) or 5. 3 µg/mL unconjugated goat anti-rat secondary antibody (Caltag Laboratories) or goat anti-mouse (Jackson Immunoresearch) diluted in 1∶10 SuperBlock for 1 hr. at room temperature (for light microscopy [LM]). Sections were then washed in 1x PBS followed by 1x PBST and incubated in 7. 5 µg/mL AP-conjugated donkey anti-goat tertiary antibody (Jackson Immunoresearch) in 1x PBST for 1 hr for LM. Sections were then washed and developed with Fast Red (Sigma) staining kit and counter-stained with hematoxylin & eosin (H&E) for LM. Sections were then mounted (fluorescent mounting medium for IF or aqueous mounting medium for LM; Dako) and coverslipped. Sections were imaged using an Olympus BX61TRF fluorescent microscope, a Zeiss Axiophot light microscope, or a Leica SP5 confocal microscope (where indicated). FDCM1: Boundaries of lymphoid follicles were identified by H&E counterstaining in LM images of AP-developed spleens and lymph nodes immunostained with FDCM1. Follicles containing FDCM1-stained germinal centers were classified as “FDCM1-positive, ” whereas follicles devoid of FDCM1 staining were classified as “FDCM1-negative. ” The total number of FDCM1-postive or negative follicles per treatment group was expressed as a percentage of the total follicles. A total of 55 follicles were scored for WT-Ig spleens, 39 for WT-LTβR-Ig spleens, 111 for TNFR1−/−-Ig spleens, 27 for TNFR1−/−-LTβR-Ig spleens, 19 for WT-Ig mLNs, 10 for WT-LTβR-Ig mLNs, 13 for TNFR1−/−-Ig mLNs, and 10 for TNFR1−/−-LTβR-Ig mLNs. MadCam1: Boundaries of lymphoid follicles were identified by H&E counterstaining in LM images of AP-developed spleens immunostained with MadCam1. Follicles containing MadCam1-stained germinal centers were classified as “MadCam1-positive, ” whereas follicles devoid of MadCam1 staining were classified as “MadCam1-negative. ” The total number of MadCam1-postive or negative follicles per treatment group was expressed as a percentage of the total follicles. For mLNs, the total number of MadCam1-positive vessels was scored per organ and averaged. A total of 80 follicles were scored for WT-Ig spleens, 37 for WT-LTβR-Ig spleens, 137 for TNFR1−/−-Ig spleens, 39 for TNFR1−/−-LTβR-Ig spleens, n = 2 mLNs for all treatment groups. PNAd/PrPSc co-stains: Total numbers of HEVs and PrPSc deposits were counted on PNAd pre-stained histoblots from TNFR1−/−-Ig mLNs. HEVs that overlapped with PrPSc deposits were scored as “PrPSc-postive, ” whereas HEVs with no PrPSc were scored as “PrPSc-negative. ” Likewise, PrPSc deposits that overlapped with HEVs were scored as “PNAd-positive, ” whereas PrPSc deposits that were not associated with HEVs were scored as “PNAd-negative. ” For HEVs, values represent the percentage of total HEVs that were either PrPSc-positive (black) or negative (white). For PrPSc deposits, values represent the percentage of total PrPSc deposits that were associated with HEVs (black) or not (white). A total of 82 HEVs and 112 PrPSc deposits were scored. 7 µM frozen sections on glass coverslips were dried and then fixed in pre-chilled 100% acetone or ethanol at −20°C for 10 min. Sections were dried for 1 min. , re-hydrated in 1x PBS for 10 min. , blocked in SuperBlock, and then incubated in 20 µg/mL MadCam1 or 6. 25 µg/mL FDCM1 in 1x PBS overnight at 4°C. Sections were then washed in 1x PBS and incubated with 0. 2–2 µg/mL Alexa Fluor 594-conjugated goat anti-rat secondary antibody for 1 hr. Sections were then washed in 1x PBS and incubated with 30 µM pentameric formic thiophene acetic acid (p-FTAA; [44]) in 1x PBS for 30 min. Sections were then washed in 1x PBS, mounted with fluorescent mounting medium (Dako), coverslipped, and imaged using an Olympus BX61TRF fluorescent microscope. Sections from uninfected WT spleens were used as negative controls for non-specific pFTAA staining (Supp. Fig. S6). TNFR1−/−-Ig mesenteric lymph node sections (10 µm) on nitrocellulose were soaked in 1x TBST for 1 hr. , blocked in 5% (w/v) milk in 1x TBST for 1 hr. , and then probed with 0. 5 µg/mL PNAd antibody diluted in 1% (w/v) milk in 1x TBST overnight at 4°C. Histoblots were then washed in 1x TBST and probed with 0. 5 µg/mL AP-conjugated goat anti-rat (Biosource # ARI3405) secondary antibody in 1% (w/v) milk in 1x TBST. Histoblots were then washed in 1x TBST and developed with Fast Red (Sigma) for 20 min. Histoblots were then washed in 1x TBST, digested with 20 µg/mL PK for 4 hrs at 37°C, washed and further processed as described above for standard histoblots.
Prions are unique infectious agents thought to be composed entirely of an abnormal conformer of the endogenous prion protein. Prions cause a severe neurological disorder in humans and other animals known as prion disease. Though prion disease can arise spontaneously or from genetic mutations in the gene encoding the prion protein, many cases of prion disease arise due to peripheral exposure to the infectious agent. In these cases, prions must journey from the gastrointestinal tract and/or the bloodstream to the brain. Prions often colonize secondary lymphoid organs prior to invading the nervous system via neighboring peripheral nerves. Prion accumulation in follicular dendritic cells found in germinal centers of lymphoid organs is thought to be a crucial step in this process. However, prion colonization of lymph nodes, in contrast to spleen, does not depend on follicular dendritic cells, indicating that other mechanisms must exist. Here, we identify the signaling pathway required for follicular dendritic cell-independent prion colonization of lymph nodes, which also controls the differentiation of high endothelial venules, the primary entry point for lymphocytes into lymph nodes. Importantly, prions could be found within these structures, indicating that high endothelial venules are required for prion entry and accumulation in lymph nodes.
medicine
neurobiology of disease and regeneration
immunology
microbiology
lymphoid organs
neuroscience
infectious diseases
biology
pathogenesis
immune system
clinical immunology
neurological disorders
neurology
molecular cell biology
Lymphotoxin, but Not TNF, Is Required for Prion Invasion of Lymph Nodes