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J Immunol. 2012 November 1; 189(9): 4520–4527.
Published online 2012 September 21. doi:  10.4049/jimmunol.1201579
PMCID: PMC3478448

Genetic Depletion of Complement Receptors CD21/35 Prevents Terminal Prion Disease in a Mouse Model of Chronic Wasting Disease


The complement system has been shown to facilitate peripheral prion pathogenesis. Mice lacking complement receptors CD21/35 partially resist terminal prion disease when infected i.p. with mouse-adapted scrapie prions. Chronic wasting disease (CWD) is an emerging prion disease of captive and free-ranging cervid populations that, similar to scrapie, has been shown to involve the immune system, which probably contributes to their relatively facile horizontal and environmental transmission. In this study, we show that mice overexpressing the cervid prion protein and susceptible to CWD (Tg(cerPrP)5037 mice) but lack CD21/35 expression completely resist clinical CWD upon peripheral infection. CD21/35-deficient Tg5037 mice exhibit greatly impaired splenic prion accumulation and replication throughout disease, similar to CD21/35-deficient murine prion protein mice infected with mouse scrapie. TgA5037;CD21/35−/− mice exhibited little or no neuropathology and deposition of misfolded, protease-resistant prion protein associated with CWD. CD21/35 translocate to lipid rafts and mediates a strong germinal center response to prion infection that we propose provides the optimal environment for prion accumulation and replication. We further propose a potential role for CD21/35 in selecting prion quasi-species present in prion strains that may exhibit differential zoonotic potential compared with the parental strains.


Chronic wasting disease (CWD) is the only recognized naturally occurring transmissible spongiform encephalopathy (TSE), affecting captive and free-ranging cervids (1) in North America and captive cervids in South Korea. Similar to other TSEs, CWD is caused by prions, unusual infectious agents devoid of instructional nucleic acid (2) and characterized by the accumulation of misfolded prion protein (PrPRES), a proteinase K (PK) resistant form of the normal cellular prion protein, PrPC. CWD and the sheep TSE scrapie can be transmitted relatively efficiently compared with other TSEs, probably contributing to their higher prevalences (3, 4). Prions have been detected in nervous and lymphoid tissue, muscle, blood, saliva, urine, and feces (513). Of particular interest are lymphoid tissues because they contain prions often before the CNS, implicating the lymphoid system as an initial site of extracerebral prion accumulation and replication. Lymphoid follicles or inflammatory foci accumulate and replicate prions primarily on follicular dendritic cells (FDCs) that express relatively large amounts of PrPC (1418). FDCs originate from perivascular precursor cells (19) and trap immune complexes on their elaborate projections and present them to B cells, which can be positively selected, activated, undergo Ig affinity maturation, and become plasma cells. FDCs may retain Ag on their cell surfaces for prolonged periods, maximizing presentation to B cells and consequently affecting the humoral immune response.

FDC depletion significantly impairs prion replication, and FDC-specific PrPC expression has been shown to be essential for optimal peripheral prion infection (14, 17, 18, 20). B cells, although replicating little prion, also play an essential role in peripheral prion pathogenesis (21, 22). This requirement presumably relates to the ability of B cells to supply FDCs with critical cytokines important in FDC maturation and maintenance, but they may also be involved in lymphotropic and/or intranodal prion trafficking.

Substantial evidence supports a significant role for the complement system in expediting peripheral prion disease by mediating prion interaction with FDCs and B cells. Complement activation leads to asymmetrical cleavage of both C3 and C4 bound to pathogens. Complement receptors CD21/35 expressed on B cells and FDCs trap opsonized pathogens by binding cleaved C3 and C4 opsonins. Mice express CD21 and CD35 only on B cells and FDCs from alternatively spliced transcripts generated from a single gene, whereas humans express them on more cell types from separate genes (23, 24). Although complement-mediated Ag trapping enhances both innate and adaptive immune responses to microbial pathogens, it actually exacerbates prion pathogenesis. Elimination of complement receptors CD21/35 reduced prion trapping, replication, and disease (17). Interestingly, depletion of CD21/35 has a greater impact on disease progression than deleting their ligand sources, C3 and C4, alluding to a role for CD21/35 in peripheral prion pathogenesis independent of their endogenous ligands. Genetic depletion of C1q also delays prion disease at high doses and prevents disease at low doses after i.p. infection (25, 26), and C1q has been shown to bind prions in vitro (27, 28).

In this study, we show that complete elimination of the complement receptors CD21/35 in transgenic mice susceptible to CWD significantly delays splenic prion accumulation and blocks progression to terminal disease upon inoculation with CWD prions. To assess the kinetics of prion accumulation in the spleen we developed a semiquantitative prion amplification scoring system based on protein misfolding cyclic amplification (PMCA), which allowed us to evaluate prion replication and/or accumulation at 15, 30, 70, and 140 d postinoculation (dpi). Mice deficient in CD21/35 show a significant impairment in prion retention and replication compared with CD21/35-sufficient mice. We also observed significant germinal center (GC) formation during scrapie prion infection that was dependent on CD21/35 and PrPC expression on FDCs. Lipid raft flotation experiments show movement of CD21/35 into lipid rafts on B cells upon prion infection. Overall, these data demonstrate that CD21/35-mediated prion trapping on FDCs and possibly B cells marks an important event in lymphoid prion pathogenesis that promotes terminal prion disease in these mouse models.

Materials and Methods


Prnpo/oCD21/35−/−, C3/C4−/−, Tg5037, and TgA20 mice were made as previously described (17, 29, 30). Prnpo/oCD21/35−/− mice were crossed with TgA20 or Tg5037 mice to produce TgA20;Prnpo/oCD21/35−/− (TgA20;CD21/35−/−) and Tg5037;Prnpo/oCD21/35−/− (Tg5037;CD21/35−/−) mice. All mice were bred and maintained at Laboratory Animal Resources, accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International, in accordance with protocols approved by the Institutional Animal Care and Use Committee at Colorado State University. Bone marrow chimeric mice were produced as previously described (17).

Preparation of inoculum

Brain homogenates (10%) were prepared in PMCA buffer (4 mM EDTA, 150 mM NaCl in PBS) from E2 homogenates derived from a terminally diseased elk brain. RML5 prions were prepared as previously described (17). Homogenates (10%) were diluted 1:10 (E2 and RML5) or 1:1000 (RML5) in 320 mM sucrose supplemented with 100 U/ml penicillin and 100 μg/ml streptomycin (Life Technologies) in PBS immediately prior to inoculation.

Inoculations, clinical scoring, and dissections

Mice were inoculated i.p. with 100 μl inoculum using a 28-gauge insulin syringe. Mice were monitored for clinical symptoms of prion disease, including tail rigidity, impaired extensor reflex, akinesia, tremors, ataxia, and weight loss. Mice with any four of these symptoms or paralysis were scored terminally sick and euthanized.

Mice were inoculated with inocula described above and euthanized at indicated time points by CO2 inhalation. Brains and spleens were collected and divided sagittally. One brain hemisphere and half a spleen were fixed in 4% paraformaldehyde in PBS for histology and one was homogenized and used for sodium phosphotungstic acid (NaPTA) precipitation, PMCA, or PK digestion.

NaPTA precipitation of PrPRES

NaPTA precipitation was performed exactly as described previously (17). Briefly, gross cellular debris was removed by centrifugation at 80 × g and 500 μl supernatant was mixed 1:1 with 4% sarkosyl in PBS. Samples were incubated for 15 min at 37°C with constant agitation, then incubated with 50 U/ml benzonase and 12.75 mM MgCl2 for 30 min at 37°C with constant agitation. Prewarmed NaPTA stock solution (pH 7.4) was added to a final concentration of 0.3% and the sample was incubated at 37°C for 30 min with constant agitation and centrifuged at 37°C for 30 min at maximum speed in an Eppendorf microcentrifuge. The pellet was resuspended in 30 μl 0.1% sarkosyl in PBS and digested with 20 μg/ml PK for 30 min at 37°C.

PMCA, PK digestion and Western blotting

PMCA was performed and quantified as previously described (13) with slight modifications. Samples were sonicated at 70–85% maximum power for 40 s in a microplate horn sonicator (Qsonica, Framingham, MA), followed by a 30-min incubation at 37°C repeated for 24 h per round for up to five rounds total. PK digestion and Western blotting were performed as previously described (31), except that spleen homogenates were digested with 10 μg/ml PK. PrP was detected using HRP-conjugated Bar244 Ab (Bertin Pharma, Paris, France).

Histochemistry and immunohistochemistry

Slides were prepared and stained as previously described (31). Briefly, 10-μm sections were cut from paraffin-embedded spleen tissue and mounted onto glass slides. Splenic follicles were stained with rat anti-mouse IgM (02031D; BD Pharmingen) followed by goat anti-rat IgG (H+L) followed by alkaline phosphatase-conjugated donkey anti-goat IgG (705-055-147; Jackson ImmunoResearch Laboratories) and visualized with fast blue (Polysciences, Warrington, PA). GCs were stained with biotin-conjugated peanut agglutinin (PNA) ABComplex/HRP (Dako) and visualized with 3-amino-9-ethylcarbazole (A5754; Sigma-Aldrich).

Splenocyte isolation and flotation assays

Individual whole spleens from at least five mice per group were ground through a nylon mesh to release lymphocytes into single-cell suspensions, which were then centrifuged 5 min at 200 × g and washed twice with ice-cold PBS. The remaining splenic tissue was digested for 20 min at 37°C in 1 mg/ml collagenase, 0.5 mg/ml dispase, and 40 μg/ml DNase I (Roche, Mannheim, Germany) with agitation. Supernatants were collected and the remaining tissue was digested for another 20 min. The samples were pooled and centrifuged 5 min at 200 × g. Cell pellets were washed twice with ice-cold PBS, combined with corresponding lymphocyte pellets, and incubated 30 min on ice in 400 μl 1% Triton X-100 in TNE buffer (25 mM Tris-HCl [ph 7.4], 150 mM NaCl, 5 mM EDTA, 5 mM DTT) containing protease inhibitors (Complete Mini tablet; Roche). Samples were centrifuged 10 min at 1000 × g at 4°C to pellet debris, and supernatants were transferred to new tubes. Lysates (133 μl) were mixed with 267 μl 60% (w/v) OptiPrep solution (Axis-Shield, Oslo, Norway) and pipetted to the bottom of 13 × 51-mm UltraClear centrifuge tubes (Beckman Coulter, Palo Alto, CA). Two hundred-microliter aliquots of 35, 30, 25, and 20% OptiPrep were gently layered on top of the lysates. Tubes were centrifuged 12 h at 4°C in an S55S Sorvall M150 SE rotor at 120,000 × g. Two hundred-microliter fractions were collected from the top of the tube. SDS-PAGE loading buffer was added to aliquots of each fraction with or without PK digestion (20 μg/ml for 30 min) and subjected to 4–12% gradient PAGE.

GC counting and statistical and phylogenetic analyses

Follicles were counted in three nonconsecutive sections from five distinct areas from at least five spleens as IgM+ B cells forming characteristic follicular foci. GCs were counted as PNA+ B cells (brown stain) within follicles (IgM+ blue stain). We derived percentages of follicles containing GCs by dividing the number of GCs by the number of follicles and multiplying by 100. Statistical analyses were performed using Excel (Microsoft) and Prism software (GraphPad Software). CD21 sequence alignment and phylogenetic anyalysis was performed using Geneious (BioMatters, Auckland, New Zealand).


Absence of CD21/35 protects mice from CWD

To create a mouse deficient in complement receptors CD21/35 and susceptible to CWD prions, mice deficient in both CD21/35 (CD21/35−/−) and mouse PrPC (Prnpo/o) were crossed to Tg(cerPrP)5037 mice that express high levels of elk, but no mouse, PrPC. Offspring were then screened and the resulting Tg5037;CD21/35−/− and Tg5037 littermates were inoculated with 100 μg brain homogenate from an elk terminally infected with CWD prions (E2). Tg5037;CD21/35−/− mice showed complete resistance to CWD prions, with no mice showing any clinical signs at the end of the study, whereas infected Tg5037 mice died of CWD at a median time of 301 dpi (Fig. 1A).

Mice deficient in CD21/35 show resistance to CWD prion infection. (A) Tg5037 (blue line, n = 8) and Tg5037;CD21/35−/− (red line, n = 11) mice were inoculated with 100 μg brain homogenate from an elk terminally infected with CWD ...

We next examined terminally sick Tg5037 and dpi-matched Tg5037;CD21/35−/− mice for characteristic signs of CWD neuropathology (Fig. 1B–G). Minimal astrogliosis and vacuolization with no PrPRES deposition were observed in nonclinical Tg5037;CD21/35−/− mice at 285 dpi (Fig. 1B, ,1D,1D, ,1F).1F). However, significant vacuolization, astrogliosis, and PrPRES deposition were observed in terminally sick Tg5037 mice at 285 dpi (Fig. 1C, ,1E,1E, ,1G).1G). Western blot and densitometric analyses revealed PrPRES in only 3 of 11 brains from Tg5037;CD21/35−/− mice, significantly less than in brains from terminally sick Tg5037 mice, all of which contained PrPRES and, when compiled together, contained 3.5-fold more PrPRES (Fig. 1H, ,1I1I).

Absence of CD21/35 delays prion propagation in the spleen

We next used a semiquantitative prion amplification assay to estimate prion loads in spleens of CWD-infected Tg5037 and Tg5037;CD21/35−/− mice. PMCA, a technique used to amplify prions in vitro, takes advantage of a prion’s ability to self-propagate, using seeded protein fibrilization (Fig. 2A). We employed PMCA to amplify and quantify minute amounts of PrPRES from spleen homogenates from mice at various intervals postinfection. Tg5037 mice at 15 dpi show a significant difference in the amount of splenic PrPRES (75.49 ± 4.43 relative PMCA units [rpu]; Fig. 2B, ,2C)2C) compared with Tg5037;CD21/35−/− mice (25.56 ± 4.24 rpu). Using our standard curve for this assay generated previously, we estimate that splenic PrPRES load in Tg5037 mice is ~20,000 pg/g spleen tissue (Fig. 2C). The PMCA score for Tg5037;CD21/35−/− mice falls just out of the dynamic range of our assay (29 rpu), so we can only estimate the load to be <100 pg/g. We detected much less PrPRES in spleens from Tg5037 mice at 30 dpi (17.17 ± 0.16 rpu) whereas Tg5037;CD21/35−/− spleens showed only a slight decrease in PrPRES load (21.88 ± 0.15 rpu). Both scores fall just below the range of the PMCA score standard curve. Accumulation of PrPRES differed drastically between the two groups at 30 dpi. PrPRES load increased significantly more in Tg5037 mice from 70 (37.50 ± 0.15 rpu, 250 pg/g) to 140 dpi (55.56 ± 0.11 rpu, 2000 pg/g) to terminal disease (71.00 ± 0.2 rpu, 10,000 pg/g). We detected significantly less PrPRES in spleens from nonclinical Tg5037;CD21/35−/− at 70 (20.00 ± 0.14 rpu, <100 pg/g) and 140 dpi (41 ± 0.16 rpu, 500 pg/g) and at dpi matched to sick Tg5037 mice (47.00 ± 0.50 rpu, 900 pg/g).

Tg5037;CD21/35−/− mice show delayed prion accumulation in the spleen. (A) Schematic representation of PMCA. Alternating cycles of incubation and sonication of cellular PrP seeded with PrPRES with substrate followed by PK digestion and ...

CD21/35 mediate a strong GC response during prion infection

Increased PrPRES retention could be mediated via formation of GCs. Ag trapping by Fcγ receptors and CD21/35 on FDCs, as well as concomitant B cell signaling through the BCR and the CD21/CD19/CD81 coreceptor, stimulates lymphoid follicles to generate GCs (3235) containing arborized FDCs. Prolonged Ag presentation by FDCs and additional signals (CD40/CD40L) activates B cells to become plasma or memory B cells (36, 37). CD21/35−/− and C3−/− mice have reduced size and numbers of GCs before antigenic stimulation (38, 39), but high doses of Ag increase GCs to near wild-type (WT) levels. We observed a similar phenomenon in spleens of mice overexpressing PrPC (TgA20, Tg5037, TgA20;CD21/35−/−, and Tg5037;CD21/35−/− mice), which exhibited significant GC formation during prion infection independent of CD21/35 expression (data not shown). This most likely occurred because increased PrPC expression increases prion replication and concomitant antigenic stimulation of GC formation, just as high doses of microbial Ags can.

In mouse models expressing normal physiological levels of PrPC and CD21/35, scrapie prion infection causes abnormal GC reactions characterized by hypertrophic FDC dendrites, PrPRES accumulation, and increased maturation and numbers of B cells (40). We therefore investigated the role of CD21/35 in this process by analyzing GC formation in mice expressing normal WT mouse PrPC levels, with or without CD21/35 expression, after infecting them with RML5 mouse-adapted scrapie prions. We discovered that i.p. inoculation of high and low doses of prions, but not uninfected (mock) brain homogenate, stimulated significant GC formation in spleens of WT, but not CD21/35−/−, mice (Fig. 3A–D, Table I; n ≥ 5). Interestingly, prion infection, but not 108 CFU heat-killed Escherichia coli or DNP-keyhole limpet hemocyanin (data not shown), induced GC formation in C3/4−/− mice, suggesting that CD21/35 can mediate prion-induced GC reactions independent of its endogenous ligands. Moreover, we detected significant amounts of CD21/35 in PrPRES preparations enriched by NaPTA precipitation from spleen homogenates of infected mice at 60 dpi, but very little CD21/35 from NaPTA precipitates from mock-infected spleens (Fig. 3E). We recovered far less PrPRES from CD21/35−/− spleens, consistent with our PMCA data from Tg5037;CD21/35−/− spleens.

Prion infection stimulates GC formation and increases CD21/35 presence in prion-enriched spleen preparations. (AE) Mice (n ≥ 10) were inoculated i.p. with 0.1% prion-infected brain homogenate, sacrificed 60 dpi, and frozen spleen sections ...
Table I.
GC formation in peripheral prion disease

CD21/35 translocate to lipid rafts on B cells upon prion infection

GC formation by the traditional primary immune response requires CD21/35 expression on B cells but not on FDCs (35). Closer examination of GC formation in bone marrow (BM) chimeric mice infected with prions divulged a pattern of GC formation predominantly dependent on both CD21/35 and PrPC expression on FDCs (Table II; n ≥ 5), suggesting that CD21/35 Ag presentation is more important than CD21/35 signaling to mediate this reaction. To resolve this discrepancy in GC formation we assessed whether prion infection alters localization of CD21/35 on the plasma membrane. In a typical primary immune response, C3d- or C4d-opsonized Ag binds to specific short consensus repeats of CD21/35, inducing CD19-mediated palmitoylation of the tetraspanin CD81 (41, 42) that moves the entire complex into lipid rafts and cholesterol- and sphingolipid-rich microdomains of the plasma membrane (43). This offers an attractive model of prion replication because PrPC resides in lipid rafts via its GPI anchor, and translocation of CD21/35 to the same rafts upon prion infection could effectively bring PrPRES to PrPC. We therefore monitored CD21/35 translocation to lipid rafts on splenocytes from infected mice by lysing them in cold Triton X-100 and subjecting the cleared lysate to density gradient centrifugation. CD21/35 reside predominantly outside lipid rafts on splenocytes from mock-infected WT mice, as indicated by its presence in detergent-soluble fractions (Fig. 4A). In contrast, PrP was detected in detergent-insoluble fractions containing lipid rafts that float to the top of the density gradient and colocalize with the raft marker flotillin. Upon prion infection, a significant amount of CD21/35 moved into detergent-insoluble fractions (Fig. 4B). CD21/35 were present in the same fractions as flotillin and PK-resistant PrPRES, but not the IgH, indicating that CD21/35 translocation occurred independent of the BCR. Furthermore, whereas a T cell-dependent Ag failed to stimulate CD21/35 translocation to lipid rafts in C3/4−/− mice (Fig. 4C), prion infection induced a significant amount of CD21/35 translocation (Fig. 4D). These data strongly suggest that CD21/35 can interact with prions independent of its endogenous ligands, which could explain why CD21/35−/− mice exhibit a more significant delay in disease progression than do C3/4−/− mice. Although the lack of CD21/35 expression does not completely prevent PrPRES accumulation in lipid rafts, it significantly decreases PrPRES load (Fig. 4E). Flotation assays on spleens from BM chimeric mice revealed that FDCs appear to express only the short form of the complement receptor CD21, and that prion infection fails to translocate CD21 to lipid rafts on FDCs (Fig. 4F). Thus, prion infection provokes CD21/35 translocation on B cells, which express all members of the CD21/CD19/CD81 coreceptor complex, but not on FDCs, which only express CD21.

Table II.
GC formation in BM chimeric mice infected with prions
Prion infection stimulates CD21/35 translocation to lipid rafts on B cells in CD21/35-expressing mice. Splenocytes (2 × 107) were lysed in ice-cold Triton X-100 and cleared lysates were centrifuged at the bottom of a density gradient (see Materials ...


We investigated the role of the complement receptors CD21/35 in CWD prion accumulation, replication, and disease progression. We observed a complete rescue from terminal CWD of Tg5037 mice lacking CD21/35. Only 3 of 11 nonclinical Tg5037;CD21/35−/− mice displayed detectable, yet reduced, prion neuropathology and PrPRES deposition in their brains. These results reveal a more dramatic outcome than earlier studies showing only a partial rescue of CD21/35-deficient mice from scrapie infection, despite those mice expressing only WT (i.e., 5-fold less) PrPC levels. This could reflect differences between mouse and cervid CD21 expression, as are apparent between mouse and human CD21. However, little is known about cervid CD21. The gene has yet to be cloned, so comparative analyses with murine CD21/35 are impossible at present. We can, however, compare CD21 sequence homology and phylogeny among other species that are susceptible to TSEs. For example, sheep, which are susceptible to scrapie, a TSE that closely resembles CWD in transmission efficiency, and lymphotropism, express a CD21 molecule that shares 65% sequence identity with murine CD21/35, including their ligand binding domains (Fig. 5A). This may explain the similar lymphotropic characteristics of murine and ovine scrapie. Ovine CD21 also shares 65% identity with human CD21/35. Overall, CD21/35 from these three species share 52% identity and 64% similarity. In contrast, bovine CD21, which is 40% larger than the other three CD21/35 molecules (~1400 compared with ~1000 aa, respectively), shares <20% similarity to the other three CD21/35 molecules. Phylogenetic analysis reveals a clustering of murine, ovine, and human CD21/35 proteins, with bovine CD21 much more distantly related (Fig. 5B). Interestingly, bovine spongiform encephalopathy has been shown to have little or no lymphotropic characteristics (4447), perhaps owing to the vastly different CD21 molecule that bovids express.

Sequence and phylogenetic analyses of CD21/35. (A) Sequence alignment of ligand binding domains of bovine, human (shown in lowercase letters), mouse, and sheep CD21. Dashes represent gaps in sequence alignment, revealing additional amino acids in bovine ...

These results indicate a significant role in prion pathogenesis for CD21/35, the importance of which may vary by prion strain. Complement components C1q and C3 have recently been shown to exhibit similar strain preferences in vitro and in vivo (48). We are currently investigating other prion strains to determine the contribution of CD21/35 to prion pathogenesis in those infection models. Interestingly, cross-species prion transmission was recently shown to result in a higher infection rate of the lymphoreticular system than the CNS in the xenohost (49). This cross-species infection resulted in distinct lymphotropic and neurotropic strains with differential host ranges. These strains may result from tissue-specific strain selection or mutation. The higher efficiency of prion infection in the spleen (which harbors CD21/35-expressing FDCs and B cells) compared with the brain (which lacks them) alludes to a critical role for CD21/35 in prion retention, replication, and possibly strain selection in trans-species prion infection. The lack of CD21/35 that delays peripheral prion accumulation might further limit the lymphoid replication of neurotropic prion strains, resulting in delayed or abrogated disease progression. If so, this would have profound implications for prion xenotransmission and possible therapeutic approaches aimed at CD21/35. For example, targeting CD21/35 to slow the spread of neurotropic prions could be an attractive alternative to most prion disease therapeutics developed to date that target the CNS, which can complicate drug delivery. Interfering with CD21/35-mediated prion strain selection could also mitigate emergence of new prion strains with expanded host ranges and prevent a breach of the species barrier similar to the one that likely caused the bovine spongiform encephalopathy and subsequent new-variant Creutzfeldt-Jakob disease outbreak 15 y ago in the United Kingdom.

To study the kinetics of extraneural CWD prion accumulation, we amplified PrPRES from spleens of CWD prion-infected Tg5037 and Tg5037;CD21/35−/− mice at various time points throughout infection. At 15, 70, and 140 dpi and at terminal disease, prion accumulation was significantly lower in CD21/35-deficient mice. The extremely high prion load detected at 15 dpi most likely reflects increased retention of prion inocula early after infection. This delay in extraneural prion accumulation strongly correlates with abrogation of prion neuropathology and terminal disease. These results coincide with our previous data from scrapie mouse models (17), further strengthening evidence that CD21/35 play an integral part in prion accumulation in peripheral lymphoid organs that ultimately facilitates neuroinvasion.

Furthermore, we show that CD21/35 are present in prion preparations enriched from spleen homogenates by NaPTA precipitation. We also demonstrate GC formation in spleens during prion infection primarily dependent on CD21/35 and PrPC expression on FDCs. It may seem surprising that CD21/35 expression on FDCs, rather than B cells, correlates with prion-induced GC formation, because CD21/CD19/CD81 B cell coreceptor ligation helps activate B cells to form GCs. However, maximal B cell activation and GC formation require signaling from both the BCR and B cell coreceptor (32, 33). In this study, we show that although prion infection stimulates CD21/35 translocation to lipid rafts on B cells, signaling appears to be suboptimal for GC formation in the absence of concomitant BCR translocation. We observed a strong dependence on both PrP and CD21/35 expression on FDCs for a strong GC response. Paradoxically, CD21/35 translocation did not occur on FDCs, which are the major prion trappers and replicators but lack other B cell coreceptor components required for CD21/35 movement. One could therefore argue that GC formation represents an artifact, rather than being a driver of splenic prion replication. Elimination of GCs had no effect on peripheral prion replication and disease progression in mice infected i.p. with RML5 (50), supporting this interpretation. However, GC-deficient mice infected intracranially with 139A mouse-adapted scrapie prions exhibited a significant delay to terminal disease (51). Thus, distinct prion strains may differentially influence GC formation and subsequent prion pathogenesis. Additionally, this discrepancy further highlights potential preferences of distinct tissues for different prion strains. CD21/35-expressing cells within GCs may facilitate this selection process in the lymphoid system. Increased retention of prions on FDCs could induce a persistent state of prion presentation to adjacent B cells sufficient to cause an atypical GC response (40). FDCs may coax B cells to linger there, providing increased lymphotoxin signaling to FDCs that may promote formation of hypertrophic dendrites that efficiently retain and replicate prions. Consistent with their role as long-lived, long-term APCs, FDCs may also present prions to neighboring PrPC-expressing B cells that could induce CD21/35 translocation and move prions proximal to PrpC in lipid rafts and promote further prion replication and GC formation.

Taken together, these data support a principal role of CD21/35 in peripheral prion pathogenesis by trapping PrPRES on both B cells and FDCs. CD21/35 expression on FDCs remains of paramount importance in this process, with B cells playing a lesser, but still important, role. We have recently shown that few prion-bearing B cells transport prions from infection sites to draining lymph nodes, but their presence increased dramatically within lymph nodes, indicating a prominent role for B cells in intranodal prion trafficking (52). We propose that CD21/35 mediate this and other crucial processes in lymph node prion trapping and replication and we are currently testing this hypothesis.


We thank Ed Hoover and Steve Dow for helpful advice and discussion of the project and data.

This work was supported by National Institute of Neurological Disorders and Stroke Grant R01 NS56379.

Abbreviations used in this article:

bone marrow
chronic wasting disease
days postinoculation
follicular dendritic cell
germinal center
sodium phosphotungstic acid
proteinase K
protein misfolding cyclic amplification
peanut agglutinin
prion protein
cellular prion protein
prion disease-associated, misfolded prion protein
relative PMCA unit
transmissible spongiform encephalopathy


The authors have no financial conflicts of interest.


1. Williams E. S., Young S. 1982. Spongiform encephalopathy of Rocky Mountain elk. J. Wildl. Dis. 18: 465–471. [PubMed]
2. Prusiner S. B. 1982. Novel proteinaceous infectious particles cause scrapie. Science 216: 136–144. [PubMed]
3. Miller M. W., Williams E. S. 2003. Prion disease: horizontal prion transmission in mule deer. Nature 425: 35–36. [PubMed]
4. Ryder S., Dexter G., Bellworthy S., Tongue S. 2004. Demonstration of lateral transmission of scrapie between sheep kept under natural conditions using lymphoid tissue biopsy. Res. Vet. Sci. 76: 211–217. [PubMed]
5. Gajdusek D. C., Gibbs C. J. J., Jr., Alpers M. 1967. Transmission and passage of experimenal “kuru” to chimpanzees. Science 155: 212–214. [PubMed]
6. Mould D. L., Dawson A. M., Rennie J. C. 1970. Very early replication of scrapie in lymphocytic tissue. Nature 228: 779–780. [PubMed]
7. Bosque P. J., Ryou C., Telling G., Peretz D., Legname G., DeArmond S. J., Prusiner S. B. 2002. Prions in skeletal muscle. Proc. Natl. Acad. Sci. USA 99: 3812–3817. [PubMed]
8. Angers R. C., Browning S. R., Seward T. S., Sigurdson C. J., Miller M. W., Hoover E. A., Telling G. C. 2006. Prions in skeletal muscles of deer with chronic wasting disease. Science 311: 1117–1117. [PubMed]
9. Mathiason C. K., Powers J. G., Dahmes S. J., Osborn D. A., Miller K. V., Warren R. J., Mason G. L., Hays S. A., Hayes-Klug J., Seelig D. M., et al. 2006. Infectious prions in the saliva and blood of deer with chronic wasting disease. Science 314: 133–136. [PubMed]
10. Seeger H., Heikenwalder M., Zeller N., Kranich J., Schwarz P., Gaspert A., Seifert B., Miele G., Aguzzi A. 2005. Coincident scrapie infection and nephritis lead to urinary prion excretion. Science 310: 324–326. [PubMed]
11. Haley N. J., Seelig D. M., Zabel M. D., Telling G. C., Hoover E. A. 2009. Detection of CWD prions in urine and saliva of deer by transgenic mouse bioassay. PLoS ONE 4: e4848. [PMC free article] [PubMed]
12. Tamgüney G., Miller M. W., Wolfe L. L., Sirochman T. M., Glidden D. V., Palmer C., Lemus A., DeArmond S. J., Prusiner S. B. 2009. Asymptomatic deer excrete infectious prions in faeces. Nature 461: 529–532. [PMC free article] [PubMed]
13. Pulford B., Spraker T. R., Wyckoff A. C., Meyerett C., Bender H., Ferguson A., Wyatt B., Lockwood K., Powers J., Telling G. C., et al. 2012. Detection of PrPCWD in feces from naturally exposed Rocky Mountain elk (Cervus elaphus nelsoni) using protein misfolding cyclic amplification. J. Wildl. Dis. 48: 425–434. [PubMed]
14. 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. [PubMed]
15. 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. Nat. Med. 5: 1308–1312. [PubMed]
16. Heikenwalder M., Zeller N., Seeger H., Prinz M., Klöhn P. C., Schwarz P., Ruddle N. H., Weissmann C., Aguzzi A. 2005. Chronic lymphocytic inflammation specifies the organ tropism of prions. Science 307: 1107–1110. [PubMed]
17. Zabel M. D., Heikenwalder M., Prinz M., Arrighi I., Schwarz P., Kranich J., von Teichman A., Haas K. M., Zeller N., Tedder T. F., et al. 2007. Stromal complement receptor CD21/35 facilitates lymphoid prion colonization and pathogenesis. J. Immunol. 179: 6144–6152. [PubMed]
18. McCulloch L., Brown K. L., Bradford B. M., Hopkins J., Bailey M., Rajewsky K., Manson J. C., Mabbott N. A. 2011. Follicular dendritic cell-specific prion protein (PrP) expression alone is sufficient to sustain prion infection in the spleen. PLoS Pathog. 7: e1002402. [PMC free article] [PubMed]
19. Krautler N. J., Kana V., Kranich J., Tian Y., Perera D., Lemm D., Schwarz P., Amulik A., Browning J. L., Tallquist M., et al. 2012. Follicular dendritic cells emerge from ubiquitous perivascular precursors. Cell 150: 194–206. [PubMed]
20. Prinz M., Heikenwalder M., Junt T., Schwarz P., Glatzel M., Heppner F. L., Fu Y. X., Lipp M., Aguzzi A. 2003. Positioning of follicular dendritic cells within the spleen controls prion neuroinvasion. Nature 425: 957–962. [PubMed]
21. 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. Nature 390: 687–690. [PubMed]
22. 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. Nat. Med. 4: 1429–1433. [PubMed]
23. Zabel M. D., Weis J. H. 2001. Cell-specific regulation of the CD21 gene. Int. Immunopharmacol. 1: 483–493. [PubMed]
24. Roozendaal R., Carroll M. C. 2007. Complement receptors CD21 and CD35 in humoral immunity. Immunol. Rev. 219: 157–166. [PubMed]
25. 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., et al. 2001. Complement facilitates early prion pathogenesis. Nat. Med. 7: 488–492. [PubMed]
26. 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. Nat. Med. 7: 485–487. [PubMed]
27. Mitchell D. A., Kirby L., Paulin S. M., Villiers C. L., Sim R. B. 2007. Prion protein activates and fixes complement directly via the classical pathway: implications for the mechanism of scrapie agent propagation in lymphoid tissue. Mol. Immunol. 44: 2997–3004. [PubMed]
28. Sim R. B., Kishore U., Villiers C. L., Marche P. N., Mitchell D. A. 2007. C1q binding and complement activation by prions and amyloids. Immunobiology 212: 355–362. [PubMed]
29. Fischer M., Rülicke T., Raeber A., Sailer A., Moser M., Oesch B., Brandner S., Aguzzi A., Weissmann C. 1996. Prion protein (PrP) with amino-proximal deletions restoring susceptibility of PrP knockout mice to scrapie. EMBO J. 15: 1255–1264. [PubMed]
30. Angers R. C., Seward T. S., Napier D., Green M., Hoover E., Spraker T., O’Rourke K., Balachandran A., Telling G. C. 2009. Chronic wasting disease prions in elk antler velvet. Emerg. Infect. Dis. 15: 696–703. [PMC free article] [PubMed]
31. Meyerett C., Michel B., Pulford B., Spraker T. R., Nichols T. A., Johnson T., Kurt T., Hoover E. A., Telling G. C., Zabel M. D. 2008. In vitro strain adaptation of CWD prions by serial protein misfolding cyclic amplification. Virology 382: 267–276. [PubMed]
32. Fang Y., Xu C., Fu Y. X., Holers V. M., Molina H. 1998. Expression of complement receptors 1 and 2 on follicular dendritic cells is necessary for the generation of a strong antigen-specific IgG response. J. Immunol. 160: 5273–5279. [PubMed]
33. Fischer M. B., Goerg S., Shen L., Prodeus A. P., Goodnow C. C., Kelsoe G., Carroll M. C. 1998. Dependence of germinal center B cells on expression of CD21/CD35 for survival. Science 280: 582–585. [PubMed]
34. Aydar Y., Balogh P., Tew J. G., Szakal A. K. 2003. Altered regulation of FcγRII on aged follicular dendritic cells correlates with immunoreceptor tyrosine-based inhibition motif signaling in B cells and reduced germinal center formation. J. Immunol. 171: 5975–5987. [PubMed]
35. Croix D. A., Ahearn J. M., Rosengard A. M., Han S., Kelsoe G., Ma M., Carroll M. C. 1996. Antibody response to a T-dependent antigen requires B cell expression of complement receptors. J. Exp. Med. 183: 1857–1864. [PMC free article] [PubMed]
36. Kawabe T., Naka T., Yoshida K., Tanaka T., Fujiwara H., Suematsu S., Yoshida N., Kishimoto T., Kikutani H. 1994. The immune responses in CD40-deficient mice: impaired immunoglobulin class switching and germinal center formation. Immunity 1: 167–178. [PubMed]
37. Xu J., Foy T. M., Laman J. D., Elliott E. A., Dunn J. J., Waldschmidt T. J., Elsemore J., Noelle R. J., Flavell R. A. 1994. Mice deficient for the CD40 ligand. Immunity 1: 423–431. [PubMed]
38. Haas K. M., Hasegawa M., Steeber D. A., Poe J. C., Zabel M. D., Bock C. B., Karp D. R., Briles D. E., Weis J. H., Tedder T. F. 2002. Complement receptors CD21/35 link innate and protective immunity during Streptococcus pneumoniae infection by regulating IgG3 antibody responses. Immunity 17: 713–723. [PubMed]
39. Fischer M. B., Ma M., Goerg S., Zhou X., Xia J., Finco O., Han S., Kelsoe G., Howard R. G., Rothstein T. L., et al. 1996. Regulation of the B cell response to T-dependent antigens by classical pathway complement. J. Immunol. 157: 549–556. [PubMed]
40. McGovern G., Brown K. L., Bruce M. E., Jeffrey M. 2004. Murine scrapie infection causes an abnormal germinal centre reaction in the spleen. J. Comp. Pathol. 130: 181–194. [PubMed]
41. Cherukuri A., Carter R. H., Brooks S., Bornmann W., Finn R., Dowd C. S., Pierce S. K. 2004. B cell signaling is regulated by induced palmitoylation of CD81. J. Biol. Chem. 279: 31973–31982. [PubMed]
42. Cherukuri A., Shoham T., Sohn H. W., Levy S., Brooks S., Carter R., Pierce S. K. 2004. The tetraspanin CD81 is necessary for partitioning of coligated CD19/CD21-B cell antigen receptor complexes into signaling-active lipid rafts. J. Immunol. 172: 370–380. [PubMed]
43. Cherukuri A., Dykstra M., Pierce S. K. 2001. Floating the raft hypothesis: lipid rafts play a role in immune cell activation. Immunity 14: 657–660. [PubMed]
44. Bradley R. 1999. BSE transmission studies with particular reference to blood. Dev. Biol. Stand. 99: 35–40. [PubMed]
45. Iwata N., Sato Y., Higuchi Y., Nohtomi K., Nagata N., Hasegawa H., Tobiume M., Nakamura Y., Hagiwara K., Furuoka H., et al. 2006. Distribution of PrP(Sc) in cattle with bovine spongiform encephalopathy slaughtered at abattoirs in Japan. Jpn. J. Infect. Dis. 59: 100–107. [PubMed]
46. Middleton D. J., Barlow R. M. 1993. Failure to transmit bovine spongiform encephalopathy to mice by feeding them with extraneural tissues of affected cattle. Vet. Rec. 132: 545–547. [PubMed]
47. Terry L. A., Marsh S., Ryder S. J., Hawkins S. A., Wells G. A., Spencer Y. I. 2003. Detection of disease-specific PrP in the distal ileum of cattle exposed orally to the agent of bovine spongiform encephalopathy. Vet. Rec. 152: 387–392. [PubMed]
48. Hasebe R., Raymond G. J., Horiuchi M., Caughey B. 2012. Reaction of complement factors varies with prion strains in vitro and in vivo. Virology 423: 205–213. [PMC free article] [PubMed]
49. Béringue V., Herzog L., Jaumain E., Reine F., Sibille P., Le Dur A., Vilotte J. L., Laude H. 2012. Facilitated cross-species transmission of prions in extraneural tissue. Science 335: 472–475. [PubMed]
50. Heikenwalder M., Federau C., Boehmer L. V., Schwarz P., Wagner M., Zeller N., Haybaeck J., Prinz M., Becher B., Aguzzi A. 2007. Germinal center B cells are dispensable in prion transport and neuroinvasion. J. Neuroimmunol. 192: 113–123. [PubMed]
51. Burwinkel M., Schwarz A., Riemer C., Schultz J., van Landeghem F., Baier M. 2004. Rapid disease development in scrapie-infected mice deficient for CD40 ligand. EMBO Rep. 5: 527–531. [PubMed]
52. Michel B., Meyerett-Reid C., Johnson T., Ferguson A., Wyckoff C., Pulford B., Bender H., Avery A., Telling G., Dow S., Zabel M. D. 2012. Incunabular immunological events in prion trafficking. Sci Rep 2: 440. [PMC free article] [PubMed]

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