Costimulatory CD40 plays an essential role in autoimmune diseases, including experimental autoimmune encephalomyelitis (EAE), a murine model of human multiple sclerosis (MS). However, how CD40 drives autoimmune disease pathogenesis is not well defined. Here, we used a conditional knockout approach to determine how CD40 orchestrates a CNS autoimmune disease induced by recombinant human myelin oligodendrocyte glycoprotein (rhMOG). We found that deletion of CD40 in either dendritic cells (DCs) or B cells profoundly reduced EAE disease pathogenesis. Mechanistically, CD40 expression on DCs was required for priming pathogenic Th cells in peripheral draining lymph nodes and promoting their appearance in the CNS. By contrast, B cell CD40 was essential for class-switched MOG-specific Ab production, which played a crucial role in disease pathogenesis. In fact, passive transfer of MOG-immune serum or IgG into mice lacking CD40 on B cells but not DCs reconstituted autoimmune disease, which was associated with inundation of the spinal cord parenchyma by Ig and complement. These data demonstrate that CD40 supports distinct effector programs in B cells and DCs that converge to drive a CNS autoimmune disease and identify targets for intervention.

We have characterized the role of the costimulatory CD40 molecule in autoimmune pathogenesis, studying experimental autoimmune encephalomyelitis (EAE), a murine model of human multiple sclerosis (MS). MS is an inflammatory demyelinating disease of the CNS, characterized by progressive neurologic deficits and disability (1). Myelin breakdown in the white matter is accompanied by infiltrates of multiple lymphoid and myeloid cell types (2, 3). At present, the mechanism of the disease is not fully understood, and there is no curative treatment (4). Immunotherapies that block T cell growth signaling by targeting CD25 (daclizumab), impeding lymphocyte trafficking to the CNS by downregulating S1PR (fingolimod), or targeting B cells with anti-CD20 mAb (rituximab) have been approved for the treatment of MS (5, 6). Additional clinical trials are targeting inflammatory cytokines with anti–IL-17A (secukinumab) or anti–GM-CSF (MOR103) Ab (7, 8), collectively suggesting that multiple components of the immune response may play a role in MS pathogenesis.

Analysis of the immune mechanisms underlying MS has been informed by the murine EAE model, which approximates the pathological features of MS. EAE can be induced by immunization with CNS Ags, including the myelin oligodendrocyte glycoprotein (MOG). In the EAE model, Ag-specific, differentiated IFN-γ–producing Th1 cells (9), IL-17–producing Th17 cells (10), IFN-γ and IL-17 double-positive cells (11), and GM-CSF–producing Th cells (12) are primed by APCs in the peripheral lymphoid organs. These pathogenic T cells traffic across the blood–brain barrier and can be restimulated by local APCs in the CNS to recruit additional immune cells that mediate tissue damage (13, 14). In addition to this role of pathogenic Th cells, B cell infiltration, Abs to myelin components, and activated complement are routinely detected in patients with MS (15, 16), and B cell–deficient mice are resistant to rhMOG-induced EAE (1719), indicating that both T and B cells are required for rhMOG-induced EAE.

Induction of autoimmune diseases, including EAE, has also been shown to require costimulatory interactions, including those mediated by the B7/CD28 (20, 21) and CD40/CD40L pathways (2225). In the complete absence of CD40 or when treated with CD40L-blocking Ab, mice are completely protected from EAE induction by MOG35-55 peptide (20, 26). However, the mechanisms mediating the CD40 role in EAE, including requirements for cell type–specific expression of CD40, have not been identified. To elucidate the mechanisms of CD40 involved in EAE as an informative model of autoimmune disease, we have analyzed the cellular requirements for CD40 expression in clinical EAE induction, in peripheral T cell priming to MOG, in T cell infiltration in the CNS, and in autoantibody production, using conditional CD40 deletion and bone marrow (BM) chimera strategies. We have identified distinct and complementary mechanisms mediating requirements for CD40 expression by B cells and by dendritic cells (DCs) in EAE pathogenesis.

Animal experiments were approved by the National Cancer Institute Animal Care and Use Committee. All mice were maintained in accordance with National Institutes of Health guidelines. Similar numbers of male and female 6–12-wk-old mice were used in all experiments. CD40flox (CD40fl/fl) mice were generated on a C57BL/6 background. The Bioengineering Core of the University of Colorado–Denver made the construct. In brief, the first exon of CD40 was flanked by two loxP sites. The pLFNFL cloning vector was used for the construction of CD40 conditional knockout vector, which was generated through the replacement of PGK-Puro in the PFlexible resource (Wellcome Sanger Institute) with a PGK-neo selection cassette by restricted digestion and ligation. The left (2808 bp) and right (2668 bp) homologous arms of CD40 amplified by PCR from C57BL/6 genomic DNA were cloned at the EcoRI and NotI sites of pLFNFL, respectively, so they flanked the loxP-FRT-neo-FRT-loxP cassette. The left and right homologous sequences correspond to the genomic sequence upstream and downstream of exon 1 of CD40, respectively. Next, a 594-bp genomic sequence encompassing exon 1 of CD40, generated by PCR, was cloned at the HindIII site located between the loxP and FRT sites upstream of neo. Finally, a thymidine kinase (TK) gene was cloned at the ClaI site through blunt-end ligation at a position proximal to the left homology arm. The resulting construct (pLFNFL-TK-Cd40) was linearized by KpnI digestion and purified by chloroform and ethanol precipitation and then introduced by electroporation into murine B6/129 hybrid EC7.1 embryonic stem cells in the Transgenic and Gene Targeting Core Laboratory at the University of Colorado–Denver. Two 96-well plates of recombinants were screened for CD40 targeting events by long-range PCR. A correctly targeted clone (A12) was identified and was microinjected into C57BL/6 blastocysts to produce chimeric founders at the Transgenic Animal Core at the University of Colorado–Denver. The FRT-flanked neo selection cassette was deleted by crossing to a Flp-deleter strain.

CD19-cre [B6.129P2(C)-Cd19tmi(cre)Cgn/J, JAX-008068] and CD11c-cre [B6.Cg-Tg(Itgax-cre)1-1Reiz/J, JAX-006785] mice were purchased from The Jackson Laboratory. For selective deletion of the floxed CD40 gene in B cells or DCs, CD19-cre or CD11c-cre mice were crossed with CD40fl/fl mice. Littermates heterozygous for the cre cassette (CD19-cre/CD40fl/fl, CD19-wt/CD40fl/fl or CD11c-cre/CD40fl/fl, CD11c-wt/CD40fl/fl) were used in experiments. C57BL/6 mice were purchased from Charles River Laboratories (strain code 027), and µMT mice (B6.129S2-Ighmtm1Cgn/J, JAX-002288) were purchased from The Jackson Laboratory. BM chimera mice were prepared as previously described by reconstitution of 950-rad irradiated host mice with 6 × 106 total T cell–depleted BM cells i.v. (27). Mixed chimeras were generated by reconstitution with 6 × 106 total T cell–depleted BM cells from donor mice at 1:1 ratio.

The rhMOG protein extracellular domain (rhMOG30-154) was expressed in H5 insect cells in the Protein Expression Laboratory, Leidos Biomedical Research, Frederick National Laboratory for Cancer Research, Frederick, MD. EAE induction was performed by mixing rhMOG (200 µg/mouse) in CFA (Difco Laboratories, DF3114-33-8) containing Mycobacterium tuberculosis H37Ra (Difco Laboratories, DF0639-60-6) and injecting s.c. into the abdomen bilaterally (50 µl emulsion in each side). Pertussis toxin (120 ng) (List Biological Laboratories, 180) diluted in PBS was administered i.p. on days 0 and 2 after immunization. Control mice receiving PBS in CFA were received injections with the same amount of pertussis toxin. The clinical severity of EAE was scored daily by an observer who was unaware of mouse genotypes, using a grading scale of 0–5 as previously described (28): 0, asymptomatic; 1, flaccid tail; 2, hind-limb weakness and impaired righting ability; 3, hind-limb paralysis; 4, front- and hind-limb paralysis; 5, moribund or death.

Serum was collected by cardiac puncture from B6 mice 3 wk after immunization with rhMOG (200 µg/mouse)/CFA or keyhole limpet hemocyanin (KLH; 200 µg/mouse)/CFA (LGC Biosearch Technologies, N-5060) as described above. Mice from the indicated groups received injections i.p. with 400 µl serum starting at the time of EAE induction and continuing every 2 d for the following weeks. IgG was purified from serum using Protein G Sepharose 4 Fast Flow resin (Cytiva, 17-0618-05) and Pierce IgG Elution Buffer (Thermo Fisher Scientific, 21009). Purified IgG was buffer exchanged to PBS and sterile filtered, and Ab integrity was confirmed by SDS-PAGE analysis. Purified IgG was injected in the same amount as contained in the 400 µl serum. This protein G purification removed >95% of IgG from serum as determined by ELISA, and the IgG-depleted serum was also used in transfer experiments.

Inguinal and axillary draining lymph nodes (DLNs) were harvested, mixed, and digested with 2.5 mg/ml collagenase D (Sigma-Aldrich, 11088882001) and 1 mg/ml DNase I (Sigma-Aldrich, 10104159001) for 30 min at 37°C. Tissues were filtered through 70-μm mesh strainers (BD Biosciences, 352350) to generate a single-cell suspension. For isolation of CNS-infiltrating cells, brain and spinal cord tissues were minced and digested with the same buffer as peripheral lymph nodes. Tissues were filtered through 70-μm mesh strainers and centrifuged through a Percoll (Sigma-Aldrich, P4937) density gradient (38% and 70%). Mononuclear cells in the interphase were removed, washed, and resuspended in culture medium for analysis by flow cytometry.

Cells were washed with FACS buffer (Corning, HBSS without calcium, magnesium, and Phenol Red) containing 0.2% BSA (Sigma-Aldrich, A3059) and 0.05% azide (Sigma-Aldrich, S2002), treated with anti-FcR (Leino, 24G2, C381), and then stained with specific Abs at 4°C for 30 min. We used antimouse CD4-FITC (BioLegend, RM4-4, 100510), CD4-BV785 (BioLegend, RM4-4, 100552), CD8α-PE-Cy7 (BioLegend, 53-6.7, 100722), B220-AF594 (BioLegend, RA3-6B2, 103254), CD11c-PE (BioLegend, N418, 117308), MHCII-BV421 (BioLegend, M5/114.15.2, 107632), CD40-allophycocyanin (BioLegend, 3/23, 124612), IFN-γ–allophycocyanin (BioLegend, XMG1.2, 505810), anti–IL-17A–BV785 (BioLegend, TC11-18H10.1, 506928), and GM-CSF–PE (BioLegend, MP1-22E9, 505406) Abs. For intracellular cytokine staining, mononuclear cells from lymph nodes and the CNS were stimulated overnight with rhMOG (10 µg/ml), with monensin (BioLegend, 420701) added in the last 4 h, and cells were fixed and permeabilized with the BD Fix/Perm kit (BD Biosciences, 554715) according to the manufacturer’s instructions and then stained with intracellular Abs for 30 min. Dead cells were excluded using the Zombie Aqua Fixable Viability Kit (BioLegend, 423102). Data were collected with an LSR II or LSRFortessa flow cytometer (BD Biosciences) and analyzed with FlowJo software.

MOG-specific IgG was measured by ELISA. rhMOG (10 µg/ml) was coated on ELISA plates (Thermo Fisher Scientific, Immulon 4HBX, 2855) overnight. The plates were then washed with ELISA wash buffer (0.5% Tween in PBS), serially diluted sera were applied to the plates, and plates were incubated for 2 h at room temperature. Antimouse IgG HRP (SouthernBiotech, 1030-05) was used to detect MOG-specific IgG. After a wash step, KPL ABTS substrate (Seracare, 5120-0041) was added to the wells, and the enzyme reaction was stopped by ABTS HRP Stop Solution (Seracare, 5150-0017). OD at 405 nm was measured with a SpectraMax iD3 plate reader.

Mice were perfused transcardially with PBS followed by 2% paraformaldehyde (Electron Microscopy Sciences) in PBS. The spinal column was then dissected and fixed in 2% paraformaldehyde overnight at 4°C. The columns were washed twice with PBS, and spinal cords were then carefully dissected and placed in 30% sucrose overnight at 4°C. Spinal cords were then frozen in tissue freezing media (VWR). Ten-micrometer sections were cut using a Leica CM1860 cryostat. Tissue sections were blocked with five drops of Background Buster (Innovex, NB306) per 1 ml of PBS with 0.5% Triton X-100 (Sigma, T8787) at room temperature for 1 h and then incubated with primary Ab in PBS containing 0.1% Triton X-100. Primary Abs included anti–CD4-BV421 (RM4-5 BioLegend, 100544; 1:500), anti–CD45-FITC (104; BioLegend, 109806; 1:500), anti–B220-PE (RA3-6B2; Thermo Fisher Scientific,12-0452-82; 1:500), anti-C3 (2/11; Hycult Biotech, HM1065-100UG; 1:50), and donkey antimouse IgG–Alexa Fluor 647 (Thermo Fisher Scientific, A-31571; 1:500). Tissue sections were then washed three times with PBS and incubated with secondary Ab in PBS containing 0.1% Triton X-100 for 2 h at room temperature if needed to detect the anti-C3 primary. Donkey antirat IgG–Alexa Fluor 594 (Thermo Fisher Scientific, A-21209; 1:500) was used as a secondary Ab. Last, sections were washed three times with PBS, and FluoroSave reagent (MilliporeSigma, 345789) was then added to each slide. Sections were coverslipped and imaged with an Olympus FV1200 confocal microscope. Images were analyzed using Imaris version 8.1. Using the Imaris surface function, a surface was drawn around the spinal cord section, avoiding folds and tears in the section. The area of the surface was then calculated. To quantify CD4+ and B220+ cells, the spots function was used to calculate the number of segmented CD4+ and B220+ spots within the spinal cord area. For each individual experiment, the number of spots per area was normalized to the highest value per group of mice collected, and significant outliers (two from KLH serum and one from EAE serum in Fig. 7D) were removed with the Grubbs test for outliers prior to combining the data. For C3 and IgG quantification, the surface function in Imaris was used to calculate the total segmented C3+ or IgG+ surface within the spinal cord area. Data were normalized and combined as described above.

GraphPad Prism 8 software was used. One-way ANOVA followed by the Dunnett test against the immunized CD40fl/fl group was performed for multiple comparisons. For statistical analysis of EAE, a nonparametric Mann-Whitney U test was performed for single comparison or one-way ANOVA followed by the Dunnett test against the immunized CD40fl/fl group was performed for multiple comparisons.

We studied the induction of EAE by immunization of C57BL/6 (B6) mice with rhMOG, previously shown to require both T and B cells (29, 30), paralleling the apparent roles of these cell types in human MS (5). This protocol induced severe disease peaking at 12–14 d in wild-type (WT) B6 mice. B cell–deficient (µMT) mice were completely resistant to rhMOG-induced disease (Fig. 1A), consistent with a previous report (30). CD40-knockout (CD40−/−) mice were similarly resistant to EAE induced by rhMOG (Fig. 1A), also consistent with previous studies reporting that CD40-knockout mice were resistant to MOG35-55 peptide–induced EAE (31).

FIGURE 1.

Maximal rhMOG-induced EAE requires CD40 expression on both B cells and DCs. (A) Clinical score of B6, μMT, or CD40-deficient (CD40−/−) mice immunized with rhMOG to induce EAE. (B) BM chimeras were established by injecting donor BM cells from CD40 WT (C57BL/6) or CD40-knockout (CD40−/−) mice into irradiated WT hosts. EAE was induced in the chimera mice by rhMOG immunization at 8 wk after establishment. (C) BM chimera were established by injecting donor BM cells from WT mice into irradiated WT or CD40−/− mouse hosts. EAE was induced in the chimera mice by rhMOG immunization at 8 wk after establishment. (D) Clinical scores of mice from CD19creCD40fl/fl, CD11ccreCD40fl/fl, CD19creCD11ccreCD40fl/fl, and no cre littermate control (CD40fl/fl) immunized with rhMOG to induce EAE. Data in A and D are combined from three independent experiments (mean ± SEM of n = 14 mice per group). Data in B and C are combined from two independent experiments (mean ± SEM of n = 10 mice per group). Medians of the total clinical score during days 11–27 were compared by two-tailed nonparametric Mann-Whitney U test. ****p < 0.0001.

FIGURE 1.

Maximal rhMOG-induced EAE requires CD40 expression on both B cells and DCs. (A) Clinical score of B6, μMT, or CD40-deficient (CD40−/−) mice immunized with rhMOG to induce EAE. (B) BM chimeras were established by injecting donor BM cells from CD40 WT (C57BL/6) or CD40-knockout (CD40−/−) mice into irradiated WT hosts. EAE was induced in the chimera mice by rhMOG immunization at 8 wk after establishment. (C) BM chimera were established by injecting donor BM cells from WT mice into irradiated WT or CD40−/− mouse hosts. EAE was induced in the chimera mice by rhMOG immunization at 8 wk after establishment. (D) Clinical scores of mice from CD19creCD40fl/fl, CD11ccreCD40fl/fl, CD19creCD11ccreCD40fl/fl, and no cre littermate control (CD40fl/fl) immunized with rhMOG to induce EAE. Data in A and D are combined from three independent experiments (mean ± SEM of n = 14 mice per group). Data in B and C are combined from two independent experiments (mean ± SEM of n = 10 mice per group). Medians of the total clinical score during days 11–27 were compared by two-tailed nonparametric Mann-Whitney U test. ****p < 0.0001.

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To elucidate the role of CD40 costimulation in EAE, we next analyzed the identity of CD40-expressing cell types required for EAE induction. Using BM chimeras, we found that WT hosts receiving CD40−/− BM donor cells were also completely resistant to rhMOG-induced EAE (Fig. 1B). However, CD40−/− hosts receiving WT BM donor cells developed EAE comparable to that in WT control hosts (Fig. 1C), indicating that CD40 expression on hematopoietically derived cells but not radioresistant cells, such as microglia, is necessary for rhMOG-induced EAE. To identify the cell types that mediate CD40 function in EAE induction, we selectively eliminated CD40 expression on either B cells or DC by crossing CD19-cre or CD11c-cre mice, respectively, to CD40fl/fl mice. The cell type specificity of conditional CD40 ablation was confirmed by flow cytometric analysis (Supplemental Fig. 1A). Strikingly, elimination of CD40 expression on either B cells or DCs resulted in significantly decreased EAE severity, indicating that CD40 expression on both B cells and DCs plays a role in EAE pathogenesis (Fig. 1D). Moreover, elimination of CD40 expression on both B cells and DCs by the combined effects of CD19-cre and CD11c-cre resulted in essentially complete elimination of EAE, indicating that CD40 expression on both B cells and DCs is required for EAE induced by rhMOG (Fig. 1D). Selective elimination of CD40 expression on B cells in vivo was also achieved by generating radiation BM chimeras in which BM from B cell–deficient µMT mice was mixed with BM from CD40−/− mice and transferred into lethally irradiated B6 recipients. These chimeric mice, with no CD40 expression on B cells but intact CD40 expression on all other cell types, were completely protected from rhMOG-induced EAE, confirming the requirement for CD40 on B cells (Supplemental Fig. 1B).

An early event following MOG immunization is the priming and differentiation of peripheral T cells, which are thought to subsequently play a role in mediating CNS events. We therefore analyzed immune cell populations in axillary and inguinal DLNs by flow cytometry 6 d after rhMOG immunization (Fig. 2A). Unimmunized naive CD40−/− mice and WT mice had equivalent numbers of lymph node CD4 and CD8 T cells, B cells, and DCs (Supplemental Fig. 2), consistent with a previous report (32). EAE induction by rhMOG immunization increased the total number of DLN cells, including T cells, B cells, and DCs in CD40fl/fl mice compared with unimmunized naive mice. In CD40−/− mice, CD4+ and CD8+ T cell numbers after immunization were significantly reduced, with approximately twofold fewer than those in immunized CD40fl/fl mice. Other immune cell populations, including B220+ B cells, and CD11c+ MHCII+ DCs were present in comparable numbers in immunized WT and CD40−/− mice (Fig. 2B). We next studied the DLN responses of mice with selective deletion of CD40 on B cells or DCs. Immunized mice with selective elimination of CD40 on B cells resembled immunized WT mice, whereas mice with deficient expression of CD40 on DCs resembled CD40−/− mice, with substantially lower numbers of CD4+ and CD8+ T cells compared with immunized non-cre control mice, demonstrating that CD40 expressed by DCs, but not CD40 expressed by B cells, is required for expansion of CD4 cell numbers in DLNs following MOG immunization (Fig. 2B).

FIGURE 2.

Immune cell populations in DLNs from the indicated mice 6 d after EAE induction or from naive CD40fl/fl mice. (A) Gating strategy by FACS. (B) Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons according to FACS analysis in A. *p < 0.05, ****p < 0.0001.

FIGURE 2.

Immune cell populations in DLNs from the indicated mice 6 d after EAE induction or from naive CD40fl/fl mice. (A) Gating strategy by FACS. (B) Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons according to FACS analysis in A. *p < 0.05, ****p < 0.0001.

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IFN-γ–producing Th1 cells and IL-17–producing Th17 cells have been shown to be pathogenic CD4+ T cells that can trigger EAE (9). In addition, several groups reported that production of the cytokine GM-CSF by CD4+ T cells is essential for their encephalitogenicity (6, 12, 33, 34). We therefore isolated DLNs from the indicated mice 6 d after EAE induction or control PBS treatment, and we stimulated cells in vitro with rhMOG overnight. IFN-γ–, IL-17–, and GM-CSF–producing Th cells were analyzed by intracellular staining. These cytokines were undetectable or detected at very low frequency in CD4+ T cells from control PBS/CFA-immunized mice (Fig. 3A). IL-17–, IFN-γ–, and GM-CSF–producing CD4+ T cells in DLNs of rhMOG-immunized WT mice were present at frequencies substantially above those in PBS/CFA-immunized WT mice, as were the numbers of what have been reported to be highly pathogenetic IL-17+ IFN-γ+ CD4 T cells (11) (Fig. 3A). The frequencies of these inflammatory cytokine-producing CD4+ T cells in DLNs were markedly reduced in CD40−/− mice (Fig. 3B), showing that induction and differentiation of these Th cell populations is dependent on CD40 expression. Notably, the selective deletion of CD40 on DCs resulted in a profound decrease in cytokine-producing Th cells equivalent to that observed in complete CD40 knockouts, paralleling the failure of these mice to manifest clinical EAE. In contrast, selective deletion of CD40 on B cells, which was equally protective against clinical EAE, had no effect on the induction of these cytokine-producing Th cells (Fig. 3). Thus, CD40 expression by DCs, but not B cells, is important in peripheral priming and differentiation of CD4+ Th cells that may play a role in EAE pathogenesis.

FIGURE 3.

Cytokine-producing CD4+ T cells in DLNs from indicated mice 6 d after EAE induction or control PBS induction. (A) CD4+ T cells were analyzed for cytokine production by intracellular staining after overnight culture with rhMOG (10 µg/ml), with monensin added in the last 4 h. (B) Statistical results showing frequency according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. **p < 0.01, ***p < 0.001, ****p < 0.0001.

FIGURE 3.

Cytokine-producing CD4+ T cells in DLNs from indicated mice 6 d after EAE induction or control PBS induction. (A) CD4+ T cells were analyzed for cytokine production by intracellular staining after overnight culture with rhMOG (10 µg/ml), with monensin added in the last 4 h. (B) Statistical results showing frequency according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. **p < 0.01, ***p < 0.001, ****p < 0.0001.

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We also studied the presence of immune cell populations in the CNS (Fig. 4A). At the peak of their disease, the total number of CD4+ T cells in the CNS of rhMOG-immunized WT mice was increased compared with unimmunized mice. MOG-immunized CD40−/− mice, which did not develop disease, correspondingly had very few CD4+ T cells recoverable from the brain and spinal cord (Fig. 4B). Mice selectively deficient in CD40 on B cells or DCs, which in both cases were highly protected from EAE, also had significantly reduced CD4+ T cells in the CNS when compared with WT mice, but with numbers that were greater than those in CD40−/− mice (Fig. 4B). Frequencies of IL-17–, IFN-γ–, and GM-CSF–producing CD4+ T cells isolated from the CNS were measured in each strain. Substantial proportions of CD4+ T cells in CD40fl/fl mice were IL-17+, IFN- γ+, IL-17+IFN-γ+, and GM-CSF+ (Fig. 5A, 5B), whereas the small numbers of CD4+ T cells isolated from the CNS of CD40−/− mice had significantly lower frequencies of cytokine-positive populations. Conditional deletion of CD40 from B cells or DCs had similar effects in decreasing the total number of CD4+ T cells (Fig. 4B). However, CD40 deletion from DCs, but not B cells, resulted in substantially reduced percentages of these cytokine-producing cells (Fig. 5A, 5B), paralleling the effects in DLNs earlier after immunization (Fig. 3A, 3B). Thus, CD40 expression on DCs but not on B cells is important for priming and for the selective presence of pathogenic cytokine-producing CD4+ T cells, suggesting that the requirement for CD40 on B cells may be mediated through a distinct mechanism in EAE pathogenesis.

FIGURE 4.

Immune cell populations in the CNS from indicated mice 14 d after EAE induction or from naive CD40fl/fl mice. (A) Gating strategy by FACS. (B) Statistical analysis of results according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. **p < 0.01, ***p < 0.001, ****p < 0.0001.

FIGURE 4.

Immune cell populations in the CNS from indicated mice 14 d after EAE induction or from naive CD40fl/fl mice. (A) Gating strategy by FACS. (B) Statistical analysis of results according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. **p < 0.01, ***p < 0.001, ****p < 0.0001.

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FIGURE 5.

Cytokine-producing CD4+ T cells in the CNS from indicated mice 14 d after EAE induction. (A) CD4+ T cells were analyzed for cytokine production by intracellular staining after overnight culture with rhMOG (10 µg/ml), with monensin added in the last 4 h. (B) Statistical results showing frequency according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. ****p < 0.0001.

FIGURE 5.

Cytokine-producing CD4+ T cells in the CNS from indicated mice 14 d after EAE induction. (A) CD4+ T cells were analyzed for cytokine production by intracellular staining after overnight culture with rhMOG (10 µg/ml), with monensin added in the last 4 h. (B) Statistical results showing frequency according to FACS analysis in A. Data are combined from two independent experiments (mean ± SD of n = 6 per group). One-way ANOVA followed by Dunnett test against immunized CD40fl/fl group was performed for multiple comparisons. ****p < 0.0001.

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We previously identified a requirement for CD40 expression on B cells, but not on DCs, in high-affinity class-switched Ab responses to model Ags (35). We therefore considered that a similar function of CD40 on B cells in generation of pathogenic Ab might contribute to EAE pathology. We found that immunization of B6 mice with rhMOG resulted in high titers of anti-MOG IgG Abs as detected by ELISA and that this response was absent in CD40-knockout and B cell–deficient µMT mice (Supplemental Fig. 3A). We further observed that elimination of CD40 expression on B cells resulted in essentially complete abrogation of anti-MOG IgG production. Although elimination of CD40 expression on DCs resulted in a reduction in clinical EAE similar to that resulting from CD40 deletion on B cells, these mice mounted anti-MOG Ab responses equivalent to WT (Supplemental Fig. 3B). To determine whether the CD40 dependence of the IgG anti-MOG response was reflected in an effect on germinal center (GC) B cell response, we also assessed the induction of GC B cells in DLNs following MOG immunization. MOG immunization increased the number of GC B cells in CD40fl/fl mice compared with PBS/CFA-immunized or naive CD40fl/fl mice. In parallel with what was observed for serum anti-MOG IgG levels, CD40 expression on B cells but not DCs was required for GL7+ CD38 GC B responses in the DLNs (Supplemental Fig. 3C, 3D).

We next tested the possibility that the pathogenic role of CD40 expression on B cells was mediated by its function in Ab production. Serum from MOG-immunized or control KLH-immunized B6 mice was transferred to mice with conditional deletion of CD40 from either B cells or DCs, and these mice were immunized with rhMOG. Transfer of MOG-immune serum but not KLH-immune serum fully reconstituted the susceptibility of B cell conditional CD40-knockout mice to induction of clinical EAE (Fig. 6A). In contrast, MOG-immune serum did not reconstitute the susceptibility of mice that are deficient in CD40 expression on DCs or of CD40−/− mice (Fig. 6B). To test whether the effect of immune serum was Ab mediated, we purified the IgG from the same MOG-immune serum and injected purified IgG into B cell CD40-deficient mice. Like MOG-immune serum, purified IgG reconstituted the EAE susceptibility of B cell CD40-deficient mice, whereas mice with the same genotype receiving control IgG from KLH-immunized mice or mice receiving IgG-depleted MOG-immune serum still developed reduced EAE (Fig. 6C). These results indicate that CD40 on B cells plays a role in EAE pathogenesis through mediation of Ab response to MOG, whereas, in contrast, the role of CD40 on DCs is independent of Ab response.

FIGURE 6.

Pathogenic anti-MOG Ab reconstituted EAE susceptibility of B cell CD40-deficient mice. (A and B) Serum was collected from B6 mice 3 wk after EAE induction and injected into the indicated mice at days 0, 3, 7, and 10 after EAE induction (400 µl serum per injection). (C) IgG-depleted EAE serum or IgG purified from EAE (EAE IgG) or KLH/CFA-immunized mice (ctrl IgG) was injected in the same amount as contained in the 400 µl serum (1 mg). Medians of the total clinical score during days 9–21 were compared by two-tailed nonparametric Mann-Whitney U test. Data are combined from three independent experiments (mean ± SEM of n = 12 mice per group). ****p < 0.0001.

FIGURE 6.

Pathogenic anti-MOG Ab reconstituted EAE susceptibility of B cell CD40-deficient mice. (A and B) Serum was collected from B6 mice 3 wk after EAE induction and injected into the indicated mice at days 0, 3, 7, and 10 after EAE induction (400 µl serum per injection). (C) IgG-depleted EAE serum or IgG purified from EAE (EAE IgG) or KLH/CFA-immunized mice (ctrl IgG) was injected in the same amount as contained in the 400 µl serum (1 mg). Medians of the total clinical score during days 9–21 were compared by two-tailed nonparametric Mann-Whitney U test. Data are combined from three independent experiments (mean ± SEM of n = 12 mice per group). ****p < 0.0001.

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Last, we examined the role of CD40-mediated Ab production from B cells in driving spinal cord inflammation during EAE. In symptomatic EAE control mice, the spinal cord parenchyma was infiltrated by both CD4+ T cells and B cells that sometimes aggregated in clusters (Fig. 7A, 7B). There were also massive deposits of IgG and complement in the spinal cords of these mice that would explain their observed symptoms (Fig. 7C). Importantly, removal of CD40 from B cells markedly reduced infiltration of the spinal cord parenchyma by CD4+ T cells and B cells (Fig. 7A, 7B). The few cells that were found in the spinal cords of these mice localized primarily to the meningeal surface. There was also very little IgG or C3 in the parenchyma following removal of CD40 from B cells, despite administration of serum from KLH-immunized mice (Fig. 7C). By contrast, administration of EAE serum to CD40-deficient mice reconstituted the heavy deposits of IgG and C3 observed in control EAE mice and facilitated infiltration of the spinal cord parenchyma by CD4+ T cells and B cells (Fig. 7A7D). These data demonstrate that Ab and C deposition in the spinal cord parenchyma is an important part of the pathological mechanism induced by CD40 expression on B cells.

FIGURE 7.

Spinal cord inflammation and Ig/complement distribution in serum-treated B cell CD40-deficient mice. (A) Representative confocal images of spinal cord sections stained with anti-CD4 (green) and anti-B220 (red) in the denoted groups of mice. (B) Magnified confocal images from the spinal cords in A showing immune cell clustering in the denoted groups. CD4 (green), B220 (red), and CD45 (blue). (C) Representative confocal images of spinal cord sections stained with anti-C3 (green) and anti-IgG (red). (D) Quantification of spinal cord CD4, B220, C3, and IgG staining in CD19creCD40fl/fl mice that received EAE serum and serum from KLH-immunized mice. Data are combined from three independent experiments (mean ± SEM, n = 9 for EAE serum group, n = 11 for KLH serum group) and analyzed using an unpaired t test. *p < 0.05, ***p < 0.001, ****p < 0.0001.

FIGURE 7.

Spinal cord inflammation and Ig/complement distribution in serum-treated B cell CD40-deficient mice. (A) Representative confocal images of spinal cord sections stained with anti-CD4 (green) and anti-B220 (red) in the denoted groups of mice. (B) Magnified confocal images from the spinal cords in A showing immune cell clustering in the denoted groups. CD4 (green), B220 (red), and CD45 (blue). (C) Representative confocal images of spinal cord sections stained with anti-C3 (green) and anti-IgG (red). (D) Quantification of spinal cord CD4, B220, C3, and IgG staining in CD19creCD40fl/fl mice that received EAE serum and serum from KLH-immunized mice. Data are combined from three independent experiments (mean ± SEM, n = 9 for EAE serum group, n = 11 for KLH serum group) and analyzed using an unpaired t test. *p < 0.05, ***p < 0.001, ****p < 0.0001.

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CD40 costimulation was shown to play an important role in the development of autoimmune diseases, both in mice and in humans (36). However, how CD40 endows different disease-relevant immune cell types with their ability to elicit effector programs that ultimately cause a CNS autoimmune disease such as EAE or MS is not fully understood. In this study, we assessed the role of CD40 on specific cell types using a conditional knockout approach to delete CD40 from two cell types, B cells and DCs, required for development of a CNS autoimmune disease initiated by myelin protein immunization. Importantly, EAE disease severity was profoundly reduced by deletion of CD40 from DCs or B cells, demonstrating that both cell types use this costimulatory protein to induce disease. The programs initiated by CD40 were, however, distinct between DCs and B cells. CD40 specifically equipped DCs with the ability to prime pathogenic, polyfunctional (IFN-γ+, IL-17+, and GM-CSF+) Th cells and promote their migration into the CNS. By contrast, B cell expression of CD40 was required for production of autoreactive, MOG-specific Abs, which were able to reconstitute disease when passively transferred into mice with CD40-deficient B cells but not DCs. In fact, these Abs in combination with complement were found to completely inundate the spinal cord parenchyma during the peak of disease, providing an explanation for the development of symptoms. These data support a model whereby CD40 facilitates the development of distinct effector programs via B cells and DCs that converge and result in a CNS autoimmune disease. These findings identify targets for intervention to ameliorate the disease process.

A pathogenic role of Th1 and Th17 cells has been well established in mediating multiple inflammatory autoimmune diseases, including EAE (10). Adoptive transfer of in vitro cultured Th1 or Th17 cells induces EAE in recipient mice (9, 37). Th17 cells can also produce IFN-γ mediated by T-bet and Runx1 or Runx3, and this developmental flexibility has been linked to the pathogenicity of Th17 cells in multiple autoimmune diseases, including EAE (11). In addition to the signature proinflammatory cytokines of the Th1 and Th17 cells, both populations produce GM-CSF under polarizing conditions, and GM-CSF is required for EAE pathogenesis, as evidenced by a reduction in disease severity in mice treated with anti–GM-CSF Abs (12). Although previous studies analyzed the in vitro requirements for induction of these pathogenic Th1-, Th17-, and GM-CSF–producing Th cells (38), the in vivo requirements for their induction were not elucidated. Here, we found that CD40 expression on DCs, but not on B cells, is required for optimal priming of pathogenic Th cells in peripheral DLNs during EAE and for the appearance of these cells in the CNS. IFN-γ–, IL-17–, and GM-CSF–positive Th cells were significantly reduced in DC CD40 conditional knockout mice to a degree comparable to that observed in CD40-knockout mice. These data demonstrate an important role of CD40 on DCs in priming pathogenic Th cells, which is consistent with previous studies showing that CD40−/− DCs are impaired in their ability to promote the development of IFN-γ–producing Th1 (35) and IL-17–producing Th17 cells (39). Although CD11c-cre mice are widely used as a DC-specific targeting strategy, CD11c-cre may also affect macrophages or other myeloid-derived cells. Activated macrophages share many features with DCs, including CD11c and CD40 expression (40), and it has been reported that myeloid-specific deletion of CD40 by LysM-cre also resulted in a significant reduction in EAE severity and reduced CNS inflammation (41, 42).

In parallel with impaired peripheral priming in response to MOG, CD4+ T cells isolated from the CNS of DC CD40 conditional knockout mice expressed significantly lower levels of inflammatory cytokines that are likely important for inflaming the CNS and promoting the recruitment of additional immune cells. In support of this conclusion, we detected fewer CNS mononuclear cells from DC CD40 conditional knockout mice and milder disease. Anti–IL-17A Ab (secukinumab) was shown to reduce lesions detected by magnetic resonance imaging in patients with MS (7), and anti–GM-CSF Ab (MOR103) is being studied in a phase I clinical trial of patients with MS (43). Our results demonstrate that deletion of CD40 signaling on DCs significantly impairs the priming of IL-17– and GM-CSF–producing Th cells in vivo and may therefore provide a promising therapeutic target in patients with MS. Interestingly, treatment of mice with anti-CD154 at the time of PLP139-151/CFA immunization has been reported to inhibit clinical disease for up to 100 d after immunization (44). Moreover, although B cell depletion in mice treated with anti-CD20 Ab was shown to slightly reduce the Th1 and Th17 cell response to MOG protein immunization in vivo (45), our findings indicate that this reduction in Th cells was not regulated by CD40 on B cells. B cell CD40 conditional knockout mice developed IFN-γ–, IL-17–, and GM-CSF–producing Th cells comparable to WT mice in the periphery, but they showed much milder EAE symptoms, suggesting that CD40 on B cells has a role other than the priming of pathogenic Th cells.

In addition to immune cell infiltrates, Abs and complement components are also found in inflammatory active lesions of patients with MS (46), and the complement system has been shown to contribute to the pathology of EAE by triggering demyelination and modifying the Ag-specific T and B cell responses (4749). Previous work from our laboratory has explored the cellular requirements for CD40-CD40L and B7-CD28 expression for GC and Ab responses to model Ags. These studies demonstrated that CD40 expression on B cells is essential for the humoral immune response to T-dependent Ag in a cell-intrinsic manner, whereas there is no requirement for CD40 expression by DCs for these responses (35, 50). Anti-MOG serum or Ab has been reported to have a pathogenic role in the development of EAE in mice (17, 51, 52), and mAb against human CD40 has been reported to prevent EAE in the common marmoset (53). We found that deletion of CD40 from B cells but not DCs eliminated production of anti-MOG IgG. These data are consistent with studies in cytidine deaminase–deficient mice, which are defective in Ig somatic hypermutations and class switching, showing decreased susceptibility to rhMOG-induced EAE, without a defect in MOG-specific T cells (52, 54). Importantly, we were able to fully reconstitute disease in B cell (but not DC) CD40 conditional knockout mice by passively transferring immune serum containing high titers of anti-MOG IgG Abs. These Abs together with complement, B cells, and CD4+ T cells were found throughout the spinal cord parenchyma of mice reconstituted with EAE serum but not control KLH serum. These data demonstrate that the major role played by CD40 on B cells is to induce a pathogenic autoantibody response. These Abs even appear to facilitate recruitment of B cells and CD4+ T cells to the spinal cord parenchyma, because their numbers were greatly reduced in mice treated with KLH serum. Anti-CD20 mAbs that target B cells were shown to be effective in treating patients with MS (5). We propose, on the basis of our data, that the therapeutic benefit of this treatment is mediated in part by inhibition of autoantibody production. In fact, Ab-producing plasma cells and complement were recently found at the leading edge of chronic active MS lesions, further implicating humoral immunity as a driver of lesion development (55). Our studies do not exclude additional roles for B cells in the development of CNS autoimmune diseases (56), but they do highlight the potential effectiveness of inhibiting or modulating the autoreactive humoral response as a therapeutic option for at least some patients with MS.

Collectively, our findings support a model in which CD40 induces complementary effector programs that contribute to EAE pathogenesis. Efficient priming of pathogenic proinflammatory Th cells and their eventual CNS infiltration requires CD40 expression on DCs, but not B cells. This infiltration appears to be necessary but not sufficient for EAE development. Our finding that passive transfer of EAE serum or IgG fully restores disease in B cell CD40 conditional knockout mice is an important one, especially in light of how these Abs inundate the spinal cord parenchyma, fostering complement deposition and adaptive immune cell infiltration. CD40 maximizes the development of CNS autoimmune disease via induction of complementary programs, but autoantibodies are nevertheless a major driving force in disease pathogenesis. Targeting Ab-producing plasma cells may therefore be a key to stopping diseases such as MS once they are underway. Our CD40 data also demonstrate that there are multiple points of potential therapeutic intervention during the development of CNS autoimmune diseases that involve distinct arms of the adaptive immune system.

We thank Vanja Lazarevic, Masashi Watanabe, and Karen Hathcock for their thoughtful comments and review of the manuscript. We thank Peter J. Koch for ES cell injections; Christopher McNees for purified IgG quality control; and Bernardo Rosa, Elena Kuznetsova, and Tarra Dumas from the NCI animal facility for assistance with caring for and scoring experimental mice.

This work was supported by the intramural research programs of the National Cancer Institute and the National Institute of Neurological Disorders and Stroke, National Institutes of Health, and support to D.W. provided by National Institutes of Health Grant R01DK07501-03S2.

fl/fl

Y.L., M.X., C.E.D., R.Z., and C.T.M. conducted experiments; Y.L. and C.E.D. analyzed the data; D.W. provided the founder CD40 mice; Y.L., D.B.M., and R.J.H. interpreted the data and supervised the research; Y.L., D.B.M., and R.J.H. wrote and edited the manuscript.

Richard J. Hodes is a Distinguished Fellow of AAI.

The online version of this article contains supplemental material.

Abbreviations used in this article:

     
  • BM

    bone marrow

  •  
  • DC

    dendritic cell

  •  
  • DLN

    draining lymph node

  •  
  • EAE

    experimental autoimmune encephalomyelitis

  •  
  • GC

    germinal center

  •  
  • KLH

    keyhole limpet hemocyanin

  •  
  • MOG

    myelin oligodendrocyte glycoprotein

  •  
  • MS

    multiple sclerosis

  •  
  • WT

    wild type

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The authors have no financial conflicts of interest.

Supplementary data