Histamine H₂ receptor signaling × environment interactions determine susceptibility to experimental allergic encephalomyelitis

Abstract

Histamine and its receptors are important in both multiple sclerosis and experimental allergic encephalomyelitis (EAE). C57BL/6J (B6) mice deficient for the histamine H₂ receptor (H₂RKO) are less susceptible to EAE and exhibit blunted Th1 responses. However, whether decreased antigen-specific T-cell effector responses in H₂RKO mice were due to a lack of H₂R signaling in CD4⁺ T cells or antigen-presenting cells has remained unclear. We generated transgenic mice expressing H₂R specifically in T cells on the H₂RKO background, and, using wild-type B6 and H₂RKO mice as controls, induced EAE either in the presence or absence of the ancillary adjuvant pertussis toxin (PTX), which models the effects of infectious inflammatory stimuli on autoimmune disease. We monitored the mice for clinical signs of EAE and neuropathology, as well as effector T-cell responses using flow cytometry. EAE severity and neuropathology in H₂RKO mice expressing H₂R exclusively in T cells become equal to those in wild-type B6 mice only when PTX is used to elicit disease. EAE complementation was associated with frequencies of CD4⁺IFN-γ⁺ and CD4⁺IL-17⁺ cells that are equal to or greater than those in wild-type B6, respectively. Thus, the regulation of encephalitogenic T-cell responses and EAE susceptibility by H₂R signaling in CD4⁺ T cells is dependent on gene × environment interactions.—Saligrama, N., Case, L. K., Krementsov, D. N., Teuscher, C. Histamine H₂ receptor signaling × environment interactions determine susceptibility to experimental allergic encephalomyelitis.

Histamine (2-[4-imidazole]-ethylamine) is a ubiquitously distributed biogenic monoamine that is synthesized, stored, and secreted mainly by mast cells and basophils (1). Histamine mediates diverse physiological processes, including brain function, secretion of pituitary hormones, neurotransmission, cell proliferation and differentiation, hematopoiesis, embryonic development, wound healing and regeneration, and the regulation of gastrointestinal, cardiovascular, and secretory functions (2–6). In addition, histamine is well known for its regulation of innate and adaptive immune responses (7–9). Histamine exerts its effects through binding to 4 different 7-transmembrane G-protein-coupled receptors (GPCRs): histamine receptors (HRs) H₁R, H₂R, H₃R, and H₄R, named according to the chronological order of their discovery (10).

H₂R couples to second messenger signaling pathways via the stimulatory G protein, Gα_s (10), leading primarily to activation of adenylate cyclase and increased cAMP (11–18). In addition, the activation of H₂R leads to phospholipid methylation (19), an increase in the slow inward Ca²⁺ current (20), stimulation of phospholipase C, intracellular Ca²⁺ mobilization (21, 22), and inhibition of phospholipase A₂ activation (23). Furthermore, histamine binding to H₂R leads to the activation of c-Fos (8, 24, 25), c-Jun (20), protein kinase C, and p70S6kinase (26). Although it is not known how histamine acting through the H₂R activates multiple second messenger signaling pathways, this is commonly observed among Gα_s-coupled receptors (27).

Because of its ubiquitous expression and the capacity to activate multiple signaling pathways, H₂R regulates diverse cellular functions, including innate and adaptive immune responses (1). Immature and mature dendritic cells (DCs) express H₂R (28), and H₂R signaling in DCs affects their maturation, cytokine production, and capacity to influence T-helper (Th) cell polarization (29, 30). In addition, histamine acting through H₂R is known to regulate Th1 and Th2 effector functions. Compared with Th1 cells, Th2 cells express greater levels of H₂R, and Th1 and Th2 responses are negatively regulated by H₂R activation (31). Splenocytes from H₂R-deficient (H₂RKO) mice pretreated with histamine and stimulated with anti-CD3 exhibit enhanced production of interferon-γ (IFN-γ), interleukin (IL)-4, and IL-13 (9). Th2 cells pretreated with histamine and stimulated with anti-CD3 produce high levels of IL-10, which can be inhibited by H₂R antagonist (32). In addition, H₂R stimulation of Th2 cells can enhance IL-13 production (33). Furthermore, H₂R can augment the suppressive activity of transforming growth factor-β on T cells (34), indicative of its role in tolerance and in inhibiting autoimmune and/or inflammatory responses. Of particularly interest with respect to multiple sclerosis (MS), H₂R signaling is required for ultraviolet B irradiation (280–320 nm)-induced systemic suppression of antigen-specific T-cell responses (35, 36).

Studies on MS suggest that the histaminergic system plays an important role in the pathophysiology of the disease. Histamine levels in the cerebrospinal fluid (CSF) of patients with both remitting and progressive MS were found to be 60% higher than those in the CSF of control subjects or those with related neurological diseases (37). Several reports support a role for H₂R in MS and experimental allergic encephalomyelitis (EAE), the principal autoimmune model of MS (38). Th1 and/or Th17 cells secreting IFN-γ and IL-17, respectively, are necessary and sufficient to induce neuropathology (39). Treatment of lymphocytes from patients with MS with cimetidine, a H₂R-specific antagonist, leads to increased antibody-dependent cell-mediated cytotoxic killing of oligodendrocytes (40), suggesting a protective effect of endogenous histamine signaling specifically through H₂R. Furthermore, the administration of dimaprit, a H₂R selective agonist, significantly reduced myelin oligodendrocyte glycoprotein 35–55 (MOG_35–55)-induced EAE in C57BL/6J mice, suggesting an antipathogenic role for H₂R in EAE (41). In SJL/J mice with EAE, mononuclear cells within brain lesions express high levels of H₂R protein, and Th1 cells reactive to proteolipid protein expressed less H₂R than Th2 cells (42, 43). Moreover, studies of pial vessels, the location where autoimmune lesions are initiated by encephalitogenic T cells in EAE (44), revealed increased permeability mediated by H₂R and elevation of Ca²⁺ (45). Taken together, these data signify a need for a better understanding of the effects of H₂R in the pathogenesis of MS and EAE.

We previously reported that H₂RKO mice were less susceptible to MOG_35–55-induced EAE than wild-type (WT) mice (46). In ex vivo recall assays, splenocytes from H₂RKO mice produced significantly less IFN-γ than those from WT mice with no detectable difference in the production of IL-4. Furthermore, analysis of cytokine and chemokine production by peritoneal exudate macrophages from H₂RKO mice revealed a significant reduction in the secretion of IL-12 and IL-6, and an increase in monocyte chemoattractant protein 1 (MCP-1). These results suggested that the blunted Th1 response and reduced EAE severity seen in H₂RKO mice was due to dysregulated antigen-presenting cell (APC) functions, including cytokine production (46). However, in vitro studies have also suggested a role for T-cell intrinsic H₂R signaling in regulating T-cell effector responses (31). To date, no study has examined the affect of H₂R signaling directly in T cells on EAE pathogenesis. Therefore, we generated transgenic mice expressing H₂R specifically in T cells on the H₂RKO background to test the hypothesis that H₂R signaling in T cells directly contributes to EAE susceptibility and T-cell effector responses. Reexpression of H₂R exclusively in T cells complemented EAE severity and neuropathology of H₂RKO mice to that of WT mice only when pertussis toxin (PTX) is used as an ancillary adjuvant, in association with an increase in the frequency of CD4⁺IFN-γ⁺ and CD4⁺IL-17⁺ T cells. Because PTX is used to model environmental factors that influence disease susceptibility (47, 48), our findings support a role for T-cell intrinsic H₂R signaling × environment interactions acting on APCs in the pathogenesis of autoimmune inflammatory disease of the central nervous system (CNS).

MATERIALS AND METHODS

Animals

C57BL/6J mice (referred to as either B6 or WT) were purchased from The Jackson Laboratory (Bar Harbor, ME, USA). B6.129P-Hrh2^tm1Wat (H₂RKO; ref. 49) mice were maintained at the University of Vermont (Burlington, VT, USA) where they were backcrossed to B6 mice for >10 generations. The genetic background of H₂RKO mice was assessed by the DartMouse Speed Congenic Core Facility at the Geisel School of Medicine at Dartmouth (Lebanon, NH, USA). DartMouse uses the Illumina GoldenGate Genotyping Assay (Illumina, San Diego, CA, USA) to interrogate 1449 single-nucleotide polymorphisms (SNPs) spread throughout the genome. The raw SNP data were analyzed using DartMouse's SNaP-Map and Map-Synth software, allowing for the determination of the SNP allele at each location. The SNP analysis revealed that the H₂RKO mice are 98% B6, with minor carryover (2%) from 129 only at the retained Hrh2 locus.

For transgenic mouse generation, a hemagglutinin (HA) tag was added to Hrh2 by deleting the methionine and adding the HA tag to the N terminus using the TOPO cloning vector (Invitrogen, Carlsbad, CA, USA). The construct was then subcloned downstream of the dlck promoter. The linear DNA fragment containing the dlck promoter, the HA-Hrh2 construct, and the human growth hormone (hGH) intron and polyadenylation signal was injected directly into fertilized B6 eggs at the University of Vermont transgenic/knockout facility. Mice were screened for the hGH gene by PCR using hGH forward (5′-TAGGAAGAAGCCTATATCCCAAAGG-3′) and hGH reverse (5′-ACAGTCTCTCAAAGTCAGTGGGG-3′) primers. Proinsulin forward (5′-CTAGTTGCAGTAGTTCTCCAG-3′) and proinsulin reverse (5′-CCTGCCTATCTTTCAGGTC-3′) primers were used as internal controls. Two founders were generated and crossed to H₂RKO mice to establish H₂RKO-dlck^Hrh2 transgenic mice (H₂RKO-Tg1 and H₂RKO-Tg2). The experimental procedures used in this study were approved by the Animal Care and Use Committee of the University of Vermont.

Real-time quantitative RT-PCR (qRT-PCR)

CD4⁺ T cells were isolated from spleen and lymph nodes by negative selection using an EasySep mouse CD4⁺ T-cell enrichment kit (Stemcell Technologies, Vancouver, BC, Canada). Total RNA was extracted from CD4⁺ T cells using RNeasy RNA isolation reagent (Qiagen, Valencia, CA, USA). cDNA generated from 1 μg of total RNA was used in qRT-PCR using the SYBR Green method. The sequences of Hrh2 primers used were as follows: forward, 5′-TGGCACGGTTCATTCC-3′, and reverse, 5′-GCAGTAGCGGTCCAAG-3′. β2m was used as the reference gene, and relative mRNA levels were calculated using the comparative threshold cycle (C_T) method.

Ca²⁺ imaging in HEK293T cells

HEK293T cells were transfected with pcDNA3.1 containing the HA-Hrh2 construct or the empty vector, and at 24 h post-transfection, the cells were loaded with Fluo-4 (10 μM) for 7 min at 35°C in the presence of pluronic acid (2.5 μg/ml). Ca²⁺ was imaged using a Revolution confocal system with an electron-multiplying charge-coupled device camera (Andor Technology, Belfast, UK) mounted on an upright microscope with a ×60 water-immersion objective (1.0 numerical aperture; Nikon, Tokyo. Japan). After addition of 100 μM histamine, images were acquired at 15 frames/s with Revolution TL acquisition software (Andor Technology). Fluo-4 fluorescence was excited by a krypton-argon laser (488 nm), and emitted fluorescence was collected at 495 nm. The images were processed using custom-designed software (supplied by A. Bonev, University of Vermont), and the fractional fluorescence was evaluated by dividing the fluorescence of a region of interest (ROI; 5×5 pixels/box) by the average fluorescence of 10 images from the same ROI.

Induction and evaluation of EAE

Mice were immunized for the induction of EAE using either a 1× or 2× immunization protocol. For the 2× protocol, mice were injected subcutaneously in the posterior right and left flank with an emulsion containing 100 μg of MOG_35–55 and an equal volume of complete Freund's adjuvant (CFA; Sigma-Aldrich, St. Louis, MO, USA) supplemented with 200 μg of Mycobacterium tuberculosis H37Ra (Difco Laboratories, Detroit, MI, USA). Then 1 wk later, all mice received an identical injection of MOG_35–55 plus CFA. For the 1× protocol, mice were immunized with an emulsion containing 200 μg of MOG_35–55 and an equal volume of CFA containing 200 μg of M. tuberculosis H37Ra. On the day of immunization, each mouse received 200 ng of PTX (List Biological Laboratories, Campbell, CA, USA) by intervenous injection.

Mice were scored daily for clinical signs of EAE beginning on d 5 after injection as follows: 0, no clinical expression of disease; 1, flaccid tail without hind-limb weakness; 2, hind limb weakness; 3, complete hind-limb paralysis and floppy tail; 4, hind-limb paralysis accompanied by a floppy tail and urinary or fecal incontinence; and 5, moribund. Clinical quantitative trait variables were assessed as described previously (50). Histopathological evaluations were performed as described previously (51).

Cytokine and proliferation assays

For ex vivo cytokine assays, mice were immunized using the 1× and 2× immunization protocols, spleens and draining lymph nodes (DLNs) were harvested on d 10, and single-cell suspensions were prepared (1×10⁶ cells/ml) in RPMI 1640 (5% fetal bovine serum) and restimulated with 50 μg/ml of MOG_35–55. Cell culture supernatants were recovered after 72 h and assayed for IFN-γ, IL-6, granulocyte macrophage-colony-stimulating factor (GM-CSF), and IL-17 by enzyme-linked immunosorbent assays (ELISAs) using anti-IFN-γ, anti-IL-6, anti-GM-CSF, and anti-IL-17 monoclonal antibodies (mAbs) and their respective biotinylated mAbs (BD Biosciences-Pharmingen, San Jose, CA, USA).

For proliferation assays, 5 × 10⁵ cells/well in RPMI 1640 were plated on standard 96-well U-bottom tissue culture plates and stimulated with 0, 1, 2, 10, and 50 μg of MOG_35–55 for 72 h at 37°C. During the last 18 h of culture, 1 μCi of [³H]thymidine (PerkinElmer, Waltham, MA, USA) was added. Cells were harvested onto glass fiber filters, and thymidine uptake was determined with a liquid scintillation counter.

For intracellular cytokine staining, spleen and DLNs were stimulated for 4 h with phorbol 12-myristate 13-acetate (PMA) and ionomycin in the presence of brefeldin A (BD Biosciences). Cells were labeled with Live/Dead UV-Blue dye (Invitrogen) followed by surface staining (CD45 from Invitrogen; CD4, CD8, TCRγδ, CD11b, CD11c, CD25, and TCRβ from BD Biosciences). Afterward, cells were fixed, permeabilized, and stained for intracellular IL-17A, IFN-γ (BioLegend, San Diego, CA, USA), and Foxp3 (eBioscience, San Diego, CA, USA).

Protein kinase A (PKA) in vitro activity assay

The in vitro PKA activity assay was performed essentially as described previously (52). In brief, 2 × 10⁶ T cells (purified by negative selection from H₂RKO and H₂RKO-Tg1 mice as described previously in ref. 53), were stimulated with amthamine dihydrobromide (Tocris Bioscience, Ellisville, MO, USA) for the times indicated, and then washed 2 times with ice-cold phosphate-buffered saline. Cells were harvested in 30 μl of PKA activity buffer [50 mM Tris (pH 7.5), 0.5 mM EDTA, 50 mM β-glycerophosphate, 1.5 mM EGTA, 1 mM dithiothreitol (DTT), 0.1% Triton X-100, 25 μM 3-isobutyl-1-methylxanthine, and Protease Inhibitor Cocktail (Roche, Indianapolis, IN, USA)] and lysed by bath sonication (10 min at 4°C, Sonifier 2150; Branson Ultrasonics Corp., Danbury, CT, USA). The protein concentration was assayed using a standard BCA protocol (Pierce Chemical, Rockford, IL, USA). For each reaction, 100 ng of soluble PKA substrate (GFP227RRRRSII) in PKA activity buffer was mixed with 5 μg of whole-cell extract in 75 μl of PKA activity buffer containing a kinase reaction mix (15 mM MgCl₂, 200 μM ATP, and 0.5 mM DTT, final concentrations). The samples were incubated at 30°C for 20 min, after which the reaction was stopped by the addition of 5× Laemmli sample buffer. Samples were then subjected to SDS-PAGE and immunoblot analysis using anti-phospho-PKA substrate and antitubulin antibodies (to ensure equal loading).

Immune profiling

Single-cell suspensions of thymocytes, lymph node cells, and splenocytes were prepared, and the red blood cells were lysed with Gey's solution. Total numbers of cells were counted using a hematology analyzer (Advia 120; Bayer/Siemens, Tarrytown, NY, USA). For flow cytometric analysis, the cells were washed twice and incubated for 30 min on ice with the desired fluorochrome-conjugated mAbs or isotype control Ig at 0.5 μg/10⁶ cells. For the identification and phenotypic analysis of T regulatory (T_R) cells (CD4⁺CD8⁻TCRβ⁺Foxp3⁺), the following surface anti-mouse mAbs were used: anti-CD4 (MCD0417; Caltag Laboratories, Burlingame, CA, USA); anti-CD8 and anti-CD25 (53-6.7 and PC61; BD Pharmingen); anti-TCRβ, and anti-Foxp3 staining set (H57-5987 and FJK-16s; eBioscience). Viable cells were selected for flow cytometric analysis (LSR II; BD, Franklin Lakes, NJ, USA) based on forward and side scatter properties, and analysis was performed using FlowJo software (TreeStar Software, Ashland, OR, USA).

Statistics

Statistical analyses, as indicated in the figure legends, were performed using GraphPad Prism 5 software (GraphPad Software Inc., San Diego, CA, USA).

RESULTS

Transgenic expression of H₂R in H₂RKO T cells

Previously, we showed that decreased EAE susceptibility in H₂RKO mice is associated with an attenuated Th1 immune response and decreased proinflammatory cytokine production by H₂RKO macrophages (46). However, whether the blunted Th1 response and reduction in EAE severity was caused by a lack of H₂R signaling directly in CD4⁺ T cells or indirectly through its effect in APCs has remained unclear. In this study, we tested the hypothesis that H₂R signaling directly in CD4⁺ T cells is necessary and sufficient to complement EAE susceptibility to WT levels in H₂RKO mice. To this end, we generated transgenic mice expressing HA-H₂R under the control of the dlck promoter. To demonstrate that HA-H₂R was properly expressed on the cytoplasmic membrane, we transiently transfected HEK293T cells with either an empty pcDNA3.1 vector or the vector containing the HA-H₂R construct, stained the cells using anti-HA mAb, and confirmed H₂R expression by flow cytometry (Fig. 1A). Moreover, the activation of H₂R by histamine in HEK293T cells transfected with HA-H₂R increased Ca²⁺ levels (Fig. 1B), demonstrating that the HA-H₂R construct is functionally competent to signal. This construct was used to generate 2 transgenic lines expressing Hrh2 under the control of the dlck promoter directly on the B6 background and crossed to H₂RKO mice (H₂RKO-Tg1 and H₂RKO-Tg2). The expression of the HA-H₂R transgene was confirmed in CD4⁺ T cells by qRT-PCR using Hrh2-specific primers. Compared with WT, H₂RKO-Tg1 and H₂RKO-Tg2 CD4⁺ T cells exhibited ∼20-fold greater Hrh2 expression (Fig. 1C). To confirm that the Tg-H₂R was functional at the protein level in T cells, we performed in vitro assays using T cells isolated from H₂RKO and H₂RKO-Tg1 mice. H₂R activates the adenylate cyclase/cAMP system in a variety of tissues (26). As a surrogate for cAMP levels, we measured the activity of the cAMP-dependent PKA. Purified T cells from H₂RKO and H₁RKO-Tg1 mice were stimulated in vitro with increasing doses of amthamine, a highly specific agonist of H₂R (54), and PKA activity was measured as described in Materials and Methods. Although H₂RKO cells failed to respond to H₂R stimulation, as expected, H₂RKO-Tg1 T cells showed strong induction of PKA activity, indicative of functional signaling by the Tg-H₂R (Fig. 1D).

Figure 1.

In vitro expression and function of HA-H₂R construct and selective expression of H₂R in H₂RKO CD4⁺ T cells. A) HEK293T cells were transfected with empty pcDNA3.1 (control) and pcDNA3.1-HA-H₂R plasmids, and expression of HA-H₂R was determined by flow cytometry using an anti-HA mAb. GFP, green fluorescent protein. B) HEK293T cells transfected with either empty pcDNA3.1 (control) or pcDNA3.1-HA-H₂R plasmids were loaded with the Ca²⁺-sensing dye Fluo-4, and the change in Fluo-4 fractional fluorescence after stimulation with histamine was measured by real-time confocal microscopy. n = 8/time point and condition with data shown as means ± sem. Significance of differences in fluorescence was determined by 2-way ANOVA followed by Bonferroni post hoc multiple comparison test. ****P < 0.0001. C) Hrh2 transgene expression in CD4⁺ T cells were determined by qRT-PCR and analyzed using the comparative C_T method with β₂-microglobulin as an endogenous control. Receptor expression levels are normalized to WT levels. n = 2/strain with the data shown as means ± sem. D) Purified T cells from H₂RKO mice and H₂RKO-Tg1 mice were stimulated in vitro with increasing doses of amthamine dihydrobromide (0.1, 1, and 10 μM, as indicated) for 5 min at 37°C and lysed. Recombinant PKA substrate was added to the lysates, and its phosphorylation (pPKA-S) was detected using a phospho-specific immunoblotting (see Materials and Methods). Tubulin is shown as a loading control.

To determine whether the transgenic expression of H₂R in T cells inherently influenced immune cell composition, we determined the frequency of different cell types in the central and peripheral immune compartments of naive WT, H₂RKO, H₂RKO-Tg1, and H₂RKO-Tg2 mice. Because the number and frequency of all immune cells tested did not differ among the 2 transgenic founder lines, immune profiling data from the 2 lines were combined. There were no striking differences in the frequency of the immune cell subtypes in the thymus, lymph nodes, or spleen among the 3 strains (Supplemental Fig. S1). Therefore, the lack of systemic H₂R expression and the reexpression of H₂R exclusively in T cells do not affect the development or maintenance of the immune cells studied.

Transgenic expression of H₂R in H₂RKO T cells fails to complement EAE severity in 2× immunized mice

To assess whether H₂R expression in CD4⁺ T cells would complement EAE severity in H₂RKO mice to that of WT mice, adult WT, H₂RKO, H₂RKO-Tg1, and H₂RKO-Tg2 mice were immunized with 100 μg of MOG_35–55 in CFA on d 0 and 7 (2× protocol) and scored for clinical signs of EAE through d 30 postimmunization. Consistent with our previous findings, the severity of the clinical disease course differed significantly between WT and H₂RKO mice, with H₂RKO mice exhibiting less severe disease (Fig. 2A and ref. 46). However, the EAE disease course of both transgenic lines was also significantly less severe than the WT and comparable to H₂RKO mice (WT>H₂RKO=H₂RKO-Tg1=H₂RKO-Tg2; Fig. 2A). This finding was confirmed by the analysis of additional clinical quantitative trait variables, including cumulative disease score, day of onset, peak score, number of days affected, and overall severity index, which showed that WT > H₂RKO = H₂RKO-Tg1 = H₂RKO-Tg2, the only exception being the incidence, which did not differ significantly among the strains (Table 1). Furthermore, histopathological analysis of the CNS revealed significantly less pathology in the spinal cords of H₂RKO, H₂RKO-Tg1, and H₂RKO-Tg2 mice than in those of WT mice (WT>H₂RKO=H₂RKO-Tg1=H₁RKO-Tg2; Fig. 2B), with no significant difference in brain pathology (data not shown). These results indicate that selective reexpression of H₂R in T cells does not influence EAE severity in H₂RKO-Tg mice immunized with the 2× protocol.

Figure 2.

Transgenic expression of H₂R in H₂RKO T cells does not complement EAE severity elicited by 2× immunization. A) WT (n=21), H₂RKO (n=28), H₂RKO-Tg1 (n=17), and H₂RKO-Tg2 (n=8) mice were immunized using the 2× immunization protocol. Clinical scores after immunization were recorded, and significance of differences between clinical courses was calculated by regression analysis with the best fit curve shown and 2-way ANOVA followed by Bonferroni post hoc multiple comparison test. B) Significance of differences observed in spinal cord histopathology between WT (n=10), H₂RKO (n=19), H₂RKO-Tg1 (n=12), and H₂RKO-Tg2 (n=8) was determined using the Kruskal-Wallis test followed by Dunn's post hoc multiple comparison test. Data are shown as means ± sem. *P < 0.05; **P < 0.01; ***P < 0.001.

Table 1.

Clinical disease traits after immunization of C57BL/6J, H₂RKO, and H₂RKO-dlck^Hrh2 Tg mice with 2× MOG_35–55 + CFA

Strain	Incidencea	CDS	DA	SI	PS	DO
B6	18/21, 86	22.4 ± 3.2	10.5 ± 1.2	2.0 ± 0.1	2.0 ± 0.2	18.4 ± 0.8
H2RKO	10/28, 36	7.0 ± 2.2	4.0 ± 1.1	1.7 ± 0.2	0.7 ± 0.2	19.8 ± 1.3
dlckHrh2 Tg1	8/17, 47	6.4 ± 2.0	5.2 ± 1.4	1.1 ± 0.1	0.7 ± 0.2	20.6 ± 0.6
dlckHrh2 Tg2	3/8, 38	4.1 ± 2.0	4.1 ± 2.0	1.0 ± 0	0.4 ± 0.2	20 ± 0
Overall	χ2 = 13.2, 3; P = 0.004	H = 19.4; P = 0.0002	H = 15.4; P = 0.002	H = 12.6; P = 0.006	H = 19.2; P = 0.0003	H = 0.3; P = NS
Post hoc	B6 > H2RKO = Tg1 = Tg2	B6 > H2RKO = Tg1 = Tg2	B6 > H2RKO = Tg1 = Tg2	B6 > H2RKO = Tg1 = Tg2	B6 > H2RKO = Tg1 = Tg2	B6 = H2RKO = Tg1 = Tg2

CDS, cumulative disease score over 30 d of experiment; DA, days affected; SI, severity index (cumulative disease score/days affected); PS, peak score; DO, day of onset in affected animals; NS, not significant. Values are shown as means ± sem. Significance of differences for the trait values among the strains was assessed by χ² analysis (overall incidence) or the Kruskal-Wallis test (H), followed by Dunn's post hoc multiple comparisons; P values are as indicated.

^aPercentage affected. Animals were considered affected if clinical scores ≥1 were apparent for ≥2 consecutive days.

Because H₂R is reported to play a role in T-cell polarization and cytokine production (55), we investigated whether the transgene had an effect on the MOG_35–55-specific immune responses in 2× immunized animals. Splenocytes and DLN cells were isolated on d 10 postimmunization and assayed for proliferation and cytokine production after restimulation with MOG_35–55 for 72 h. The proliferative responses of WT, H₂RKO, and H₂RKO-Tg T cells were not significantly different (Fig. 3A). However, production of IFN-γ, IL-17, IL-6, and GM-CSF by H₂RKO T cells was significantly lower than that for the WT and was not complemented to WT levels in H₂RKO-Tg mice (Fig. 3B−D), except for GM-CSF, which was partially complemented to WT levels (Fig. 3E). Therefore, as with clinical EAE, these results show that H₂R signaling in T cells does not affect T-cell effector responses elicited using the 2× immunization protocol.

Figure 3.

Ex vivo MOG_35–55-specific T-cell proliferative responses and cytokine profiles in 2× immunized WT, H₂RKO, and H₂RKO-Tg mice. A) MOG_35–55-specific T-cell proliferative responses were evaluated by [³H]thymidine incorporation. Mean ± sd counts per minute were calculated from triplicate wells of n = 10/strain. Significance of differences in proliferation was determined by 2-way ANOVA. B−E) IFN-γ (B), IL-17 (C), IL-6 (D), and GM-CSF (E) production by MOG_35–55-stimulated DLN cells and splenocytes from WT (n=10), H₂RKO (n=22), and H₂RKO-Tg mice (n=24) were quantified by ELISA and are shown as means ± sem. Significance of differences observed in cytokine production was determined by 1-way ANOVA followed by Bonferroni post hoc multiple comparison test. ***P < 0.001; ****P < 0.0001.

To examine whether the selective expression of H₂R in T cells had an effect on the frequency of effector T-cells, splenocytes and DLN cells were isolated on d 10 after 2× immunization, stimulated with PMA and ionomycin for 4 h in the presence of brefeldin A and analyzed by flow cytometry. The 3 strains had a similar frequency of IFN-γ⁺ and IL-17⁺ CD4⁺ and CD8⁺ T cells (Fig. 4A, B). We did, however, detect a significant decrease in the frequency of CD4⁺Foxp3⁺ T_R cells in H₂RKO mice that was complemented to WT levels in H₂RKO-Tg mice (Fig. 4C). These data show that the expression of H₂R selectively in T cells plays a role in the development and/or maintenance of induced T_R cells. The decrease in T_R frequency in EAE-resistant H₂RKO mice is probably a downstream effect of overall diminished T-cell responsiveness and is most likely not directly responsible for disease severity.

Figure 4.

Frequency of effector T cells in the DLNs of 2× immunized WT, H₂RKO, and H₂RKO-Tg mice. Single-cell suspensions of splenocytes and DLN cells were prepared from 2× immunized mice on d 10 postimmunization, stimulated with PMA and ionomycin for 4 h in the presence of brefeldin A, stained, and analyzed by flow cytometry. A, B) Percentage of IFN-γ⁺ and IL-17⁺ cells gated on TCRαβ⁺CD4⁺ (A) and TCRαβ⁺CD8⁺ (B) of WT (n=10), H₂RKO (n=22), and H₂RKO-Tg mice (n=24). C) Frequency of CD4⁺CD8⁻TCRβ⁺Foxp3⁺ T_R cells in the DLNs was determined by intracellular Foxp3 staining using the mouse/rat Foxp3 staining set (eBioscience). Data are shown as means ± sem. Significance of differences observed was determined by 1-way ANOVA followed by Bonferroni post hoc multiple comparison test. ***P < 0.001; ****P < 0.0001.

EAE severity and neuropathology in H₂RKO mice are complemented by expression of H₂R exclusively in T cells and environmental stimuli

PTX, a major virulence factor of Bordetella pertussis, is used as an ancillary adjuvant to elicit tissue-adjuvant models of organ-specific autoimmune diseases including EAE (56). In addition, we have shown that PTX is able to override genetic checkpoints and alter disease susceptibility in a gene × PTX specific fashion and is therefore an environmental factor that can be used to model gene × environment interactions in EAE susceptibility (47, 48). We studied EAE susceptibility of WT, H₂RKO, H₂RKO-Tg1, and H₂RKO-Tg2 mice elicited using the 1× immunization protocol, which includes PTX. The mice received on d 0 the same amount of MOG_35–55 in CFA as in the 2× protocol and 200 ng of PTX by intravenous injection. As observed with the 2× protocol, EAE severity was significantly reduced in H₂RKO mice compare with that in WT mice. However, in contrast to the 2× protocol, the reexpression of H₂R specifically in T cells complemented disease severity of H₂RKO mice to that of WT mice (WT>H₂RKO=H₂RKO-Tg1=H₂RKO-Tg2; Fig. 5A). Furthermore, an analysis of EAE-associated clinical quantitative trait variables, including cumulative disease score, peak score, day of onset, number of days affected, and overall severity index, confirmed this observation (Table 2). In addition, histopathological analysis of the CNS revealed a reduction in the spinal cord pathology of H₂RKO mice compared with that in WT mice and the 2 transgenic lines (WT>H₂RKO=H₂RKO-Tg1=H₂RKO-Tg2; Fig. 5B), with no significant difference in brain pathology (data not shown). These results demonstrate that a H₂R-PTX interaction in H₂RKO-Tg mice is required to complement EAE susceptibility to that of WT animals.

Figure 5.

Transgenic expression of H₂R in H₂RKO T cells complements EAE severity elicited by 1× immunization. A) WT (n=30), H₂RKO (n=66), H₂RKO-Tg1 (n=42), and H₂RKO-Tg2 (n=19) mice were immunized using the 1× immunization protocol. Clinical scores after immunization were recorded, and the significance of differences between clinical courses was calculated by regression analysis with the best fit curve shown and 2-way ANOVA followed by Bonferroni post hoc multiple comparison test. B) Significance of differences observed in spinal cord histopathology between WT (n=8), H₂RKO (n=31), H₂RKO-Tg1 (n=14), and H₂RKO-Tg2 (n=5) mice was determined using the Kruskal-Wallis test followed by Dunn's post hoc multiple comparison test. Data are shown as means ± sem. **P < 0.01; ***P < 0.001; ****P < 0.0001.

Table 2.

Clinical disease traits after immunization of C57BL/6J, H₂RKO, and H₂RKO-dlck^Hrh2 Tg mice with 1× MOG_35–55 + CFA + PTX

Strain	Incidencea	CDS	DA	SI	PS	DO
B6	28/30, 94	46.3 ± 3.7	17.8 ± 1.1	2.7 ± 0.1	2.9 ± 0.2	12.0 ± 0.7
H2RKO	56/66, 85	25.0 ± 1.9	13.8 ± 0.8	1.8 ± 0.1	1.8 ± 0.1	14.6 ± 0.5
dlckHrh2 Tg1	40/42, 95	46.4 ± 3.1	18.1 ± 0.8	2.5 ± 0.1	3.0 ± 0.2	12.0 ± 0.4
dlckHrh2 Tg2	17/19, 90	44.1 ± 4.7	18.5 ± 1.6	2.4 ± 0.1	2.5 ± 0.3	10.3 ± 0.5
Overall	χ2 = 3.54, 3; P = 0.32	H = 44.4; P < 0.0001	H = 31.8; P < 0.0001	H = 45.9; P < 0.0001	H = 38.5; P < 0.0001	H = 33.3; P < 0.0001
Post hoc	B6 = H2RKO = Tg1 = Tg2	B6 = Tg1 = Tg2 > H2RKO	B6 = Tg1 = Tg2 > H2RKO	B6 = Tg1 = Tg2 > H2RKO	B6 = Tg1 = Tg2 > H2RKO	B6 = Tg1 = Tg2 > H2RKO

CDS, cumulative disease score over 30 d of experiment; DA, days affected; SI, severity index (cumulative disease score/days affected); PS, peak score; DO, day of onset in affected animals; NS, not significant. Values are shown as means ± sem. Significance of differences for the trait values among the strains was assessed by χ² analysis (overall incidence) or the Kruskal-Wallis test (H), followed by Dunn's post hoc multiple comparisons; P values are as indicated.

^aPercentage affected. Animals were considered affected if clinical scores ≥1 were apparent for ≥2 consecutive days.

To investigate the effects of the H₂R-PTX interaction on the immune response associated with EAE complementation, we studied the MOG_35–55-specific proliferative and cytokine recall responses of CD4⁺ T cells elicited by 1× immunization. Unexpectedly, the transgenic expression of H₂R in T cells did not complement the proliferative capacity of these cells to WT levels nor did it restore the deficiency in IL-17 production observed in H₂RKO cells (Fig. 6A, B). In contrast to IL-17 production, T cells from 1× immunized mice produced similar levels of IFN-γ, IL-6, and GM-CSF regardless of the presence or absence of H₂R expression (Fig. 6C−E).

Figure 6.

Ex vivo MOG_35–55-specific T-cell cytokine profiles and proliferative response in 1× immunized WT, H₂RKO, and H₂RKO-Tg mice. A) MOG_35–55-specific T-cell proliferative responses were evaluated by [³H]thymidine incorporation. Mean counts per minute ± sd were calculated from triplicate wells of n = 10/strain. The significance of the observed differences in proliferation was determined by stepwise 2-way ANOVA. B−E) IFN-γ (B), IL-17 (C), IL-6 (D), and GM-CSF (E) production by MOG_35–55-stimulated DLN cells and splenocytes from WT (n=10), H₂RKO (n=10), and H₂RKO-Tg mice (n=32) were quantified by ELISA and are shown as means ± sem. Significance of differences observed in cytokine production was determined by 1-way ANOVA followed by Bonferroni post hoc multiple comparison test. **P<0.01.

We next examined the frequency of effector T cells. Splenocytes and DLN cells were isolated on d 10 after 1× immunization, and cytokine production was analyzed by intracellular staining and flow cytometry. There was a reduction in the frequency of CD4⁺IFN-γ⁺ cells in H₂RKO mice that was restored to WT levels by the transgenic expression of H₂R in T cells. Furthermore, the frequency of CD4⁺IL-17⁺ T cells was significantly greater in H₂RKO-Tg mice than in WT and H₂RKO mice (Fig. 7A). The frequencies of CD8⁺IFN-γ⁺ and CD8⁺IL-17⁺ T cells as well as that of T_R cells were comparable among the strains (Fig. 7B, C). The discrepancy between the levels of cytokine production observed (Fig. 6) and the frequency of cytokine-positive cells (Fig. 7) is mainly attributed to the experimental conditions and methods used for cytokine detection. In Fig. 6, cytokine production reflects the accumulation/consumption of cytokines over a period of 72 h. In Fig. 7, the frequency of cytokine-positive cells reflects the capacity of individual cells to produce cytokines immediately after harvesting from immunized mice. Therefore, these data suggest that the H₂R-PTX interaction influencing EAE susceptibility is probably at the level of the development and frequency of CD4⁺IFN-γ⁺ and CD4⁺IL-17⁺ T cells rather than on the secretion of these cytokines by CD4⁺ effector cells.

Figure 7.

Frequency of effector T cells in the DLNs of 1× immunized WT, H₂RKO, and H₂RKO-Tg mice. Single-cell suspensions of spleen and DLN cells were prepared from 1× immunized mice on d 10 postimmunization, stimulated with PMA and ionomycin for 4 h in the presence of brefeldin A, stained, and analyzed by flow cytometry. A, B) Percentage of IFN-γ⁺ and IL-17⁺ cells gated on TCRαβ⁺CD4⁺ (A) and TCRαβ⁺CD8⁺ (B) of WT (n=10), H₂RKO (n=10), and H₂RKO-Tg mice (n=32). C) Frequency of CD4⁺CD8⁻TCRβ⁺Foxp3⁺ T_R cells in the DLNs was determined as described in the legend to Fig. 5. Data are shown as means ± sem. Significance of differences observed was determined by 1-way ANOVA followed by Bonferroni post hoc multiple comparison test. *P < 0.05; **P < 0.01.

DISCUSSION

Previously, we reported that H₂RKO mice are significantly less susceptible to MOG_35–55-induced EAE than WT mice (46). Decreased disease susceptibility in H₂RKO mice was associated with an attenuated Th1 response and decreased production of proinflammatory cytokines, including IL-12 and IL-6 by APCs. Furthermore, H₂RKO APCs produced increased levels of MCP-1, a cytokine that negatively regulates Th1 immune responses (46, 57). In this study, we extended our previous findings by determining whether T-cell intrinsic H₂R signaling is necessary and sufficient to restore EAE susceptibility to that of WT mice, as has been observed previously with H₁R (50), by generating transgenic mice expressing H₂R exclusively in T cells.

We found that the EAE disease course, disease-associated quantitative trait variables, and pathology of H₂RKO mice were not complemented by the selective transgenic expression of H₂R in T cells when immunized using the 2× protocol. Similarly, ex vivo MOG_35–55-specifict cytokine production by H₂RKO-Tg CD4 T cells from 2× immunized mice was not complemented to that of WT CD4 T cells. In contrast, the EAE disease course, disease-associated clinical trait variables, pathology, and frequency of IFN-γ⁺ and IL-17⁺ CD4⁺ T cells were complemented in H₂RKO-Tg mice to WT levels on 1× immunization, which includes the use of PTX as an ancillary adjuvant that activates APCs (58–63). Therefore, we have identified a H₂R-PTX interaction that is essential for the optimal generation of pathogenic CD4⁺ T-cell responses and EAE susceptibility. Notable, this is in contrast to H₁R signaling in T cells, which is necessary and sufficient for full EAE susceptibility elicited by immunization with both the 1× and 2× protocols (50).

PTX, a major virulence factor of B. pertussis, is the causative agent for whooping cough. The holotoxin is a hexameric protein that conforms to the A-B model of bacterial exotoxins (64). The A promoter (S1-subunit) is an ADP-ribosyltransferase, which ribosylates the α-subunits of heterotrimeric Gα_i/o proteins, resulting in uncoupling from GPCRs. The B oligomer, which consists of 4 subunits (S2−S5) binds cell surface receptors on a variety of mammalian cells and is capable of activating an intracellular signal transduction cascade (65). However, for the induction of EAE, the active holotoxin is required (66). Although H₃R and H₄R couple via Gα_i/o proteins, H₁R and H₂R couple to the second messenger signaling pathways via Gα_q/11 and Gα_s proteins (10, 26, 67), suggesting that PTX does not influence H₂R signaling directly. However, the effect of PTX on EAE may be explained by its inhibitory effects on various Gα_i/o protein-coupled receptors present on T cells or APCs. In this regard, T cells from mice deficient in Gα_i2 exhibit a Th1-biased effector response in association with spontaneous autoimmune disease (68–71). The cytokine hyperresponsiveness in Gα_i2-knockout mice was subsequently shown to be due to abnormal T-cell receptor signaling (72). Although this finding suggests a direct role for PTX in regulating T-cell effector responses and some studies have suggested a direct effect of PTX on T cells (73), including increased expression of CD28 paralleled by up-regulation of CD69 and induction of IFN-γ, granzyme B, and IL-17 (74), the preponderance of the data suggests that PTX acts indirectly on T cells by activating APCs and up-regulating CD80, CD86, and major histocompatibility complex class II expression (58–63).

The finding that H₂RKO-Tg mice exhibit disease severity similar to that of H₂RKO mice in the absence of PTX suggests that H₂R signaling in T cells alone is not sufficient to generate encephalitogenic T-cell responses in this model. This probably reflects the critical role of H₂R signaling in APCs, consistent with other findings on the role of H₂R in APCs (29, 30). However, in the presence of PTX, H₂R signaling in T cells alone is in fact sufficient to restore EAE severity to WT levels. This suggests that PTX overrides the requirement for H₂R signaling in APCs for the generation of encephalitogenic T cells. Taken together, these results predict a model that requires two H₂R-dependent signals for the generation of encephalitogenic T cells: signal 1, an H₂R-dependent signal in T cells and signal 2, an H₂R-dependent signal in APCs, which is complemented by PTX in H₂RKO APCs. The finding that PTX can override signal 2 is not surprising, given the known functions of PTX in enhancing APC function (58–63) and the findings that PTX can override many genetic checkpoints in EAE resistance (47, 48). Because PTX inhibits Gα_i/o protein-coupled signaling, our model also predicts the presence of a Gα_i/o-coupled receptor that is inhibitory for APC function in EAE. Several lines of evidence suggest that in macrophages PTX-sensitive Gα_i/o proteins regulate not only GPCR signaling, but also Toll-like receptor (TLR) signaling (75). In addition, the expressions of GPCR kinases and arrestins, 2 major protein families that primarily regulate receptor desensitization of almost all GPCRs (76, 77), are differentially regulated by TLRs at the transcriptional, post-transcriptional, and post-translational levels (78). Clearly, this pathway is a likely target of PTX action in our model and may be an important pathway in integrating other environmental risk factors, including exposure to infectious agents, which is thought to play an adjuvant-like role in CNS autoimmune disease (79).

In this regard, Frei et al. (80) has recently shown that cell intrinsic crosstalk between H₂R and TLR signaling modifies APC and DC cell functional responses to microbial ligands. Clearly, noncell autonomous crosstalk between H₂R signaling in T cells and APCs, as well as cell autonomous crosstalk between H₂R and signaling pathways activated by environmental risk factors in APCs and DCs, has the potential to play a central role in regulating innate and adaptive immune responses in both health and disease.