Designing mimetic of the interface functional groups of known receptor-ligand complexes is an attractive strategy for developing potential therapeutic agents that interfere with target protein-protein interactions. The CD80/CD86-CD28/CD152 costimulatory interactions transmit signals for CD4+ T cell activation and suppression and are critically involved in the initiation, progression, and reactivation of the immunopathology in multiple sclerosis. Differences in the pattern, levels, and kinetics of expression of CD80/CD86 molecules in conjunction with differences in the strength of the signals delivered upon binding CD28 or CD152 determine the outcome of the immune response. A temporal up-regulation of surface expression of CD80 relative to CD86 on APCs and CNS-infiltrating cells has been shown to correlate with disease progression in experimental autoimmune encephalomyelitis an animal model for multiple sclerosis. Hence blockade of the CD80 costimulatory axis has therapeutic potential in multiple sclerosis. In this study, we report the efficacy of a novel CD80-blocking agent CD80-competitive antagonist peptide (CD80-CAP) in suppressing clinical disease and relapse in experimental autoimmune encephalomyelitis. The CD80-CAP mediates protection by inhibiting proinflammatory cytokines and skewing toward anti-inflammatory response presumably by enhancing the expression of glucocorticoid-induced leucine zipper in activated CD4+ T cells.

Multiple sclerosis (MS)3 is the most common inflammatory disease of the CNS affecting ∼2.5 million individuals worldwide, and is second only to trauma as the cause of acquired neurological disability in young adults (1). The relapsing-remitting form of experimental autoimmune encephalomyelitis (EAE) is an animal model that shares many clinical and histopathological features of MS and is often used as a preclinical ground to test the efficacy of potential therapeutic agents (2).

Considerable evidence suggests that the T cell-mediated immune responses are critical in the pathogenesis of MS and EAE (2). Complete activation of T cells requires a primary signal delivered via TCR binding an antigenic peptide contained within the class II MHC on an APC and a second costimulatory signal. Two structurally and functionally well-characterized T cell costimulatory molecules are the CD28 and the CTLA-4 (CD152), both of which bind the same ligands, CD80 and CD86 on the APC. Upon binding the B7 ligands, CD28 costimulation amplifies TCR signaling and promotes T cell proliferation, differentiation, and survival. Activated T cells express CD152, which bind the B7 ligands and down-regulates T cell response by interfering with the TCR signals, facilitating development of Ag-specific tolerance (2, 3).

The B7-CD28/CD152 interactions are integrally involved in regulating T cell functions at several stages in EAE and MS (4). These include the initial activation of peripheral T cells, homing and reactivation of autoreactive T cells within the CNS (2, 4, 5, 6). Although myelin basic protein (MBP)-specific PL/J TCR transgenic mice (that express a Vα4 Vβ8 TCR on T cells) in a RAG1−/− background develop spontaneous EAE, CD28−/− transgenic MBP-positive RAG1−/− mice are free of disease providing a genetic evidence for CD28 costimulation in the initiation of EAE (7). The CD80−/−/CD86−/− mice exhibit markedly reduced clinical disease and inflammatory histopathology after adoptive transfer of encephalitogenic T cells, implying a critical role for B7-CD28 interactions in the effector phases of EAE (5).

Biophysical and computational analyses of contributions by CD28 costimulation for T cell activation suggests that the amount of signaling via CD28 is proportional to the number of CD80 and CD86 ligands available for interaction (8). Although CD86 is constitutively expressed on resting APCs, the expression of CD80 is up-regulated following activation (9). Thus, although due to its higher expression CD86 may be the dominant ligand for CD28 following initial contact with the T cells, the elevated levels of CD80 following activation makes it the dominant ligand subsequently. Hence, selective blockade of CD80-receptor interaction represents an attractive strategy to modulate T cell-mediated immune responses.

Structurally, CD28 and CD152 are members of the Ig superfamily (IgSF) with one Ig variable domain (10). Analogous to the complementarity-determining regions (CDRs) that form the Ag-combining sites of the Ab, the high specificity of the Ig superfamily receptor-ligand interaction is attributed to the sequence and structure of the CDR-like regions unique to each Ig superfamily protein. Hence, CDR-like regions provide ideal templates for the design of mimetic that can potentially perturb-specific Ig superfamily receptor-ligand interactions (11). CD28 and CD152 share a highly conserved hydrophobic motif localized in the ligand binding CDR-3-like region that adopts a polyproline type II (PPII) helical conformation (12, 13). Integrating interface residue preference and propensity to form PPII helix, we designed and characterized a selective CD80-binding hexapeptide referred as CD80-competitive antagonist peptide (CD80-CAP) that possess optimum PPII helical content, competitively inhibit B7-receptor interactions and suppress T cell activation (14, 15).

In the present study, we investigated the potential of the CD80-CAP to suppress EAE and characterized its mechanism of action. Our results indicate that the treatment with the CD80-CAP ameliorated clinical disease in relasping EAE (R-EAE). The suppression was mediated by the inhibition of Th1 cytokines (IFN-γ, TNF-α, IL-12) and skewing toward Th2 (IL-10, TGF-β) response presumably secondary to the enhanced expression of glucocorticoid-induced leucine zipper (GILZ) and GATA-3, the canonical Th2 transcription factor in CD4+ T cells (16).

The 6- to 8-wk-old SJL/J (H-2s), BALB/c, and B10.PL female mice were purchased from Charles River Breeding Laboratory or The Jackson Laboratory and housed under specific pathogen-free conditions at the Bioresearch facility of the Indiana University School of Dentistry (Indianapolis, IN). All animal protocols were approved by Indiana University Institutional Animal Care and Use Committee.

CD80-CAP is a hexapeptide designed to mimic the PPII helical orientation of the CD28/CD152 receptor in the binding interface with CD80. The control peptides include sequence control consisting of scrambled residues of the CD80-CAP and a structural control for the PPII helix (15). The CD80-CAP and the control peptides were synthesized on Rink amide resin by solid-phase peptide synthesis using F-moc methodology at the Biochemistry and Biophysics facility (Indiana University School of Medicine, Indianapolis, IN), as described (15). The free N-terminal group of the terminal amino acid residue was acetylated. All peptides were purified by semipreparative RP-HPLC, and the identity of the purified peptide was confirmed by MALDI-TOF mass spectrometry. The proteolipid protein (PLP)139–151 (HSLGKWLGHPDKF) or MBP NAc1–11 (AASQKRPSQRHG) peptide used for induction of EAE were commercially prepared by GenScript and supplied at 95% purity.

To investigate the in vivo tolerance of the CD80-CAP, a standard LPS-induced lethal endotoxemia mouse model was used (17, 18). BALB/c mice were i.p. injected with LPS (100 μg). Groups of mice were administered immediately i.v. sterile PBS/the CD80-CAP (25 μg/kg)/the control peptide in PBS. Serum collected after 1.5, 4, and 8 h later was assessed for concentrations of TNF-α, IFN-γ, and IL-12, respectively by sandwich ELISA using mAb pairs and recombinant cytokine standards the manufacturer’s instructions (BD Pharmingen).

By active immunization.

The 8- to 10-wk-old female mice were immunized with 100 μg of PLP139–151 (SJL/J) or MBP NAc1–11 (B10.PL) in PBS emulsified 1:1 in CFA supplemented with 200 μg of Mycobacterium tuberculosis H37RA (Difco Laboratories) distributed over four sites on the lateral hind flanks s.c. Mice were i.p. injected with 100 ng of pertussis toxin (List Biological Laboratories) on the day of immunization and 2 days later. Animals were observed daily for clinical signs and scored as follows: 0, no clinical signs; 1, limp tail or waddling gait with tail tonicity; 2, waddling gait with limp tail (ataxia); 2.5, ataxia with partial paralysis of one limb; 3, partial hind-limb paralysis; 3.5, full paralysis of one limb with partial paralysis of the second limb; 4, full paralysis of two limbs; 4.5 moribund; and 5, death (19). As opposed to the B10.PL mice that often develop monophasic EAE, PLP139–151 immunized SJL/J mice develop relapsing-remitting form of EAE (R-EAE). For treatment of EAE, mice were injected i.v. with 500 μg of the CD80-CAP or the control peptide or vehicle (reconstituted in sterile PBS) on the day of immunization and 21 days later in some experiments.

By adoptive transfer.

Naive 7- to 8-wk-old female donor B10.PL mice were s.c. primed with 100 μg of MBP NAc1–11 in CFA. Groups of mice were treated with vehicle or 500 μg of the CD80-CAP or the control peptide. Because the efficacy of the CD80-CAP is based on the ability to adopt optimal PPII orientation in the context of the CD80 binding interface, we used the structural control peptide for comparative analysis (14, 15). Single cell suspensions of draining lymph node cells (LNC) and splenocytes harvested 10 days after disease induction were restimulated in vitro for 48 h with MBP NAc1–11 (40 μg/ml) in RPMI 1640 medium containing 10% FCS, 25 mM HEPES, 2 mM l-glutamine, 50 U/ml penicillin, 50 μg/ml streptomycin, and 5 × 105 M 2-ME in round-bottom 96-well plates. CD4+CD25 effector T cells were isolated by magnetic cell sorting using an AutoMACS according to the manufacturer’s instructions (Miltenyi Biotec). At first, CD4+ T cells were enriched by negative depletion of CD8+ T cells and B cells using anti-CD8α and anti-CD45RA microbeads. Then CD4+ cells were stained with biotinylated mAb against CD25 (clone OX39) followed by streptavidin microbeads. CD4+CD25 cells were isolated by negative selection. The purity of the CD4+CD25 cells was >95%. Groups of naive B10.PL mice were immunized with MBP NAc1–11 peptide and transferred 1 × 106 CD4+CD25 cells isolated from the mice treated with vehicle/CD80-CAP/control peptide in a final volume of 100 μl.

In separate experiments, splenocytes isolated 10 days postimmunization from mice induced EAE and administered vehicle/the CD80-CAP/the control peptide were stimulated similarly ex vivo with MBP-NAc1–11. CD4+CD25+ cells were isolated using the CD4+CD25+ regulatory T cell isolation kit (catalog no. 130-091-041; Miltenyi Biotec) as per the recommended protocol. Groups of naive B10.PL mice were immunized with MBP NAc1–11 peptide and transferred 0.5 × 106 CD4+CD25+ cells. All animals were observed daily and the clinical disease was recorded. Serum from blood collected by tail vein puncture 8 days posttransfer was assessed for IFN-γ, TNF-α, and TGF-β by ELISA.

Draining lymph nodes and spleen were harvested 10 or 45 days postimmunization from SJL/J mice induced R-EAE and treated with vehicle/500 μg of the CD80-CAP/the control peptide. Single cell suspensions were obtained and CD4+ T cells were separated by magnetic cell sorting using anti-CD4 microbeads following the recommended protocol (Miltenyi Biotec). CD4+ LNC/splenocytes were cultured in 96-well plates at 5 × 105 cells/well and restimulated with PLP139–151 peptide at a final concentration of 20 μg/ml in HL-1 medium (BioWhittaker). Irradiated splenocytes from syngenic mice were used as APC. CD4+ splenocytes were pulsed with [3H]thymidine (New England Nuclear) at 1 μCi/well on day 3 of culture for the final 18 h. Mean incorporation of thymidine in DNA was measured in triplicate wells by liquid scintillation counting (model LS 5000; Beckman Instruments). Supernatants from cultures of LNC were harvested at 24, 48, or 72 h of culture. Cytokines (IFN-γ, IL-12, and IL-10) were determined by sandwich ELISA. In separate experiments, draining LNC and splenocytes isolated 10 days postimmunization from B10.PL mice induced EAE were assessed similarly for proliferative and cytokine responses to MBP-NAc1–11 in the presence of increasing concentrations (125–500 μM) of the CD80-CAP or the control peptide.

For CFSE analysis, single cell suspension of splenocytes obtained from naive SJL/J mice were incubated with 5 μM CFSE (Molecular Probes) in RPMI 1640 for 10 min and washed three times with ice-cold PBS (20). The cells were then resuspended in complete RPMI 1640 and cultured in the presence of PLP139–151 at 40 μg/ml and varying concentrations of the CD80-CAP or control peptide as described. Cells were harvested at the end of 24 h (day 1), 48 h (day 2), and 72 h (day 3), stained with PE-conjugated anti-CD4 mouse mAb or rat anti-mouse IgG2a istoype, washed, and subsequently fixed in PBS/2% paraformaldehyde. CD4+ splenocytes were then analyzed for cell division on a FACSCalibur flow cytometer using CellQuest Pro software (BD Biosciences). Stimulation index was determined by the ratio of proliferated to unproliferated cells (21).

Draining LNC isolated 10 days postimmunization from SJL or B10.PL mice induced EAE with PLP139–151 or MBP Nac1–11, respectively, were restimulated in vitro with the immunogen (40 μg/ml) for 48 h as described. CD4+ T cells were isolated by magnetic sorting. Total cellular RNA from pure populations of CD4+ LNC was isolated using Qiagen kit (Invitrogen) following the manufacturer’s protocol. The 2–4 μg of total RNA was reverse transcribed using iScript cDNA synthesis kit (Bio-Rad). The concentration of the cDNA was measured at 260 and 280 nm by the Gensys5 model UV-visible spectrophotometer (Thermoelectronic). Real-time PCR was performed by using the SYBR Green/ROX quantitative PCR master mix (SA Biosciences), according to the manufacturer’s recommendations on the ABI Prism 7000 Sequence Detection System (PerkinElmer Applied Systems). Each reaction contains 2 × 12.5 μl of SYBR Green Master Mix, 1 μl of 10 μM primers, 50 ng of the cDNA, to a total volume of 25 μl. The thermal cycling conditions included an initial denaturation step at 50°C for 2 min, 95°C for 10 min; 40 cycles at 95°C for 15 s, primer specific annealing temperature for 30 s and extension at 72°C for 30 s. The following primers include: GAPDH (forward) 5′-AGAAGGACTATAACCCTGGCTC-3′, (reverse) 5′-CTTCCACGATCCCGAAGTTTT-3′; β2-microglobulin (forward) 5′-ATGGCTCGCTCGGTGACCCT-3′, (reverse) 5′-TTCTCCGGTGGGTGGCGTGA-3′; T-bet (forward) 5′-TCCCATTCCTGTCCTTCA-3′, (reverse) 5′-GCTGCCTTCTGCCTTT C-3′; GATA-3 (forward) 5′-ACCACGGGAGCCAGGTATG-3′, (reverse) 5′-CGGAGGGTAAACGGACA GAG-3′; and GILZ (forward) 5′-TGACTGCAACGCCAAAGC-3′, (reverse) 5′-TCCATGGGGGTCTGATACA T-3′. The primers generated fragments of 372 bp for T-bet, 170 bp for GATA-3, 110 bp for β2-microglobulin (22), 111 bp for GAPDH, and 102 bp for GILZ (23). The melting point analysis was conducted by heating the amplicon from 65–95°C, allowing the characteristic melting point to be found for each product. The length of the product was confirmed by electrophoresis. The gene-specific threshold cycle (Ct) for each sample (ΔCt) was corrected by subtracting the threshold cycle for the housekeeping gene β2-microglobulin/GAPDH. Untreated controls were chosen as the reference samples, and the ΔCt for all experimental samples were subtracted by the ΔCt for the control samples (ΔΔCt) (25). The magnitude of change in the mRNA was expressed as 2−ΔΔCt. Experiments were performed in triplicate.

LNC and splenocytes were incubated with various mAb (PE-anti-CD25, FITC-anti-CD4 (BD Pharmingen) and FITC-anti-FoxP3 (eBioscience)) at 0.5 μg/ml final concentration, fixed in 1% paraformaldehyde and analyzed on a FACSCalibur flow cytometer (BD Biosciences). For analysis of intracellular CD152 and FoxP3, cells were permeabilized briefly with 0.5% saponin, washed, and stained with PE-anti-CD152 mAb or FITC-anti-FoxP3 (19). Isotype-matched Ab and IgG block to avoid nonspecific binding to Fc receptors were used as controls.

For the mean clinical score, proliferation, cytokine, flow cytometric, and mRNA assays, a one-way ANOVA with Tukey’s posthoc was performed to determine differences between the groups. Results were considered statistically significant at p < 0.05.

CD80/CD86-CD28 costimulatory blockade has been shown to suppress proliferation of TCR-stimulated T cells (2, 14, 19). To determine the functional potential of the CD80-CAP in EAE, we first evaluated the efficacy of the CD80-CAP to inhibit responses of in vivo primed T cells. Draining LNC restimulated in vitro with MBP NAc1–11 in the presence of the CD80-CAP at 500 μM concentration (mean Δcpm of 7965 ± 608) exhibited significantly reduced proliferation (Fig. 1,A), IL-12 (Fig. 1,D), and IFN-γ (Fig. 1,E) secretion as compared with the vehicle (mean Δcpm of 13477 ± 3770) or the control peptide (mean Δcpm of 12590 ± 1454) treated cells (Fig. 1,A). The inhibitory effect was almost completely lost at 125 μM concentration of the CD80-CAP. Treatment with the anti-CD80 plus anti-CD86 also resulted in significant inhibition of proliferative responses (mean Δcpm 5880 ± 550) and cytokine secretion (Fig. 1, A, D, and E). Treatment with the CD80-CAP (mean Δcpm 10100 ± 2095) or the control peptide (mean Δcpm 9367 ± 1371) alone in the absence of MBP NAc1–11 did not exhibit any inhibition (data not shown). Visualization of cell division by the shift of CFSE peak to the left showed that fewer CD4+ splenocytes isolated from the PLP139–151 primed SJL/J mice, underwent division following in vitro restimulation with the immunogen in the presence of the CD80-CAP at 500 μM concentration as compared with the untreated or cells cultured in the presence of control peptide (Fig. 1, B and C).

Following in vitro evaluation, the preclinical toxicity of the CD80-CAP was assessed in a mouse model of endotoxemia by LPS injection, which induces a rapid increase in plasma concentrations of TNF-α within 1 to 1.5 h (18). The sharp increase in plasma TNF-α was inhibited by the CD80-CAP (25 μg/kg) administered immediately following LPS (100 μg) injection (Fig. 2,A). Continued suppression of acute response was evident in the decreased plasma concentrations of IFN-γ (Fig. 2,A) and IL-12 (Fig. 2 B) at 8 and 12 h postadministration of LPS, respectively, in the mice treated with CD80-CAP. These results suggest that the CD80-CAP is nontoxic in vivo.

The biological potential of the CD80-CAP during Ag priming in vivo was then investigated in the relapsing remitting model of EAE. SJL mice immunized with PLP139–151 peptide were administered i.v. vehicle or 500 μg of the CD80-CAP/the control peptide on the day of EAE induction. The development of clinical disease was delayed in groups of CD80-CAP treated mice (day 14) as compared with the vehicle (day 10) or control peptide (day 10) treated mice (Fig. 3,A). Furthermore, the severity of clinical disease was significantly lower in mice treated with CD80-CAP (mean clinical score 4.73 ± 0.47) as compared with the vehicle treated mice (mean clinical score 9.8 ± 0.1) or the control peptide treated mice (9.79 ± 0.8 and 9.61 ± 0.614). The average clinical score per day was significantly lower in mice treated with CD80-CAP as compared with the control groups (Fig. 3,B). The decreased severity correlated with reduced inflammatory cytokines, namely IFN-γ and TNF-α and elevated anti-inflammatory cytokine TGF-β in the serum of CD80-CAP treated mice (Fig. 3 C).

The PLP139–151-induced EAE in SJL mice follows a relapsing remitting course secondary to epitope spreading (24). To determine the effects on the number or severity of clinical relapse, we administered a second dose of the CD80-CAP/control peptide (500 μg) on day 21 following disease induction when most animals had gone through one episode of clinical disease. No significant difference was observed between the mice administered additional dose of the CD80-CAP (maximum clinical score 1.33 ± 0.3; n = 4 mice) as compared with the mice treated with a single dose on the day of immunization (maximum clinical score 1.375 ± 0.7; n = 5 mice) (Fig. 3,D). However, mice that received CD80-CAP twice continued to exhibit significantly lower average clinical score per day as compared with the mice receiving similar regimen of vehicle and control peptide (Fig. 3 E). The protection offered by the CD80-CAP lasted for the entire period of observation.

To determine whether the protection mediated by the CD80-CAP is due to modulation of T cell responses, we assessed the functional responses of CD4+ T cells isolated from splenocytes of mice induced R-EAE and treated with PBS/CD80-CAP/control peptide on the day of immunization. Proliferative responses to PLP139–151 was significantly decreased in CD4+ splenocytes from mice treated with the CD80-CAP (mean Δcpm 22841 ± 2500) as compared with the vehicle (mean Δcpm 31500 ± 2174) or the control peptide (mean Δcpm 28564 ± 3602.7) treated mice (Fig. 4,A). The average cpm of unstimulated CD4+ splenocytes was 9037 ± 704.5 (vehicle), 4559 ± 1134 (CD80-CAP), and 8522 ± 634 (control peptide) treated mice (data not shown). No significant difference was observed in the proliferative responses of splenocytes to the irrelevant Ag OVA between the different treatment groups (data not shown). In addition, CD4+ LNC isolated 45 days postimmunization from SJL mice induced EAE and treated with CD80-CAP (mean Δcpm = 9520.5 ± 1663.8) on the day of immunization and 21 days later (clinical score shown in Fig. 3,B), exhibited significantly lower proliferative responses to PLP139–151 as compared with the cells from mice receiving similar treatment regimen of vehicle (mean Δcpm 15521 ± 1976)/control peptide (mean Δcpm 21353 ± 8525) (Fig. 4 B). The average cpm of unstimulated CD4+ LNC were 5992.2 ± 687 (vehicle), 4518 ± 398 (CD80-CAP), and 5486 ± 578 (control peptide) treated mice (data not shown).

We next evaluated the effect of the CD80-CAP on T cell cytokine secretion. Upon restimulation with the PLP139–151, LNC derived from mice induced EAE and treated with the CD80-CAP on the day of immunization secreted significantly reduced Th1 cytokines IFN-γ (7.77 ± 11.5 pg/ml) and IL-12 (117.3 ± 17.4 pg/ml) as compared with the vehicle (69.3 ± 28.7 pg/ml and 368.7 ± 26.2 pg/ml, respectively) or the control peptide (73.4 ± 16.4 pg/ml and 783.2 ± 85.5 pg/ml, respectively) treated mice (Fig. 5, A and B). In addition, LNC from mice treated with CD80-CAP (1848 ± 241 pg/ml) secreted higher IL-10 in response to the PLP139–151 as compared with the vehicle (1293 ± 144 pg/ml) or the control peptide (1628.5 ± 430.8 pg/ml) treated mice (Fig. 5 C). Similar observations of reduced Th1 cytokine secretion following costimulatory blockade has been reported previously (2, 6, 8, 19, 25).

We next determined whether the CD80-CAP facilitates Th1 to Th2 skewing by modulating the specific transcription factors T-bet and GATA-3, respectively (26). CD4+ T cells isolated 10 days postimmunization from mice induced EAE and treated with the CD80-CAP/control peptide/vehicle on the day of disease induction and restimulated in vitro with the PLP139–151 were evaluated for T-bet and GATA-3 mRNA. Quantitative real-time PCR showed that although the levels of T-bet mRNA were equivalent, GATA-3 mRNA was significantly up-regulated in the CD4+ LNC from the CD80-CAP treated mice as compared with the cells from the vehicle or the control peptide treated mice (Fig. 5, D and E).

We next investigated the effect of the CD80-CAP in passive EAE. Naive mice adoptively transferred CD4+CD25 T effector cells from the CD80-CAP treated mice exhibited significantly reduced mean clinical score compared to the mice injected effector T cells from the vehicle or the control peptide treated mice (Fig. 6,A). Peripheral blood collected 8 days posttransfer exhibited reduced proinflammatory cytokines IFN-γ (Fig. 6,B) and TNF-α (Fig. 6,C) and elevated TGF-β (Fig. 6 D) levels in the CD80-CAP treated mice as compared with the control or vehicle treated mice.

CD28-B7 interactions have been shown to differently affect CD4+CD25+ natural regulatory T cells depending upon the nature of the ligand, using CD80/CD86 and the experimental system (27). Treatment with the CD152-Ig has been shown to significantly up-regulate FoxP3 expression in CD4+CD25+ regulatory T cells in experimental autoimmune myocarditis (28). In this experiment, we investigated the effects of the CD80-CAP treatment on CD4+CD25+ regulatory T cells isolated from the splenocytes of mice induced active EAE. We observed that the CD4+CD25+ regulatory T cells from the CD80-CAP treated mice exhibited significantly higher expressions of FoxP3 (Fig. 7, A and C) and CD152 (Fig. 7, B and C) as compared with that from vehicle or control peptide treated mice.

Previously transfer of the CD4+CD25+ regulatory T cells from naive mice has been shown to suppress EAE (29). In this experiment, we observed that the adoptive transfer of CD4+CD25+ regulatory T cells from mice induced EAE and administered CD80-CAP/control peptide/vehicle into groups of naive mice immunized with MBP NAc1–11 resulted in minimal clinical disease in all mice (data not shown). However, interestingly the mice receiving the regulatory T cells isolated from the CD80-CAP treated mice exhibited significantly lower proinflammatory cytokines; TNF-α and IFN-γ, and an elevated amount of anti-inflammatory cytokine, TGF-β in the circulating blood (Fig. 7 D).

The anti-inflammatory potential of glucocorticoids widely used in the management of inflammatory autoimmune diseases like MS, have been largely attributed to its ability to suppress proinflammatory or Th1 cytokines (30). Recently it is reported that the Th1 to Th2/(anti-inflammatory) cytokine skewing by glucocorticoids is mediated by the transcriptional up-regulation of GILZ in activated T cells (16). Glucocorticoids have also been shown to inhibit CD80 expression on APC (31). We investigated whether the suppression of Th1 cytokine and skewing toward Th2 cytokine by the CD80-CAP may be mediated by a similar mechanism. CD4+ T cells from B10.PL mice induced EAE and restimulated in vitro with MBP NAc1–11 in the presence of dexamethasone, anti-CD80, CD80-CAP, or control peptide were assessed for GILZ expression by Western blot (Fig. 8,A) analysis and quantitative real-time PCR (Fig. 8, B and C). We observed that the GILZ expression was significantly up-regulated in the CD4+ T cells stimulated in the presence of dexamethasone or anti-CD80 (Fig. 8,C). Importantly, presence of the CD80-CAP at 500 μM concentration also significantly up-regulated GILZ expression in the Ag-stimulated CD4+ T cells. Stimulation of CD4+ T cells in the presence of the control peptide had no significant effect on the GILZ expression (Fig. 8).

The critical role played by the CD80-CD86/CD28-CD152 pathway in regulating T cell functions at initiation, progression, and reactivation of the immunopathology in EAE and MS is well documented (5, 7). Differences in the pattern, levels, and kinetics of expression of CD80/CD86 molecules in conjunction with differences in the strength and quality of the signals delivered upon binding CD28 or CD152 determines the outcome of the immune response (8). A temporal up-regulation of surface expression of CD80 relative to CD86 on splenic APCs as well as CNS-infiltrating cells has been shown to correlate with disease progression in EAE (4, 6). In this study, we report the therapeutic potential of CD80 costimulatory blockade by a synthetic peptide inhibitor, the CD80-CAP, in R-EAE.

Structurally, the CD80-CAP was designed to mimic the ligand binding domain of CD28/CD152 that engages the hydrophobic pocket of CD80 adopting a PPII helical conformation (10, 12). Previously we have shown that the CD80-CAP competitively inhibits binding of CD152/CD28 with the CD80 and interferes weakly with the CD86-receptor interactions (15). Similar short peptides derived from the interface of other receptor-ligand complexes as conformational mimics have been shown to specifically interfere with the intermolecular binding and inhibit protein-protein interactions (11).

The mouse model of LPS endotoxemia, which induces a rapid increase in plasma concentration of inflammatory cytokines is often used to evaluate the preclinical toxicity of biotechnological agents (17). Our observations of reduced plasma TNF-α, IFN-γ, and IL-12 concentrations in mice receiving CD80-CAP suggested that the CD80-CAP was well tolerated and capable of suppressing inflammatory response.

The advantage of costimulatory blockade in suppressing inflammation is evident in the observed beneficial effects of the CD152-Ig, a fusion protein that blocks B7-CD28/CD152 interactions, in human clinical trials of MS and rheumatoid arthritis (32). In addition to the APC, CD80 is also expressed on activated CD4+ T cells. It has been suggested that the CD80 expressing CD4+ T cells preferentially migrate to the CNS (4). In contrast to the intact mAb, interference of CD80: receptor interactions with the anti-CD80 Fab protected mice against R-EAE (24). Since the CD80-CAP prevents the CD80 interactions by steric hindrance, the observed suppression of R-EAE following CD80-CAP administration is consistent with these findings. The reduced ability of the effector T cells from the CD80-CAP treated mice to transfer disease suggests that the CD80 blockade preferentially inhibits Ag-activated T cells. Although we did not evaluate the functional responses to all encephalitogenic epitopes of PLP, the prolonged suppression of clinical disease even at 7 wk after the CD80-CAP administration and the reduced serum proinflammatory cytokines following adoptive transfer of CD4+CD25 effector T cells from the CD80-CAP treated mice suggest that the protection mediated by the CD80-CAP is relatively long-lasting.

Modulation of CD80/CD86 signaling has been shown to variably affect central and peripheral regulatory T cells (27, 33, 34). Although absence of both CD80 and CD86 significantly inhibits thymic regulatory T cell development, absence of CD80 alone does not critically affect the homeostasis of peripheral regulatory T cells (34, 35, 36). In addition, as compared with CD80 expression, CD86 expressing dendritic cells are more effective in stimulating the proliferation of natural regulatory T cells (27). This may explain our observations of increased FoxP3 expression on CD4+CD25+ T cells in CD80-CAP treated mice because selective inhibition of CD80 binding may allow for increased CD86-receptor interactions between the APC and the peripheral regulatory T cells. CD152 expression has also been shown to play a critical role in peripheral regulatory T cell homeostasis (37). In this context it is interesting to note that the CD4+CD25+ T cells in the CD80-CAP treated mice exhibited increased expression of both FoxP3 and CD152. It has been suggested that polyclonal expansion of regulatory T cells may interfere with the effector T cell migration and prevent EAE (38).

Suppression of Th1 cytokines and skewing toward Th2 response is another mechanism of immunomodulation mediated by the CD28 costimulatory blockade in ameliorating autoimmune diseases (1, 2, 5, 32). The therapeutic effects of glucocorticoids widely used in the management of chronic inflammatory conditions such as MS are largely attributed to the inhibition of NF-κB dependent transcription of proinflammatory/Th1 cytokines (30). GILZ is one of the genes up-regulated by glucocorticoids (23). It binds the p65 subunit of NF-κB, prevents its nuclear translocation inhibiting transactivation of proinflammatory target genes. GILZ expression is down-regulated in activated T cells (39, 40). Furthermore overexpression of GILZ suppresses T-bet, the canonical Th1 transcription factor and up-regulates the Th2 transcriptional factors GATA-3 and STAT6 (16). It has been suggested that the glucocorticoids may also suppress T cell responses by reducing CD80 costimulation (31). We observed that the CD80-CAP treatment elevated GILZ and GATA-3 expression and suppressed T-bet and IFN-γ expression in CD4+ T cells. Collectively our results suggest that the enhanced GILZ expression in activated CD4+ T cells following costimulatory blockade may represent one mechanism that mediate skewing of proinflammatory/Th1 cytokine response to anti-inflammatory Th2 profile, facilitating suppression of autoimmune disease. In this context it is interesting to note that GILZ overexpressing transgenic mice are protected against transfer colitis, a Th1 cytokine mediated model of human inflammatory bowel disease (41).

The integral role played by the CD80-CD86/CD28/CD152 molecules in T cell development, proliferation, differentiation, regulation, and homeostasis imply that the manipulation of this pathway will likely modulate more than one mechanisms of T cell response (9, 35). The therapeutic effects of potential agents that interfere with this pathway in MS will depend on the molecular profile of the pathogenic T cells at the time of administration. Designing mimetic of the interface functional groups of known receptor-ligand complexes is an attractive strategy for developing potential therapeutic agents that interfere with target protein-protein interactions (11, 42). In this study, we show that the CD80 costimulatory blockade by a synthetic hexapeptide suppress R-EAE and that multiple mechanisms, not mutually exclusive, may be involved in mediating the protection. The selective inhibition of CD80-receptor interactions, the rapid binding kinetics, the low molecular weight, and the ability to suppress activated T cell responses in vivo are significant advantages of the CD80-CAP as potential therapeutic agent in MS. Furthermore, the CD80-CAP may provide a significant lead for the design and development of CD80-specific small molecule inhibitors (43).

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported by Grant RG3723A1 from the National Multiple Sclerosis Society (to M.S.).

3

Abbreviations used in this paper: MS, multiple sclerosis; EAE, experimental autoimmune encephalomyelitis; GILZ, glucocorticoid-induced leucine zipper; CDR, complementarity-determining region; LNC, lymph node cell; PPII, polyproline type II; CAP, competitive antagonist peptide; PLP, proteolipid protein; MBP, myelin basic protein; NAc, N-acetylated.

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