Recent evidence indicates that leptin modifies T cell immunity, and may provide a key link between nutritional deficiency and immune dysfunction. To study the influence of leptin on autoimmunity, susceptibility to experimental autoimmune encephalomyelitis induced by immunization with a myelin-derived peptide was examined in leptin-deficient, C57BL/6J-ob/ob mice, with or without leptin replacement, and in wild-type controls. Leptin replacement converted disease resistance to susceptibility in the C57BL/6J-ob/ob mice; this was accompanied by a switch from a Th2 to Th1 pattern of cytokine release and consequent reversal of Ig subclass production. Our findings suggest that leptin is required for the induction and maintenance of an effective proinflammatory immune response in the CNS.

The obese gene product, leptin, is a hormone belonging to the helical cytokine family derived primarily from adipocytes, described as a central mediator of the neuroendocrine pathways involved in the control of food intake, basal metabolism, and reproductive function (1, 2). Recently, leptin has also been shown to modulate the cognate T cell-mediated immune response by signaling through the long isoform of the leptin receptor (ObRb) expressed on the cell surface of mature CD4+ T lymphocytes (3). Specifically, leptin reverses the immunodeficiency and lymphoid organ atrophy, induced by acute starvation in rodents (3, 4). Leptin-deficient obese mice (C57BL/6J-ob/ob) display immune dysfunction similar to that observed in starved animals and malnourished humans, both conditions associated with low circulating leptin concentrations (4, 5, 6). C57BL/6J-ob/ob mice exhibit impaired cell-mediated immunity and thymic atrophy (3, 4, 7). Furthermore, chronic leptin deficiency in these animals leads to reduced in vitro secretion, upon Ag stimulation, of the classical Th1-type proinflammatory cytokines such as IL-2 and IFN-γ and an increased production of IL-4, typical of the Th2 regulatory phenotype (3).

Experimental autoimmune encephalomyelitis (EAE)3 is an animal model of human multiple sclerosis, which can be induced in susceptible strains of mice by immunization with self Ags, derived from CNS myelin (8). The disease is characterized by the generation of autoreactive T cells that traffic to the brain and the spinal cord and initiate injury to CNS myelin, resulting in a chronic or relapsing-remitting paralysis according to the Ag and the strain of mice used (8, 9). Direct evidence for the role of CD4+ T cells in EAE induction has come from adoptive transfer studies, in which myelin Ag-reactive Th1 CD4+ cell lines or clones induce encephalomyelitis and demyelination leading to paralysis following transfer (10). Th1 cytokines are present in inflammatory EAE lesions in the CNS, whereas Th2 cytokines are absent, suggesting that Th1 cytokines play a role in the pathogenesis of the disease (8, 11). Recovery from EAE in mice is associated with an increase in the presence of Th2 cytokines in the CNS (12). Furthermore, IL-4 administration during EAE reduces both the intensity and the progression of the disease (13).

To determine the contribution of leptin to the pathogenesis of the multiple sclerosis-like disease provoked by the immunodominant myelin oligodendrocyte glycoprotein (MOG) peptide, MOG35–55 (14), we tested disease susceptibility in naturally leptin-deficient C57BL/6J-ob/ob mice before and after chronic leptin administration, comparing it with wild-type controls under the same experimental conditions. Both leptin-deficient and control mice are on the C57BL/6J-susceptible genetic background (H-2b) for MOG35–55 peptide-induced EAE (14). We report in this work that, upon immunization with MOG35–55 peptide or after adoptive transfer of pathogenic MOG35–55-specific CD4+ Th1 cells, mice lacking circulating leptin do not develop any neurological impairment. Conversely, only after chronic leptin administration C57BL/6J-ob/ob mice become susceptible to the antigenic peptide or to adoptive transfer of pathogenic T cells. In wild-type controls, leptin supplementation causes a more severe and chronic disease than in the untreated group. These findings show for the first time that EAE may not be induced in the absence of leptin, and that chronic leptin administration enables leptin-deficient mice to develop EAE actively induced by MOG35–55 peptide or adoptively transferred by MOG35–55-specific T cells.

Female C57BL/6J wild-type and C57BL/6J-ob/ob leptin-deficient obese mice 8–10 wk old were obtained from Charles River Italy (Calco, Italy) and from Harlan Italy (Corezzana, Italy). Experiments were performed under an approved protocol in accordance with the animal use guidelines of the Istituto Superiore di Sanita (Rome, Italy). Wild-type and leptin-deficient mice were age matched for individual experiments and were group-housed two to six mice per standard cage according to the different experimental protocol, with a 12-h light/dark cycle. Paralyzed mice were afforded easier access to food and water to prevent dehydration.

The peptide used in this study is the immunodominant MOG35–55 peptide (MEVGWYRSPFSRVVHLYRNGK) (14). It was synthesized by PRIMM s. r. l. (Milan, Italy); purity was assessed by HPLC (>97% pure), and amino acid composition was verified by mass spectrometry. MOG35–55 peptide batches for in vivo and in vitro assays were all from one preparation, initially solubilized in LPS-free saline solution at 4 mg/ml concentration, and stored at −80°C.

Mouse recombinant leptin (rleptin) was purchased from R&D Systems Europe (Oxon, U.K.); purity was >97%, as determined by SDS-PAGE and visualized by silver staining analysis. The endotoxin level was <0.1 ng/μg of leptin, as determined by the Limulus amebocyte lysate method. Mice comprised three groups (n = 6–11 per group) for C57BL/6J-ob/ob leptin-deficient obese mice (all housed in pairs) and two groups (n = 6–10 per group) for C57BL/6J normal age- and sex-matched control mice (housed two to six mice/cage). For the active disease induction, mice were injected i.p. with rleptin or PBS starting 10 days before the immunization and continuing over a period of 40 days; for adoptively induced disease, mice were treated starting 3 days before the transfer of T cells and continuing over a period of 30 days. Of the groups of leptin-deficient mice, one was injected with 200 μl of PBS twice daily (at 10:00 a.m. and 6:00 p.m.) and allowed to feed ad libitum; the second group was injected with murine rleptin (0.5 μg/g initial body weight twice daily in 200 μl volume i.p., for a total of 1 μg/g/day of rleptin); and the third group was pair fed to the food intake of the rleptin-treated mice and received twice daily injections of PBS according to the same schedule (4). Of the two groups of C57BL/6J wild-type mice, one was injected with PBS twice daily and allowed to feed ad libitum; the second was injected twice daily with rleptin according to the same schedule of obese mice. All mice were weighed and their food intake was recorded daily.

For actively induced EAE, mice were immunized s.c. in the flank with 100 μl of CFA (Difco Laboratories, Detroit, MI) emulsified with 200 μg of MOG35–55 peptide on days 0 and 7, and with 200 ng of pertussis toxin (Sigma, St. Louis, MO) i.p., on days 0, 1, 7, and 8. Control mice (n = 5 mice per group) were injected with CFA emulsified with PBS plus pertussis toxin, according to the same schedule (14). For adoptively transferred EAE (14), 9–10 female donor C57BL/6J mice (6–8 wk old) were primed s.c. with 300 μg of MOG35–55 peptide in CFA distributed over four sites. After 9–10 days, draining lymph nodes (axillary and inguinal) and spleens were harvested, homogenized into a single cell suspension, and cultured separately in vitro in 24-well plates (Falcon; Becton Dickinson, Franklin Lakes, NJ) (8 × 106 cells/well) with RPMI 1640 medium (Life Technologies, Gaithersburg, MD) supplemented with 10% FCS (Life Technologies), 2 mM l-glutamine (Life Technologies), 0.1 mM nonessential amino acids (Life Technologies), 1 mM sodium pyruvate (Life Technologies), 50 μM 2-ME (Sigma), 100 U/ml penicillin, 100 μg/ml streptomycin (Life Technologies), and 25 μg/ml of MOG35–55 peptide. After 4 days in culture and addition to medium of 2 U/ml of rIL-2 (Roche Biochemicals, Monza, Italy), the cells were harvested and centrifuged over a Ficoll gradient (Pharmacia Biotech, Uppsala, Sweden) to remove debris. Recipient syngeneic naive female leptin-deficient or wild-type control mice in the different conditions of treatment after 500 rad irradiation were i.v. injected with 2.5 × 107 T cells in a final volume of 500 μl of PBS. Mice also received 200 ng of pertussis toxin immediately after cell transfer and 1 day later. Naive C57BL/6J-ob/ob and C57BL/6J control mice (n = 5 mice per group) for adoptive transfers, after 500 rad irradiation, were injected with PBS alone or T cells from unprimed animals plus pertussis toxin, according to the same schedule.

Individual mice were observed daily for clinical signs of disease for up to 40 days after immunization and up to 30 days after adoptive transfer. Mice were weighed and scored daily according to the clinical severity of symptoms on a scale of 0 to 6 (14) by a “blinded” to mice identity experimenter (A.D.G.), with 0.5 points for intermediate clinical findings: grade 0, no abnormality; grade 0.5, partial loss of tail tonicity, assessed by inability to curl the distal end of the tail; grade 1, reduced tail tone or slightly clumsy gait; grade 2, tail atony, moderately clumsy gait, impaired righting ability, or any combination of these signs; grade 3, hind limb weakness or partial paralysis; grade 4, complete hind limb paralysis or fore limb weakness; grade 5, tetraplegia or moribund state; grade 6, death. The data were plotted as daily mean clinical score for all animals in a particular treatment group. Scores of asymptomatic mice (score = 0) were included in the calculation of the daily mean clinical score for each group. The brains and spinal cords were dissected between 25 and 35 days after immunization and fixed in 10% Formalin. Paraffin-embedded sections of 5 μm thickness were cut from optic nerve, forebrain, cerebellum, hind brain, cervical, thoracic, lumbar, and sacral spinal cord regions and stained with hematoxylin-eosin and Luxol fast blue (Sigma) for evidence of inflammation and demyelination. Sections from 4–10 segments per mouse were examined blindly by one investigator (A.D.T.) using a published scoring system for inflammation and demyelination, respectively (15).

DTH responses to MOG35–55 peptide during induction of disease were also quantitated using a time-dependent (12–72 h) footpad-swelling assay (16). Briefly, mice previously sensitized with MOG35–55 in CFA were challenged by s.c. injection of 25 μg of MOG35–55 (in 50 μl PBS) into the right hind footpad. PBS alone was injected into the left footpad to serve as control for measurements. As negative control, we used unimmunized mice (sensitized with CFA alone). Footpad thickness was measured 12, 24, 48, and 72 h after challenge by a “blinded” to sample identity experimenter (A.D.G.), using a caliper-type engineer’s micrometer. The footpad-swelling response was calculated as the thickness of the right footpad (receiving Ag) minus the baseline thickness of the left footpad (receiving PBS).

Spleen and lymph node cells were obtained from mice 35 days after MOG35–55 sensitization, dissociated into single cell suspension, and cultured for proliferation assays in flat-bottom 96-well microtiter plates (Falcon) at a density of 5 × 105 viable cells/well in a total volume of 200 μl of RPMI 1640 medium (Life Technologies), supplemented with 2% FCS (Life Technologies), 2 mM l-glutamine (Life Technologies), 0.1 mM nonessential amino acids (Life Technologies), 1 mM sodium pyruvate (Life Technologies), 50 μM 2-ME (Sigma), 100 U/ml penicillin, and 100 μg/ml streptomycin (Life Technologies). Cells were cultured at 37°C in 100% humidity and 5% CO2 in the presence or absence of varying concentrations of MOG35–55 peptide (from 0 to 50 μg/ml peptide). As control for proliferation, anti-CD3 Ab stimulation (2C11 hybridoma supernatant, diluted 1/100) was also performed 48–60 h after initiation of culture cell supernatants (100 μl) were removed from single well and frozen at −80°C for cytokine assay. IFN-γ and IL-4 were measured by ELISA developed in our laboratory using cytokine-specific capture and detection Abs (PharMingen, San Diego, CA), according to the manufacturer’s instructions (Abs R4-6A2 and XMG1.2 for detection of IFN-γ; Abs BVD4-1D11 and BVD6-24G2 for the detection of IL-4). Standard curves for each assay were generated using recombinant mouse cytokines (IFN-γ and IL-4; PharMingen), and the concentration of the cytokines in the cell supernatants was determined by extrapolation from the appropriate standard curve. The lower limits of detection for each assay were: <2 pg/ml for IFN-γ; <0.6 pg/ml for IL-4. The remaining cells were incubated for an additional 16 h, pulsed with 0.5 μCi/well of [3H]thymidine (Amersham Pharmacia Biotech, Piscataway, NJ), harvested on glass-fiber filters using a Tomtec (Orange, CT) 96-well cell harvester, and counted in a 1205 Betaplate liquid scintillation counter (Wallac, Gaithersburg, MD). Results are expressed as mean cpm ± SD from duplicate cultures.

Serum samples obtained from tail veins during the period of observation of the animals (1 day before the immunization, and then 7, 25, and 35 days after the immunization). All samples were tested for MOG35–55-specific total IgG (1/100 dilution), IgG1 (1/100 dilution), and IgG2a (1/100 dilution) on MOG35–55 peptide-coated 96-well ELISA plates (Corning Glass, Corning, NY). For total IgG measurements, Mouse ExtrAvidin Staining Kit (Sigma) and for IgG1/IgG2a subclasses biotin anti-IgG1 (clone A85-1) and anti-IgG2a (clone Igh-1b) mouse Abs (PharMingen) were used (17). Briefly, 100 μl of MOG35–55 peptide was added to 96-well ELISA plates at final concentration of 10 μg/ml in carbonate buffer, pH 8.2. After 16 h at 4°C, the plates were washed in PBS, blocked with 200 μl of PBS/10% FCS for 2 h, and repeatedly washed. Diluted sera in PBS-Tween/10% FCS were added at 100 μl/well for 2 h at room temperature. After five washes, anti-mouse subclasses-specific biotin-conjugated Abs at 2 μg/ml in PBS-Tween/10% FCS were added for 45 min. After six washes, 1/1000 diluted ExtrAvidin-peroxidase (Sigma) was added for 30 min. The reaction was developed with Sigma-Fast OPD (o-phenylenediaminedihydrochloride, peroxidase substrate) (Sigma) and read after 30 min at 450 or 492 nm after stopping with 1 M HCl in an ELISA plates reader (Bio-Rad Laboratories, Hercules, CA).

Analyses were performed using Mann-Whitney U test (for unpaired two group analyses) and Kruskal-Wallis ANOVA test (for three or more group analyses). Results are expressed as mean ± SD; p values <0.05 were considered to be statistically significant.

We first tested the ability of MOG35–55 peptide to induce EAE in leptin-deficient C57BL/6J-ob/ob mice and wild-type age- and sex-matched C57BL/6J controls with and without leptin treatment. None of the PBS-treated leptin-deficient mice ad libitum fed or pair fed to the leptin-treated group developed any sign of clinically evident disease after active immunization with MOG35–55 peptide (Table I; Fig. 1,a). rLeptin administration, starting 10 days before the immunization and continuing over a period of 40 days, restored disease susceptibility in C57BL/6J-ob/ob mice, comparable with that of the PBS-treated C57BL/6J wild-type controls (Table I; Fig. 1, a and b). Furthermore, C57BL/6J mice treated with rleptin exhibited a significantly more severe and chronic disease than PBS-treated animals, as indicated by a more severe clinical score and mortality (Table I; Fig. 1,b). None of the animals that survived EAE showed signs of recovery at the termination of the experiments (40 days). The possibility that the adjuvants used for the immunization were pathogenic in these mice was ruled out by the fact that none of the C57BL/6J-ob/ob or C57BL/6J mice (n = 5 for each group) injected with CFA, alone and with pertussis toxin, developed disease (data not shown). Consistent with these results, upon histological examination of the CNS tissues from both the leptin-deficient PBS- or PBS-pair fed-treated mice, no perivascular infiltrates or signs of demyelination were found in the brain and spinal cord (Fig. 2, a-c). The absence of inflammatory foci in the C57BL/6J-ob/ob animals eliminated the occurrence of silent disease. In contrast, the rleptin C57BL/6J-ob/ob-treated group showed extensive mononuclear cell infiltration throughout the brain and spinal cord with signs of demyelination (Fig. 2, a-c). The frequency and the degree of inflammation and demyelination were comparable between the C57BL/6J-ob/ob treated with rleptin and the wild-type PBS-treated control group (Fig. 2,c). As for the clinical score, administration of rleptin to C57BL/6J wild-type controls significantly increased the numbers of inflammatory foci and demyelination when compared with that of PBS-treated mice (Fig. 2 c). No cellular infiltration or demyelination was observed in control animals receiving adjuvants and pertussis toxin only (data not shown).

Table I.

Neurological impairment in leptin-deficient and wild-type control mice immunized with MOG35–55 peptide and treated or not with rleptina

MiceIncidenceOnset (range) (days)MortalityDuration (days)Clinical ScorebInitial Body Weight (g)Final Body Weight (g)
C57BL/6J-ob/ob PBS 0/11 (0.0%) 0.0 (0–0) 0/11 (0.0%) 0.0 0.0 49.3 ± 4.5 58.3 ± 6.4 
C57BL/6J-ob/ob PBS-pair fed 0/11 (0.0%) 0.0 (0–0) 0/11 (0.0%) 0.0 0.0 47.4 ± 5.5 33.1 ± 2.8 
C57BL/6J-ob/ob rleptin 10/11 (90.9%) 20.0 ± 1 (19–22) 0/11 (0.0%) 21.0c 1.5 ± 0.9c 46.7 ± 4.3 22.6 ± 2.7c 
C57BL/6J PBS 9/10 (90.0%) 19.5 ± 1.5 (18–24) 1/10 (10.0%) 22.0d 1.9 ± 1.6d 20.5 ± 1.7 19.5 ± 2.3 
C57BL/6J rleptin 10/10 (100.0%) 16.0 ± 2.7 (13–19) 2/10 (20.0%) 27.0e 3.9 ± 1.1e 21.1 ± 2.3 17.4 ± 2.5e 
MiceIncidenceOnset (range) (days)MortalityDuration (days)Clinical ScorebInitial Body Weight (g)Final Body Weight (g)
C57BL/6J-ob/ob PBS 0/11 (0.0%) 0.0 (0–0) 0/11 (0.0%) 0.0 0.0 49.3 ± 4.5 58.3 ± 6.4 
C57BL/6J-ob/ob PBS-pair fed 0/11 (0.0%) 0.0 (0–0) 0/11 (0.0%) 0.0 0.0 47.4 ± 5.5 33.1 ± 2.8 
C57BL/6J-ob/ob rleptin 10/11 (90.9%) 20.0 ± 1 (19–22) 0/11 (0.0%) 21.0c 1.5 ± 0.9c 46.7 ± 4.3 22.6 ± 2.7c 
C57BL/6J PBS 9/10 (90.0%) 19.5 ± 1.5 (18–24) 1/10 (10.0%) 22.0d 1.9 ± 1.6d 20.5 ± 1.7 19.5 ± 2.3 
C57BL/6J rleptin 10/10 (100.0%) 16.0 ± 2.7 (13–19) 2/10 (20.0%) 27.0e 3.9 ± 1.1e 21.1 ± 2.3 17.4 ± 2.5e 
a

Mice were immunized with 200 μg of MOG35–55 peptide emulsified in CFA and observed daily for 40 days. Treatment with PBS or rleptin was performed starting 10 days before the immunization and continuing over a period of 40 days. None of the control mice (n = 5 for each group) immunized with CFA alone with pertussis toxin developed disease (data not shown). Data are cumulated and averaged from two separate experiments and they are presented as mean ± SD.

b

Data are presented as mean of the group clinical score of all the animals of that group.

c

p < 0.001 compared with C57BL/6J-ob/ob PBS and C57BL/6J-ob/ob PBS-pair fed.

d

NS compared with C57BL/6J-ob/ob rleptin.

e

p < 0.01 compared with C57BL/6J PBS.

FIGURE 1.

Leptin enables MOG35–55 peptide and adoptively transferred MOG-specific CD4+ T cells to induce EAE in disease-resistant C57BL/6J-ob/ob mice. a, Mean clinical course and severity of EAE disease in C57BL/6J-ob/ob leptin-deficient obese mice respectively treated with PBS, PBS-pair fed, or rleptin. None of the leptin-deficient mice developed clinical symptoms. Only the rleptin-treated group developed clinical signs of disease. Data are representative of two independent experiments with similar results (n = 6 mice per group/trial) and are presented as mean of the group clinical score of all the animals of that group ± SD. b, Mean clinical course and severity in wild-type control C57BL/6J PBS- or rleptin-treated mice. PBS-treated mice showed a similar disease score and severity to leptin-deficient mice after rleptin administration. rLeptin-treated wild-type animals had a more severe clinical course than PBS-injected animals. Data are representative of two independent experiments with similar results (n = 5 mice per group). In active induction of disease, mice were treated with rleptin starting 10 days before immunization until day 40. c, Adoptive transfer of EAE disease in C57BL/6J-ob/ob mice was present only after rleptin administration. Leptin-deficient mice treated with PBS or PBS-pair fed were resistant to EAE induction even when passively transferred with 2.5 × 107 MOG35–55-specific encephalitergic T cells (n = 6 mice per group). d, Adoptive transfer of EAE disease in C57BL/6J wild-type mice showed classical disease progression in PBS-injected mice and an increase in disease severity and progression in rleptin-treated mice (n = 5 mice per group). In the adoptive transfer of disease, mice were injected with rleptin starting 3 days before the transfer until day 25. Doses of rleptin used were 1 μg/g/day of initial body weight.

FIGURE 1.

Leptin enables MOG35–55 peptide and adoptively transferred MOG-specific CD4+ T cells to induce EAE in disease-resistant C57BL/6J-ob/ob mice. a, Mean clinical course and severity of EAE disease in C57BL/6J-ob/ob leptin-deficient obese mice respectively treated with PBS, PBS-pair fed, or rleptin. None of the leptin-deficient mice developed clinical symptoms. Only the rleptin-treated group developed clinical signs of disease. Data are representative of two independent experiments with similar results (n = 6 mice per group/trial) and are presented as mean of the group clinical score of all the animals of that group ± SD. b, Mean clinical course and severity in wild-type control C57BL/6J PBS- or rleptin-treated mice. PBS-treated mice showed a similar disease score and severity to leptin-deficient mice after rleptin administration. rLeptin-treated wild-type animals had a more severe clinical course than PBS-injected animals. Data are representative of two independent experiments with similar results (n = 5 mice per group). In active induction of disease, mice were treated with rleptin starting 10 days before immunization until day 40. c, Adoptive transfer of EAE disease in C57BL/6J-ob/ob mice was present only after rleptin administration. Leptin-deficient mice treated with PBS or PBS-pair fed were resistant to EAE induction even when passively transferred with 2.5 × 107 MOG35–55-specific encephalitergic T cells (n = 6 mice per group). d, Adoptive transfer of EAE disease in C57BL/6J wild-type mice showed classical disease progression in PBS-injected mice and an increase in disease severity and progression in rleptin-treated mice (n = 5 mice per group). In the adoptive transfer of disease, mice were injected with rleptin starting 3 days before the transfer until day 25. Doses of rleptin used were 1 μg/g/day of initial body weight.

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FIGURE 2.

Inflammatory and demyelinated lesions in C57BL/6J-ob/ob leptin-deficient and wild-type control mice treated with PBS, PBS-pair fed, or rleptin. a, Brain parenchyma from ad libitum fed PBS-, PBS-pair fed-, and rleptin-treated C57BL/6J-ob/ob mice, showing lymphocytic cuffing around blood vessels with extensive infiltration of mononuclear cells consisting of lymphocytes and macrophages only after treatment with rleptin. Hematoxylin and eosin (H&E). Magnification, ×100 and ×400 for insets. b, Spinal cord parenchyma according to a. Hematoxylin and eosin (H&E). Magnification, ×100 and ×400 for insets. c, Summary of the inflammatory and demyelinated lesions in brain and spinal cord of leptin-deficient and wild-type mice. Results are expressed as the mean histological score ± SD (n = 4–10 sections/mouse). ∗, p < 0.05 compared with C57BL/6J-ob/ob PBS or PBS-pair fed; ∗∗, p < 0.05 compared with C57BL/6J PBS.

FIGURE 2.

Inflammatory and demyelinated lesions in C57BL/6J-ob/ob leptin-deficient and wild-type control mice treated with PBS, PBS-pair fed, or rleptin. a, Brain parenchyma from ad libitum fed PBS-, PBS-pair fed-, and rleptin-treated C57BL/6J-ob/ob mice, showing lymphocytic cuffing around blood vessels with extensive infiltration of mononuclear cells consisting of lymphocytes and macrophages only after treatment with rleptin. Hematoxylin and eosin (H&E). Magnification, ×100 and ×400 for insets. b, Spinal cord parenchyma according to a. Hematoxylin and eosin (H&E). Magnification, ×100 and ×400 for insets. c, Summary of the inflammatory and demyelinated lesions in brain and spinal cord of leptin-deficient and wild-type mice. Results are expressed as the mean histological score ± SD (n = 4–10 sections/mouse). ∗, p < 0.05 compared with C57BL/6J-ob/ob PBS or PBS-pair fed; ∗∗, p < 0.05 compared with C57BL/6J PBS.

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In separate experiments, we also found that leptin was able to affect the adoptive transfer of disease by Ag-specific T lymphocytes, derived from susceptible C57BL/6J mice previously sensitized to MOG35–55 and injected into the tail vein of naive mice irradiated with 500 rad. As for actively induced disease, leptin-deficient mice treated with PBS or PBS-pair fed mice showed complete resistance to EAE induction when adoptively transferred with 2.5 × 107 MOG35–55-specific encephalitogenic T cells (Table II; Fig. 1,c). Treatment of recipient mice with rleptin, starting 3 days before the transfer and continuing until day 25, converted leptin-deficient mice to a state of susceptibility for the adoptively induced disease (Table II; Fig. 1,c). Wild-type C57BL/6J PBS-treated mice showed a frequency of disease and clinical scores similar to that of C57BL/6J-ob/ob treated with rleptin. As with the actively induced disease, there was a statistically significant increase in clinical score and cellular infiltrates in the C57BL/6J mice injected with rleptin, when compared with C57BL/6J PBS-treated mice (Table II; Fig. 1 d). Histological examination of brain and spinal cord showed that adoptive transfer of encephalitogenic T cells produced perivascular and parenchymal infiltration in the CNS tissues of the C57BL/6J-ob/ob mice treated with rleptin, whereas no infiltration was seen in either C57BL/6J-ob/ob PBS or PBS-pair fed animals (data not shown), confirming resistance to the adoptively transferred disease. The possibility that the passive transfers were pathogenic in these mice was ruled out by the fact that none of the C57BL/6J-ob/ob or C57BL/6J mice (n = 5 for each group) injected with PBS or T cells from unprimed animals and with pertussis toxin developed disease (data not shown).

Table II.

Neurological impairment in leptin-deficient and wild-type control mice treated or not with rleptin and adoptively transferred with MOG35–55 peptide-specific wild-type T cellsa

MiceIncidenceOnset (range) (days)MortalityDuration (days)Clinical ScoreInitial Body Weight (g)Final Body Weight (g)
C57BL/6J-ob/ob PBS 0/6 (0.0%) 0.0 (0–0) 0/6 (0.0%) 0.0 0.0 50.7 ± 1.9 53.0 ± 0.7 
C57BL/6J-ob/ob PBS-pair fed 0/6 (0.0%) 0.0 (0–0) 0/6 (0.0%) 0.0 0.0 51.5 ± 1.7 37.9 ± 0.5 
C57BL/6J-ob/ob rleptin 6/6 (100.0%) 7.1 ± 1.8 (5–10) 0/6 (0.0%) 18.0b 1.9 ± 0.5b 50.4 ± 2.0 24.8 ± 1.2b 
C57BL/6J PBS 6/6 (100.0%) 8.3 ± 2.0 (5–10) 0/6 (0.0%) 18.0c 1.8 ± 0.4c 21.2 ± 1.0 21.6 ± 2.0 
C57BL/6J rleptin 6/6 (100.0%) 5.2 ± 0.9 (4–7) 1/6 (16.7%) 22.0d 2.8 ± 0.4d 21.5 ± 1.5 18.9 ± 0.8d 
MiceIncidenceOnset (range) (days)MortalityDuration (days)Clinical ScoreInitial Body Weight (g)Final Body Weight (g)
C57BL/6J-ob/ob PBS 0/6 (0.0%) 0.0 (0–0) 0/6 (0.0%) 0.0 0.0 50.7 ± 1.9 53.0 ± 0.7 
C57BL/6J-ob/ob PBS-pair fed 0/6 (0.0%) 0.0 (0–0) 0/6 (0.0%) 0.0 0.0 51.5 ± 1.7 37.9 ± 0.5 
C57BL/6J-ob/ob rleptin 6/6 (100.0%) 7.1 ± 1.8 (5–10) 0/6 (0.0%) 18.0b 1.9 ± 0.5b 50.4 ± 2.0 24.8 ± 1.2b 
C57BL/6J PBS 6/6 (100.0%) 8.3 ± 2.0 (5–10) 0/6 (0.0%) 18.0c 1.8 ± 0.4c 21.2 ± 1.0 21.6 ± 2.0 
C57BL/6J rleptin 6/6 (100.0%) 5.2 ± 0.9 (4–7) 1/6 (16.7%) 22.0d 2.8 ± 0.4d 21.5 ± 1.5 18.9 ± 0.8d 
a

Mice were adoptively transferred with 2.5 × 107 MOG35–55 peptide specific highly encephalitergic T cells obtained from draining lymph nodes and spleens of C57BL/6J wild-type donors, previously immunized with MOG35–55 peptide. Mice were observed daily for 30 days. Treatment with PBS or rleptin was performed starting 3 days before the adoptive transfer and continuing over a period of 25 days. None of the control mice (n = 5 for each group) adoptively transferred with PBS or T cells from unprimed animals and with pertussis toxin developed disease (data not shown). Data are cumulated and averaged from one experiment and they are presented as mean ± SD.

b

p < 0.01 compared with C57BL/6J-ob/ob PBS and C57BL/6J-ob/ob PBS-pair fed.

c

NS compared with C57BL/6J-ob/ob rleptin.

d

p < 0.05 compared with C57BL/6J PBS.

To determine the nature of the in vivo T cell response against MOG35–55 peptide in leptin-deficient mice, DTH reactions (16) were performed in all groups of mutant and wild-type control mice treated or not with rleptin. Seven days after priming with the MOG35–55 peptide emulsified in CFA, mice were challenged with 25 μg of MOG35–55 injected intradermally in the footpad. The degree of local footpad swelling was measured as a readout for the DTH reaction. Typical DTH kinetics were observed with footpad swelling peaking between 24 and 48 h and subsiding after 72–96 h. DTH responses to the MOG35–55-priming epitope were absent in both leptin-deficient C57BL/6J-ob/ob PBS- and PBS-pair fed-treated mice, whereas the ones treated with rleptin exhibited a significant DTH response (Fig. 3,a), similar to that observed in PBS-treated wild-type mice (Fig. 3,b). C57BL/6J mice injected with rleptin showed a statistically significant increase in the DTH response when compared with the PBS-treated group (Fig. 3 b). None of the control mice injected with CFA alone developed any significant anti-MOG35–55 peptide DTH reaction (not shown).

FIGURE 3.

Effect of leptin treatment on anti-MOG35–55 DTH reaction in leptin-deficient and wild-type control mice measured as footpad swelling. a, Seven days after sensitization with MOG35–55 in CFA, mice were challenged by injection of MOG35–55 into the footpad and then assessed for footpad-swelling assay 12, 24, 48, and 72 h later. Data are representative of two independent experiments with similar results, showing the means ± SD of footpad-swelling responses. ∗, p < 0.001 compared with C57BL/6J-ob/ob PBS or PBS-pair fed. b, DTH reaction in wild-type mice treated with rleptin or PBS. Data are representative of two independent experiments, showing the means ± SD of footpad-swelling responses. ∗∗, p < 0.05 compared with C57BL/6J PBS.

FIGURE 3.

Effect of leptin treatment on anti-MOG35–55 DTH reaction in leptin-deficient and wild-type control mice measured as footpad swelling. a, Seven days after sensitization with MOG35–55 in CFA, mice were challenged by injection of MOG35–55 into the footpad and then assessed for footpad-swelling assay 12, 24, 48, and 72 h later. Data are representative of two independent experiments with similar results, showing the means ± SD of footpad-swelling responses. ∗, p < 0.001 compared with C57BL/6J-ob/ob PBS or PBS-pair fed. b, DTH reaction in wild-type mice treated with rleptin or PBS. Data are representative of two independent experiments, showing the means ± SD of footpad-swelling responses. ∗∗, p < 0.05 compared with C57BL/6J PBS.

Close modal

We examined whether the absence of leptin in C57BL/6J-ob/ob mice affected the activation and cytokine secretion of MOG35–55-specific T cells in vitro (18). The T cell response to MOG35–55 peptide was tested on draining lymph node and spleen cells, taken from all groups of mice 35 days after immunization and cultured in the presence or absence of different concentrations of Ag. As shown in Fig. 4, a–c, lymph node cells derived from leptin-deficient PBS- or PBS-pair fed-treated mice showed very low levels of proliferation and IFN-γ production, but consistent amounts of IL-4 when exposed to MOG35–55 peptide. rLeptin administration in these mice increased the proliferative response to a level 11-fold higher than untreated obese control mice (Fig. 4, a and d, and b and e, respectively), whereas IFN-γ secretion was increased 21-fold and IL-4 was inhibited 11-fold when compared with obese PBS or PBS-pair fed groups of mice (Fig. 4, b and c). Proliferation, IFN-γ, and IL-4 levels in rleptin-treated obese mice were comparable with those observed in C57BL/6J PBS-treated mice. Viability and capacity of T cells to respond to a polyclonal TCR-mediated stimulation were also assessed by anti-CD3-induced activation (Fig. 4, a, b, and c, inset graphs). In wild-type C57BL/6J mice, rleptin did not alter proliferative responses during either MOG35–55- or anti-CD3-induced proliferation (Fig. 4,d, and inset graph filled symbol), while rleptin did increase IFN-γ and reduce IL-4 secretion (Fig. 4, e and f, and inset graph filled symbol). Similar results were also observed using spleen cells as the responder population (not shown).

FIGURE 4.

MOG35–55 peptide-specific and anti-CD3 proliferation and cytokine profiles in leptin-deficient and wild-type mice treated with rleptin or PBS. a–c, Ag-specific proliferative, IFN-γ, and IL-4 responses, respectively, of lymph node cells from C57BL/6J-ob/ob mice treated with PBS, PBS-pair fed, or rleptin. d–f, Ag-specific proliferative, IFN-γ, and IL-4 responses, respectively, of lymph node cells from C57BL/6J wild-type mice treated with PBS and rleptin. Insets for each graph represent parallel polyclonal TCR-dependent anti-CD3-induced T cell proliferation, IFN-γ and IL-4 secretion of the same lymph node cells for all the single conditions tested. Lymph node cells (5 × 105/well) were taken 35 days after immunization and cultured in the absence or presence of varying concentrations of MOG35–55 peptide. Data are presented as means ± SD of [3H]thymidine incorporation (cpm) for proliferation and as means ± SD of pg/ml cytokines secretion, all tested in duplicate. These data are representative of two independent experiments with similar results. In all cases, SD was less than 10% of the values. Similar results were obtained also with spleen cells as responders (not shown).

FIGURE 4.

MOG35–55 peptide-specific and anti-CD3 proliferation and cytokine profiles in leptin-deficient and wild-type mice treated with rleptin or PBS. a–c, Ag-specific proliferative, IFN-γ, and IL-4 responses, respectively, of lymph node cells from C57BL/6J-ob/ob mice treated with PBS, PBS-pair fed, or rleptin. d–f, Ag-specific proliferative, IFN-γ, and IL-4 responses, respectively, of lymph node cells from C57BL/6J wild-type mice treated with PBS and rleptin. Insets for each graph represent parallel polyclonal TCR-dependent anti-CD3-induced T cell proliferation, IFN-γ and IL-4 secretion of the same lymph node cells for all the single conditions tested. Lymph node cells (5 × 105/well) were taken 35 days after immunization and cultured in the absence or presence of varying concentrations of MOG35–55 peptide. Data are presented as means ± SD of [3H]thymidine incorporation (cpm) for proliferation and as means ± SD of pg/ml cytokines secretion, all tested in duplicate. These data are representative of two independent experiments with similar results. In all cases, SD was less than 10% of the values. Similar results were obtained also with spleen cells as responders (not shown).

Close modal

Serum levels of MOG35–55 peptide-specific Ab were also tested 7, 25, and 35 days after immunization using an ELISA. We measured MOG35–55-specific total IgG and IgG1/IgG2a subclasses typical of an in vivo switch toward a Th2 or a Th1 response, respectively (19, 20). Significant levels of anti-MOG35–55 IgG were present in all injected mice, regardless of their leptin phenotype (Fig. 5, a and b). rLeptin treatment significantly increased total IgG titers on days 7 and 25 for leptin-deficient mice, and on days 7, 25, and 35 in wild-type animals. More specifically, IgG1 levels were higher in C57BL/6J-ob/ob PBS or PBS-pair fed animals compared with those found after rleptin administration (Fig. 5,c). In wild-type animals, IgG1 levels were slightly but significantly reduced by rleptin treatment (Fig. 5,d). In contrast, peptide-specific IgG2a levels were absent at all the time points in leptin-deficient mice, but were markedly increased after rleptin administration (Fig. 5,e). In wild-type control mice, IgG2a levels were also increased by rleptin injection, although to a lesser degree (Fig. 5 f).

FIGURE 5.

MOG35–55-specific Ab response in MOG35–55 peptide-sensitized leptin-deficient and wild-type animals before and after treatment with rleptin. a and b, MOG35–55-specific total IgG titer in serum from C57BL/6J-ob/ob mice treated with PBS, PBS-pair fed, rleptin, or C57BL/6J PBS and rleptin treated, respectively. c and d, MOG35–55-specific IgG1 subclass titer in serum of all group of mice according to the same schedule as a and b. e and f, MOG35–55-specific IgG2a subclass titer in serum of all groups of mice according to the same schedule as a and b. Serum was collected 1 day before (preimmune) and 7, 25, and 35 days after the sensitization and tested by ELISA (see Materials and Methods) with MOG35–55-coated plates. Each bar represents the mean absorbance values ± SD of serum from three to six mice per group, each tested in duplicate at 1/100 dilution. ∗, p < 0.01 compared with C57BL/6J-ob/ob PBS or PBS-pair fed. ∗∗, p < 0.05 compared with C57BL/6J PBS.

FIGURE 5.

MOG35–55-specific Ab response in MOG35–55 peptide-sensitized leptin-deficient and wild-type animals before and after treatment with rleptin. a and b, MOG35–55-specific total IgG titer in serum from C57BL/6J-ob/ob mice treated with PBS, PBS-pair fed, rleptin, or C57BL/6J PBS and rleptin treated, respectively. c and d, MOG35–55-specific IgG1 subclass titer in serum of all group of mice according to the same schedule as a and b. e and f, MOG35–55-specific IgG2a subclass titer in serum of all groups of mice according to the same schedule as a and b. Serum was collected 1 day before (preimmune) and 7, 25, and 35 days after the sensitization and tested by ELISA (see Materials and Methods) with MOG35–55-coated plates. Each bar represents the mean absorbance values ± SD of serum from three to six mice per group, each tested in duplicate at 1/100 dilution. ∗, p < 0.01 compared with C57BL/6J-ob/ob PBS or PBS-pair fed. ∗∗, p < 0.05 compared with C57BL/6J PBS.

Close modal

This study examines the role of leptin in the activation of autoreactive T cells in vivo, using leptin-deficient C57BL/6J-ob/ob mice before and after rleptin administration. The data presented in this work provide evidence that, under the experimental conditions used, leptin is required for the induction and progression of autoimmune-mediated demyelination induced by MOG35–55 in C57BL/6J strain of mice. Leptin deficiency had a protective role in that C57BL/6J-ob/ob animals were resistant to both the MOG35–55-induced and adoptively transferred disease; conversely, systemic administration of rleptin rendered these mice susceptible to EAE induction. In wild-type mice, rleptin treatment increased disease severity and inflammatory cell infiltration in the brain in both the MOG35–55-induced and adoptively transferred EAE. The conversion of resistant leptin-deficient mice to a state of susceptibility by rleptin administration implies that this molecule is influential in MOG35–55-induced EAE. The absence of disease in the leptin-deficient mice was accompanied by the almost complete absence of a measurable DTH response to the MOG35–55 peptide; the DTH reaction was fully restored by rleptin replacement. The presence or absence of leptin also determined different patterns of in vitro T cell responses in both leptin-deficient and wild-type rleptin-treated mice. In fact, in C57BL/6J-ob/ob mice injected with rleptin, the proliferative response and IFN-γ levels were induced, whereas IL-4 was suppressed when compared with obese PBS or PBS-pair fed controls. In wild-type animals, we observed mainly an increase in IFN-γ production after rleptin administration. Taken together, these findings indicate that activation and differentiation toward a Th1 phenotype of MOG35–55-specific T cells were hindered in leptin-deficient mice, whereas the switch to a Th2-type response was promoted. The presence of leptin restored the capacity of MOG35–55 to induce a classical Th1 pathogenic response in leptin-deficient mice and enhanced the inflammatory response in wild-type animals. Specific Ab titers reflected the nature of the T cell response in C57BL/6J-ob/ob mice after rleptin administration. Despite an apparent normal MOG35–55-specific IgG response, the pattern of IgG subclass production was markedly different in the presence or absence of leptin. The response was dominated by IgG1 in the absence of leptin, with almost no IgG2a production. This pattern was reversed by leptin administration, consistent with the reversal of Th cell polarization.

Active immunization of leptin-deficient mice with MOG35–55 leads to the generation of IL-4-secreting T cells and IgG1 Abs that are not able to induce a clinically evident disease and brain infiltration (21, 22). Furthermore, the resistance of C57BL/6J-ob/ob mice to adoptively induced disease and the absence of significant DTH reaction after transfer of CD4+ T cells (data not shown) suggest a level of resistance downstream of the generation of encephalitogenic T cells. Because leptin has been shown to affect endothelial cell function (23), expression of adhesion molecules such as ICAM-1 and VLA-2 on CD4+ T cells (3, 24), and survival of thymocytes (4), the interplay between one or more of these mechanisms may be responsible for resistance of leptin-deficient mice to adoptive transfers. Furthermore, the adoptive transfer experiments show for the first time that the presence of leptin is required for the expansion, differentiation, and maintenance of activated Th1 pathogenic T cells in the peripheral immune compartment to mediate tissue injury and disease progression.

It is clear that leptin is a pleiotropic molecule with effects on multiple biological systems, of which the immune system is but one. Leptin influences the neuroendocrine system at several levels, including the hypothalamic-pituitary-adrenal, thyroid, gonadal, and growth hormone axes (2). Therefore, it is possible that the interplay between these endocrine systems and the immune response may have influenced, indirectly, the pattern of disease susceptibility and evolution observed in this study. The data described in this work indicate that the influence of leptin on T cell immunity is sufficiently profound to control susceptibility to autoimmune disease. These findings suggest that the immune effects of leptin deficiency in the context of nutritional deficiency may be far reaching, and conversely that antagonism of the leptin axis may have potential in the field of immunotherapy.

This work is dedicated to the memory of Dr. Antonino Di Tuoro, as a tribute to his unique human and scientific qualities. We are particularly indebted to A. Coppola for histological analysis. We also thank G. Sequino for technical advice in the animal facility; G. Ruggiero for ELISA reader support; P. Reynolds for reading the manuscript; and F. Perna for photographic assistance.

1

This work was supported by the Fondazione Italiana Sclerosi Multipla (FISM); G.M. is a Universita di Napoli “Federico II,” Fondo Sociale Europeo Fellow, Italy; V.S. is a Consiglio Nazionale delle Ricerche Fellow, Italy; G.M.L. and J.K.H. are Medical Research Council, Clinical Training Fellows, United Kingdom.

3

Abbreviations used in this paper: EAE, experimental autoimmune encephalomyelitis; DTH, delayed-type hypersensitivity; MOG, myelin oligodendrocyte glycoprotein; rleptin, recombinant leptin.

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