| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
*
Cattedra di Immunologia, Dipartimento di Biologia e Patologia Cellulare e Molecolare, and
Dipartimento di Scienze Biomorfologiche, Universita di Napoli "Federico II," Napoli, Italy;
Laboratorio di Immunologia Cellulare, Azienda Ospedaliara "V. Monaldi," Napoli, Italy;
Centro di Endocrinologia e Oncologia Sperimentale-Consiglio Nazionale delle Ricerche (CEOS-CNR), Napoli, Italy; and
¶
Department of Immunology and
|| Endocrine Unit, Imperial College School of Medicine, Hammersmith Hospital, London, United Kingdom
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
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.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
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.
Antigens
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.
Leptin administration
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.
Induction of active and adoptive disease
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 x 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 x 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.
Clinical and histological assessment
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).
Induction of delayed-type hypersensitivity (DTH) (footpad-swelling assay)
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).
Proliferation assays and cytokine analysis
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
x 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- 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-; Abs
BVD4-1D11 and BVD6-24G2 for the detection of IL-4). Standard curves for
each assay were generated using recombinant mouse cytokines (IFN-
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-; <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.
ELISA for MOG35–55-specific Abs
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).
Statistical analyses
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.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
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. 1a). 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. 1b). 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. 2c). 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. 2c). No cellular infiltration or
demyelination was observed in control animals receiving adjuvants and
pertussis toxin only (data not shown).
|
|
|
; 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. 1c). 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. 1d). 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).
|
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. 3a), similar to that observed
in PBS-treated wild-type mice (Fig. 3b). C57BL/6J mice
injected with rleptin showed a statistically significant increase in
the DTH response when compared with the PBS-treated group (Fig. 3b). None of the control mice injected with CFA alone
developed any significant anti-MOG35–55
peptide DTH reaction (not shown).
|
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- 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- 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-, 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. 4d, and inset graph filled symbol), while rleptin did
increase IFN- 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).
|
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. 5c). In
wild-type animals, IgG1 levels were slightly but significantly reduced
by rleptin treatment (Fig. 5d). 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. 5e). In wild-type control mice, IgG2a
levels were also increased by rleptin injection, although to a lesser
degree (Fig. 5f).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
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- 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.
|
Acknowledgments |
|---|
|
Footnotes |
|---|
2 Address correspondence and reprint requests to Dr. Robert I. Lechler, Department of Immunology, Imperial College School of Medicine, Hammersmith Hospital, Du Cane Road, London W12 ONN, U.K.
3 Abbreviations used in this paper: EAE, experimental autoimmune encephalomyelitis; DTH, delayed-type hypersensitivity; MOG, myelin oligodendrocyte glycoprotein; rleptin, recombinant leptin.
Received for publication December 7, 2000. Accepted for publication March 5, 2001.
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and IL-18. Proc. Natl. Acad. Sci. USA 97:2367.
expression of encephalitogenic T cells. Eur. J. Immunol. 25:1951.[Medline]
enhances secretion of IgG2a from IgG2a-committed LPS-stimulated murine B cells: implications for the role of IFN- in class switching. Cell. Immunol. 135:95.[Medline]
This article has been cited by other articles:
|
|
 |
|
  H. Nave, G. Mueller, B. Siegmund, R. Jacobs, T. Stroh, U. Schueler, M. Hopfe, P. Behrendt, T. Buchenauer, R. Pabst, et al. Resistance of Janus Kinase-2 Dependent Leptin Signaling in Natural Killer (NK) Cells: A Novel Mechanism of NK Cell Dysfunction in Diet-Induced Obesity Endocrinology, July 1, 2008; 149(7): 3370 - 3378. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. J. Mattapallil, A. Augello, C. Cheadle, D. Teichberg, K. G. Becker, C.-C. Chan, J. J. Mattapallil, G. Pennesi, and R. R. Caspi Differentially Expressed Genes in MHC-Compatible Rat Strains That Are Susceptible or Resistant to Experimental Autoimmune Uveitis Invest. Ophthalmol. Vis. Sci., May 1, 2008; 49(5): 1957 - 1970. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Thewissen and J. W. Cohen Tervaert Leptin and ANCA-associated vasculitis: a healthy link? Rheumatology, April 1, 2008; 47(4): 387 - 389. [Full Text] [PDF]
|
|
|
|
 |
|
 J. Y. Park, A. Y. Chong, E. K. Cochran, D. E. Kleiner, M. J. Haller, D. A. Schatz, and P. Gorden Type 1 Diabetes Associated with Acquired Generalized Lipodystrophy and Insulin Resistance: The Effect of Long-Term Leptin Therapy J. Clin. Endocrinol. Metab., January 1, 2008; 93(1): 26 - 31. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. Taleb, O. Herbin, H. Ait-Oufella, W. Verreth, P. Gourdy, V. Barateau, R. Merval, B. Esposito, K. Clement, P. Holvoet, et al. Defective Leptin/Leptin Receptor Signaling Improves Regulatory T Cell Immune Response and Protects Mice From Atherosclerosis Arterioscler. Thromb. Vasc. Biol., December 1, 2007; 27(12): 2691 - 2698. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. Kumpers, F. Gueler, S. Rong, M. Mengel, I. Tossidou, I. Peters, H. Haller, and M. Schiffer Leptin is a coactivator of TGF-beta in unilateral ureteral obstructive kidney disease Am J Physiol Renal Physiol, October 1, 2007; 293(4): F1355 - F1362. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. Batra, J. Pietsch, I. Fedke, R. Glauben, B. Okur, T. Stroh, M. Zeitz, and B. Siegmund Leptin-Dependent Toll-Like Receptor Expression and Responsiveness in Preadipocytes and Adipocytes Am. J. Pathol., June 1, 2007; 170(6): 1931 - 1941. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. J. Ryan, G. R. McLemore Jr, and S. T. Hendrix Insulin Resistance and Obesity in a Mouse Model of Systemic Lupus Erythematosus Hypertension, November 1, 2006; 48(5): 988 - 993. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Margalit, Z. Shalev, O. Pappo, M. Sklair-Levy, R. Alper, M. Gomori, D. Engelhardt, E. Rabbani, and Y. Ilan Glucocerebroside Ameliorates the Metabolic Syndrome in OB/OB Mice J. Pharmacol. Exp. Ther., October 1, 2006; 319(1): 105 - 110. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 G. Palmer, M. Aurrand-Lions, E. Contassot, D. Talabot-Ayer, D. Ducrest-Gay, C. Vesin, V. Chobaz-Peclat, N. Busso, and C. Gabay Indirect Effects of Leptin Receptor Deficiency on Lymphocyte Populations and Immune Response in db/db Mice. J. Immunol., September 1, 2006; 177(5): 2899 - 2907. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Otero, R. Lago, R. Gomez, C. Dieguez, F. Lago, J. Gomez-Reino, and O. Gualillo Towards a pro-inflammatory and immunomodulatory emerging role of leptin Rheumatology, August 1, 2006; 45(8): 944 - 950. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 P. Plaisancie, R. Ducroc, M. E. Homsi, A. Tsocas, S. Guilmeau, S. Zoghbi, O. Thibaudeau, and A. Bado Luminal leptin activates mucin-secreting goblet cells in the large bowel Am J Physiol Gastrointest Liver Physiol, April 1, 2006; 290(4): G805 - G812. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. A. Oral, E. D. Javor, L. Ding, G. Uzel, E. K. Cochran, J. R. Young, A. M. DePaoli, S. M. Holland, and P. Gorden Leptin Replacement Therapy Modulates Circulating Lymphocyte Subsets and Cytokine Responsiveness in Severe Lipodystrophy J. Clin. Endocrinol. Metab., February 1, 2006; 91(2): 621 - 628. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C.-H. Lee, Y.-G. Chen, J. Chen, P. C. Reifsnyder, D. V. Serreze, M. Clare-Salzler, M. Rodriguez, C. Wasserfall, M. A. Atkinson, and E. H. Leiter Novel Leptin Receptor Mutation in NOD/LtJ Mice Suppresses Type 1 Diabetes Progression: II. Immunologic Analysis Diabetes, January 1, 2006; 55(1): 171 - 178. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
S. Musio, B. Gallo, S. Scabeni, M. Lapilla, P. L. Poliani, G. Matarese, H. Ohtsu, S. J. Galli, R. Mantegazza, L. Steinman, et al. A Key Regulatory Role for Histamine in Experimental Autoimmune Encephalomyelitis: Disease Exacerbation in Histidine Decarboxylase-Deficient Mice J. Immunol., January 1, 2006; 176(1): 17 - 26. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. W. Wieland, S. Florquin, E. D. Chan, J. C. Leemans, S. Weijer, A. Verbon, G. Fantuzzi, and T. van der Poll Pulmonary Mycobacterium tuberculosis infection in leptin-deficient ob/ob mice Int. Immunol., November 1, 2005; 17(11): 1399 - 1408. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
C.-H. Lee, P. C. Reifsnyder, J. K. Naggert, C. Wasserfall, M. A. Atkinson, J. Chen, and E. H. Leiter Novel Leptin Receptor Mutation in NOD/LtJ Mice Suppresses Type 1 Diabetes Progression: I. Pathophysiological Analysis Diabetes, September 1, 2005; 54(9): 2525 - 2532. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Bruno, S. Conus, I. Schmid, and H.-U. Simon Apoptotic Pathways Are Inhibited by Leptin Receptor Activation in Neutrophils J. Immunol., June 15, 2005; 174(12): 8090 - 8096. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. P. McGillis White Adipose Tissue, Inert No More! Endocrinology, May 1, 2005; 146(5): 2154 - 2156. [Full Text] [PDF]
|
|
|
|
