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The Journal of Immunology, 2006, 177: 2899-2907.
Copyright © 2006 by The American Association of Immunologists, Inc.

Indirect Effects of Leptin Receptor Deficiency on Lymphocyte Populations and Immune Response in db/db Mice1

Gaby Palmer*,{ddagger}, Michel Aurrand-Lions*, Emmanuel Contassot*,§, Dominique Talabot-Ayer*,{ddagger}, Dominique Ducrest-Gay*, Christian Vesin{dagger}, Véronique Chobaz-Péclat, Nathalie Busso and Cem Gabay2,*,{ddagger}

* Department of Pathology and Immunology and {dagger} Department of Cellular Physiology and Metabolism, University of Geneva School of Medicine, Geneva, Switzerland; {ddagger} Division of Rheumatology and § Department of Dermatology, Louis-Jeantet Skin Cancer Laboratory, University Hospital, Geneva, Switzerland; and Department of Medicine, Division of Rheumatology, University Hospital, Lausanne, Switzerland


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Leptin-deficient ob/ob and leptin receptor (Ob-rb)-deficient db/db mice display a marked thymic atrophy and exhibit defective immune responses. Lymphocytes express leptin receptors and leptin exerts direct effects on T cells in vitro. In addition, ob/ob and db/db mice display multiple neuroendocrine and metabolic defects, through which leptin deficiency may indirectly affect the immune system in vivo. To study the relative contributions of direct and indirect effects of leptin on the immune system in a normal environment, we generated bone marrow chimeras (BMCs) by transplantation of leptin receptor-deficient db/db, or control db/+, bone marrow cells into wild-type (WT) recipients. The size and cellularity of the thymus, as well as cellular and humoral immune responses, were similar in db/db to WT and db/+ to WT BMCs. The immune phenotype of db/db mice is thus not explained by a cell autonomous defect of db/db lymphocytes. Conversely, thymus weight and cell number were decreased in the reverse graft setting in WT to db/db BMCs, indicating that expression of the leptin receptor in the environment is important for T cell development. Finally, normal thymocyte development occurred in fetal db/db thymi transplanted into WT hosts, indicating that direct effects of leptin are not required locally in the thymic microenvironment. In conclusion, direct effects of leptin on bone marrow-derived cells and on thymic stromal cells are not necessary for T lymphocyte maturation in normal mice. In contrast, leptin receptor deficiency affects the immune system indirectly via changes in the systemic environment.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Leptin is a peptide hormone that plays an important role in the regulation of body weight by inhibiting food intake and stimulating energy expenditure. Leptin is mainly produced by adipose tissue and its levels are regulated by a variety of factors, including food intake and the endocrine system (for review, see Ref. 1). Leptin-deficient (ob/ob) mice and leptin receptor-deficient (db/db) mice exhibit severe hereditary obesity (2, 3). However, leptin has many other effects in addition to the regulation of body weight. Consistently, ob/ob and db/db mice are not only obese, but display also hormonal imbalances, abnormalities in thermoregulation, infertility, and evidences of hemopoietic and immune defects (4, 5, 6, 7, 8). Similar alterations have been described in leptin-deficient humans (9). Leptin, therefore, seems to represent an important link between fat mass and metabolism on one hand, and the regulation of neuroendocrine function, reproduction, and immunity in contrast. In particular, leptin has been recently (10) linked to an increased susceptibility to autoimmune diseases.

ob/ob and db/db mice display a marked reduction in the size and cellularity of the thymus (11, 12, 13, 14). ob/ob mice also exhibit defective cellular and humoral immune responses and are protected from immune-mediated inflammation in various disease models, such as experimental colitis, autoimmune encephalomyelitis, Con A-induced hepatitis, or Ag-induced arthritis (15, 16, 17, 18, 19). Similarly, starvation and malnutrition, two conditions characterized by low leptin levels, are associated with alterations of the immune response and thymic atrophy, which can be reversed by leptin administration (12, 20).

The long-signaling leptin receptor isoform (Ob-rb or Leprb) is expressed in T and B cells, and several studies (19, 20, 21, 22) have described direct effects of leptin on lymphocytes. Leptin was shown (12, 20, 21, 23, 24), for instance, to modulate T cell proliferation, to promote Th1 responses, and to protect thymocytes from corticosteroid-induced apoptosis in vitro. These observations led to the suggestion that direct effects of leptin on lymphocytes might account for immune defects observed in ob/ob and db/db mice. However, the connection between leptin deficiency and immune deficiencies in vivo is likely to be more complex. Ob/ob and db/db mice indeed display multiple neuroendocrine and metabolic modifications, including the activation of the hypothalamic-pituitary-adrenal axis and hypercorticosteronemia, hyperglycemia, and diabetes, which may indirectly affect the immune system. Similarly, leptin deficiency after starvation in rodents is linked to increased glucocorticoid levels, and decreased levels of thyroid and growth hormone, each of which may mediate immune suppression (25). Both direct and indirect effects of leptin on the immune system are thus likely to account for the immune defects observed in leptin- and leptin receptor-deficient rodents. In the present study, we used bone marrow chimeras (BMCs)3 and fetal thymus transplantation to examine independently the respective contributions of leptin receptor-deficient db/db bone marrow-derived cells, of the abnormal systemic db/db environment and of db/db thymic stromal cells to the immune phenotype observed in db/db mice.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice

Male C57BL/6 congenic B6.Cg-m+/+ Leprdb/db (db/db) mice, male and female B6.Cg-m+/+ Leprdb/+ (db/+) mice, andC57BL/6 congenic B6.SJL-Ptprca Pep3b/BoyJ (CD45.1) mice, carrying the differential CD45.1 allele of the leukocyte common Ag, were obtained from The Jackson Laboratory. Male C57BL/6 mice were obtained from Janvier. All animals were housed under conventional conditions. Water and standard laboratory chow were provided ad libitum. Institutional approval was obtained for all animal experiments.

Generation of BMCs

For the generation of db/+ to wild-type (WT) and db/db to WT BMCs, 4- to 6-mo-old male CD45.1 recipients were irradiated at 800 rad using a 137Cs source and reconstituted 24 h later with 2 x 106 total bone marrow cells from adult (> 8-wk-old) male db/+ or db/db donors. For the generation of WT to db/+ and WT to db/db BMCs, 3-mo-old male db/+ recipients were irradiated at 800 rad. Because we anticipated altered sensitivity to irradiation in db/db mice, 3-mo-old male db/db mice were irradiated at 700, 800, or 900 rad. Irradiated mice were reconstituted 24 h later with 2 x 106 total bone marrow cells from male CD45.1 donors. After irradiation, all mice were kept on drinking water supplemented with neomycin sulfate for 4–6 wk before returning to regular water.

Measurement of serum leptin and corticosterone levels

Serum levels of leptin were measured using a commercial Quantikine ELISA kit from R&D Systems. The detection limit for this test is 22 pg/ml. Serum corticosterone levels were determined by RIA (Diagnostic Systems Laboratories) as previously described (26).

Analysis of lymphocyte populations

Male db/+ and db/db mice were sacrificed at 10 wk and 6 mo of age. Db/+ to WT and db/db to WT BMCs from three independent graft experiments were analyzed 5, 7, and 12 wk after the graft. WT to db/+ and WT to db/db BMCs were sacrificed 10 wk after the graft. Blood was collected on citrate and diluted into PBS/1.25 mM EDTA. Thymus, spleen, inguinal, and mesenteric lymph nodes (LN) were dissected and cell suspensions were prepared by gently grinding the organs through a 70-µm cell strainer (BD Biosciences). Cells were washed and resuspended in FACS buffer (PBS and 0.2% BSA) containing saturating amounts of Abs. Flow cytometry was performed with the following Abs: FITC-conjugated anti-CD45.2 (27), PE-conjugated anti-CD4, CyChrome-conjugated anti-B220, biotinylated anti-CD8 followed by streptavidin coupled to allophycocyanin, and PE-conjugated anti-CD11b (Mac-1). All Abs for flow cytometry were obtained from BD Biosciences. In blood and spleen samples, RBC were lysed using the BD FACS Lysing Solution (BD Biosciences). Cells were analyzed on a FACSCalibur (BD Biosciences) using CellQuest software.

Immune response to methylated BSA (mBSA)

db/+ and db/db mice were immunized at 10 wk of age. db/+ to WT and db/db to WT BMCs were immunized 15 wk after the graft. Mice were injected intradermally at the base of the tail with 100 µg of mBSA (Fluka) and emulsified in CFA (Difco) containing 5 mg/ml Mycobacterium tuberculosis. Heat-killed Bordetella pertussis organisms (0.2 x 109; Berna Biotech) were injected i.p. as an additional adjuvant. On day 7, a booster injection of 100 µg of mBSA in IFA (Difco) was given at the base of the tail. Mice were sacrificed 28 days after the first immunization. Draining LN and spleen cells were seeded at 4 x 105 cells/well in 96-well plates in 200 µl of RPMI 1640 medium containing 100 IU/ml penicillin, 100 µg/ml streptomycin, 5 x 10–5 M 2-ME, and 1% autologous mouse serum. Cells were incubated at 37°C in 5% CO2 for 48 h without or with 10 µg/ml mBSA or 5 µg/ml Con A (Amersham Biosciences). During the final 18 h of incubation, [3H]thymidine was added at 1 µCi/well. Cells were harvested and radioactivity was counted to determine [3H]thymidine incorporation into DNA as a measure of cell proliferation. Serum levels of total anti-mBSA Abs were measured as described elsewhere (19).

RT-PCR

Six- to 8-wk-old male C57BL/6 mice were injected with 200 µg/g hydrocortisone or saline and sacrificed 24 h later. db/db to WT BMCs were sacrificed 15 wk after the graft. The thymus was dissected and total RNA was isolated using the TRIzol reagent (Invitrogen Life Technologies). Total RNA (3 µg) was digested with DNase I (Promega) and reverse-transcribed using AMV-RT (Promega) and random hexamer primers. PCR amplification (40 cycles for Ob-rb, 30 cycles for beta-actin) was performed using TaqDNA polymerase (Qiagen) and the following primers: Ob-rb forward primer 5'-GAATTGTTCCTGGGCACAAG-3'; Ob-rb reverse primer 5'-GGGACCATCTCATCTTTATT-3'; beta-actin forward primer 5'-CCAAGGCCAACCGCGAGAAGATGAC-3' and beta-actin reverse primer 5'-AGGGTACATGGTGGTGCCGCCAGAC-3'. Annealing temperatures were 57°C for Ob-rb or 60°C for beta-actin. The absence of DNA contamination in RNA preparations was tested by including RNA samples, which had not been reverse-transcribed and distilled water was used as a negative control for PCR amplification. The identity of the amplified products was confirmed by DNA sequencing.

Western blotting

The thymus of 4- to 8-wk-old male C57BL/6 mice was dissected and one lobe was cut into small pieces and homogenized in TNT buffer (50 mM Tris (pH 7.4), 150 mM NaCl, 1 mM PMSF, 10 µg/ml aprotinin, 4 µg/ml pepstatin, and 0.5% Triton X-100). The other lobe was forced through a 70-µm cell strainer to obtain a single-cell suspension corresponding to total thymocytes. Isolated thymocytes, as well as the tissue remaining on the membrane of the cell strainer, which was considered to represent a thymic stromal fraction, were homogenized in TNT buffer. All samples were sonicated briefly and cleared by centrifugation at 13,000 rpm at 4°C for 15 min. Thymic epithelial MTE-4.14 cells (28) were grown to confluence in 6-well plates and lysed in 200 µl of Laemmli buffer. Human COLO 320DM whole cell lysate (Santa Cruz Biotechnology) was used as a positive control for OB-R expression. Samples were heated in Laemmli buffer, fractionated by SDS-PAGE, and transferred to a Porablot membrane (Macherey-Nagel). The membrane was blocked in PBS containing 5% blotto (Santa Cruz Biotechnology) and immunoblotting was performed with a rabbit anti-Ob-r polyclonal Ab (H300, dilution 1/300; Santa Cruz Biotechnology) or with a rabbit anti-actin polyclonal Ab (A2066, dilution 1/500; Sigma-Aldrich;). Immunoreactive bands were visualized by ECL (Amersham Pharmacia Biotech) using an anti-rabbit HRP-labeled secondary Ab (dilution 1/10,000; Santa Cruz Biotechnology).

Fetal thymus transplantation

Thymic lobes were obtained from day 14.5 WT, db/+, and db/db embryos derived from heterozygous db/+ intercrosses. Each embryo was genotyped using two distinct PCR to amplify the WT and the db allele of the Lepr gene, respectively. The following primers were used: WT forward 5'-GATGTTTACATTTTGATGGAGGG-3'; WT reverse 5'-GTCACACCATTATCATTTTATG-3'; db forward 5'-AGAACGGACACTCTTTGAAG –3'; and db reverse 5'-CCATAGTTTAGGTTTGTTTA-3'. PCR amplification (40 cycles) was performed using TaqDNA polymerase (Qiagen) and an annealing temperature of 55°C. A small incision was made in the peritoneal cavity of anesthetized 10- to 13-wk-old male CD45.1 recipient mice. Their left kidneys were exposed under a stereomicroscope and the two thymic lobes from an individual fetus were positioned under the kidney capsule. The wound was closed with surgical sutures and the recipient mice were kept on drinking water supplemented with neomycin sulfate for 2 wk before returning to regular water. Recipient mice were sacrificed 6 wk after the graft. Blood was collected on citrate and the grafted thymi were recovered for flow cytometric analysis.

Statistical analysis

Significance of differences was calculated by Student’s unpaired t test or ANOVA. A difference between experimental groups was considered significant when the p value was <0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Lymphocyte populations in db/db mice

We examined lymphocyte populations in male db/db mice in a C57BL/6 genetic background at 10 wk and 6 mo of age. As in previous studies (19, 20), examining the immune phenotype of db/db mice, we used heterozygous db/+ littermates as controls. At both time points, db/db mice were severely obese and displayed increased serum leptin and corticosterone levels (Table I), which are characteristic features of leptin receptor deficiency in mice (3, 29). In 10-wk-old male db/db mice, thymus weight and cell number were markedly decreased as compared with db/+ controls (Fig. 1A), whereas the respective percentages of CD4 and CD8 double-positive (DP), CD4+, CD8+, and double negative (DN) thymocytes were similar in db/+ and db/db mice (Fig. 1B), as previously reported for db/db mice in the C57BL/KS background (14). Peripheral lymph nodes were smaller in db/db mice as compared with db/+ controls, but there were no differences in CD4+, CD8+, or B220+ cell populations between db/+ and db/db mice (Fig. 1C). Similarly, CD4+, CD8+, B220+, and Mac-1+ cell populations in peripheral blood were comparable in db/+ and db/db mice (Fig. 1D). In addition, differential blood cell counts confirmed similar numbers of white blood cells (db/+, 2930 ± 580 cells/µl; db/db, 2200 ± 300 cells/µl) and lymphocytes (db/+, 2350 ± 470 cells/µl; db/db, 1780 ± 250 cells/µl) in the two groups of mice. Finally, spleen weight was decreased in db/db mice (db/+, 87 ± 9 mg; db/db, 52 ± 3 mg, p < 0.05), although splenocyte numbers were not different between the two groups, in accordance with data previously reported (30) for leptin-deficient ob/ob mice.


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  		Table I. Phenotype of db/+ and db/db micea

 

Figure 1
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  		FIGURE 1. Lymphocyte populations in db/db mice. A, Thymus weight and cell number are shown for 10-wk-old db/+ (n = 4; {square}) and db/db (n = 4, {blacksquare}) mice. Results are shown as mean ± SEM. *, p < 0.05 vs db/+ mice. B, Thymocyte populations, as assessed by flow cytometry, are shown for 10-wk-old db/+ (n = 4; {square}) and db/db (n = 4; {blacksquare}) mice. Data are expressed as a percentage of total thymocytes and results are shown as mean ± SEM. DP, CD4, and CD8 DP cells; DN, CD4, and CD8 DN cells. No significant differences were observed between the two groups. C, CD4+, CD8+, and B220+ LN cell populations, as assessed by flow cytometry, are indicated for 10-wk-old db/+ (n = 4; {square}) and db/db (n = 4; {blacksquare}) mice. Data are expressed as a percentage of total LN cells and results are shown as mean ± SEM. No significant differences were observed between the two groups. D, CD4+, CD8+, B220+, and Mac-1+ peripheral blood cell populations, as assessed by flow cytometry, are indicated for 10-wk-old db/+ (n = 4; {square}) and db/db (n = 4; {blacksquare}) mice. Data are expressed as a percentage of total blood cells and results are shown as mean ± SEM. No significant differences were observed between the two groups.

 
At 6 mo of age, thymus weight and cell number were again low in db/db mice, although the difference with db/+ mice was no longer significant, due to involution of the thymus with age in the control group (thymus cell number, db/+, 20.0 ± 3.9 mio cells; db/db, 12.7 ± 0.7 mio cells). It is interesting to note that thymocyte numbers in WT C57BL/6 mice are generally around 60 mio# cells at 6 mo of age, suggesting accelerated thymic involution in db/+ mice. There were no differences in the percentages of DP, CD4+, CD8+, and DN thymocytes, of CD4+, CD8+, and B220+ spleen cells, or of CD4+, CD8+, B220+, and Mac1+ peripheral blood cell populations between db/+ and db/db mice at 6 mo of age (data not shown).

Lymphocyte populations in db/db to WT BMCs

To study the importance of direct effects of leptin on lymphocytes in a normal environment, we generated BMCs by transplantation of db/db bone marrow cells into lethally irradiated WT recipient mice. We used recipient mice carrying the differential CD45.1 allele of the leukocyte-common Ag. Because donor db/+ and db/db mice carry the CD45.2 allele, the use of this allelic marker allowed for the distinction of leukocytes of donor and recipient origin. We conducted three independent bone marrow transfer experiments and analyzed BMCs at various time points after the graft. Body weight, serum leptin, and corticosterone levels were within the normal range for both db/+ and db/db BMCs 12 wk after graft transplantation (Table II), suggesting that transplantation of db/db bone marrow does not modify the metabolic and endocrine environment in recipient mice. In particular, the hypothalamic-pituitary-adrenal axis did not seem to be affected. Twelve weeks after the graft, donor-derived CD45.2+ cells represented >80% of total leukocytes in all organs tested (>98% in the thymus), indicating successful repopulation of the host by donor bone marrow-derived cells (Fig. 2A). The chimerism was similar in db/+ to WT and in db/db to WT BMCs in the different organs tested.


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  		Table II. Phenotype of db/+ to WT and db/db to WT BMCsa

 

Figure 2
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  		FIGURE 2. Lymphocyte populations in db/db to WT BMCs. A, Percentages of donor-derived CD45.2+ cells (chimerism) in thymus, LN spleen, and blood 12 wk after the graft are shown for db/+ to WT (n = 4; {square}) and db/db to WT (n = 6; {blacksquare}) BMCs. Data are expressed as a percentage of total cells and results are shown as mean ± SEM. No significant differences were observed between the two groups. B, Thymus weight and cell number are shown for db/+ to WT (n = 4{square}) and db/db to WT (n = 6; {blacksquare}) BMCs 12 wk after the graft. Results are shown as mean ± SEM. No significant differences were observed between the two groups. C, Representative flow cytometric analyses of thymocytes stained for CD4 and CD8 are shown for db/+ to WT (left panel) and db/db to WT (right panel) BMCs. D, Thymocyte populations, as assessed by flow cytometry, are shown for db/+ to WT and db/db to WT BMCs 12 wk after the graft. Data are expressed as a percentage of donor-derived CD45.2+ cells. DP, CD4 and CD8 DP cells; DN, CD4 and CD8 DN cells. No significant differences were observed between the groups.

 
Thymus weight and cell numbers were similar in db/+ to WT and db/db to WT BMCs (Fig. 2B), and DP, CD4+, CD8+, and DN cell populations were comparable in both groups (Fig. 2, C and D). Thymus cell numbers were also identical in both groups at an earlier time point (7 wk) after the graft (db/+: 52.7 ± 9.4 x 106 cells, n = 3; db/db: 53.1 ± 5.5 x 106 cells, n = 3), as was the chimerism (thymus db/+: 83.5 ± 10.5%, n = 3; db/db: 78.1 ± 1.9%, n = 3). Taken together, these data indicate that the thymic atrophy observed in db/db mice (Fig. 1A) is not explained by a cell autonomous defect of db/db thymocytes.

The size of peripheral LN was similar in db/+ to WT and db/db to WT BMCs 5, 7 and 12 wk after the graft and T and B lymphocyte populations were comparable in both groups. Finally, CD4+, CD8+ and B220+ lymphocyte populations in blood were also similar in db/+ to WT and db/db to WT BMCs 12 wk after the graft (data not shown).

Immune response to mBSA in db/db mice and in db/db to WT BMCs

To assess the importance of direct effects of leptin on lymphocytes for the response to an antigenic challenge in vivo, we compared the immune response to mBSA in db/+ and db/db mice and in db/+ to WT and db/db to WT BMCs. Ag-specific proliferation, induced by stimulation of LN and spleen cells from immunized animals with mBSA in vitro, was decreased in db/db as compared with db/+ control mice (Fig. 3A). In contrast, Ag-induced LN and spleen cell proliferation was not distinctly reduced in db/db to WT BMCs, as compared with db/+ to WT BMCs (Fig. 3B). In fact, basal and mBSA-stimulated thymidine uptake was even increased in cells isolated from db/db to WT BMCs.


Figure 3
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  		FIGURE 3. Cellular immune response to mBSA in db/db mice and in db/db to WT BMCs. A, Proliferation of LN and spleen cells stimulated with mBSA (10 µg/ml; Figure 3) or left unstimulated (control; {square}) was assessed for immunized db/+ (n = 4) and db/db (n = 4) mice by measuring [3H]thymidine incorporation into DNA. Results are shown as mean ± SEM. *, p < 0.05 vs db/+ cells. Fold increases in proliferation in mBSA-stimulated/control cells were: 28.17 ± 2.82 in db/+ LN cells, 14.38 ± 2.15 in db/db LN cells, 2.97 ± 0.39 in db/+ spleen cells, and 1.14 ± 0.03 in db/db spleen cells. B, Proliferation of LN and spleen cells stimulated with mBSA (10 µg/ml; Figure 3) or left unstimulated (control; {square}) was assessed for immunized db/+ to WT (n = 3) and db/db to WT (n = 3) BMCs. Results are shown as mean ± SEM. *, p < 0.05 vs db/+ cells. Fold increases in proliferation in mBSA -timulated/control cells were 44.6 ± 3.52 in LN cells isolated from db/+ to WT BMCs, 24.38 ± 2.35 in LN cells isolated from db/db to WT BMCs, 1.74 ± 0.16 in spleen cells isolated from db/+ to WT BMCs, and 4.87 ± 0.38 in spleen cells isolated from db/db to WT BMCs.

 
We also assessed the production of anti-BSA IgG in db/+ and db/db mice. Anti-BSA Ab titers tended to be lower in db/db mice, although the difference in anti-BSA IgG levels between the groups was not significant (db/+, 44.3 ± 13.6 arbitrary units (AU), n = 4; db/db, 19.4 ± 8.7 AU, n = 4). In db/+ to WT and db/db to WT BMCs, anti-BSA IgG levels were similar (db/+ to WT BMCs, 87.3 ± 29.3 AU, n = 3; db/db to WT BMCs, 88.0 ± 48.1 AU, n = 3). In a second independent graft experiment, BMCs were immunized 9 wk after the graft and anti-BSA IgG, assessed at day 26 after the first immunization, were again similar (db/+ to WT BMCs, 88.9 ± 8.2 AU, n = 4; db/db to WT BMCs, 86.2 ± 26.3 AU, n = 4).

Lymphocyte populations in WT to db/db BMCs

To test the importance of indirect effects of leptin receptor deficiency on lymphocyte populations, we performed the reverse WT to db/db bone marrow transplantation experiment, which allowed us to examine normal bone marrow-derived cells in the abnormal environment present in db/db recipients. WT to db/+ BMCs were used as controls. Db/db mice suffered high mortality after irradiation, and, despite standard reconstitution, one mouse of four died after irradiation with 700 rad, two of four with 800 rad, and three of four with 900 rad. Ten weeks after the graft, the respective body weights of WT to db/+ BMCs and of the surviving WT to db/db BMCs were significantly different (WT to db/+ BMCs, 28.4 ± 0.5 g, n = 5; WT to db/db BMCs, 50.8 ± 1.8 g, n = 6; p < 0.05), and thus comparable to nonirradiated db/+ or db/db mice, respectively (Table I). Donor-derived cells represented >80% of total leukocytes in all organs tested in both groups, although in thymus and spleen the chimerism appeared to be slightly lower in WT to db/db than in WT to db/+ BMCs (Fig. 4A).


Figure 4
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  		FIGURE 4. Lymphocyte populations in WT to db/db BMCs. A, Percentages of donor-derived CD45.2– cells (chimerism) in thymus, LN, spleen, and blood 10 wk after the graft are shown for WT to db/+ (n = 5; {square}) and WT to db/db (n = 6; {blacksquare}) BMCs. We observed no differences in chimerism in WT to db/db BMCs upon irradiation with 700, 800, or 900 rad and pooled data are thus shown for all surviving WT to db/db BMCs. Data are expressed as a percentage of total cells and results are shown as mean ± SEM. *, p < 0.05 vs WT to db/+ BMCs. B, Thymus weight and cell number are shown for WT to db/+ (n = 5; {square}) and WT to db/db (n = 6; {blacksquare}) BMCs 10 wk after the graft. Results are shown as mean ± SEM. *, p < 0.05 vs WT to db/+ BMCs. C, Thymocyte populations, as assessed by flow cytometry, are shown for WT to db/+ and WT to db/db BMCs 10 wk after the graft. Data are expressed as a percentage of donor-derived CD45.2– cells. DP, CD4 and CD8 DP cells; DN, CD4- and CD8-negative cells. No significant differences were observed between the groups.

 
Thymus weight and cell number were significantly decreased in WT to db/db BMCs as compared with WT to db/+ BMCs (Fig. 4B), whereas DP, CD4+, CD8+, and DN thymocyte populations were similar in both groups (Fig. 4C). Exposure of WT thymocytes to the db/db environment in WT to db/db BMCs thus resulted in a nonselective thymic lymphopenia.

Peripheral LNs were smaller in WT to db/db as compared with WT to db/+ BMCs and, interestingly, the percentages of both CD4+ and CD8+ T cells were reduced in WT to db/db BMCs, suggesting impaired or delayed T cell repopulation in peripheral LN in the db/db environment (Table III).


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  		Table III. LN cell populations in BMCsa

 
OB-R expression in the thymus

Although previous studies (24) described the presence of Ob-rb mRNA in thymus, Ob-rb-expressing cell types had not been characterized. Treatment of mice with hydrocortisone drastically decreased thymocyte numbers, preferentially depleting DP cells (Fig. 5A) as previously described (31). However, Ob-rb mRNA expression in the thymus was not decreased after hydrocortisone injection, suggesting that cells other than DP thymocytes express substantial amounts of Ob-rb (Fig. 5A). This hypothesis was further substantiated by the observation that both db/db donor bone marrow-derived thymocytes and WT recipient-derived thymic stromal cells express Ob-rb mRNA in the thymus of db/db to WT BMCs (Fig. 5B). Finally,, Western blotting analysis revealed expression of the Ob-rb protein in whole thymus, isolated thymocytes, as well as in a crude thymic stromal fraction (Fig. 5C). Consistently, RT-PCR and Western blotting also showed expression of Ob-rb mRNA and protein in the murine thymic epithelial cell line MTE-4.14 (28) (Fig. 5C and data not shown).


Figure 5
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  		FIGURE 5. Expression of OB-R in the thymus. A, Ob-Rb mRNA expression was examined in the thymus 24 h after injection of saline (control) or 200 µg/g hydrocortisone to WT C57BL/6 mice. Left panels, Representative flow cytometric analyses of thymocytes from control (top) or hydrocortisone (bottom)-injected mice stained for CD4 and CD8. Total thymocyte numbers were 45 x 106 cells (control) and 4 x 106 cells (hydrocortisone). The right panels show Ob-rb (top) and actin (bottom) mRNA expression as assessed by RT-PCR. Lane 1, Control thymus; lane 2, hydrocortisone thymus; lane 3, negative control, PCR performed on non-reverse-transcribed RNA from control thymus. The position of molecular mass markers (bp) is indicated on the right. B, Expression of WT and db alleles of Ob-rb mRNA was examined by RT-PCR on thymus RNA obtained from two db/db to WT BMCs. A schematic representation of the two alleles with the location of the PCR primers (arrows) is shown on the left. The db mutation (*) leads to insertion of an additional 106-bp exon ({blacksquare}), between exons 17 ({square}) and 18b (Figure 5) of the mouse lepr gene. This additional exon contains a stop codon causing premature termination of the Ob-rb protein in the db mutant (42 ). Amplification of the WT and db alleles with the indicated primers yields PCR products of 170 and 276 bp, respectively. The right panel shows the expression of mRNA encoding both WT (recipient-derived) and db (donor-derived) alleles in the thymus of two different db/db to WT BMCs (lanes 1 and 2). C, Expression of the Ob-rb protein was examined by Western blotting in thymocytes, whole thymus, a thymic stromal fraction, and in the thymic epithelial cell line MTE-4.14 (upper panel). The membrane was stripped and reblotted with anti-beta-actin Abs (lower panel). The positions of molecular mass standards are indicated on the right. Lane 1, Human COLO 320DM cell lysate (positive control); lane 2, total thymocytes; lane 3, whole thymus; lane 4, thymic stromal fraction; and lane 5, MTE- 4.14 total cell lysate.

 
Analysis of thymocyte development in db/db thymi grafted into WT recipients

Expression of Ob-rb in the thymic stromal fraction and in thymic epithelial cells suggested that leptin might exert indirect effects on thymocytes by acting locally on the thymic environment. To investigate the importance of direct effects of leptin on thymic stromal cells in a normal environment, we examined thymocyte development in fetal db/db thymi transplanted into WT CD45.1 recipients. db/+ and WT fetal thymi were used as controls. We observed a similar efficiency of engraftment for all three genotypes (4 of 6 WT thymi, 8 of 11 db/+ thymi, and 4 of 6 db/db thymi successfully engrafted). Six weeks after transplantation, grafted WT, db/+, and db/db thymi were of similar size and were equally well colonized by recipient T cells. Indeed, the chimerism (percentage of CD45.2– recipient T cells) was > 99% in the 16 grafted thymi and was similar for all genotypes. Recipient DP, CD4+, CD8+, and DN T cell populations were generated in normal proportions in db/db thymi as compared with control WT and db/+ thymic grafts (Fig. 6). Finally, circulating donor (CD45.2+) and recipient (CD45.2–) derived CD4+ and CD8+ T cell populations in the blood of recipient mice were similar for all genotypes (data not shown). Leptin signaling in thymic stromal cells is thus not required for T cell development in a normal environment.


Figure 6
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  		FIGURE 6. Lymphocyte populations in db/db fetal thymi transplanted into WT recipients. A, WT, db/+ and db/db fetal thymi were grafted under the kidney capsule of WT CD45.1 recipient mice. T cell populations in the grafts were analyzed 6 wk after transplantation. Representative flow cytometric analyses of recipient-derived (CD45.2–) T cells stained for CD4 and CD8 are shown for WT (left panel), db/+ (middle panel), and db/db (right panel) fetal thymus grafts. B, Recipient-derived (CD45.2–) T cell populations in thymus grafts, as assessed by flow cytometry, are shown for WT (n = 4; {square}), db/+ (n = 8; Figure 6), and db/db (n = 4, {blacksquare}) fetal thymi grafted under the kidney capsule of WT CD45.1 mice. Data are expressed as a percentage of total thymocytes and results are shown as mean ± SEM. DP, CD4 and CD8 DP cells; DN, CD4 and CD8 DN cells. No significant differences were observed between the groups.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
db/db mice in a C57BL/6 genetic background displayed a marked nonselective thymic lymphopenia, as previously reported for C57BL/KS db/db mice (14, 24). A similar nonselective T lymphopenia was also described in leptin receptor-deficient fa/fa rats (32), and in ob/ob mice (33). In contrast, one study reported (12) a decrease in the percentage of DP thymocytes, as well as an increase in CD4+ and DN cells, along with a decrease in the total cell number in the thymus of ob/ob mice. The reason for these discrepant findings in ob/ob mice remains unclear, but might be related to differences in age or gender of the mice studied. Similarly, variable phenotypes have been reported (32, 33, 34) for blood cell populations in leptin and leptin receptor-deficient rodents, lymphocyte counts being sometimes similar to those in WT control mice and sometimes decreased. Furthermore, one study (33) showed an increased number of circulating monocytes in ob/ob mice. In the present study, we observed similar lymphocyte and Mac-1+ myeloid cell populations in the blood of db/+ and db/db mice, both at 10 wk and 6 mo of age.

In marked contrast to db/db mice, the size and cellularity of the thymus were not different between db/+ or db/db to WT BMCs, indicating that the thymic atrophy observed in db/db donors does not reflect a cell autonomous effect of db/db lymphocytes. In fact, all examined organs showed normal lymphocyte populations in db/db to WT BMCs at three different time points after the graft. The results of these experiments thus indicate that direct effects of leptin on bone marrow-derived cells are not required for lymphocyte maturation in a normal environment, at least in our experimental settings.

The Ag-specific in vitro proliferative response of LN and spleen cells was decreased in db/db mice as compared with db/+ controls, as reported earlier for db/db mice in the C57BL/KS background (19). Proliferation induced by nonspecific mitogens was previously reported either to be normal in lymphocytes isolated from db/db mice (13) or to be impaired in lymphocytes obtained from db/db mice or fa/fa rats (32, 35). In the present study, proliferation induced by the nonspecific mitogen Con A was similar in db/+ and db/db cells (data not shown). In db/db to WT BMCs, the Ag-specific proliferative response of LN and spleen cells was not reduced as compared with db/+ to WT BMCs. Both basal and mBSA-stimulated thymidine uptake were even rather increased in db/db to WT BMCs, indicating that direct effects of leptin are not required to sustain T cell proliferation in our experimental conditions. This observation is in contrast with recent studies (17, 36) suggesting that leptin, produced by activated T cells themselves, stimulates their proliferation in an autocrine loop. However, this issue probably deserves further investigation, since, in another study (23), leptin was shown on the contrary to decrease in vitro proliferation of mature T cells isolated from mouse spleen. Anti-mBSA IgG titers tended to be decreased in immunized C57BL/6 db/db mice, confirming our previous observations (19) in C57BL/KS db/db mice. Similarly, decreased humoral immune responses have been previously observed (19, 37) in ob/ob mice, although in one report (37) this reduction was not observed in all experiments. In db/db to WT BMCs, anti-mBSA IgG titers were identical to those measured in db/+ to WT BMCs. Altogether, our observations thus suggest that, in a normal environment, direct effects of leptin on bone marrow-derived cells are dispensable for the induction of cellular and humoral immune responses.

On the contrary, exposure of WT bone marrow-derived cells to the abnormal environment prevalent in db/db mice resulted in a significant reduction in thymus weight and cellularity. Furthermore, T cell populations were decreased in the LN of WT to db/db BMCs, suggesting that the db/db environment may also affect T cell differentiation in the periphery. These observations indicate that leptin receptor deficiency has indirect effects on the immune system, mediated by as yet undefined changes in the environment.

We observed Ob-rb expression in a thymic stromal fraction and in thymic epithelial cells, suggesting that leptin might exert indirect effects on thymocytes not only via systemic modifications, but also by acting locally on the thymic environment and in particular on the thymic epithelium. To investigate the importance of direct effects of leptin on thymic stromal cells, we examined thymocyte development in fetal db/db thymi transplanted into WT recipient mice. T cell differentiation was normal in db/db thymus grafts, indicating that leptin signaling in thymic stromal cells is not required for T cell development in a normal environment.

Systemic modifications influencing the immune system might include various neuroendocrine perturbations observed in db/db mice, including the activation of the hypothalamic-pituitary-adrenal axis and hypercorticosteronemia, decreased activity of the sympathetic nervous system, and/or altered production of different neuropeptides, which are all known to have immunomodulatory effects (38). Excessive corticosteroids favor thymic atrophy; however, the relative proportions of DP, CD4+, CD8+, and DN thymocyte populations were not affected in db/db mice, which is different from the preferential depletion of DP thymocytes induced by corticosteroids (31), suggesting a noncorticosteroid-related mechanism. Consistently, Montez et al. (30) recently described a model of leptin deficiency induced by high-dose leptin treatment of WT mice until fat mass is depleted and, as a consequence, endogenous leptin production is reduced. At this point, exogenous leptin is abruptly withdrawn, inducing a state of leptin deficiency in otherwise normal mice. In this model, a marked nonselective thymic atrophy developed, despite the fact that corticosterone levels were not significantly altered. Interestingly, leptin replacement did not correct the immune phenotype in this model, in which high-dose leptin treatment had induced a state of partial leptin resistance, whereas refeeding restored the thymic cellularity within 2 days, indicating that factors affected by the feeding status, but distinct from leptin, influence thymic cellularity in leptin-resistant mice. Furthermore, it has been recently demonstrated (39, 40) that effects of leptin deficiency on bone metabolism and on the hepatic innate immune system, for instance, are also indirect and, in both cases, are related to the decreased sympathetic tone and beta-adrenergic signaling prevailing in ob/ob mice. Interestingly, we recently observed that chronic administration of the beta-adrenergic receptor antagonist propranolol decreased thymus weight in normal C57BL/6 mice (G. P.almer D. Talabot-Ayer, and C. Gabay, our unpublished data).

Based on our data, we cannot exclude that db/db cells might be even more sensitive to the abnormal environment present in db/db mice than WT cells. In fact, this might represent one explanation as to why thymic atrophy was less severe in WT to db/db BMCs than in db/db mice. It has been reported (12, 24), for instance, that leptin acts directly on T cells to protect them from corticosteroid-induced apoptosis, and it was suggested that this effect is mediated by activation of Stat3 and induction of Bcl-xL. Although our data clearly demonstrate that direct effects of leptin on thymocytes are dispensable in a normal environment, antiapoptotic effects of leptin might still be beneficial in vivo for thymocytes exposed to the high corticosterone levels prevalent in db/db mice. However, Stat3 appears as an unlikely mediator for such effects, because Stat3 signaling in response to leptin is apparently not essential for thymic development in vivo. Indeed, it has been recently reported (41) that s/s mice, which carry a targeted mutation of Ob-rb at Tyr1138 and thus cannot activate Stat3 signaling in response to leptin, display increased thymic cellularity, indicating that disruption of Stat3 activation by Ob-rb actually results in the stimulation of thymocyte production.

In conclusion, our results indicate that direct effects of leptin on bone marrow-derived cells are not required for lymphocyte maturation and immune response in a normal environment. Similarly, direct effects of leptin on thymic stromal cells are also dispensable for T cell development. In contrast, leptin receptor deficiency affects the immune system indirectly, likely via changes in the systemic environment.


   Acknowledgments
 
We thank Pierre-François Piguet for help with the bone marrow transplantation experiments, Sandrine Duffey for expert technical assistance, and Jean-Marc Waldburger and David Moulin for helpful discussions.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
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 grants from the Jean and Linette Warnery Foundation, the de Reuter Foundation, the Academic Society of Geneva, and the Swiss National Science Foundation (Grant 3200–107592/1 to C.G.; 3100–067896.02 to M.A.L.; and 310000–112551 to N.B.). Back

2 Address correspondence and reprint requests to Dr. Cem Gabay, Division of Rheumatology, University Hospital, 26 av. Beau-Séjour, 1211 Geneva 14, Switzerland. E-mail address: Cem.Gabay{at}hcuge.ch Back

3 Abbreviations used in this paper: BMC, bone marrow chimera; WT, wild type, LN, lymph node; DN, double negative; DP, double positive; mBSA, methylated BSA; AU, arbitrary units. Back

Received for publication July 19, 2005. Accepted for publication June 16, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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