Leptin Promotes Differentiation and Survival of Human Dendritic Cells and Licenses Them for Th1 Priming ¹

Leptin, the product of the obese (ob) gene, is a 16-kDa nonglycosylated protein, mainly secreted by adipocytes (1, 2). Structurally, leptin is a member of the helical cytokine family, which includes IL-6, IL-11, IL-12, G-CSF, and has a similar structure to IL-2 (3, 4).

Leptin is released into the circulation, and its plasma levels correlates with total body fat mass and changes in energy balance (5, 6, 7) and is regulated dynamically, which is reduced by fasting (8, 9) and raised by inflammatory mediators such as IL-1 and LPS (10, 11).

The leptin receptor (Ob-R) is encoded by the diabete (db) gene (12, 13) and is a member of the class I cytokine receptor family (14). The Ob-R mRNA is spliced alternatively, giving rise to six different splice forms of the receptor (15, 16). Several in vitro studies have demonstrated that leptin acts directly on the receptor (17), and only the long isoform (Ob-Rb) is thought to be of prime importance in leptin signaling (13), which involves the JAK-STAT pathway for the majority of cytokines (18). Leptin receptor is expressed at the highest relative density in the hypothalamus but also in peripheral tissues such as the kidney, lung, adrenal gland (19), and hemopoietic cells (20). Leptin receptor is also expressed in peripheral T cells (21), NK cells (22), monocyte-macrophages (23), and B cells (24).

The most important role for leptin is considered to be its inhibitory effect on appetite. However, both leptin-deficient (ob/ob) and leptin receptor-deficient (db/db) mice are not only obese. They also develop a complex syndrome characterized by abnormal alterations of hematopoiesis and lymphopoiesis (25, 26). Similar alterations have been found in leptin-deficient and leptin receptor-deficient humans, although those cases are very rare (27, 28). Naturally leptin-deficient obese ob/ob mice display many abnormalities similar to those observed in starved animals and malnourished humans (29), including impaired cell-mediated immune functions (30, 31, 32).

Leptin has been shown to influence CD4⁺ T lymphocyte proliferation and cytokine secretion, shifting immune responses toward the Th1 phenotype and suppressing Th2 responses (33, 21). It has been shown that leptin has an immunomodulatory role on NK cell (22), macrophage (23, 34), and monocyte (35) function, acting as a proinflammatory signal and inducing activation and TNF-α and IL-6 production. More recently, it has been reported that leptin indirectly activates human neutrophils via induction of TNF-α (36).

Dendritic cells (DCs) ³ are the most potent APCs and play a crucial role in the generation and regulation of immunity (37, 38, 39). Their priming ability is acquired upon maturation and is characterized by the activation of different transcriptional factors, leading to the modulation of genes involved in cytoskeleton rearrangement, Ag processing, control of migration, and regulation of inflammatory responses (40, 41, 42, 43). Regulated migration of DCs is central to the induction of physiological immune responses, and this process necessitates plasticity of the cytoskeleton. To date, the majority of studies has focused on the effect of leptin on T lymphocytes, monocytes, NK cells, and polymorphonuclear neutrophils (29, 44, 45). However, the effect of leptin on DCs has not been studied. Because DCs are instrumental in the development of immune responses and prime naive T cells determining their polarization toward Th1 or Th2, the present study was designed to evaluate the expression of leptin receptor on immature and mature DCs and the impact of leptin on their phenotypical, morphological, and functional developmental program and survival.

Human recombinant leptin was purchased from R&D Systems. Lyophilized leptin was dissolved in sterile water as recommended by the manufacturer. Aliquots were prepared and stored at −70°C. A dose-response titration curve was performed to assess the optimal leptin concentration. In all experiments, leptin was added at the beginning of the culture period and left throughout at the concentration of 10 nM. This dose was chosen because of the optimal stimulation and the incorporation of the range of human serum levels. The recombinant leptin was monitored for endotoxin contamination using the Limulus assay (BioWhittaker). Endotoxin levels were <0.03 endotoxin U/ml.

PBMCs were isolated by Ficoll-Hypaque (Flow Laboratories) gradient separation of buffy coats obtained from healthy volunteer blood donors by the Transfusion Center of Università Degli Studi “La Sapienza” Rome. DCs were generated from monocytes purified from PBMCs by positive selection using magnetic cell separation columns and CD14 Microbeads (Miltenyi Biotec). Highly enriched monocytes (>95% CD14⁺) were cultured at 6 × 10⁵/ml in RPMI 1640 medium supplemented with 15% heat-inactivated FCS, l-glutamine, and penicillin-streptomycin, 250 ng/ml GM-CSF (PeproTech), and 500 U/ml IL-4 (R&D Systems) at 37°C for 5 days. Differentiation to DCs was assessed both by morphologic observation and the detection of specific surface markers by flow cytometry. These cells were CD14⁻, CD1a⁺, HLA-DR^int, HLA-ABC^int, CD80^low, and CD86 ^low, consistent with an immature DC phenotype. Untreated immature DCs were used as controls. After 5 days of culture, leptin and/or 200 ng/ml LPS (Escherichia coli serotype 0111:B4, Sigma-Aldrich) were added to immature DCs. LPS-treated DCs were CD83⁺, HLA-DR^high, and HLA-ABC^high, consistent with a mature DC phenotype. Naive CD45RA⁺ T cells were purified from PBMCs by monocytes depletion using magnetic cell separation columns and CD14 microbeads, followed by positive selection using magnetic cell separation columns and CD45RA microbeads (Miltenyi Biotec). The resulting cell preparations were at least 99% viable by trypan blue dye exclusion. The purity of the monocytes and CD45RA⁺ T cells was verified as >90% by direct staining for membrane expression of CD14 and CD45RA/CD3, respectively (anti-CD14, anti-CD3, and anti-CD45RA mAbs were purchased from BD Pharmingen).

Cell staining was performed using mouse mAbs FITC or PE conjugate. The following mAbs were used: CD14 (IgG2a, PE, clone M5E2); CD1a (IgG1, FITC, clone HI149); HLA-DR (IgG2a, FITC, clone G46-6); HLA-ABC (IgG1, FITC, clone G46-2.6); CD80 (IgG1, FITC, clone L307.4); CD86 (IgG1, FITC, clone FUN-1); and CD83 (IgG1, PE, clone HB15e) (all from BD Pharmingen). Samples were analyzed using a FACScan flow cytometer and CellQuest software (BD Biosciences).

RT-PCR for detection of mRNA expression of GAPDH, IL-1β, TNF-α, IL-10, IL-6, IL-12p35, IL-12p40, IL-8, MIP-1α, Bcl-2, and Bcl-x_L was performed. Total cellular RNA was extracted using the RNeasy kit (Qiagen) after 24 h of culture. RNA was reverse transcribed into cDNA and amplified by PCR using the Access RT-PCR System (Promega), according to the manufacturer’s instruction. IL-1β, IL-6, and TNF-α set of primers were purchased from BD Clontech, and IL-12p35 and IL-12p40 set of primers were purchased from Stratagene. GAPDH, IL-8, IL-10, MIP-1α, Bcl-x_L, and Bcl-2 set of primers were synthesized by M-Medical-GENENCO (Cornaredo). For OB-Rb mRNA amplification, a two-round PCR was performed as described in Ref. 46 . Briefly, 300 ng of total RNA were reverse transcribed into cDNA and amplified by PCR using the One-Step RT-PCR kit (BD Clontech), according to the manufacturer’s instruction. For the second-round PCR, 5 μl of the product generated from the first-round PCR were used as template. The first round set of primers were as follows: sense, 5′-AATTGTTCCTGGGCACAAGG-3′, and antisense, 5′-CACAAATCTGAAGGTTTCTTC-3′. The second-round primers were as follows: sense, 5′-AAGATGTTCCGAACCCCAAG-3′, and antisense, 5′-CAAATTTGGACTCTGGTTTCT-3′. To avoid contamination, in all experiments a control RT-PCR without RNA was conducted. For densitometric analysis, the reaction products were size-separated by electrophoresis on 1.8% agarose gel (Bio-Rad). Ethidium bromide-stained gel was analyzed on an UV transilluminator and photographed with positive/negative 665 type film (Polaroid). The negative was scanned by densitometry (Bio-Rad Multi Analyst) as described previously (47). The cytokine mRNA levels were normalized to GAPDH mRNA levels.

Analysis of supernatant cytokine content was performed both on immature and mature DCs and treated or untreated with leptin. Culture supernatants were collected after 24 h of treatment, and IL-1β, IL-6, IL-10, IL-12, and TNF-α contents were measured using a sandwich ELISA (R&D Systems for IL-1β, IL-12, and TNF-α; Euroclone for IL-6; and Pierce for IL-10), according to the manufacturer’s instructions.

Heterologous DCs were used for the MLR experiments. DCs were stimulated for 24 h with leptin, LPS, and LPS + leptin and were then washed extensively and suspended in RPMI 1640 medium supplemented with 10% human serum, l-glutamine, and penicillin/streptomycin, irradiated (3000 rad from a ¹³⁷Cs source), and added in graded doses to 1 × 10⁵ responder T cells in 96 flat-bottom microplates (Corning Glass). Responder cells were heterologous PBMCs. After 5 days, cultures were pulsed for 18 h with 0.5 μCi/well [³H]thymidine (5 Ci/mM; Valeant Pharmaceuticals). Cells were then harvested onto glass fiber filters, and [³H]thymidine incorporation was measured by liquid scintillation spectroscopy.

Naive CD45RA⁺ T cells were cultured in RPMI 1640 medium supplemented with 15% FCS at 10⁶ cells/well in the presence of 25 × 10⁴ untreated or 24 h of leptin-, LPS-, or LPS + leptin-treated autologous or heterologous DCs. Parallel cultures of T cells were cultivated with the supernatants of untreated or 24 h of leptin-, LPS-, or LPS + leptin-treated autologous or heterologous DCs. On day 6, the cultures were stimulated with 30 U/ml IL-2 (R&D Systems). On day 10, T cells were stimulated with 25 ng/ml PMA plus 1 μg/ml ionomycin (Sigma-Aldrich) and 1 μg/ml brefeldin A (Sigma-Aldrich) for the last 4 h of culture. Cells were then fixed with ice-cold 4% paraformaldehyde for 10 min at 4°C and permeabilized with 1 ml of 0.5% Triton X-100 (Sigma-Aldrich) for 5 min at room temperature and stained with PE-labeled anti-human-IFN-γ (R&D Systems) and FITC-labeled anti-human-IL-4 mAb (R&D Systems) and analyzed by flow cytometry.

To monitor NF-κB activation, we used the NF-κB p65/NF-κB p50 transcription factor assay kits (Alexis Biochemicals). Briefly, we used a 96-well plate coated with oligonucleotide containing NF-κB consensus binding site. The activated NF-κB contained in cell extracts from immature and mature DCs, untreated or leptin-treated for 30 min or 1 h, specifically binds to this oligonucleotide. By using an Ab directed against the NF-κB p65 or p50 subunit, the NF-κB complex bound to the oligonucleotide is detected. A secondary Ab conjugated to HRP provides color development, and absorbance is quantified on a spectrophotometer at 450 nm. As a positive control, we used a HeLa cell extract, and to monitor the specificity of the assay, we used NF-κB wild-type and mutated consensus oligonucleotides, according to manufacturer’s instructions. Moreover, localization of p65/NF-κB was detected by static cytometry after 30 min of leptin treatment. Cells were fixed in acetone/methanol 1/1 (v/v) for 10 min at room temperature, stained with polyclonal NF-κB p65 (Santa Cruz Biotechnology), and successively with FITC-labeled anti-rabbit Ab (Sigma-Aldrich). Finally, all samples were plated on coverslips by centrifugation at 2000 rpm for 15 min with Cytospin (Thermo Shandon), mounted with glycerol:PBS (2:1), and analyzed with a Nikon Microphot fluorescence microscope or with intensified video microscopy (IVM) by a charge-coupled device camera (Carl Zeiss).

Untreated and leptin-treated immature and mature DCs were fixed with 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) containing 3% (w/v) sucrose at room temperature for 20 min, plated on poly-l-lysine-coated coverslips for 20 min at room temperature, and postfixed in 1% OsO₄ for 30 min. After washing in buffer, cells were dehydrated through graded ethanols, critical point dried in CO₂, and gold coated by sputtering. The samples were examined with a Cambridge 360 scanning electron microscope.

For actin detection, untreated and leptin-treated immature and mature DCs were fixed with 3.7% formaldehyde in PBS for 10 min at room temperature, permeabilized with 0.5% (v/v) Triton X-100 (Sigma-Aldrich) in PBS for 5 min at room temperature, and, after washing in PBS, stained with fluorescein-phalloidin (Sigma-Aldrich), a toxin capable of direct binding to F-actin.

For OB-Rb detection, cells were stained with anti-OB-Rb polyclonal Ab (Santa Cruz Biotechnology) for 30 min at 4°C, washed in PBS, and incubated with anti-goat polyclonal IgG fluorescein-linked whole Ab (Sigma-Aldrich) at 4°C for 30 min. After washing in PBS, cells were fixed with cold 3.7% formaldehyde in PBS. To analyze the nuclei, all samples were stained with Hoechst 33258 (Sigma-Aldrich) at 37°C for 15 min as described previously (48).

Finally, all samples were plated on coverslips by centrifugation at 2000 rpm for 15 min with Cytospin (Thermo Shandon), mounted with glycerol:PBS (2:1) and analyzed with a Nikon Microphot fluorescence microscope or with IVM by a charge-coupled device camera (Carl Zeiss).

For detection of total STAT3 and OB-Rb protein levels, immature and mature DCs were left untreated or treated with leptin for 24 h and lysed in cold lysis buffer (20 mM HEPES, 50 mM NaCl, 10 mM EDTA, 2 mM EGTA, and 0.5% Nonidet P-40 containing protease and phosphatase inhibitors) for 20 min on ice. Protein concentration of the supernatant was calculated using the Bio-Rad protein assay. Aliquots of 60 μg were loaded on 8% SDS-PAGE and analyzed by Western blot analysis using anti-STAT-3 and anti-OB-Rb polyclonal Ab (Santa Cruz Biotechnology). Immunoreactive bands were visualized by using a secondary HRP-conjugated anti-goat Ab and Opti-4CNT Detection kit (Bio-Rad). For phosphorylated (p)-STAT3 immunoprecipitation, cells were lysed in cold RIPA buffer (150 mM NaCl, 1% Nonidet P-40, 0,5% sodium deoxycholate, 1% SDS, and 50 mM Tris (pH 8) containing protease and phosphatase inhibitors) for 30 min on ice and then centrifuged at 10,000 × g for 10 min. Lysates were precleared by 10 min of incubation with 2 μl of protein A/G agarose conjugated bead slurry (Santa Cruz Biotechnology), followed by 10 min of centrifugation at 14,000 × g. The cell lysates protein concentration was then determined, and 1 mg of total cellular protein was incubated with 2 μg of anti-STAT3 polyclonal Ab for 2 h at 4°C. The samples were then adsorbed to protein A/G agarose-conjugated suspension for 2 h at 4°C with rocking. The immunoprecipitates were then collected by centrifugation, washed with RIPA buffer, and resolved by SDS-PAGE and transferred to nylon membranes for immunoblot analysis using anti-p-Tyr mAb (Santa Cruz Biotechnology). Immunoreactive bands were visualized as described above. Densitometric analysis was performed as described previously (47).

Untreated and leptin-treated immature and mature DCs were exposed to UVB (1200 J/m²) from a Philips TL 20W/12 lamp. Leptin treatment was performed 30 min before and during UVB exposure and for the following 24 h. The dishes were placed without covers 10 cm below the lamp. To eliminate UV radiation not in the UVB range, a Kodak filter (type Kodacell TL 401) was used. In these conditions, the UVB radiant flux density was measured with an Osram Centra radiometer.

FITC-annexin V/propidium iodide (PI) double staining was used to detect apoptosis both spontaneous and induced by a UVB treatment as described before. Immature and mature DCs, untreated or 24 h leptin-treated, were harvested and washed twice with ice-cold PBS, and specific binding of FITC-annexin V and staining with PI was performed with an apoptosis detection kit (BD Pharmingen), according to the manufacturer’s instructions. The cells were then analyzed by flow cytometry. Late apoptosis, spontaneous and UVB induced, was also detected by Hoechst staining. Briefly, after 24 h of leptin treatment, cells were fixed with 3.7% formaldehyde in PBS for 10 min at room temperature, stained with Hoechst 33258 (Sigma-Aldrich) at 37°C for 15 min, and analyzed with a Nikon Microphot fluorescence microscope.

Statistical analysis was calculated using a two-tailed Student’s t test. A value of p < 0.05 was considered as statistically significant.

To determine the expression of leptin receptor, levels of OB-Rb transcripts were detected on immature or mature DCs left untreated or treated with leptin for 24 h. We found that both immature and mature DCs express the leptin receptor (Fig. 1,A). Notably, OB-Rb mRNA levels were comparable between immature and mature DCs and, in both cases, were up-regulated by leptin treatment (Fig. 1,A). These findings were confirmed by analyzing the OB-Rb protein levels by Western blot analysis (Fig. 1,B). The presence of Ob-Rb receptor on immature and mature DCs was also detected by indirect immunohistochemistry staining (Fig. 1,C). The presence of OB-Rb on human DCs is an indication of a possible responsiveness of these cells to leptin stimulation. It is known that a major consequence of leptin binding to its receptor is activation of JAK/STAT pathway that leads to STAT3 phosphorylation (49). Therefore, we evaluated the functionality of OB-Rb in DCs, analyzing levels of p-STAT3. As shown in Fig. 1 D, 30 min of leptin treatment scored a 1.6-fold increase of p-STAT3 level in immature DCs compared with control DCs. LPS or LPS + leptin stimulation scored 1.4- and 1.9-fold increase, respectively. These results indicate that signaling upon OB-Rb binding by leptin is activated.

To investigate whether leptin induces phenotypic differentiation of human DCs, immature and maturing DCs were cultured with leptin for 24 h and then analyzed for surface molecule expression.

Despite some variability between donors, there was no consistent modulation of any marker in response to leptin (Fig. 2). In particular, leptin did not induce changes in immature (Fig. 2,A) and mature (Fig. 2 B) DCs on the surface expression of CD1a and HLA class I and II molecules involved in the presentation of lipidic and antigenic peptides, respectively. Similarly leptin did not have an impact on the expression of costimulatory/signaling molecules CD86 and CD80 and on the maturation Ag CD83. Therefore, leptin does not induce phenotypical maturation of DCs and does not interfere with LPS-induced maturation.

Because the stimuli that induce DC differentiation generally stimulate secretion of cytokines and chemokines by these cells, we analyzed the pattern of the production of these mediators by DCs both at protein secretion and mRNA synthesis levels.

Fig. 3 shows that a 24-h treatment with leptin triggered a statistically significant increase in the production of IL-1β, IL-6, and IL-12 on immature and mature DCs, whereas TNF-α secretion was up-regulated only in immature DCs. In contrast, leptin down-regulated IL-10 secretion both by immature and mature DCs.

We then investigated how cytokine mRNA synthesis would be affected by leptin (Fig. 4). The semiquantitative RT-PCR analysis performed on immature and mature DCs showed that IL-1β, IL-6, and TNF-α mRNA transcription was up-regulated after 24 h of leptin treatment, whereas mRNA for IL-10 was down-regulated. Moreover, leptin up-regulated IL-12p40 and IL-12p35 mRNA transcription in mature DCs.

We also analyzed the expression profile of the inflammatory chemokine MIP-1α and IL-8 mRNA. We found that in immature and mature DCs leptin up-regulated MIP-1α transcript levels, while leptin down-regulated IL-8 transcript levels after 24 h of treatment (Fig. 4).

Because leptin up-regulated the production of cytokines and chemokines that are critical for the T cell stimulatory function of DCs, we asked whether leptin-treated DCs could be able to stimulate T lymphocytes. To this end, we set up a heterologous MLR between PBMCs and irradiated heterologous DCs left untreated or treated with leptin (Fig. 5,A), LPS, or LPS + leptin (Fig. 5,B) for 24 h. As shown in Fig. 5, despite the comparable levels of presentation and costimulatory/signaling molecules observed in leptin-treated DCs, the ability of leptin-treated immature DCs to induce T cell proliferation was higher than untreated DCs. Notably, these results are referred to the same donors showing a variable expression of MHC-class I and II molecules (Fig. 2). Simultaneous treatment with leptin and LPS resulted in a more vigorous response than observed with any other single stimulus.

DC maturation is accompanied by morphological changes, including acquisition of high cellular motility associated with loss of adhesive structures and cytoskeleton reorganization (40). In the present study, we investigated whether the leptin-induced DC functional differentiation occurs with morphological remodelling. Fig. 6 A shows that 24 h of leptin-, LPS-, and LPS + leptin-treated DCs developed a series of evident modifications of cell surface structures as detected by SEM analysis. Although untreated immature DCs appeared round and with small microvillous structures on the cell surface, leptin-treated immature DCs developed a polarized morphology (uropod formation), which is typical of cells that undergo migration, activation, and cell-to-cell interaction. Similar morphology was observed in mature DCs that in addition showed consistent formation of ruffles typical of mature DCs. No significant differences were visible between LPS- and LPS + leptin-treated DCs.

Cytoskeleton rearrangement is considered as a prerequisite for the occurrence of DC morphological remodeling. In particular, microfilament network plays a key role in the formation of both uropods and surface ruffling (40). Thus, we analyzed the F-actin network in leptin-, LPS-, and LPS + leptin-treated DCs. As illustrated in Fig. 6 B, while in untreated immature DCs the actin filaments appeared organized to form a ring in the subcortical region, leptin treatment induced a polarization of the actin filaments, especially in the uropod region, favoring adhesion and cell contact. Similar organization of actin filaments was detected in LPS- and LPS + leptin-treated DCs.

It is well known that leptin elicits a shift toward Th1 responses, thus we asked whether leptin-treated DCs were able to prime naive T lymphocytes and induce Th1 polarization. To test this hypothesis, we set up a MLR in which purified CD4⁺CD45RA⁺ T cells were stimulated with heterologous DCs left untreated or treated for 24 h with leptin, LPS, or LPS + leptin. We then analyzed the pattern of IL-4 and IFN-γ production by T lymphocytes primed by DCs or by the supernatants of the same DC cultures. As shown in Fig. 7,A, T cells stimulated by leptin- or LPS + leptin-treated DCs differentiated mainly into IFN-γ-producing cells compared with those stimulated by untreated or LPS-treated DCs, respectively. Similar results were obtained by stimulation with supernatants (Fig. 7 B). The same experiment was done with autologous DCs and culture supernatant, obtaining the same trend, although a lower rate of polarization (data not shown). These results show that leptin-treated DCs drive T cells polarization toward the Th1 phenotype. Moreover, leptin added together with LPS, a known Th1-polarizing stimuli, further strengthens its polarizing effects.

It has been demonstrated that leptin has antiapoptotic effects on T cells and monocytes by up-regulating prosurvival signaling (50, 51). Therefore, we evaluated whether leptin protects DCs from spontaneous and induced apoptosis. To this aim, immature DCs were left untreated or treated for 24 h with leptin, LPS, or LPS + leptin, and quantitative evaluation of apoptosis was performed by flow cytometry by double staining with PI and annexin V. As shown in Fig. 8,A, leptin significantly reduced the percentage of annexin V⁺/PI⁻ (early apoptosis) and annexin V⁺/PI⁺ DCs (late apoptosis) after 24 h. We then analyzed the effect of leptin on induced apoptosis. As apoptotic stimulus, we used UVB radiation that is known to trigger apoptosis of human DCs very efficiently and cause immunosuppression (52). We found that leptin significantly counteracts UVB-induced apoptosis in immature DCs, reducing the percentage of annexin V⁺/PI⁻ and annexin V⁺/PI⁺ cells after 24 h (Fig. 8 B). In LPS-treated DCs, leptin exerted a weaker antiapoptotic effect. These results show that leptin promotes DCs survival and partially protects them from induced apoptosis process.

After 24 h of treatment with leptin, bcl-2 and bcl-x_L mRNA levels in immature and mature DCs were determined by semiquantitative RT-PCR. Consistent with the observed antiapoptotic effect of leptin, we observed the up-regulation of bcl2 and bcl-x_L mRNA transcription. Leptin scored a 1.3-fold increase of bcl-2 and a 1.7-fold increase of bcl-x_L mRNA levels in immature DCs. LPS or LPS + leptin stimulation scored 1.3- and 1.9-fold increase, respectively, of bcl-x_L mRNA levels, whereas bcl-2 transcripts were not affected by leptin (Fig. 9 A).

Because NF-κB is a transcription factor involved in cell survival signals, we analyzed the rates of NF-κB activation in total lysates from untreated or leptin-, LPS-, or LPS + leptin-treated DCs by measuring the levels of both NF-κB p50 and p65 subunits capable of binding an oligonucleotide containing the NF-κB consensus binding site. As shown in Fig. 9 B in immature DCs, leptin induced a 1.6-fold increase of active p50 levels after 30 min of treatment and 1.7-fold increase after 1 h, whereas the p65 levels scored a 1.3-fold increase after 30 min and 1.4-fold increase after 1 h. In mature DCs NF-κB p50 and p65 subunits were not affected by leptin treatment. As a positive control, we showed levels of activation of NF-κB in HeLa cell extracts. The assay was specific because incubation of HeLa cell extract in the presence of a nonbound wild-type consensus oligonucleotide abolished binding of both subunits, whereas with the mutated consensus oligonucleotide, the binding was equal to control HeLa cells.

Cells from the same culture were also analyzed by static cytometry to detect p65 subunit translocation to the nucleus (Fig. 9,C). As shown, after 30 min of treatment of immature DCs with leptin, positive nuclei for p65 NF-κB subunit are increased. The results obtained indicate that after leptin treatment, 80% of immature DCs showed nuclear p65 subunit localization. By contrast, in untreated immature DCs, only 28% of cells showed nuclear p65. In mature DCs, positive cells were 90%, and leptin treatment during maturation did not affect the translocation of p65 subunit (data not shown). In Fig. 9 C are depicted two fields representative of untreated and leptin-treated immature DCs.

In this study, we outline that leptin oversees the DC developmental program inducing a coordinate series of functional and morphological changes in these cells, making them more competent APCs and licensing them toward Th1 priming. We have also provided evidence that leptin promotes DC survival up-regulating antiapoptotic gene expression.

In the present study, we show, for the first time, the presence of Ob-Rb on DCs, a cellular population widely acknowledged as a master for induction and regulation of immune responses. Specifically, we found that immature and mature DCs express comparable levels of Ob-Rb as determined both by mRNA and protein analysis. Interestingly, the expression of Ob-Rb was up-regulated by leptin treatment. Therefore, we checked whether the leptin receptor expressed on DCs was active biologically in terms of the ability to transduce signaling pathways, and we found that leptin, upon binding to its receptor, triggers STAT-3 phosphorylation.

The presence of a functionally Ob-Rb on DCs led us to investigate the impact of leptin on phenotypical, morphological, and functional differentiation of these cells. In immature and mature DCs, we found no consistent modulation of surface molecules known to be critical for their APC function, i.e., CD1a, CD80, CD83, and CD86 in response to leptin. The effect of leptin on the expression of presentation molecules HLA-ABC and HLA-DR was variable among the different donors. Despite the lack of phenotypic modulation, leptin turned out to induce functional consequences in immature and mature DCs up-regulating cytokine and chemokine production, as well as T cell stimulatory capacity. Specifically, we found that leptin induces a significant up-regulation of IL-12 in mature DCs and of IL-1β, IL-6, and TNF-α mRNA synthesis levels in immature and mature DCs. The up-regulation of cytokine production induced by leptin might sustain the activation of cells in charge of innate and adaptive immunity. In particular, the up-regulation of TNF-α can promote the activation and function of neutrophils, and the up-regulation of IL-12 can favor NK cells activation while TNF-α, IL-6, and IL-1β up-regulation can activate monocytes/macrophages. In contrast, the leptin-induced up-regulation of cytokine production by DCs might promote adaptive immunity through the activation of bystander T cells. As expected, we actually found that leptin significantly up-regulates T cell stimulatory capacity of both immature and mature DCs, despite the comparable levels of presentation and costimulatory/signaling molecules observed in leptin-treated DCs. We found that IL-10 production was down-regulated significantly by leptin. This, together with the observed up-regulation of IL-12 production, implies a bias toward Th1 immune responses. It has been recognized that cells belonging to the myelomonocytic differentiation pathway, including macrophages and DCs, have a key role in polarized innate and adaptive immune responses. They act promoting polarization toward type I or II immune responses and expressing specialized and polarized effector functions. Therefore, we have verified whether leptin licenses DCs for Th1 priming. We indeed found that leptin-pulsed DCs support a significant increase in the production of the Th1-type cytokine IFN-γ by naive T cells. Notably, a similar trend was observed using leptin-pulsed DC supernatants, indicating that both cell-to-cell contact and mainly soluble factors are involved in the Th1 polarization. Experiments are in progress to assess the mediators responsible for this phenomenon. These data are in line with previous results demonstrating that leptin promotes Th1 cytokine production by T cells (45). In the present study, we show that leptin by a direct effect on DCs drives the Th1 cell differentiation.

During type I and II immune responses, master cytokines regulate chemokine production, which appears to be also involved in immune responses polarization (53). Chemokines can support selective recruitment of polarized T cells and specific I or II effector cells expressing distinct panels of chemokine receptors (54). Therefore, we checked whether leptin impacts on chemokine production, and we found that leptin significantly up-regulates MIP-1α production by immature and mature DCs. This is relevant considering that the MIP-1α receptor CCR5 is expressed preferentially on polarized type I T cells as well as on NKs, eosinophils, monocytes, and type I macrophages (M1). Macrophages can be classified as M1 or M2, as reported by Mantovani et al. (54), in analogy to Th1 and Th2 nomenclature, and are polarized by the cytokines IFN-γ or TNF-α and IL-10, respectively. Polarized macrophages differ in terms of effector functions as well as chemokines production and receptor expression. M1 express CCR5 and are potent effector cells integrated in Th1 responses, whereas M2 express IL-8Rs CXCR1 and CXCR2 and tune Th2 responses. This fits with our data showing that leptin down-regulates the production of IL-8 by immature and mature DCs, while leptin up-regulates MIP-1α production, contributing to type I response polarization.

It has been demonstrated that DCs, unlike other APCs, actively reorganize their actin cytoskeleton during interaction with T cells (55). DC cytoskeletal rearrangement is a prerequisite for the occurrence of DC morphological remodeling, leading to the acquisition of motility, and is critical for clustering and activation of resting T cells. In the present study, we found that leptin-treated immature DCs develop a polarized morphology with the formation of uropods. Moreover, leptin treatment induces a polarization of the actin filaments, especially in the uropode region, favoring adhesion and cell contact. These results indicate that leptin triggers the morphological developmental program of DCs.

It has been demonstrated that leptin inhibits stress-induced apoptosis of T lymphocytes by the up-regulation of the bcl-x_L gene (50). Moreover, Najib and Sanchez-Margalet (51) reported that leptin promotes survival of human-circulating monocytes prone to apoptosis by activation of the p42/44 MAPK pathway. In the present study, we found that leptin promotes survival and expression of the antiapoptotic proteins bcl-2 and bcl-x_L in DCs associated with NF-κB activation. This data suggest that leptin may act as a trophic factor for the survival of DCs.

This work provides new insights about the role of leptin in immune system homeostasis and highlights the ability of leptin to modulate immunologic outcome. Because of its peculiar activating property on DCs, leptin can actually represent an invaluable modulator and supervisor of the immune system.

It is known that leptin influences both innate and adaptive immunity modulating the activity and the function of different cell types (56). DCs are components of the innate immune system and organize and transfer information from the outside world to the cells of the adaptive immune system representing, indeed, a link between the two arms of immune responses. Therefore, our findings that DCs are responsive to leptin contribute to the understanding of the mechanism by which leptin balances the immune surveillance and metabolism.

Of note, these data strengthen the previous proposal of a potential use of leptin as a therapeutic agent in several immune-mediated diseases (57, 58). In particular, clinical applications could be hypothesized in immunodeficiency associated with reduced food intake such as anorexia nervosa or HIV-1 infection where leptin levels are reduced dramatically, as well as CD4⁺ T cells number and function (59). In these patients, leptin administration could be helpful for the immunoreconstitution process. In the context of allergic diseases, immune deviation from a Th2 to a Th1 response may be a suitable regulatory strategy for treatment of these pathologies (60, 61). Therefore, the observed leptin-induced Th1 polarization could suggest its use in treatment and prevention of allergic diseases.

Recently, increasing emphasis has emerged on the development of novel adjuvant immunopotentiators able to shift and modulate natural immune responses. Th1 or Th2 polarization is a critical step that defines the efficacy of vaccine-mounted immune response to eliminate pathogens and induce protection. Therefore, our findings additionally provide a rationale for the use of leptin in vaccination protocols as an adjuvant to efficiently boost Th1 type responses in infectious diseases.

The authors have no financial conflict of interest.