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Published as doi: 10.1096/fj.07-098095.
 (The FASEB Journal. 2008;22:2012-2022.)
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Leptin as an immunological adjuvant: enhanced migratory and CD8+ T cell stimulatory capacity of human dendritic cells exposed to leptin

Benedetta Mattioli, Elisabetta Straface, Paola Matarrese, Maria Giovanna Quaranta, Luciana Giordani, Walter Malorni and Marina Viora1

Department of Drug Research and Evaluation, Istituto Superiore di Sanità, Rome, Italy

1Correspondence: Department of Drug Research and Evaluation, Istituto Superiore di Sanità, 299 Viale Regina Elena, 00161 Rome, Italy. E-mail: viora{at}iss.it


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Leptin is an adipocyte-derived hormone/cytokine that links nutrition, metabolism, and immune homeostasis and is endowed to modulate several immune responses. We previously demonstrated that both immature and mature human dendritic cells (DCs) express a functional leptin receptor, and we found that leptin activates DCs, licenses them for Th1 priming, and promotes DC survival. Moreover, we found that leptin induces rearrangement of actin microfilaments, leading to uropod and ruffle formation. Here we monitor the effects of leptin on DC migratory capacities, focusing on the intracellular signaling driving cytoskeleton rearrangement. We found that leptin increases immature DC migratory performance both by favoring cytoskeleton dynamics and by up-regulating CCR7 surface expression, thus favoring chemotactic responsiveness. We found that in immature DCs, leptin activates cofilin, favoring the turnover of actin microfilaments, and, by triggering Vav phosphorylation, promotes Rac1 activation. Finally, we found that in immature DCs, leptin up-regulates interleukin-12p70 production on CD40 stimulation and, more importantly, increases their capacity to stimulate activation of autologous CD8+ T cells. Taken altogether, the findings herein highlight the potential use of leptin as an adjuvant tool in vaccination protocols employing ex vivo-generated autologous DCs.—Mattioli, B., Straface, E., Matarrese, P., Quaranta, M. G., Giordani, L., Malorni, W., Viora, M. Leptin as an immunological adjuvant: enhanced migratory and CD8+T cell stimulatory capacity of human dendritic cells exposed to leptin.


Key Words: chemotaxis • vaccination • cytoskeleton • CTL


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
LEPTIN, THE PRODUCT OF 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, including interleukin (IL) -6, IL-11, IL-12, and granulocyte-macrophage colony-stimulating factor (3 , 4) .

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

Leptin acts by binding its specific receptor ObR (12 13 14) , which is expressed at highest relative density in the hypothalamus but also in peripheral tissues (15) and hemopoietic cells (16) . Moreover, ObR is expressed in different cell populations of the immune system (17 18 19) , and leptin has been shown to act on innate as well as acquired immunity (17 18 19 , 20 21 22 23) . We recently demonstrated that both human immature and mature DCs express leptin receptor in a biologically active form able to transduce leptin intracellular signals (24) , rendering DCs responsive to leptin. Indeed, we found that leptin acts on DCs, improving their functional activity and promoting Th1 priming and survival. Recently, Lam et al. (25) obtained similar results in an in vitro murine system, and Macia et al. (26) demonstrated that leptin-deficient obese mice show an alteration of DC function and steady-state number.

DCs are the most potent antigen-presenting cells (APCs), acknowledged as a master for induction and regulation of immune responses (27 28 29) . DCs are present in peripheral tissues, where they are poised to capture antigens. As the antigens are processed, the DCs mature and move toward the draining secondary lymphoid organs, where they present the peptides to naive T cells, thereby inducing a cellular immune response that involves both CD4+ and CD8+ T cells. Their priming ability is acquired on maturation and is characterized by the modulation of genes involved in cytoskeleton rearrangement, antigen processing, control of migration, and regulation of inflammatory responses (30 31 32 33) . DCs are also critical in launching humoral immunity and in activating natural killer cells.

A therapeutic approach for tumor vaccination, aimed to induce patient immune response against autologous tumors, involves the injection into cancer patients of DCs, generated and loaded with tumor antigens ex vivo. Many clinical trials have used monocyte-derived DCs and have demonstrated the necessity to activate them for cytotoxic T lymphocyte (CTL) induction (34 35 36) and have revealed their limited capacity to migrate to the lymph nodes (LNs) (37 , 38) , an event known to be controlled by the chemokine receptor CCR7. It is well established that DC cytoskeletal rearrangement is a prerequisite for the occurrence of DC morphological remodeling leading to the acquisition of motility. In this context, we previously demonstrated (24) that the treatment of immature DCs with leptin induces morphological changes such as actin microfilament rearrangement leading to uropod and ruffle formation.

The present study was designed to further investigate the effects of leptin on DC migratory capacities with the goal of evaluating a possible use of leptin as an adjuvant in DC-based vaccines.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Leptin
Human recombinant leptin was purchased from R&D Systems Inc. (Minneapolis, MN, USA). A dose-response titration curve was performed to assess the optimal leptin concentration. Leptin was added at the beginning of the culture period and left throughout at the concentration of 10 nM. This dose gives an optimal stimulation and falls into the range of human serum (HS) levels. Endotoxin levels were <0.03 endotoxin units/ml as assessed by Limulus assay (BioWhittaker, Inc., Walkersville, MD, USA).

Cell culture
Peripheral blood mononuclear cells (PBMCs) were isolated by Ficoll-Hypaque (Flow Laboratories, Hornby, ON, Canada) gradient separation of buffy coats obtained from healthy volunteer blood donors by the Transfusion Center of the University of Rome. DCs were generated as described previously (24) . 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 Corp., St Louis, MO, USA), was added to immature DCs. CD8+ T cells were purified from PBMC by positive selection, using magnetic cell separation columns and CD8 Microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). The purity of the monocytes and CD8+ T cells was verified as >90%. J558-CD40L-transfected cells were cultured in RPMI 1640, 10% fetal calf serum (FCS), and 0.01 M histidinol (Sigma) at a concentration of 3 x 105 cells/ml (39) .

Chemotaxis assay
The in vitro migration of untreated, leptin- or LPS-treated DCs in response to CCL19 and CCL4 chemokines (PeproTech Inc., Rocky Hill, NJ, USA) was assessed in a Transwell cell culture chamber with 3.0 µm pore size polycarbonate filters (BD Falcon, Franklin Lakes, NJ, USA). DCs (5x105) in RPMI 1640 were added to the upper chamber. CCL19 or CCL4 diluted in RPMI 1640 at the indicated doses (50–200 ng/ml), or medium alone as a control for spontaneous migration, was loaded in the lower compartment. As negative controls, untreated, leptin-treated, or LPS-treated DCs were incubated 1 h at 37°C before the migration assay with saturating amounts (5 µg/ml) of CCL19 and CCL4 to block CCR7 and CCR5 chemokine receptors, respectively.

After 4 h incubation, the cells that had migrated to the bottom chamber were collected and counted by flow cytometry in a FACScalibur, acquiring events for a fixed time period of 60 s using CellQuest software (Becton Dickinson, Franklin Lakes, NJ, USA). Each experiment was performed in duplicate. Values represent percentages of migrated cells, presented as means ± SD.

Flow cytometry
Cell staining was performed using mouse monoclonal Abs (mAbs) fluorescein isothiocyanate (FITC) or phycoerythrin (PE) -conjugated. The following mAbs were used: anti-CD14, -CD1a, -human leukocyte antigen (HLA) -DR, -HLA-ABC, -CD40, -CD80, -CD86, -CD83, -CD8, and -CCR5. CCR7 was stained using an unlabeled primary mAb, and the secondary polyclonal Abs (pAbs) immunoglobulin (Ig)G+IgM. Intracellular staining of perforin was performed using the BD Cytofix Cytoperm Plus Kit (Becton Dickinson Biosciences, San Jose, CA, USA) and anti-perforin mAb; all mAbs were purchased from Pharmingen (San Diego, CA, USA). Samples were analyzed using a FACScalibur flow cytometer and CellQuest software (Becton Dickinson).

The analysis of microfilament system dynamic was performed by quantification of filamentous (F), globular (G), and total actin as described previously (40) .

Immunofluorescence
For CCR7 surface detection, cells were stained with anti-CCR7 mAb for 30 min at 4°C, washed in PBS, incubated with anti-mouse IgG fluorescein-linked whole Ab (Sigma) for 30 min at 4°C, washed in PBS, and then fixed in cold 3.7% formaldehyde.

For intracellular P-cofilin and Rac detection, cells were fixed with 3.7% formaldehyde, permeabilized 5 min with Triton X-100 (both Sigma), stained with anti-P-cofilin pAb (Cell Signaling Technology, Danvers, MA, USA) or anti-Ras-related C3 botulinum toxin substrate 1 (Rac1) mAb (Transduction Laboratories, Lexington, KY, USA) for 30 min at 37°C, and then incubated with anti-rabbit or anti-mouse IgG fluorescein-linked whole Ab (Sigma) at 37°C for 30 min.

For actin detection, cells were stained with fluorescein-phalloidin (Sigma) as described previously (24) .

Western blot and affinity precipitation
For detection of Vav, P-Vav, cofilin, P-cofilin, and actin protein levels, immature DCs were left untreated or treated with leptin or LPS for different times, then lysed, and protein concentration was calculated using the DC Protein Assay (Bio-Rad Laboratories, Hercules, CA, USA). Samples were loaded on 8% SDS-PAGE and analyzed by Western blot using anti-Vav or anti-P-Vav pAb (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA), anticofilin or anti-P-cofilin pAb (Cell Signaling Technology), and anti-actin pAb (Sigma). Immunoreactive bands were visualized using a secondary horseradish-peroxidase-conjugated anti-rabbit or anti-mouse pAb and the Western blot Detection Kit, Chemioluminescent system (Upstate Biotechnology, Lake Placid, NY, USA). Active Rac1 levels were determined as described previously (41) . In some experiments, DCs were pretreated for 30 min at 37°C with 10 µM PP1 [4-amino-5-(4-methylphenyl)-7-(t-butyl)pyrazolo-D-3,4-pyrimidine] (Alexis Biochemicals, Lausen, Switzerland) in order to inhibit Src tyrosine kinases (42) .

Cocultures
For the DC/J558-CD40L cocultures, J558-CD40L cells cultured in RPMI 1640, 10% FCS, and 0.01 M histidinol were collected and washed to eliminate all histidinol and resuspended in RPMI 1640, 10% FCS at a concentration of 106cells/ml. DCs untreated or treated for 24 h with leptin or LPS were collected and resuspended in RPMI 1640, 10% FCS, at a concentration of 106cells/ml. After 24 h, a set of samples was used for phenotype analysis by flow cytometry, and the supernatants were used for IL-12p70 evaluation. Another set of samples was collected, washed, and resuspended in RPMI 1640, 10% FCS at a concentration of 106 cells/ml. Cells were then either recultured for the following 24 h or cocultured at a 1:5 DC:J558-CD40L ratio in a 24-well plate. After 24 h, supernatants were collected for IL-12p70 evaluation, and DC phenotype was analyzed by flow cytometry.

For the DC-CD8+ T cell cocultures, both DCs and CD8+ T cells were cultured in 10% HS at 1.5 x 106 cells/ml and cocultured at 1:1 DC:CD8+ T cell in a 24-well plate.

Autologous mixed lymphocyte reaction (MLR)
Autologous DCs were used for the MLR experiments. DCs were stimulated for 24 h with leptin or LPS and were then extensively washed and suspended in RPMI 1640 supplemented with 10% HS, L-glutamine, and penicillin/streptomycin. After irradiation (3000 rad from a 137Cs source), DCs were added in graded doses to 1 x 105 autologous responder CD8+ T cells in 96 flat-bottom microplates (Corning Costar Corp., Cambridge, MA, USA). After 5 days, cultures were pulsed for 18 h with 0.5 µCi/well of [3H]thymidine (5 Ci/mM; ICN, Irvine, CA, USA). Cells were then harvested onto glass fiber filters, and [3H]thymidine incorporation was measured by liquid scintillation spectroscopy.

Cytokine assay
Analysis of supernatant cytokine content was performed using a sandwich ELISA (Euroclone; Pero, Italy). After 24 or 48 h of DC/CD8 or DC/J558-CD40L coculture, supernatants were collected, and IFN-{gamma} or IL-12p70 contents, respectively, were measured according to the manufacturer’s instructions.

Data analysis and statistics
Statistical analysis was calculated using a 2-tailed Student’s t test. A value of P <0.05 was considered statistically significant. In the flow cytometry experiments for quantification of actin microfilaments, a comparison between single samples was performed by CellQuest software using the parametric Kolmogorov-Smirnov (K/S) test on a population of at least 2 x 105 cells. Values of P < 0.01 were considered significant. In the chemotaxis assay, 2-way ANOVA was used.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Leptin improves immature DC migratory capacity
As known, DCs developmentally regulate the expression of chemokine receptors to facilitate their migration from the peripheral tissues to regional LNs. In immature DCs, the chemotaxis is mediated by inflammatory chemokine receptor CCR5, which leads migration in response to its ligand CCL4, whereas mature DCs, after the receptor switch that occurs during maturation, express high levels of CCR7 and migrate to its ligand CCL19. The capacity of leptin to favor DC migratory capacities and chemotactic responsiveness was checked in an in vitro chemotaxis assay. To this aim, we set up experiments using a 96-transwell chemotaxis chamber. RPMI 1640 and different concentrations of the probed chemokines, CCL19 or CCL4, were loaded into the lower chamber, while untreated or 24 h leptin- or LPS-treated DCs were loaded into the upper chamber and incubated for 4 h to allow transmigration through the filter. As shown in Fig. 1 A, leptin- or LPS-treated DCs, in the absence of chemokine stimulation, scored significantly higher percentages of spontaneous migration (24±1.9 and 32±1.3%, respectively) compared with control DCs (10±1.8%). The addition of CCL19 induced an increase in the percentage of migrating leptin- and LPS-treated DCs in a dose-dependent manner, whereas control DCs were not affected. This result shows that leptin treatment, as for LPS, not only increases spontaneous migration, but the addition of CCL19 extends the gap between the percentages of migrating leptin-treated and control DCs. As negative controls, untreated, leptin-treated, or LPS-treated DCs were preincubated 1 h at 37°C with saturating amounts of CCL19 to block CCR7 receptors. As shown in Fig. 1A , this pretreatment did not affect the percentage of migrating control DCs, whereas it abrogated CCL19 chemotactic effects on leptin- and LPS-treated DCs. To verify whether the effect of CCL19 was specific, we also evaluated the effects of CCL4 on DC migration. Differently from CCL19, the addition of CCL4 did not affect the percentage of LPS-treated migrating DCs at any dose tested. On the other hand, both control and leptin-treated DCs increased the percentage of migrating cells with the addition of CCL4 in a dose-dependent manner. The percentage of migrating leptin-treated DCs was significantly higher than control DCs at all doses. As for CCL19, a negative control incubated with saturating amounts of CCL4 was set up. As shown in Fig. 1A (negative control), this pretreatment did not affect the percentage of migrating LPS-treated DCs, whereas it abrogated CCL4 chemotactic effects on control or leptin-treated DCs.


Figure 1
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  		Figure 1. Leptin improves immature DC migration and up-regulates CCR7 expression on immature DCs. A) The chemotactic responsiveness of immature DCs toward CCL19 and CCL4 was tested by performing migration assay using 24-well chemotaxis chambers with 3 µm pore polycarbonate filters. Immature DCs (5x105) suspended in medium were loaded into the upper well, and RPMI 1640 alone or containing the probed chemokines at the indicated concentrations (50–200 ng/ml) was placed in the lower wells. DCs that had migrated through the filter to the bottom chamber were collected after 4 h and counted by flow cytometry. Results are expressed as percentage of migrated cells (means±SD); results from 3 different experiments are shown. Two-way ANOVA demonstrated significant effects for treatment (P<0.0001), dose (P<0.0001) and for treatment x dose interaction (P<0.0001). B) DCs were left untreated (CTR) or were treated with leptin or LPS for 24 h. DCs were then analyzed for surface expression of CCR7 and CCR5 by flow cytometry. Histograms show 1 representative experiment of 4. Values are presented as means ± SE and express the percentage of positive cells (%) and mean fluorescence intensity (MFI) from 4 different experiments. C) Intensified video microscopy (IMV) analysis of CCR7 surface expression in untreated (CTR) and 24 h leptin- or LPS-treated DCs, is shown. While untreated DCs exhibit a lower and diffused distribution of CCR7, leptin treatment up-regulates CCR7 expression and induced a clustered distribution of the protein. In LPS-treated DCs, the CCR7 receptor appears as diffuse dot spots. Original image x1800.

Leptin up-regulates CCR7 expression on immature DCs
Results obtained from the migration test showed that leptin improves DC migration both in the presence or absence of CCL19 stimulation. Therefore, leptin could interfere both with chemotactic responsiveness, acting on the chemokine receptor CCR7 expression, and with cell mobility, acting on cytoskeleton dynamics. We first investigated CCR7 surface expression on immature DCs left untreated or treated with leptin or LPS for 24 h. As shown in Fig. 1B , leptin significantly (P<0.05) up-regulated the CCR7 expression on immature DCs. As expected, LPS reached the maximum expression level. This finding was confirmed by the analysis of CCR7 surface expression by indirect immunocytochemistry staining (Fig. 1C ). Of note, a different surface distribution was detectable between leptin- and LPS-treated DCs: leptin induced a clustered distribution, whereas in LPS-treated DCs, the expression of CCR7 receptor appeared as diffuse dot spots. On the other hand, and in accordance with the migration test of Fig. 1A , the surface expression of the chemokine receptor CCR5, known to be expressed by immature DCs, was not affected by leptin treatment, whereas, as expected, LPS treatment caused a significant (P<0.05) down-regulation of CCR5 expression (Fig. 1B ).

Leptin increases actin polymerization in immature DCs
We then investigated whether the increased migratory capacities observed in leptin-treated DCs was associated with actin microfilament remodeling. To this aim, we evaluated the effects of leptin on the G-actin (monomer) and F-actin (polymer) equilibrium by flow cytometry analysis. We found a higher amount of F-actin and total actin in leptin-treated DCs compared with control DCs after 4 h of treatment (Fig. 2 ). Statistical analyses performed using K/S test (Fig. 2 , right panels) clearly demonstrated that the increase was significant both for F-actin (P<0.001; D=0.51) and total actin (P<0.001; D=0.73). Conversely, leptin did not affect the amount of G-actin. The G/F actin ratio was significantly (P<0.01) decreased (0.95±0.5 in control DCs vs. 0.56±0.2 in 4 h leptin-treated DCs). In LPS-treated DCs, the amounts of G-, F-, and total actin were significantly increased compared with control DCs both after 2 and 4 h of treatment (P<0.01).


Figure 2
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  		Figure 2. Leptin increases actin polymerization in immature DCs. Flow cytometry analysis of actin polymerization state was performed in DCs left untreated or treated with leptin or LPS for 2 or 4 h. Representative histograms from 1 of 3 donors giving similar results are shown. Values are reported as means ± SE and represent the mean fluorescent intensity (MFI) obtained in 3 independent experiments. Right panels present the statistical analysis by K/S test from a representative experiment.

Leptin induces cofilin phosphorylation/dephosphorylation in immature DCs
Bcause cofilin represents an essential factor for enhancement of actin filament turnover, we analyzed the effect of leptin in the activation of this actin-severing protein. To this aim, we used a polyclonal Ab recognizing Ser-3 phosphorylated (P-) cofilin, corresponding to the inactive form of the protein. A transient rise in phosphorylation is involved in the initiation of actin cytoskeleton reorganization (38) , and the subsequent dephosphorylation results in turnover of F-actin. As shown in Fig. 3 A, leptin treatment of immature DCs induced a transient phosphorylation of cofilin, which reached a maximum after 30 min, followed by a dephosphorylation to basal levels after 2 h. LPS-treated DCs were also analyzed for P-cofilin levels and, as expected, the phosphorylation peak was reached earlier, after 5 min treatment, and was almost undetectable after 2 h. The levels of total cofilin remained unchanged after leptin and LPS treatments. These findings were supported by immunofluorescence analysis showing that P-cofilin was detectable as scattered small spots in 30 min leptin-treated DCs, whereas after 4 h, P-cofilin levels decreased and were similar to control cells (Fig. 3B ). Simultaneously, with the dephosphorylation of cofilin, leptin induced, after 4 h of treatment, an increase in F-actin amount and its rearrangement, (i.e., polarization) (Fig. 3B ).


Figure 3
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  		Figure 3. Leptin induces cofilin phosphorylation/dephosphorylation. A) DCs left untreated (CTR) or treated with leptin or LPS for the indicated times (5 min to 4 h), were lysed and analyzed by Western blot. P-cofilin and total cofilin levels are shown. Representative blots from 1 of 3 donors giving similar results are shown. The histogram expresses the mean ± SE fold increase over the basal level of P-cofilin in control DCs, normalized to the total cofilin levels, from 3 different experiments. *P<0.05 vs. control. B) Immunofluorescence microscopy showed an increase of P-cofilin levels after 30 min of leptin treatment, whereas F-actin was increased and polarized after 4 h treatment. Original image x1800.

Leptin promotes Rac1 activation
It is well known that the members of the small Rho GTPase family, Rho, Cdc42, and Rac1, are central regulators of cell migration via actin cytoskeleton rearrangement (43) . Among them, Rac1 is a surface ruffling-associated molecule (44) , activated at the leading edge of motile cells to regulate actin polymerization and lamellipodia formation as a driving force of cell movement. Therefore, we evaluated the capacity of leptin after 30 min, 1, 2, or 24 h to induce activation of Rac1 by measuring the levels of the active GDP-bound state by pull-down assay. As shown in Fig. 4 A, leptin treatment of immature DCs induced a significant increase of GTP-Rac1 levels after only 2 h of treatment (1.7-fold increase compared with control DCs) reaching a 3.8-fold increase after 24 h. As expected, LPS treatment induced higher levels of activation (1.8-, 1.9-, and 5.9-fold increase after 1, 2, and 24 h of treatment, respectively). Moreover, although the levels of total Rac1 were unchanged after leptin and LPS treatment, the immunofluorescence analysis revealed a different distribution of total Rac1 after 24 h of leptin and LPS treatment of DCs (Fig. 4B ). In particular, leptin promoted Rac1 translocation at the pole of the cell, (i.e., the uropod region), whereas in LPS-treated DCs, Rac1 was concentrated in the cell surface ruffling structures.


Figure 4
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  		Figure 4. Leptin promotes Rac1 activation. A) DCs left untreated (CTR) or treated with leptin or LPS for 30 min, 1, 2, or 24 h, were lysed and affinity-precipitated with glutathione S-transferase (GST) -PAK. Bound proteins were analyzed by Western blot using anti-Rac1 polyclonal Ab. Positive (CTR+) and negative (CTR–) controls were obtained incubating the lysate of untreated cells with GTP{gamma} and GDP, respectively, before GST-PAK precipitation. GTP-Racl and total Rac1 levels are shown. Representative blots from 1 of 3 donors giving similar results are shown. The histogram expresses the mean ± SE fold increase over the basal level of active Rac1 in control DCs, normalized to the total Rac1 levels, from 3 different experiments. *P < 0.05 vs. control. B) IVM analysis of total Rac1 distribution in untreated (CTR) and 24 h leptin- or LPS-treated DCs. Rac1 distribution in untreated DCs was characterized by diffused dot spots, whereas leptin promoted Rac1 translocation at the uropod region. In LPS-treated DCs, Rac1 was concentrated in the ruffles (arrows). Representative results from 1 of 3 donors giving similar results are shown. Original image x2600.

Leptin induces Vav phosphorylation
We next examined whether leptin-induced Rac1 activation in immature DCs is mediated by Vav, a guanine nucleotide exchange factor for Rho family proteins. Since Vav activity is strictly controlled by its tyrosine phosphorylation, we monitored levels of P-Vav from lysates of untreated and leptin- or LPS-treated immature DCs. We found that although total levels of Vav were unchanged, the levels of P-Vav were rapidly and transiently up-regulated by leptin treatment (Fig. 5 ), reaching a peak level after 30 min treatment. LPS treatment induced a comparable phosphorylation rate but with an anticipated time course, with a peak reached after 5 min treatment.


Figure 5
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  		Figure 5. Leptin induces Vav phosphorylation. A) DCs left untreated (CTR) or treated with leptin or LPS for the indicated times (5 min to 1 h) were lysed and analyzed by Western blot using anti-P-Vav polyclonal Ab. P-Vav and total Vav levels are shown. Representative blots from 1 of 3 donors giving similar results are shown. The histogram represents the mean ± SE fold increase over the basal level of P-Vav in control DCs, normalized to the total Vav levels, from 3 different experiments. *P < 0.05 vs. control. B) DCs pretreated for 30 min with 10 µM PP1 and left untreated (CTR) or treated with leptin for 30 min or 24 h were lysed, and P-Vav or active Rac1 levels were determined as described above. Representative blots from 1 of 3 donors giving similar results are shown.

Because tyrosine phosphorylation of Vav is required for its GEF activity, we were interested in establishing a direct correlation between the leptin-induced Rac1 activation and Vav phosphorylation. To this aim, we used the inhibitor of Src kinase PP1 (42) to block the main pathway leading to Vav phosphorylation. As shown in Fig. 5B , PP1 treatment abolished the leptin-induced up-regulation of Vav phosphorylation and concomitantly abrogated the leptin-induced Rac1 activation.

In both cases, activity values were similar to those of untreated DCs.

Leptin up-regulates IL-12p70 production in response to CD40 stimulation
A relevant limit in the preparation of DCs for antitumor therapy is related to DC exhaustion. It is known that strong maturation stimuli, such as LPS, can induce high but transient IL-12p70 production (45) . These mature DCs are, however, refractory to further stimulation and incapable of further IL-12p70 production during antigen presentation. Different protocols for the in vitro DC maturation have been used to avoid this limit, and cytokine cocktails, including tumor necrosis factor (TNF) -{alpha}, have correlated with favorable outcome (46) . It is known that stimulation of immature DCs with CD40L-transfected J558 cells (J558-CD40L), which mimic in vitro the physiological DC/T cell interaction, induces IL-12p70 production (47 , 48) . We therefore set up cocultures of DCs and J558-CD40L to test the effect of leptin on IL-12p70 production. First, we analyzed the phenotype of untreated or 24 h leptin-treated DCs. The results were compared with LPS- or TNF-{alpha} + IL-1β-treated DCs. As shown in Fig. 6 A, leptin did not modulate DC phenotype, as we previously reported (24) , whereas LPS as well as TNF-{alpha} + IL-1β triggered phenotypic maturation, as assessed by up-regulation of surface expression of HLA-ABC, HLA-DR, CD40, CD80, CD83, and CD86 molecules. It should be noted that in our system there is no leptin-induced up-regulation of CD40 surface expression. Conversely, Lam et al. (49) , in murine bone marrow-derived DCs, showed a leptin-induced up-regulation of CD40 surface expression. Next, we analyzed the phenotype of untreated or 24 h leptin-, LPS-, and TNF-{alpha} + IL-1β- treated DCs, cocultured for the following 24 h with J558-CD40L. As expected, CD40 stimulation induced phenotypic maturation in untreated DCs, which is similar in leptin-treated DCs (Fig. 6B ). The levels of expression of the principal surface markers that characterize mature DCs were comparable between leptin- and LPS- or TNF-{alpha} + IL-1β-treated DCs after 24 h coculture with J558-CD40L. Therefore, leptin did not interfere with the maturation process that occurs on CD40 stimulation and induced a higher expression of HLA-ABC and CD86 molecules compared with control. The supernatants of these same cultures were then analyzed for IL-12p70 production. As shown in Fig. 6C (C1), 24 h of leptin treatment significantly up-regulates IL-12p70 production, and LPS induces the maximum production. After 24 h of treatment with TNF-{alpha} + IL-1β, IL-12p70 production was not detectable but increased after 48 h (data not shown).


Figure 6
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  		Figure 6. Leptin does not interfere in phenotypic maturation of DCs and up-regulates IL-12p70 production in response to CD40 stimulation. A) DCs were left untreated (CTR) or treated with leptin, TNF-{alpha} + IL-1β, or LPS for 24 h. B) The treatment was followed by 24 h of coculture with J558-CD40L. DCs were then analyzed for the expression of the indicated markers by staining with PE- or FITC-conjugated mAbs. Isotype controls for direct stains exhibited mean fluorescence <5. Results are presented as means ± SE and report the mean fluorescent intensity (MFI) from 4 independent experiments. *P < 0.05 vs. control. C) Analysis of IL-12p70 supernatant concentration in the same cultures analyzed in A (C1) or B (C2) was determined by ELISA. In some experiments, DCs left untreated (CTR) or treated with leptin, TNF-{alpha} + IL-1β, or LPS for 24 h were rinsed and recultured for additional 24 h (C3). Results are presented as means ± SE and report concentration (pg/ml) from 4 independent experiments. *P < 0.05 vs. control.

In the second set of samples (Fig. 6C , C2), DCs left untreated or treated for 24 h with leptin, LPS, or TNF-{alpha} + IL-1β were washed and afterward cocultured with J558-CD40L for the following 24 h. Supernatants were then analyzed for IL-12p70 production. As expected, immature DCs produced high amounts of IL-12p70 after the CD40 stimulation, comparable to TNF-{alpha} + IL-1β-treated DCs. Leptin-treated DCs cocultured with J558-CD40L scored the highest levels of IL-12p70 production, demonstrating very efficient responsiveness to CD40 stimulation. On the other hand, LPS-treated DC cocultured with J558-CD40L produced a significantly lower amount of IL-12p70 than control DCs cocultured with J558-CD40L. Moreover, the increase in IL-12p70 production of LPS-treated DC cocultured with J558-CD40L compared with LPS treatment alone was much lower (50%) than that observed for leptin-treated DCs (99%). This difference probably results from an exhaustion of the maximum IL-12p70 production capacity of LPS-treated DC in the first 24 h. To verify that the IL-12p70 production in the DC/J558-CD40L cocultures was not due to a residual effect of the first 24 h treatments with leptin, LPS, or TNF-{alpha} + IL-1β, in some experiments the cultures were rinsed after the first 24 h and then recultured for the successive 24 h with medium alone, in absence of J558-CD40L (Fig. 6C , C3). As shown in Fig. 6C , in these conditions, only TNF-{alpha} + IL-1β-treated DCs produced detectable amounts of IL-12p70, demonstrating that the IL-12p70 produced in the DC/J558-CD40L cocultures supernatant was due to the CD40 stimulation, and leptin-treated DCs proved to be the most efficiently responsive. These results show that leptin, on CD40 stimulation, yields an efficient maturation of DCs and, more importantly, a higher IL-12p70 production compared with that obtained with conventional stimuli.

Leptin-treated immature DCs efficiently activate autologous CD8+ T cells
Tumor vaccination aims to induce patient immune responses against autologous tumor, and the activation of tumor-specific CTL is essential for vaccine efficacy. Therefore, we evaluated the capacity of leptin-treated DCs to activate autologous CD8+ T cells. To this aim, we set up an autologous MLR between CD8+ T cells and irradiated autologous DCs left untreated or treated with leptin or LPS for 24 h. As shown in Fig. 7 A, the ability of leptin-treated DCs to induce CD8+ T cell proliferation was comparable to untreated DCs, whereas treatment with LPS resulted in the maximum response. We next analyzed IFN-{gamma} and perforin production of CD8+ T cells in cocultures with untreated, leptin or LPS-treated autologous DCs. As shown in Fig. 7B , leptin-treated DCs significantly increased the IFN-{gamma} production by CD8+ T cells compared with control cells, both at 24 and 48 h. The maximum production was given by LPS-treated DCs at both time points. Similarly, CD8+ T cells showed a significant up-regulation of intracellular perforin production after stimulation with leptin-treated DCs (Fig. 7C ) compared with control DCs. At both time points, the levels were comparable to those obtained with LPS-treated DCs. Notably, the increase in IFN-{gamma} and perforin production by leptin-treated DCs is not due to an increase in CD8+ T number in the cocultures (Fig. 7A ) but to a major activation status.


Figure 7
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  		Figure 7. Leptin-treated DCs efficiently activate CD8+ autologous T cells. A) Immature DCs were left untreated (CTR) or treated with leptin or LPS for 24 h. An autologous MLR with irradiated DCs cultured at different cell numbers with 1 x 105 autologous CD8+ T cells was then set up. [H3]thymidine incorporation was measured after 5 days. Results are presented as means ± SE and expressed as counts per minutes (cpm); results from 4 different experiments are shown. *P < 0.05. B, C) DC:CD8+ T cells cocultured at a 1:1 ratio were set up in 24-well plates. After 24 and 48 h, supernatants were collected for IFN-{gamma} concentration evaluation by ELISA (B), and cells were collected for the evaluation of intracellular perforins by flow cytometry on CD8+ gated cells (C). Histograms represent 1 representative experiment of 4 and indicate the percentage of perforin-positive CD8+ cells. Marker was placed based on isotype-matched mAb. Bar histograms present means ± SE from 4 different experiments. *P < 0.05.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, we outline a novel property for leptin, demonstrating its ability to improve the migratory performance of immature DCs by activating specific pathways driving cytoskeleton rearrangement. We specifically found that leptin up-regulates CCR7 surface expression and favors the turnover of actin microfilaments interfering with cofilin-severing function. Moreover, we found that leptin, triggering Vav phosphorylation, promotes Rac1 activation. At a functional level, leptin up-regulates IL-12p70 production on CD40 stimulation in immature DCs and increases their capacity to activate autologous CD8+ T cells.

In several clinical settings, it could be appealing to take advantage of the knowledge gained on the migratory properties of DCs to design strategies for the delivery of DCs to LNs to effect immunoregulation and to exploit them as cellular adjuvants in cancer vaccination (50 , 51) . Several clinical trials have been carried out to determine the efficacy of therapeutic vaccines that use ex vivo antigen-loaded and matured DCs. The final outcome of the clinical response is significantly affected by the protocols used to generate DCs, in particular in terms of the stimuli used to induce DC maturation and, therefore, of the final DC maturation state (52) . However, only a small proportion of injected DCs subsequently migrate to the LNs, limiting the efficacy of the vaccine (53 , 54) . Moreover, despite some early encouraging results, a limited rate of objective tumor regressions has been observed in clinical studies (55) , and the need for improvement is widely acknowledged (50) .

It is well known that DC homing to the LNs is controlled by the chemokine receptor CCR7 and its ligands CCL21 and CCL19 (56) . Here, we found that leptin up-regulates CCR7 surface expression in immature DCs. This result closely fits with our previous data demonstrating that leptin activates in DCs the transcriptional factor NF-{kappa}B (24) , known to be involved in the transcriptional regulation of CCR7 (57) .

Moreover, we showed that the up-regulation of CCR7 coincides with an improvement of DC migratory performances in the presence of CCL19 chemotactic stimulus. Besides, leptin also enhances the migratory capacities of DCs in the absence of a chemotactic stimulus, which parallels the previously demonstrated ability of leptin to induce morphological changes in DCs (24) .

In addition to chemotaxis, CCR7 has been shown to control the cytoarchitecture, the survival, the migratory speed, and the maturation of DCs, regulating independent signaling modules (57 , 58) . Moreover, it has been demonstrated that the up-regulation of CCR7 expression alone is insufficient to drive DC migration (59) , which is achieved in the presence of additional stimuli, facilitating the coupling of the receptor to its signal transduction modules. Here, we demonstrate that the treatment of immature DCs with leptin is efficient in inducing the phosphorylation/dephosphorylation of cofilin in the absence of CCR7 engagement. Cofilin is an actin-binding protein that can bind both F- and G-actin and is involved in increasing the rate of actin depolymerization/polymerization and the severing of actin filaments (60 , 61) . It is inactivated by phosphorylation, and the subsequent dephosphorylation can result in an increase of filamentous actin and the appearance of veils. Cofilin disassembles F-actin from the rear of the actin network to recycle monomers to the leading edge for further rounds of polymerization. It has been shown that on LPS-induced DC maturation, cofilin is rapidly phosphorylated and then gradually dephosphorylated; simultaneously, the amount of F-actin increases (62) . We found that leptin induces phosphorylation/dephosphorylation of cofilin with a delayed time course compared with LPS, which is indeed associated with an increased amount of F-actin in the cell. In fact, accordingly, we found that leptin was able to shift the balance between polymeric F-actin and monomeric G-actin in favor of the former. This finding correlates with previous studies showing an increased actin polymerization in migrating DCs (63) and with the results obtained by O’Malley et al. (64) in hippocampal neurons, showing that leptin promotes the rapid rearrangement and destabilization of actin filaments. Moreover, Saxena et al. (65) demonstrated that leptin increases the migration capability of hepatocellular carcinoma cells, and Attoub et al. (66) showed the same effect on kidney and colonic epithelial cells on various stages of neoplastic progression.

The members of the Rho family of small guanosine triphosphatases (GTPases) are key regulators of the actin cytoskeleton in several cellular functions, including cell migration (67 , 68) . Among Rho family GTPases, Rac1 is activated at the leading edge of motile cells to regulate actin polymerization (69 , 70) . We found that, in immature DCs, leptin activates Rac-1 presumably through the induction of Vav phosphorylation/activation, since the pretreatment of DCs with PP1, which inhibits the Src kinases, the major Vav-activating pathway, inhibits the leptin-induced activation of Rac1. Overall, these findings strongly suggest that in immature DCs, leptin, on binding its receptor, improves chemotactic responsiveness by up-regulating CCR7 expression and triggers the activation of intracellular signaling pathways driving microfilament cytoskeleton rearrangement. Taking into account the previously hypothesized activity of leptin on Rho family GTPases in relation to the actin cytoskeleton (71) , we cannot rule out the possibility that such an activity could significantly contribute to functional results reported in our work. The different time course of effects observed after stimulation of DCs with leptin vs. LPS, in terms of cofilin and Vav activation, and in terms of different distribution of Rac1, reflects the different strengths of the 2 stimuli, with leptin being a milder stimulus, not able to induce full DC maturation (24) . This differentiation is useful to sustain our proposal of leptin as a candidate adjuvant for cancer vaccination protocols employing ex vivo-generated autologous DCs. In these protocols, the final outcome of the clinical response is significantly affected by the protocols used to generate DCs, in particular in terms of the stimuli used to induce DC maturation. Strong stimuli, such as LPS, are not useful because of the problem of DC exhaustion, as discussed in the Results section.

Moreover, a DC preparation that has both high migratory capacity and potent antitumor CTL-inducing capacities is essential for the development of new DC-based vaccines. However, production of IL-12 and high migratory capacities have been considered to be mutually exclusive (72) . Here, we found that leptin, on CD40 stimulation, significantly up-regulates IL-12p70 production compared with conventional stimuli such as TNF-{alpha} + IL-1β. This finding is relevant considering that a strong production of IL-12 is gainful once DCs have reached LNs, where interactions with T cells occur. More important, we found that leptin enhances immature DC capacity to activate autologous CD8+ T cells in terms of perforin and IFN-{gamma} production. The critical role of CD8+ T cells in the prevention and eradication of tumors has been highlighted by many studies (73) . A functional hallmark of CD8+ T cells is the production of IFN-{gamma}, an essential cytokine involved in the innate and adaptive immune responses against tumor development (74) that acts in various ways on host and tumor cells to favor tumor regression. Moreover, IFN-{gamma} contributes directly to perforin-mediated apoptosis by increasing target cell sensitivity. Gao et al. (75) showed that functional defects in CD8+ T cells, in terms of IFN-{gamma} and perforin production, lead to compromised CTL functions and cause high susceptibility of mice to both melanoma or lymphoma tumorigenesis. Therefore, IFN-{gamma} and perforin production are commonly used as activation and cytotoxic indicators, respectively, in the functional analysis of CD8+ T cells.

To validate the hypothesis here presented and supported by these in vitro results, we are planning in vivo experiments in mouse systems employing ex vivo-generated DCs treated with leptin and reinfused in the animal host. The capacity of leptin-pulsed DCs to reach LNs and to stimulate antitumor-specific CD8+ T cells will then be evaluated.

Overall, our data suggest that leptin could represent an optimal candidate adjuvant for cancer vaccination, which is additionally sustained by our previous results demonstrating that leptin licenses immature DCs to polarize immune response toward Th1. Therefore, leptin could be employed in all vaccination protocols requiring an efficient boost of Th1-type responses.


   ACKNOWLEDGMENTS
 
This work was supported by grants from the Italian Ministry of Health to M.V. and W.M.

Received for publication September 7, 2007. Accepted for publication December 23, 2007.


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