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Published as doi: 10.1096/fj.07-8652com.
 (The FASEB Journal. 2008;22:1032-1042.)
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(The FASEB Journal. 2008;22:1032-1042.)
© 2008 FASEB

Control of dendritic cell maturation and function by triiodothyronine

Ivan Mascanfroni*,1, María del Mar Montesinos*,1, Sebastián Susperreguy*, Laura Cervi*, Juan M. Ilarregui{dagger}, Vanesa D. Ramseyer*, Ana M. Masini-Repiso*, Héctor M. Targovnik{ddagger}, Gabriel A. Rabinovich{dagger} and Claudia G. Pellizas*,2

* Centro de Investigaciones en Bioquímica Clínica e Inmunología (CIBICI-CONICET), Departamento de Bioquímica Clínica, Facultad de Ciencias Químicas, Universidad Nacional de Córdoba, Córdoba, Argentina;

{dagger} Laboratorio de Inmunopatología, Instituto de Biología y Medicina Experimental (IBYME) CONICET y Facultad de Ciencias Exactas y Naturales, and

{ddagger} Cátedra de Genética y Biología Molecular, Facultad de Farmacia y Bioquímica, Universidad de Buenos Aires, Buenos Aires, Argentina

2Correspondence: CIBICI-CONICET, Departamento de Bioquímica Clínica, Facultad de Ciencias Químicas, Universidad Nacional de Córdoba, Haya de la Torre esq. Medina Allende, Ciudad Universitaria, 5000 Córdoba, Argentina. E-mail: claudia{at}mail.fcq.unc.edu.ar


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Accumulating evidence indicates a functional crosstalk between immune and endocrine mechanisms in the modulation of innate and adaptive immunity. However, the impact of thyroid hormones (THs) in the initiation of adaptive immune responses has not yet been examined. Here we investigated the presence of thyroid hormone receptors (TRs) and the impact of THs in the physiology of mouse dendritic cells (DCs), specialized antigen-presenting cells with the unique capacity to fully activate naive T cells and orchestrate adaptive immunity. Both immature and lipopolysaccharide-matured bone marrow-derived DCs expressed TRs at mRNA and protein levels, showing a preferential cytoplasmic localization. Remarkably, physiological levels of triiodothyronine (T3) stimulated the expression of DC maturation markers (major histocompatability complex II, CD80, CD86, and CD40), markedly increased the secretion of interleukin-12, and stimulated the ability of DCs to induce naive T cell proliferation and IFN-{gamma} production in allogeneic T cell cultures. Analysis of the mechanisms involved in these effects revealed the ability of T3 to influence the cytoplasmic-nuclear shuttling of nuclear factor-{kappa}B on primed DCs. Our study provides the first evidence for the presence of TRs on bone marrow-derived DCs and the ability of THs to regulate DC maturation and function. These results have profound implications in immunopathology, including cancer and autoimmune manifestations of the thyroid gland at the crossroads of the immune and endocrine systems.—Mascanfroni, I., Montesinos, M., Susperreguy, S., Cervi, L., Ilarregui, J. M., Ramseyer, V. D., Masini-Repiso, A. M., Targovnik, H. M., Rabinovich, G. A., Pellizas, C. G. Control of dendritic cell maturation and function by triiodothyronine.


Key Words: adaptive immunity • thyroid hormones • antigen presentation


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
THE ENDOCRINE AND IMMUNE SYSTEMS are interrelated via a bidirectional network in which hormones affect immune function, and, in turn, immune responses are reflected in neuroendocrine changes. This bidirectional communication is possible because both systems share common ligands (hormones and cytokines) and their specific receptors (1) .

Thyroid hormones (THs) play critical roles in differentiation, growth, and metabolism (2) . Despite the role assigned for THs in maintaining immune system homeostasis (3 4 5) , the study of TH effects on cells of the immune system received relatively less attention than the study of effects exerted by other hormones of the hypothalamus-pituitary-adrenal (HPA) axis (6 , 7) . Interactions between hormones from the pituitary and thyroid glands and the immune system were revealed mainly by the presence of specific receptors for thyrotropic and thyroid hormones on lymphocytes or by the frequent immune alterations associated with physiological or pathological fluctuations of THs (1 , 8) . Evidence arising from analysis of lymphocyte development and function in mice with genetic defects in the expression of THs or thyroid hormone receptors (TRs) suggested that THs may play an essential role in maintaining immune system homeostasis in response to environmental changes or stress-mediated immunosuppression (4 , 6 , 8) . Whereas studies on the effects of THs in the control of immune responses were mainly conducted on effector B and T lymphocytes, the role of THs in the initiation of adaptive immune responses still remains uncertain. Dendritic cells (DCs) are highly specialized antigen-presenting cells (APCs) that recognize, process, and present antigens to naive T cells for the induction of antigen-specific immune responses (9) . Because DCs are pleiotropic modulators of T cell activity capable of orchestrating adaptive immunity and are endowed with exquisite plasticity, manipulation of their function to favor the induction of DCs with immunogenic or tolerogenic properties could be exploited to positively or negatively regulate adaptive immune responses (10) . Several factors may influence the decision of DCs to become immunogenic or tolerogenic, including the maturational and activation status, and the cytokine milieu (including growth factors, neuropeptides, and hormones) at sites of T cell activation and inflammation (11 , 12) .

After in vitro or in vivo exposure to lipopolysaccharides (LPSs) or other bacterial products, DCs undergo activation and maturation through different signaling pathways including mitogen-activated protein kinase kinase 1/extracellular signal-regulated kinase, which favors DC survival, and the NF-{kappa}B pathway, which allows for DC maturation (13) . Signaling through NF-{kappa}B also determines the increased expression of major histocompatability complex (MHC) II and costimulatory molecules, release of proinflammatory cytokines and chemokines, and DC migration and recruitment. This coordinated process leads to sustained T cell stimulatory capacity and interleukin (IL)-12 release, which result in the induction of protective immunity (13) .

The classic genomic actions of THs are mediated by nuclear TRs that act as hormone-inducible transcription factors. Several TR{alpha} and TRβ isoforms are encoded by the TRA and TRB genes, respectively. The TR{alpha}1, TR{alpha}2, TRβ1, and TRβ3 isoforms are widely expressed, whereas TRβ2 is predominantly restricted to the hypothalamus-pituitary (HP) axis (14 , 15) . However, nongenomic actions of THs have also been described at the level of the plasma membrane, cytoskeleton, cytoplasm, and distinct organelles of mammalian cells (8 , 16) .

Despite major advances in understanding of the interplay between distinct hormones and the immune cell network (11 , 12 , 17 18 19) , the role of THs in the initiation of adaptive immunity still remains uncertain (20) . In the present study we provide the first evidence of the expression of TRs on bone marrow-derived DCs and their striking localization in the cytoplasmic compartment of immature and mature DCs. Furthermore, we demonstrate the effects of THs on DC maturation and IL-12 secretion and the capacity of these cells to stimulate T cell responses.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Mice
Female C57BL/6 (B6; H-2b) and BALB/c (H-2d) mice were obtained from Ezeiza Atomic Center (Buenos Aires, Argentina). Mice were maintained under specific pathogen-free conditions and used at 6–10 wk of age. Animal protocols complied with the Guide for the Care and Use of Laboratory Animals published by the U.S. National Institutes of Health and local institutional animal care committee guidelines.

DC preparation and culture
DCs were obtained as described by Inaba et al. (21) . Briefly, bone marrow progenitors were collected from the femurs of 4- to 6-wk-old female C57BL/6 mice and cultured in RPMI 1640 10% fetal calf serum (FCS) depleted of THs by treatment with resin AG-1-X8 (Bio-Rad Laboratories, Hercules, CA, USA) in the presence of granulocyte-macrophage colony-stimulating factor from supernatant of the J558 cell line and fed every 2 days. At day 10 of cell culture, >85% of the harvested cells expressed MHC class II, CD40, CD80, and CD11c, but not Gr-1. Immature DCs (iDCs) were stimulated with LPS (100 ng/ml; Escherichia coli strain 0111:B4; Sigma-Aldrich, St. Louis, MO, USA) for 18 h to obtain mature DCs (mDCs). Alternatively, iDCs were incubated with 3,3',5-triiodo-L-thyronine (T3) (0.05–500 nM) or T3 (5 nM) plus LPS (100 ng/ml) for 18 h. Afterward, DCs were collected and washed. T3 was purchased from Sigma-Aldrich and prepared according to the manufacturer’s recommended protocol. To rule out endotoxin contamination of the T3 preparation, the same set of experiments were performed in the presence of polymyxin B (10 µg/ml; Sigma-Aldrich). In addition, we checked the endotoxin content of the T3 preparation after reconstitution, which raised levels <0.03 endotoxin unit/ml (limit of detection), by using the Limulus amebocyte lysate assay (Sigma-Aldrich).

Flow cytometric analysis of DC phenotype
DCs were washed twice with PBS supplemented with 2% FCS and resuspended in 10% FCS-PBS. Cells were then incubated with the following fluorochrome-conjugated monoclonal antibodies (mAbs) for 30 min at 4°C: fluorescein isothiocyanate (FITC)-anti-CD11c, phycoerythrin (PE)-anti-IA/IE (MHC II), PE-anti-CD40, PE-anti-CD80, and PE-anti-CD86 (all from BD PharMingen, San Diego, CA, USA). Cells were then processed and analyzed in an Ortho Cytoron Absolute flow cytometer (Ortho Diagnostic Systems, Raritan, NJ, USA) using FlowJo software (Tree Star, Ashland, OR, USA).

Transfection of COS-7 cells with TRβ1 expression vector
COS-7 cells do not express TRs and were used as a negative control for Western blot analysis. As a positive control, COS-7 cells were transiently transfected with TRβ1 expression vector as described previously (22) . Briefly, cells were maintained in Dulbecco’s modified Eagle’s medium (Invitrogen/Life Technologies Corporation, Carlsbad, CA, USA) supplemented with 10% FCS, antibiotics, and glutamine at 37°C in a humidified atmosphere (5% CO2). Cells (3x105) were seeded in a 60-mm dish 24 h before transfection. Cells were transfected with 5 µg of pCDM8-TRβ1 expression vector by calcium phosphate coprecipitation as described previously (23) . After 2 h of incubation with the precipitate, cells were shocked with 15% glycerol in PBS. Twenty-four hours after transfection, cells were harvested and lysed as described below to obtain cell lysates for Western blot analysis.

Reverse transcription (RT) and polymerase chain reaction (PCR)
Cells were homogenized with TRIzol, and RNA extraction was performed according to the manufacturer’s recommended protocol based on the Chomczynski and Sacchi method (24) . mRNA was reverse transcribed and amplified by PCR essentially as described (22) with minor modifications. Briefly, 1 µg of total RNA was incubated with 0.1 µM degenerated oligo dT12VG primers at 65°C. After 3 min on ice, the following reagents were added: 20 U of RNase inhibitor (RNaseOUT; Promega, Madison, WI, USA), 4 µl of 5x RT buffer [250 mM Tris-HCl, pH 8.3, 75 mM KCl, 3 mM MgCl2 and 10 mM dithiothreitol (DTT)], 0.5 mM concentrations of each dNTP, and 200 U of Moloney murine leukemia virus reverse transcriptase (Promega). After 1 h at 37°C, remnant reverse transcriptase was inactivated at 95°C for 5 min. Expressions of TR{alpha}1 and TRβ1 mRNAs were normalized using the housekeeping glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA. Primers (Sigma-Aldrich, Buenos Aires, Argentina) were designed to distinguish cDNA and genomic DNA/pseudogenes (25) to amplify a 505-bp band for GAPDH, a 371-bp band for TR{alpha}1, and a 590-bp band for TRβ1 mRNAs as follows: 5'-CAGTGCCAGGAATGTCGCTTTAAG-3' (TRβ1 forward), 5'-ACTCTGGTAATTGCTGGTGTGATGAT-3' (TRβ1 reverse), 5'-TTCAGCGAGTTTACCAAGATCATCAC-3' (TR{alpha}1 forward), 5'-TTAGACTTCCTGATCCTCAAAGACCTC-3' (TR{alpha}1 reverse), 5'-GAAGGTGAAGGTCGGAGTCAACG-3' (GAPDH forward), and 5'-GATACCAAGTTGTCATGGATGACCTT-3' (GAPDH reverse).

PCR was carried out in a 20 µl final volume using 1.5 mM MgCl2, 4 µl of 5x PCR buffer, 1 U of Taq polymerase (Promega), 0.25 mM concentrations of each dNTP (Promega), and 2 µl of RT product. A negative control (sterile water instead of RT product) was included in each PCR run. The PCR amplification was performed on a ICycler PCR System (Bio-Rad). The thermal profile was 94°C for 5 min (34 cycles for TRs and 26 cycles for GAPDH); 94°C for 1 min, 56°C for 1 min, 72°C for 2 min, and 72°C for 10 min. The mass of total RNA for RT, the number of cycles for PCR, and MgCl2, primer, and dNTP concentrations were selected experimentally (data not shown). RT-PCR products were resolved by electrophoresis in 2% agarose gels followed by ethidium bromide staining.

Preparation of total, nuclear, and cytoplasmic extracts
To obtain total cell lysates of COS-7 and DCs, 3 x 106 cells were resuspended in 50–200 µl of RIPA buffer, disrupted by passages through a 25-G needle, and incubated on ice for 30 min, followed by removal of DNA and cell debris by centrifugation at 10,000 g for 20 min at 4°C. Nuclear and cytoplasmic DC extracts were obtained by subcellular fractionation essentially as described by Schreiber et al. (26) . The supernatant containing cytoplasm was collected and frozen at –70°C or used immediately. The nuclear pellet was resuspended in 50 µl of ice-cold buffer (20 mM HEPES, pH 7.9, 0.4 M NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, and 1 mM PMSF), and the tube was vigorously rocked at 4°C for 15 min on a shaking platform. The nuclear extract was centrifuged for 5 min in a Microfuge at 4°C, and the supernatant was frozen in aliquots at –70°C or used immediately. Rat liver nuclear extracts used as positive controls were obtained as described previously (22) .

Western blot analysis
Total cell lysates and nuclear and cytoplasm extracts of DCs (40 µg), COS-7 cells (40 µg; negative control), COS-7 cells overexpressing TRβ1, and rat liver nuclear extract (40 µg) (positive controls) were used for immunodetection of TR{alpha}1 (47 kDa) and TRβ1 (55 kDa). The rabbit anti-TR polyclonal antibody (Ab) (FL-408, sc-772; Santa Cruz Biotechnology, Santa Cruz, CA, USA), which cross-reacts with chicken, mouse, rat, and human TR{alpha}1 and TRβ1, was used at a 1:2000 dilution. NF-{kappa}B (p65) expression was evaluated with an anti-p65 Ab (sc-8008; Santa Cruz Biotechnology). Anti-{alpha}-tubulin (Clone B-5-1-1) and antihistone deacetylase 1 (HDAC1) Abs (Sigma-Aldrich) were used to control the purity of the subcellular fractions. Equal protein loading was checked using an anti-β-actin mAb (sc-1616; Santa Cruz Biotechnology). Western blot analysis was performed as described (22) and revealed using the enhanced chemiluminescence protocol (NEL-100; DuPont NEN Research Products, Wilmington, DE, USA).

Immunofluorescence microscopy
Bone marrow-derived DCs were generated as described above and cultured on coverslips for 3 days. After treatments, cells were fixed in 4% paraformaldehyde, permeabilized in 0.25% Triton X-100 in PBS, blocked for 1 h in PBS (pH 7.4) plus 3% BSA fraction V (Fisher), incubated with a primary Ab (mouse anti-TRβ1 sc-738; Santa Cruz Biotechnology) at a dilution of 1:100 for 1 h, washed, and further incubated with an Alexa conjugated goat anti-mouse secondary Ab (Molecular Probes, Inc., Eugene, OR, USA) for 1 h at a dilution of 1:1000. Nuclei were stained with 4,6-diamidino-2-phenylindole for 5 min, and samples were washed in PBS and mounted on glass slides using Fluoromount-G (Southern Biotechnology Associates, Birmingham, AL, USA) for examination using a Leica DM IRBE inverted microscope (Hamamatsu Corporation, Bridgewater, NJ, USA). Images were captured using Openlab 3.1 software (Improvision, Lexington, MA, USA) at a magnification of x1000.

Allogeneic T cell cultures
Allogeneic T cell cultures were performed to assess the ability of DCs to stimulate allogenic splenocytes in vitro as described (27) . Briefly, allogenic splenocytes (1x105 cells/well, responder cells) were incubated for 3 days with irradiated DCs (30 Gy, stimulator cells) at a ratio of 1:10 to 1:40 DCs/splenocyte) in 96-well round-bottom plates. On day 2, 0.5 µCi (0.0185 MBq)/well of [3H]thymidine (Amersham Life Sciences, Buckinghamshire, UK) was incorporated into each well for 18 h. Proliferation was determined as counts per minute of triplicate determinations.

Cytokine determination
IL-12p70, IL-10, and IFN-{gamma} detection was performed in cell culture supernatants using standard capture ELISAs. Coating Abs included a rat anti-mouse IL-12p70 mAb (clone C15.6; PharMingen), rat anti-mouse IL-10 mAb (clone JES5–2A5; PharMingen), and rat anti-mouse IFN-{gamma} mAb (clone R4–6A2; PharMingen). Detection Abs included biotinylated rat anti-mouse IL-12p70 mAb (clone C17.8; PharMingen), biotinylated rat anti-mouse IL-10 mAb (clone SXC-1; PharMingen), and biotinylated rat anti-mouse IFN-{gamma} mAb (clone XMG1.2; PharMingen). Streptavidin-horseradish peroxidase and 3-ethylbenzthiazoline-6-sulfonic acid (Sigma-Aldrich) were used as enzyme and substrate, respectively. Intracellular cytokine was detected by flow cytometry as described previously (28) using PE-conjugated anti-IL-12, PE-conjugated anti-IL-5, and FITC-conjugated anti-IFN-{gamma} mAbs (all from BD PharMingen). Briefly, cells were exposed to brefeldin A (10 µg/ml; Sigma) for the last 6 h of cell culture. Allogeneically activated splenocytes were labeled with PE- or FITC-conjugated anti-CD4 mAbs (BD Biosciences, San Jose, CA, USA) for 30 min. Cells were then fixed with 1% paraformaldehyde, treated with fluorescence-activated cell sorter permeabilizing solution (BD Biosciences) and stained with an optimal concentration of anticytokine mAb or an appropriate isotype control mAb (all from BD Biosciences). Cells (at least 10,000 viable cells) were then analyzed in an Ortho Cytoron Absolute flow cytometer using FlowJo software.

Statistical analysis
Statistical analysis was performed using Student’s paired t test. A Wilcoxon nonparametric test for paired data was used to determine the significance of the time-response curves. Values of P < 0.05 were considered statistically significant. To adjust the significance level for multiple comparisons, a Bonferroni correction was applied using a corrected significance level of 0.017. All experiments were performed at least in triplicate.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
TRs are highly represented in iDCs and mDCs and are confined mainly to the cytosolic compartment of these cells
To investigate the role of THs in the physiology of DCs and their influence in the initiation of adaptive immune responses, we first examined the expression and subcellular localization of TRs in bone marrow-derived DCs. Both iDCs and LPS-matured DCs (mDCs) expressed TR{alpha}1 and TRβ1 mRNA (Fig. 1 A, B), although at a lesser extent than rat liver tissue, which was used as a positive control. Moreover, expression of TRs was confirmed at the protein level by Western blot analysis of iDCs and mDCs (Fig. 1C ). We could not find significant differences in TR expression between iDCs and mDCs (Fig. 1A-C ). However, both DC subsets showed higher expression of TRβ1 than TR{alpha}1 (Fig. 1C ), similar to B and T lymphocytes (29 , 30) .


Figure 1
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  		Figure 1. Expression of TRs in iDCs and mDCs. A, B) Detection of mRNA for TR{alpha}1 (A) and TRβ1 (B) by RT-PCR analysis of total RNA of iDCs and LPS-matured DCs. Total RNA from rat liver was run in parallel as a positive control. The expression of TR{alpha}1 and TRβ1 mRNAs was normalized with the housekeeping GAPDH mRNA. RT-PCR products were resolved by electrophoresis in 2% agarose gels followed by ethidium bromide staining. C) Western blot analysis of total cell lysates of iDCs, mDCs, COS-7 cells (negative control), COS-7/TR transfected cells (positive TRβ1 control), and nuclear extracts from rat liver (positive TRβ1 and TR{alpha}1 control). Samples were separated by 10% SDS-PAGE, transferred onto nitrocellulose membranes, and blotted with anti-TR Abs. Equal loading was checked using an anti-β-actin mAb. Representative of three independent experiments with similar results are shown.

To examine the subcellular compartmentalization of TRs on iDCs and mDCs, we performed Western blot analysis of subcellular DC fractions. Strikingly, the expression of cytoplasmic TRβ1 was markedly increased compared with that of nuclear TRβ1 in both iDCs and mDCs (Fig. 2 ). The purity of subcellular fractions was checked by using {alpha}-tubulin and HDAC1 as specific markers of cytoplasmic and nuclear fractions, respectively. In addition, immunofluorescence staining showed bright cytoplasmic labeling of iDCs and mDCs, whereas nuclear staining was scarce and diffuse, in broad agreement with Western blot analysis (Fig. 3 ). Of importance, no substantial differences were observed in TRβ1 staining between both DC subsets.


Figure 2
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  		Figure 2. Subcellular compartmentalization of TRs in the nuclear and cytoplasmic fractions of iDCs and mDCs. A) Western blot analysis of nuclear and cytoplasmic fractions of iDCs and mDCs for detection of TRβ1. Anti-{alpha}-tubulin and HDAC1 Abs were used to check the purity of cytoplasmic and nuclear fractions, respectively. β-Actin was used as a control of equal protein loading (lower panel). B) Densitometric analysis of immunoreactive protein bands. Results are expressed as arbitrary units (AU) calculated from the densitometric analysis of TRβ1 signal normalized to β-actin. Data are presented as mean ± SD. *P < 0.01 vs. nucleus. Blots are representative of six independent experiments.


Figure 3
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  		Figure 3. Immunofluorescence analysis of TR localization in iDCs and mDCs. TR localization was assessed by immunofluorescence staining of iDCs and mDCs using a specific anti-TR Ab as described in Materials and Methods. Both DC subsets exhibited moderate to strong cytoplasmic staining, whereas nuclear staining was scarce. Magnification x1000.

T3 induces the maturation of bone marrow-derived iDCs
To determine the physiological relevance of TR expression in DCs, we cultured iDCs in the presence of LPS or T3 and evaluated their cell surface phenotype and functionality. Unstimulated DCs exhibited an immature phenotype characterized by marked expression of CD11c, but low levels of MHC II and the costimulatory molecules CD40, CD80, and CD86 (Fig. 4 ). As expected, LPS induced DC maturation, as demonstrated by the increased levels of MHC II, CD40, CD80, and CD86 on the surface of bone marrow-derived DCs. Remarkably, exposure to T3 resulted in a similar DC maturation phenotype (Fig. 4A , B). T3-induced DC maturation was found to be dose-dependent, showing an optimal T3 stimulating concentration of 5 nM. Interestingly, this concentration is close to that found in mice sera and was, therefore, used for further DC maturation assays. As a control, exposure of iDCs to T3 in the presence of polymyxin B did not abolish T3-induced DC maturation, disregarding the possibility of endotoxin contamination in the T3 preparation (Fig. 4B ).


Figure 4
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  		Figure 4. Effects of T3 on DC maturation. iDCs were incubated with T3 (5 nM) or LPS (100 ng/ml) for 18 h. A) Cell surface phenotype was analyzed by flow cytometric analysis of T3 or LPS-matured DCs using PE-conjugated anti-MHC II (IA/IE), anti-CD40, anti-CD80, and anti-CD86 mAbs. Polymyxin B (PolB) was added to DC cultures to check possible endotoxin contamination of T3 preparations. Representative histograms of eight independent experiments are shown. B) Results are expressed as the percentage of increase of the relative mean fluorescence intensity (rMFI). Data are the mean ± SD of eight independent experiments (*P<0.001 vs. control; Wilcoxon nonparametric test).

To investigate whether the maturation phenotype induced by T3 is associated with an increased functionality of these cells, we first examined the ability of T3 to modulate cytokine secretion by DCs. iDCs exposed to T3 showed a significant increase in IL-12 secretion (Fig. 5 A) (P<0.05), whereas the production of IL-10 was not modified (Fig. 5B ). Consistently, treatment with T3 resulted in a significant increase in the frequency of IL-12-producing CD11c+ DCs (P<0.01, Fig. 5C, D ) with no changes in the frequency of IL-10-producing CD11c+ cells (data not shown). Thus, T3-conditioned bone marrow-derived DCs exhibit a highly mature phenotype comparable to that induced by LPS.


Figure 5
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  		Figure 5. Effects of T3 in the modulation of the IL-12/IL-10 cytokine balance. A, B) DCs were stimulated with LPS (100 ng/ml), T3 (5 nM), or LPS plus T3 for 48 h. IL-12 (A) and IL-10 (B) production was determined in culture supernatants by ELISA. Data are expressed as mean ± SD (pg/ml) of three independent experiments. *P < 0.05, vs. control DCs. C, D) DCs were stimulated with T3 (5 nM) or LPS (100 ng/ml) for 18 h. For intracytoplasmic cytokine staining, cells were incubated with brefeldin A for 4 h, stained for CD11c, fixed, permeabilized, and then stained with anti-IL-12p70 Ab as described in Materials and Methods. The frequency of CD11c+ IL-12+ cells was determined by flow cytometry. Values are given as the percentage of total CD11c+ IL-12-producing cells. D) Data are expressed as mean ± SD of three independent experiments. *P < 0.01, vs. control DCs.

T3 favors the generation of mDCs with increased T cell-stimulatory capacity
The mature cell surface phenotype and the increased IL-12 production induced by T3 treatment prompted us to investigate the T cell allostimulatory capacity of T3-conditioned DCs. Proliferation of BALB/c (H-2d) splenocytes was strongly enhanced in response to irradiated T3-conditioned DCs (C57BL/6) in a wide range of DC/splenocyte ratios at all T3 concentrations tested (Fig. 6 ). This effect was dose-dependent at concentrations ranging from 0.05 to 5 nM, reaching a plateau at a T3 dose of 5 nM. No differences were found between concentrations of 5 and 500 nM. Interestingly, DCs cultured in the presence of 5 and 500 nM T3 markedly enhanced the proliferation of responder splenocytes and were significantly more potent allostimulators than DCs matured in the presence of LPS (P<0.01).


Figure 6
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  		Figure 6. Impact of T3 on the allostimulatory capacity of DCs. Bone marrow-derived DCs were stimulated with T3 (0.05, 5, and 500 nM) or LPS (100 ng/ml). After 18 h, DCs were extensively washed, irradiated, and cultured with allogenic splenocytes (1x105 cells/well) for 3 days at different stimulator/responder ratios (1:10 to 1:40 DC/splenocyte). Proliferation of allogeneic splenocytes was measured by [3H]thymidine incorporation. Data are expressed as mean ± SD (cpm) representative of six independent experiments. *P < 0.01 vs. control DCs.

The augmented allostimulatory capacity of T3-matured DCs was also reflected by the greatly enhanced production of the effector cytokine IFN-{gamma} in culture supernatants of splenocytes stimulated with T3-conditioned DCs (DC/splenocyte ratio 1:15) (Fig. 7 A). Furthermore, T3 was able to potentiate the allostimulatory capacity of LPS-matured DCs, as shown by the increased IFN-{gamma} production (Fig. 7A ). However, IL-10 was undetectable in allogeneic T cell cultures stimulated with T3-conditioned DCs (Fig. 7B ). To further explore the ability of T3-conditioned DCs to direct T cell differentiation toward a T1-type profile, naive allogeneic splenocytes were cultured with T3-stimulated DCs for 72 h at a cell ratio of 1:15 (DCs/splenocytes). Activated T cells were analyzed for intracellular IFN-{gamma} and IL-5 production by flow cytometry. Remarkably, T3-stimulated DCs, but not control DCs significantly enhanced the frequency of IFN-{gamma}-producing cells in both the CD4+ and CD4– T cell compartments (Fig. 8 A) (P<0.01). As a positive control, LPS-treated DCs induced a significant increase in the proportion of IFN-{gamma}-secreting CD4+ T cells (Fig. 8A ). However, no changes were observed in the percentage of IL-5-producing CD4+ T cells after exposure to T3-conditioned DCs (Fig. 8B ). Thus, T3 may instruct the generation of a DC1 phenotype with increased T cell stimulatory potential and the ability to direct the development of a dominant T1-type response.


Figure 7
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  		Figure 7. Influence of T3-conditioned DCs on cytokine secretion by splenocytes. DCs were stimulated with LPS (100 ng/ml), T3 (5 nM), or a combination of LPS and T3. After 18 h, DCs were extensively washed, irradiated, and cultured with allogeneic splenocytes (at an optimal DC/splenocyte ratio of 1:15) for 3 days. IFN-{gamma} (A) and IL-10 (B) production were measured in culture supernatants by ELISA. Results are expressed as mean ± SD of three independent experiments. *P < 0.01 vs. control DCs.


Figure 8
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  		Figure 8. Modulation of Th1/Th2 cytokine production by T3-conditioned DCs. Bone marrow-derived DCs were stimulated with or without T3 (5 nM) or LPS (100 ng/mL) for 18 h. A total of 3.75 x 104 irradiated DCs were used to stimulate 1 x 106 allogeneic naive splenocytes. Brefeldin A (10 µg/ml) was added during the last 6 h of culture. Cells were stained with PE–conjugated anti-CD4- or FITC-conjugated anti-CD4 mAbs and then processed for intracellular cytokine staining using FITC-anti-IFN-{gamma} (A) and PE-anti-IL-5 (B) mAb, respectively. Values in dot plots show the percentage of activated CD4+ and CD4– T cells producing each cytokine. Results are representative of three independent experiments with similar results (P<0.01, T3-conditioned DCs vs. control DCs for IFN-{gamma}; NS, T3-conditioned DCs vs. control DCs for IL-5).

Control of DC maturation by T3 involves shuttling and nuclear translocation of NF-{kappa}B
The NF-{kappa}B pathway regulates different processes associated with DC maturation and function (13) . This signaling pathway is activated by LPS or cytokines through phosphorylation of the NF-{kappa}B inhibitory protein kinases, which in turn phosphorylate the inhibitory protein I{kappa}Bs that are bound to the NF-{kappa}B transcription factors in the cytoplasm. Phosphorylated I{kappa}Bs are then degraded by the proteasome, allowing the NF-{kappa}B transcription factors to translocate to the nucleus and activate gene transcription (13) . To gain insights into the mechanisms involved in T3-induced DC maturation, we investigated the potential role of NF-{kappa}B in this process. Analysis of the cytoplasmic-nuclear shuttling of this transcription factor revealed a substantial decrease in NF-{kappa}B/p65 in the cytoplasmic fractions of T3-treated compared with control DCs. In contrast, NF-{kappa}B/p65 was increased in the nuclear fractions of T3-treated cells (Fig. 9 ). Interestingly, T3-induced NF-{kappa}B translocation was even more pronounced than that observed in LPS-matured DCs (Fig. 9) , suggesting involvement of the NF-{kappa}B signaling pathway in T3 effects. Thus, T3-induced DC maturation involves shuttling of NF-{kappa}B/p65 from the cytoplasmic compartment to the nucleus, a critical event in DC maturation and function.


Figure 9
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  		Figure 9. Control of the cytoplasmic-nuclear shuttling of NF-{kappa}B by T3. DCs were stimulated with T3 (5 nM), LPS (100 ng/ml), or a combination of T3 and LPS for 18 h. Nuclear and cytoplasmic DC extracts (40 µg) were used for immunodetection of NF-{kappa}B/p65. Anti-{alpha}-tubulin and HDAC1 Abs were used to control the purity of subcellular fractions. A) Representative Western blots of three independent experiments. B) Densitometric analysis of immunoreactive protein bands. Results are expressed as arbitrary units (AU) calculated from the densitometric profile of the NF-{kappa}B/p65 signal normalized to β-actin. Data are presented as mean ± SD of six independent experiments. *P < 0.01 vs. control DCs; {dagger}P < 0.05 vs. T3-stimulated DCs and LPS-stimulated DCs.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
Despite major advances in understanding of the interplay between distinct hormones and the immune cell network (11 , 16 , 18) , the role of THs in the initiation of adaptive immunity still remains uncertain (20) . Here we provide the first evidence of the expression of TRs on bone marrow-derived murine DCs and their striking localization in the cytoplasmic compartment of iDCs and mDCs. Furthermore, we demonstrate that THs contribute to DC maturation and IL-12 production and potentiate the T cell stimulatory capacity of these cells.

Until a few years ago, TH-mediated effects were thought to be primarily, if not solely, initiated by T3 binding to nuclear TRs attached to specific DNA sequences in the promoter region of target genes. However, extranuclear actions exerted mainly through a plasma membrane receptor for thyroxine (T4) (integrin {alpha}Vβ3) were recently described (31) . Moreover, effects mediated by the classical nuclear TRs with higher affinity for T3 than for T4, but exerted through cytosolic (32) and plasma membrane (33) mechanisms, were also reported. Albeit unusual, the preferential cytoplasmic compartmentalization of TRs at both stages of DC maturation was in accordance with TR localization in bone marrow mast cells (34) , human hypothalamic and pituitary cells (35) , human umbilical vein endothelial cells (36) , rat hepatocytes (37) , and rat thymic cells (38) . In turn, several reports indicate that TRs may shuttle rapidly between the nuclear and the cytoplasmic compartments (39 40 41) . Furthermore, the dual cytoplasmic and nuclear localization appears to be a general feature of steroid hormone receptors (39 , 42) . In addition, the higher expression of TRβ1 compared with that of TR{alpha}1 in bone marrow-derived DCs is in consonance with the isoform distribution reported in other mouse immune cells including B and T lymphocytes (29 , 30) . However, the functional relevance of this particular subcellular distribution and the isoform prevalence of TRs in DC subsets still need to be fully elucidated.

The expression of TRs has recently been reported in other APCs such as macrophages as part of a functional nuclear receptor Atlas (43) , although the effects of THs on these cells have not yet been described. Moreover, other members of the nuclear receptor superfamily have recently been identified in human and mice DCs, as the nonsteroid retinoic acid receptor (44) , the peroxisome proliferator-activated receptors {alpha}, {delta}, {gamma}1, and {gamma}2 (45) , and the steroid estrogen and glucocorticoid receptors (17 , 46) .

The endocrine system participates in regulating the differentiation and maturation of different DC subtypes, e.g., thyroid-stimulating hormone induces a stimulatory effect on phagocytosis and cytokine production in murine DCs (47) . In addition, mRNAs for estrogen receptor-{alpha} and -β have been demonstrated in CD14+ monocytes, cultured immature CD1a+ cells, and mature CD83+ cells (48) . In culture, bone marrow progenitors give rise to the generation of DCs (49) , which can be influenced by the action of androgens and estrogens (48 , 50) . Moreover, glucocorticoids inhibit the in vitro differentiation of DCs from their progenitors and impair their capacity to undergo terminal differentiation or generate proinflammatory cytokines (17) . Our results indicating a positive role for T3 in triggering T cell-mediated immunity are in broad agreement with earlier observations showing a T3-mediated stimulatory effect on mitogen activation of T cells (51 , 52) . In addition, a differentiating effect of THs and other iodinated compounds was observed in the transition of human monocytes into veiled/DCs (20) . Our study provides the first evidence of a stimulatory effect of T3 on DC maturation and function with critical implications in orchestrating protective immunity and/or inciting T helper 1 (Th1)-mediated immunopathology.

Similarly to LPS-induced DC maturation (53) , exposure to T3 results in increased secretion of IL-12p70. In this regard, previous studies reported increased synthesis of IL-12 by DCs obtained from hyperthyroid mice (54) , as well as increased amounts of IL-12 in sera from patients with Graves’ disease (55) . Although these situations cannot be directly extrapolated to the experimental conditions of our work (use of physiological concentrations of T3), further studies in vitro and in vivo are required to determine the different effects of increasing amounts of THs found in sera from patients with thyroid-related pathological conditions on the maturation and immunostimulatory capacity of DCs. As IL-12p70, a heterodimeric cytokine composed of the p40 and p35 subunits, is essential for the promotion and maintenance of Th1 differentiation (56) , T3 might critically influence the development of T1-mediated immunity in vivo. Accordingly, our results reveal that T3 does not promote the secretion of IL-10 by DCs in contrast with the tolerogenic and regulatory responses induced by glucocorticoids (57) , which share metabolic actions similar to those of THs in the context of other target tissues such as the pituitary (58) and the liver (59) . In this regard, we found that T3-conditioned DCs are capable of directing the development of a T1-type cytokine response in vitro.

Circulating iDCs migrate within peripheral tissues, suggesting that at least part of the DCs found in the bloodstream might in fact represent a circulating pool of APCs available for immediate recruitment to sites of inflammation, where their antigen sampling and processing function is required (60) . Unlike most other hormones, THs circulate at relatively constant levels throughout postnatal life in human and animal species (6) . Therefore, the effects of circulating THs in the modulation of DC function in vivo is expected, given the stimulatory effects of physiological levels of T3 on DC maturation in vitro. In addition, DCs simultaneously exposed to LPS and T3 showed a greatly enhanced capacity to activate T cell responses, suggesting that T3 might potentiate LPS-induced initiation of adaptive immune responses during infectious processes. Studies are currently being conducted to address the role of T3 in the modulation of DC physiology in vivo and the effects of these hormones in inciting and perpetuating Th1-mediated immunopathology.

The transcription of proinflammatory cytokines including IL-12p70 is controlled, at least in part, by the transcription factor NF-{kappa}B (61) . Accordingly, the increased cytoplasmic-nuclear shuttling of NF-{kappa}B after exposure of DCs to T3 may be associated with increased activity of the NF-{kappa}B pathway during T3-induced DC maturation and IL-12 production. In conclusion, our results provide the first evidence of the presence of TRs in immature and mature DCs and their preferential cytoplasmic localization. In addition, we demonstrate the effects of THs in DC differentiation and function, suggesting their possible role in controlling the initiation of adaptive immune responses. Our findings broaden our perspective of the interactions between the endocrine and immune systems, providing a novel link between THs and the initiation of T cell responses. Because thyroid-related pathological conditions are the most common endocrine dysfunctions, our observations may contribute to understanding the molecular bases of immune-mediated pathological conditions of the thyroid gland and the immunological consequences of hypo- and hyperthyroid disorders. In addition, our findings provide a novel molecular target for manipulating the immunogenic potential of DCs to positively regulate the development of protective immunity or negatively control the generation of autoimmune thyroid inflammation.


   ACKNOWLEDGMENTS
 
This work was supported by grants from Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Agencia Nacional de Promoción Ciencia y Técnica (FONCYT), Secretaría de Ciencia y Tecnología de la Universidad Nacional de Córdoba, and Third World Academy of Sciences and Fundación Sales/CONICET. I.D.M. is a research fellow of FONCYT. M.M.M., S.S., and J.M.I. are research fellows of CONICET. H.M.T., G.A.R., and C.G.P. are members of the research career of CONICET. The authors thank Dr. Mariana Matrajt, Department of Microbiology and Molecular Genetics, University of Vermont, Burlington, VT, USA, for excellent assistance in this work.


   FOOTNOTES
 
1 These authors contributed equally to this work.

Received for publication June 15, 2007. Accepted for publication October 4, 2007.


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 DISCUSSION
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