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*
Laboratory of Experimental Surgery, Surgical Department, S. Raffaele Scientific Institute, Milan, Italy;
Department of Immunology and Cell Biology, "Mario Negri" Institute, Milan, Italy;
Laboratory of Analysis, San Luigi Centre, S. Raffaele Scientific Institute, Milan, Italy; and
§
University of Milan, Milan, Italy
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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or CD40 ligand, Dex-DC expressed lower levels of CD83 and CD86 than
untreated cells. Dex-DC showed a higher endocytic activity, a lower APC
function, and a lower capacity to secrete cytokines than untreated
cells. Overall, these results indicate that DC differentiated in the
presence of Dex are at a more immature stage. Moreover, Dex also
partially blocked terminal maturation of already differentiated DC. In
conclusion, our data suggest that glucocorticoids may act at the very
first step of the immune response by modulating DC differentiation,
maturation, and function. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, IL-1) or stimulation via CD40
ligand (CD40L) or LPS, DC up regulate adhesion and costimulatory
molecules to become more a potent stimulator of T cell immunity (8, 9).
In the last years, a number of new procedures have been developed to
obtain DC starting from CD34+ hemopoietic progenitor cells
of peripheral blood and bone marrow (10, 11) by culturing with TNF-
and GM-CSF or from human blood monocytes by culturing with GM-CSF, IL-4
(8, 12, 13), or IL-13 (14). In vitro-differentiated DC show functional
and phenotypic characteristics of immature DC (i.e., high capacity
of Ag uptake and processing, low capacity to stimulate T cells
proliferation) and can be further differentiated in vitro into mature
DC with TNF-, LPS, IL-1, or CD40L. It is obviously important to
identify factors that might affect the differentiation and maturation
of DC.
Glucocorticoids (GCs) are widely used as antiinflammatory and
immunosuppressive agents in the therapy of many autoimmune and allergic
diseases and in transplantation to prevent rejection. Moreover, GCs are
hormones naturally released during the course of an immune response
(15). This production is considered a crucial negative feedback to
regulate the magnitude of the response, thus preventing potential
damage to the host (16). GCs affect the immune and inflammatory
responses in many different ways. Most importantly, GCs down-regulate,
in monocyte/macrophages, the expression of many cytokine genes at the
transcriptional and posttranscriptional levels (17), including IL-1
(18, 19, 20, 21), IL-6 (22, 23), TNF- (24), IL-10 (25), and macrophage
inflammatory protein-1 (26). In addition, GCs regulate several
immunologically relevant activities of these cells including the
secretion of PGs, bactericidal (27) and fungicidal activities (26, 28),
phagocytosis and pinocytosis (29, 30), and the expression of
surface receptors for complement and Igs. GCs also regulate cytokine
secretion in T lymphocytes and inhibit their mitogenic potential
(31, 32, 33, 34, 35). In contrast, GCs up regulate cytokine receptor
expression (36, 37, 38, 39), which correlates with enhanced cytokine effects on
target cells (17). On the basis of these observations, the activities
of GCs have been attributed primarily to their influence on
monocytes/macrophages and on T cells (40, 41, 42, 43). The aim of our work was
to study the effects of GCs on human DC differentiation and maturation
because these cells, rather than macrophages and monocytes, are
responsible for the initiation of T cell-mediated immunity. In vitro
pharmacological modifications of DC differentiation, maturation, and
function may be useful to optimize their capacity of modulating T
cell-mediated immune responses, thus offering new chances in
immunotherapeutic protocols. Here we report that exposure of
monocytes to dexamethasone (Dex) together with GM-CSF and IL-4 inhibit
their differentiation to DC. Moreover, Dex partially blocked terminal
maturation of already differentiated DC.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Human recombinant GM-CSF (sp. act. 1.1 x 104
U/µg) was obtained from Novartis (Basel, Switzerland). Human
recombinant IL-4 (sp. act. >2 x 106 U/mg) and human
recombinant TNF- (sp. act. >2 x 107 U/mg) were
obtained from PeproTech (London, U.K.). Water-soluble Dex was obtained
from Sigma (St. Louis, MO). Human recombinant IL-2 was obtained from
Chiron (Milan, Italy). Human recombinant IL-10 was obtained from
Schering-Plough (Kenilworth, NJ). LPS was obtained from Sigma
(Escherichia coli 0127:B8).
DC culture
Highly enriched monocytes (>80% CD14+) were obtained from buffy coats of 20 blood donors (through the courtesy of Centro Trasfusionale, Ospedale San Raffaele, Milan, Italy) by Ficoll and Percoll gradients and purified by adherence. Monocytes were cultured for 7 days at 1 x 106/ml in 6-well Multiwell tissue culture plates (Falcon, Becton Dickinson, Rutherford, NJ) in RPMI (Biochrom, Berlin, Germany) in 10% FCS (HyClone, Logan, UT) supplemented with 50 ng/ml GM-CSF, 10 ng/ml IL-4, and with (Dex-DC) or without (Ctr-DC) different concentrations of Dex. In the control group (GM-CSF plus IL-4), the cell yield was about 80% of input cells. All the cultures were tested for the presence of endotoxin (<0.03 U/ml; lymulus test).
DC maturation
TNF- (10 ng/ml) or LPS (1 µg/ml) was added to induce
maturation of DC. Alternatively, J558L cells transfected with CD40L
(J558LmCD40L) were used to induce CD40 triggering on DC. Untransfected
J558L cells were used for control cultures. J558L after
irradiation (10,000 rad) were seeded together with DC at a 1:1 ratio in
24-well culture plates in culture medium (1 x 106
cells/well). Cells were recovered after 48–72 h of culture.
FACS analysis
Cell staining was performed using mouse mAbs followed by FITC-conjugated affinity-purified, isotype-specific goat anti-mouse Abs (Ancell, Bayport, MN). The following mAbs were used: L243 (IgG2a, anti-MHC class II), TS1/18 (IgG1, anti-CD18), TS1/22 (IgG1, anti-CD11a), 17aba (anti-CD11b), 32.2 (anti-CD32), and IV.3 (anti-CD64) (obtained from American Type Culture Collection, Manassas, VA); HK14 (IgG2a, anti-MHC class II), UCHM-1 (IgG2a, anti-CD14), and W6/32 (IgG2a, anti-MHC I) (obtained from Sigma); SK9 (IgG2b, anti-CD1a) and L306 (IgG2a, anti-CD58) (obtained from Becton Dickinson, San Jose, CA); B73.1 (IgG2a, anti-CD16) and PAM-1 (IgG1, anti-mannose receptor (MR) produced by immunizing mice with human alveolar macrophages (44)) (a kind gift of Dr. P. Allavena, Milan, Italy); BB1 (IgM, anti-CD80), BU63 (IgG1, anti-CD86), and EA-5 (IgG1, anti-CD40) (obtained from Ancell); HB15a (IgG2b, anti-CD83) (obtained from Immunotech, Marseille, France); CBR-IC3/1 (IgG1, anti-CD50) (obtained from Alexis Corporation, Nottingham, U.K.); and DX2 (IgG1, anti-CD95) (obtained from PharMingen, San Diego, CA). Results are expressed as the percent of positive cells or as fluorescence intensity (FI), calculated according to the formula: FI = mean fluorescence (sample) - mean fluorescence (control). The comparison of fluorescence intensity between two different groups was calculated as comparative FI index (CFII) according to the formula: CFII = (FI sample B)/(FI sample A).
Endocytosis
MR-mediated endocytosis was measured as the cellular uptake of FITC-dextran (FITC-DX) and quantified by flow cytometry. Approximately 2 x 105 cells for sample were incubated in media containing FITC-DX (1 mg/ml) (m.w. 40,000; Sigma) for 0, 60, and 120 min. After incubation, cells were washed twice with PBS to remove excess dextran and fixed in cold 1% formalin. The quantitative uptake of FITC-DX by the cells was determined using FACS. At least 8000 cells per sample were analyzed. Fluid-phase endocytosis via membrane ruffling was measured as the cellular uptake of 1 mg/ml of Lucifer yellow (LY) dipotassium salt (Sigma) and quantified by flow cytometry.
MLR
DC cultured in GM-CSF plus IL-4 and with or without Dex for 7 days were extensively washed, irradiated (3000 rad from a 137Cs source), and added in graded doses to 1 x 105 responder cells in 96-well flat-bottom microtest plates (Costar, Cambridge, MA). Responder cells were purified allogeneic T cells depleted of autologous APC by passage with CD14- and CD19-coated Dynabeads (Unypath, Milan, Italy). Each group was performed in triplicate. Thymidine incorporation was measured on day 5 by a 16-h pulse with [3H]thymidine (1 µCi/well, sp. act. 5 Ci/mM; Amersham Life Science, Buckingham, U.K.).
Ag presentation assay
Tetanus toxin (TT)-responsive T cell lines were generated in our
laboratory by culturing mononuclear cells with TT (36 µg/ml;
Cannaught Laboratories, Willowdale, Ontario, Canada) for 1 mo in the
presence of IL-2. TT-responsive T cells were tested at least 2 wk after
the last PBMC stimulation and 5 days after the last addition of IL-2.
DC were obtained from the same donor by culturing monocytes. After 7
days, DC were preincubated with TT (6 µg/ml) for 12 h and with
different maturation-inducing stimuli (TNF-, CD40L) for 48 h.
Then, DC were extensively washed, irradiated (3000 rad), and cocultured
with TT autologous responsive T cell lines for 72 h in 96-well
microtiter plates, and [3H]thymidine uptake was measured
during the last 12 h of culture (1 µCi/well, sp. act. 5 Ci/mM;
Amersham Life Science).
Quantification of cytokines
After 7 days of culture with GM-CSF (50 ng/ml) and IL-4 (10
ng/ml) in the presence or absence of Dex, DC were washed twice and
cultured for 3 days at 1 x 106/ml in a 24-well
flat-bottom plate (Costar). DC were either nonstimulated or stimulated
with TNF- (10 ng/ml), J558L cells transfected with CD40L
(J558LmCD40L), untransfected J558L cells, or LPS (50 ng/ml). After 3
days, medium was collected and TNF-, IL-6, IL-1ß, and IL-12 p70
were quantified by ELISA (Benfer-Scheller, Milan, Italy).
Electron microscopy
DC were processed for electron microscopy. DC were fixed for 2 h in 2.5% glutaraldeyde in 0.1 M cacodylate buffer. Then, DC were postfixed in 1% OsO4 in cacodylate buffer at 4°C for 1 h, dehydrated in graded ethanol up to propylene-oxide, and finally embedded in an Epon-Araldite mixture. Well-preserved areas were identified by light microscopy of semithin sections (0.5 mm). Subsequently, serial ultrathin sections (80 nm) were mounted on 200-mesh copper grids, stained with uranyl acetate and lead citrate, and finally examined with a Zeiss CEM 902 electron microscope (Oberkochen, Germany).
Calculations and statistical analysis
Data were expressed as mean ± SD. Comparisons were performed by Student’s t test. Values of p < 0.05 were considered statistically significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To investigate the effect of Dex on DC differentiation from human
monocytes, we cultured monocytes in the presence of GM-CSF, IL-4
(Ctr-DC), and different concentrations (10-10 M to
10-6 M) of Dex (Dex-DC). At concentration higher than
10-8 M, Dex significantly affected cell recovery (cells
yield as percentage of Ctr cells: 10-7 M, 75 ± 6;
10-6 M, 50 ± 8; n = 20) (Fig. 1
) The standard concentration of Dex
chosen for the study was 10-8 M (cells yield as percentage
of Ctr cells: 92 ± 13; n = 20). Upon culture with
GM-CSF and IL-4, the cells became nonadherent, clustered, with abundant
cytoplasm and protruding veils typical of DC (6, 11). In the presence
of Dex 10-8 M, DC showed a more irregular external surface
with more cytoplasmic projections and endocytic vacuoles (Fig. 2). The presence of Dex during the
differentiation of DC from monocytes induced a modification of membrane
phenotype (Fig. 3). Ctr-DC expressed high
levels of CD1a and were negative or low positive for CD14 and CD16,
while Dex-DC were negative or low positive for CD1a but expressed high
levels of CD14 and CD16 (CFII Dex vs Ctr: CD1a = 0.26; CD14 =
14.9; CD16 = 6.78; n = 20) (Table I). This effect was dose and
time-of-exposure dependent. In fact, the addition of Dex at day +2 and
+6 after initiation of the 7-day culture had a lower inhibitory
activity (Figs. 4 and
5). A partial conversion to a
monocyte/macrophage phenotype was also seen when we added Dex for 7
days to differentiated immature DC. Analysis of MHC class I, MHC class
II molecules, and CD80 showed an up-regulation in Dex-DC (CFII Dex vs
Ctr: MHC I = 1.61; MHC II = 1.31; CD80 = 1.97;
n = 20), while the expression of costimulatory
molecules CD40, CD86 (Fig. 5), and CD58 was decreased (CFII Dex vs Ctr:
CD86 = 0.67; CD40 = 0.79; CD58 = 0.84; n
= 20). Furthermore, Dex up-regulated adhesion molecules like CD11a,
CD11b, CD18, and CD54 and did not influence the expression of CD50.
Ctr-DC and Dex-DC were both negative for CD68 and CD83. To evaluate
whether the Dex-induced modifications in DC were reversible, DC were
cultured for 7 days in the presence of GM-CSF, IL-4, and Dex; then Dex
was washed away and cultures were prolonged with GM-CSF and IL-4 for 7
days. We compared these cells with DC obtained after 14 days of culture
in GM-CSF, IL-4, and Dex. The modifications induced by Dex, as
evaluated by phenotypic analysis, remained constant even after 7 days
of culture without Dex (data not shown).
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Immature DC such as the cells derived by culturing monocytes with
GM-CSF and IL-4 express a potent ability to uptake external molecules,
essentially via two main mechanisms: a receptor-mediated endocytosis
and a fluid-phase endocytosis (macropinocytosis) (7). To study the
endocytic activity of DC, two fluorescent markers were used: LY, a
nonspecific fluid-phase marker, and FITC-DX, which is mainly taken up
via the MR. Efficient accumulation of FITC-DX has been shown to be
characteristic of immature DC, and neither monocytes nor macrophages
share this property (7). Dex-DC showed a vigorous endocytosis of
FITC-DX, higher than Ctr-DC (2.3-fold increase at 1 h, 2.4-fold
increase at 2 h, n = 13) (Fig. 6A). The same behavior was
seen when we used LY as a marker of fluid-phase pinocytosis (1.8-fold
increase at 1 h, 1.6-fold increase at 2 h, n
= 13) (Fig. 6B). We investigated the expression of two
receptors involved in Ag capture in DC: MR and IgG FcRII (CD32). Dex
up-regulates MR expression on DC (CFII = 1.61, n =
18, p < 0.05 vs Ctr). Similar up-regulation was
observed for IgG FcRII CD32 (CFII = 2.16, n = 18,
p < 0.05 vs Ctr). The cells cultured in the presence
or absence of Dex were both negative for IgG FcRI (CD64)
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or
CD40L
DC obtained with GM-CSF and IL-4 show functional and phenotypic
characteristics of immature DC and can be further differentiated in
vitro into mature DC. TNF- 10 ng/ml was added to induce maturation
of DC for 48 h. Alternatively, J558L cells transfected with CD40L
(J558LmCD40L) were used to induce CD40 triggering on DC. Ctr-DC exposed
to maturation-inducing stimuli showed an increase of MHC I, MHC II,
CD80, CD86, CD40, CD54, CD58, and CD83 expression and a reduction of
CD1a and MR (Table I
). Dex-DC were less sensitive to
maturation-inducing stimuli. In fact, after TNF- exposure the
up-regulation of MHC I, MHC II, CD80, CD86, CD40, and CD83 expression
was quantitatively less evident in Dex-DC than in Ctr-DC (Table I). The
same behavior was seen after the exposure to the more powerful stimulus
CD40L. Inhibition of DC maturation by Dex was confirmed also in
endocytosis assay. The endocytic activity of FITC-DX and LY, despite a
reduction induced by maturation, remained higher in Dex-DC than in
Ctr-DC (1.35-fold increase at 1 h, 2.82-fold increase at 2 h
for FITC-DX, n = 6, p < 0.05;
1.33-fold increase at 1 h, 1.85-fold increase at 2 h for LY,
n = 6, p < 0.05).
Dex down-regulates the immunostimulatory capacity of DC
DC are potent stimulators of allogeneic T cells. We tested if
Dex-DC were able to stimulate allogeneic T lymphocytes in MLR. Dex-DC
showed an impaired capacity to induce MLR (Fig. 7). In vitro exposure of DC to TNF- or
CD40L (Fig. 7) increased T cells proliferation in Ctr-DC and in Dex-DC
but the ability of Dex-DC was always lower than control DC. The
down-regulation of APC function by Dex was noted even when DC were
formalin-fixed before setting-up the MLR (data not shown).
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In view of the fact that Ag capture was increased in cells
cultured with Dex but the stimulatory capacity was impaired in MLR, we
evaluated the ability to present soluble Ag that need to be uptaken and
processed. Cells differentiated in the presence of Dex showed much
lower efficiency in presenting TT to specific autologous T cell lines
(Fig. 8). In vitro exposure of DC to
TNF- or CD40L (Fig. 8) for 48 h after Ag pulsing increased T
cell proliferation in Ctr-DC and in Dex-DC, but the latter were
less potent than Ctr-DC.
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To investigate the capacity of Dex to interfere in cytokines
production, after 7 days of culture with GM-CSF and IL-4 with or
without Dex, DC were washed, seeded in the presence of
maturation-inducing stimuli, and cultured for 3 days. Supernatants were
quantified for IL-1ß, IL-6, IL-10, IL-12 p70, and TNF-. Dex-DC
showed a reduction of TNF- and IL-1ß production in response to the
maturation-inducing stimuli CD40L, LPS, and TNF-. IL-12 p70
production was decreased when Dex-DC were exposed to TNF- or LPS,
while with CD40L, a more powerful stimulus, the secretion of IL-12 p70
was comparable in Dex-DC and in Ctr-DC (Fig. 9). IL-6 and IL-10 production was not
affected.
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To evaluate the effects of Dex on the maturation of DC induced by LPS or ligation of CD40, Dex 10-8 M was added to 7-day culture DC for 48–72 h, together with the maturation stimulus. Dex partially inhibited the LPS-induced up-regulation of costimulatory molecules (e.g., CD40, CD80, CD83, and MHC II) and their accessory cell function in MLR. In addition, DC exposed to LPS and Dex had higher levels of MR and CD32 and higher endocytic activity of FITC-DX than LPS-treated control cells (data not shown). These results indicate that Dex interferes with process of DC maturation induced by LPS and freezes the cells at an immature stage. Similar result were obtained when CD40L-transfected cells where used as maturation stimulus, though inhibition of DC maturation was less striking, as ligation of CD40 is a more potent stimulus than LPS (data not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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or CD40L. In the literature, two
reports have shown that, in mice, corticosteroids inhibit DC
maturation. Moser have reported that Dex affects culture-induced
maturation of mouse DC by selectively inhibiting CD80 and CD86
expression (45). Kitajima have shown that Dex inhibits the T
cell-mediated terminal maturation of a murine epidermal-derived
dendritic cell line by diminishing CD86 expression and cytokine
secretion (51). In our system, Dex inhibited the MHCII, CD86, CD40, and
CD83 increase induced by maturation and the secretion of some
cytokines. The observation that Dex inhibits cytokine secretion by DC
is not unexpected. In fact GCs are known to down-regulate the capacity
of monocytes and macrophages to secrete IL-1 (18, 19, 20, 21), IL-6 (22),
TNF- (24), and IL-12 (52). The inhibition of secretion of those
cytokines that play a relevant role in the induction of T cell
responses demonstrates a relevant mechanism of GC action in the first
phase of the human immune response. The effects of Dex on immature DC, which have already been differentiated for 7 days in the presence of GM-CSF and IL-4, appear to be similar but not identical to the effects of Dex included at the beginning of the culture. Overall the effect of GCs on immature DC can be summarized as follows: a partial conversion to a monocyte/macrophage phenotype, an impaired capacity to reach maturation, and a decreased ability to stimulate T cells (data not shown). These results confirm the in vitro instability of immature DC generated with GM-CSF and IL-4 (53). Palucka et al. (54) showed that upon removal of both GM-CSF and IL-4 and/or reculture with M-CSF, immature CD1a+/CD14- DC easily converted to a macrophage phenotype-expressing CD14 with a decreased ability to stimulate allogeneic T cells. Dex showed a similar action even in the presence of GM-CSF and IL-4. It is tempting to speculate that GCs may act at two different steps of DC life: 1) by inhibiting the differentiation from blood precursors, thus impairing the normal turnover of DC in tissues, and 2) by inhibiting the terminal maturation of DC into a potent APC.
On the molecular mechanisms by which GCs modulate maturation and
differentiation of DC, we can speculate a possible role of the
NF-
B-Rel transcription factor family. NF-B-Rel protein’s family
plays an important role in the expression of genes involved in the
immune response or acute-phase reaction. DC contain high levels of all
known Rel family members and express strong activity for NF-B in DNA
binding (55, 56, 57). The expression of Rel B correlates with DC
differentiation, and disruption of Rel B expression in Rel
B-/- animals blocked the development of DC (57, 58, 59).
Recently, it was shown that GCs are potent inhibitors of NF-B in
mice and cultured cells. The activation of NK-B involves the
targeted degradation of its cytoplasmic inhibitor IB and the
translocation of NF-B to the nucleus. GCs induce the transcription
of the IB gene, which results in an increased rate of IB
protein synthesis and in a reduction of the amount of NF-B that
translocates to the nucleus (60, 61). Therefore inhibition of NF-B
activity by GCs is likely to have a role in the impaired DC
differentiation. Interestingly the effects of Dex on DC maturation and
differentiation are very similar to those observed with IL-10, an
antiinflammatory cytokine. IL-10 was shown to prevent differentiation
of monocytes to DC (14, 62), to promote their maturation to macrophages
(63), to impair DC’s capacity to induce a Th1 response, to inhibit
IL-12 secretion (64, 65), to increase DC’s capacity to capture Ags
(66), and to inhibit DC’s maturation (67, 68). IL-10 also inhibits
NF-B activity via an effect on IB (69). However, we can
exclude that GC’s effects on DC are mediated by IL-10 as we show that
Dex-DC did not secrete augmented levels of IL-10.
Another important point to clarify is whether the effects described in this study are relevant only at pharmacological doses of GCs or have relevance also for physiological concentrations. In humans, the main GC in plasma is cortisol. Cortisol is secreted episodically with 8–10 bursts per day, especially in the morning, leading to a diurnal fluctuation of plasmatic concentration ranging between 0.80 and 6.90 x 10-7 M (70). However only 3%–10% of circulating cortisol is in free state. In our experiments, we used a standard concentration of 10-8 M of Dex. Because Dex has a GC potency of 30 and cortisol is arbitrarily assigned a value of 1 (71), in terms of GC potency, 10-8 M of Dex corresponds to 3 x 10 -7 M of cortisol. This value is about 4- to 125-fold higher than the normal free plasma cortisol, and so we can assume that the effects described on DC differentiation and function by GCs are not relevant in normal conditions. However, in situations of acute stress or in some pathological conditions, endogenous GCs may reach levels that are in the range of the concentration used in this study. In fact, cortisol secretion increases up to 10-fold in acute stress situations (72) like trauma, surgeries, burns, etc. DC may be an important target of the immune system-CNS regulatory loop (72). Inflammatory cytokines (e.g., IL-1, IL-2, IL-3, IL-6, and TNF) and inflammatory mediators (e.g., PGF2 and platelet-activating factor) stimulate the hypothalamic-pituitary-adrenal axis to increase cortisol plasma levels (72). These, in turn, suppress the inflammatory and immune response via a variety of mechanisms, and this study shows that this suppression may occur also by inhibiting DC differentiation and function. Several lines of evidence suggest that high cortisol levels are associated with immunosuppression in several pathological conditions, including depression, chronic alcoholism, anorexia nervosa, and bulimia (71). Moreover, very high cortisol levels were seen in some malignant neoplasms like bronchial carcinoids, medullary thyroid carcinoma, and metastatic prostatic carcinoma due to the capacity of these tumors to produce corticotropin-releasing hormone (71). Also, patients with Cushing syndrome, characterized by hypercortisolemia, are highly susceptible to opportunistic infections (73), and there is a direct correlation between the risk of infection and the degree of hypercortisolism (73). Moreover, patients receiving corticosteroid therapy for long periods show a defect of cell-mediated immunity and susceptibility to opportunistic infections. In conclusion, our data suggest that GCs modulate DC differentiation, maturation, and function and these effects could be part of the dynamic regulatory interactions between the immune and neuroendocrine system. GCs may act at the very first step of the immune response by inhibiting maturation of DC into potent APC, thereby preventing the activation of naive T cell.
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Lorenzo Piemonti, Laboratory of Experimental Surgery, S. Raffaele Scientific Institute, Via Olgettina 60, 20132 Milan, Italy. E-mail address:
3 Abbreviation used in this paper: DC, dendritic cells; Dex, dexamethasone; GC, glucocorticoid; LY, Lucifer yellow; MR, mannose receptor; TT, tetanus toxin; CD40L, CD40 ligand; Ctr, untreated; FI, fluorescence intensity; CFII, comparative FI index; DX, dextran.
Received for publication October 30, 1998. Accepted for publication March 18, 1999.
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M. Bros, F. Jahrling, A. Renzing, N. Wiechmann, N.-A. Dang, A. Sutter, R. Ross, J. Knop, S. Sudowe, and A. B. Reske-Kunz A newly established murine immature dendritic cell line can be differentiated into a mature state, but exerts tolerogenic function upon maturation in the presence of glucocorticoid Blood, May 1, 2007; 109(9): 3820 - 3829. [Abstract] [Full Text] [PDF]
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G. Penna, A. Roncari, S. Amuchastegui, K. C. Daniel, E. Berti, M. Colonna, and L. Adorini Expression of the inhibitory receptor ILT3 on dendritic cells is dispensable for induction of CD4+Foxp3+ regulatory T cells by 1,25-dihydroxyvitamin D3 Blood, November 15, 2005; 106(10): 3490 - 3497. [Abstract] [Full Text] [PDF]
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 J A Hardin Dendritic cells: potential triggers of autoimmunity and targets for therapy Ann Rheum Dis, November 1, 2005; 64(suppl_4): iv86 - iv90. [Full Text] [PDF]
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L. Freeman, M. Hewison, S. V. Hughes, K. N. Evans, D. Hardie, T. K. Means, and R. Chakraverty Expression of 11{beta}-hydroxysteroid dehydrogenase type 1 permits regulation of glucocorticoid bioavailability by human dendritic cells Blood, September 15, 2005; 106(6): 2042 - 2049. [Abstract] [Full Text] [PDF]
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C. Chauveau, S. Remy, P. J. Royer, M. Hill, S. Tanguy-Royer, F.-X. Hubert, L. Tesson, R. Brion, G. Beriou, M. Gregoire, et al. Heme oxygenase-1 expression inhibits dendritic cell maturation and proinflammatory function but conserves IL-10 expression Blood, September 1, 2005; 106(5): 1694 - 1702. [Abstract] [Full Text] [PDF]
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G. Nalbandian, V. Paharkova-Vatchkova, A. Mao, S. Nale, and S. Kovats The Selective Estrogen Receptor Modulators, Tamoxifen and Raloxifene, Impair Dendritic Cell Differentiation and Activation J. Immunol., August 15, 2005; 175(4): 2666 - 2675. [Abstract] [Full Text] [PDF]
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G.-Y. Kim, K.-H. Kim, S.-H. Lee, M.-S. Yoon, H.-J. Lee, D.-O. Moon, C.-M. Lee, S.-C. Ahn, Y. C. Park, and Y.-M. Park Curcumin Inhibits Immunostimulatory Function of Dendritic Cells: MAPKs and Translocation of NF-{kappa}B as Potential Targets J. Immunol., June 15, 2005; 174(12): 8116 - 8124. [Abstract] [Full Text] [PDF]
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J Grisar, M Aringer, M D Koller, G H Stummvoll, D Eselbock, B Zwolfer, C W Steiner, B Zierhut, L Wagner, P Pietschmann, et al. Leflunomide inhibits transendothelial migration of peripheral blood mononuclear cells Ann Rheum Dis, December 1, 2004; 63(12): 1632 - 1637. [Abstract] [Full Text] [PDF]
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 S. N. Georas Inhaled Glucocorticoids, Lymphocytes, and Dendritic Cells in Asthma and Obstructive Lung Diseases Proceedings of the ATS, November 1, 2004; 1(3): 215 - 221. [Abstract] [Full Text] [PDF]
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 J. A. Long, M. Fogel-Petrovic, D. A. Knight, P. J. Thompson, and J. W. Upham Higher Prostaglandin E2 Production by Dendritic Cells from Subjects with Asthma Compared with Normal Subjects Am. J. Respir. Crit. Care Med., September 1, 2004; 170(5): 485 - 491. [Abstract] [Full Text] [PDF]
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P. Monti, B. E. Leone, A. Zerbi, G. Balzano, S. Cainarca, V. Sordi, M. Pontillo, A. Mercalli, V. Di Carlo, P. Allavena, et al. Tumor-Derived MUC1 Mucins Interact with Differentiating Monocytes and Induce IL-10highIL-12low Regulatory Dendritic Cell J. Immunol., June 15, 2004; 172(12): 7341 - 7349. [Abstract] [Full Text] [PDF]
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R. E. Wiley, M. Cwiartka, D. Alvarez, D. C. Mackenzie, J. R. Johnson, S. Goncharova, L. Lundblad, and M. Jordana Transient Corticosteroid Treatment Permanently Amplifies the Th2 Response in a Murine Model of Asthma J. Immunol., April 15, 2004; 172(8): 4995 - 5005. [Abstract] [Full Text] [PDF]
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 V. Mirenda, I. Berton, J. Read, T. Cook, J. Smith, A. Dorling, and R. I. Lechler Modified Dendritic Cells Coexpressing Self and Allogeneic Major Histocompatability Complex Molecules: An Efficient Way to Induce Indirect Pathway Regulation J. Am. Soc. Nephrol., April 1, 2004; 15(4): 987 - 997. [Abstract] [Full Text] [PDF]
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Z. Banki, L. Kacani, B. Mullauer, D. Wilflingseder, G. Obermoser, H. Niederegger, H. Schennach, G. M. Sprinzl, N. Sepp, A. Erdei, et al. Cross-Linking of CD32 Induces Maturation of Human Monocyte-Derived Dendritic Cells Via NF-{kappa}B Signaling Pathway J. Immunol., April 15, 2003; 170(8): 3963 - 3970. [Abstract] [Full Text] [PDF]
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 J Plumb, M A Armstrong, M Duddy, M Mirakhur, and S McQuaid CD83-positive dendritic cells are present in occasional perivascular cuffs in multiple sclerosis lesions Multiple Sclerosis, April 1, 2003; 9(2): 142 - 147. [Abstract] [PDF]
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A. M. Woltman and C. van Kooten Functional modulation of dendritic cells to suppress adaptive immune responses J. Leukoc. Biol., April 1, 2003; 73(4): 428 - 441. [Abstract] [Full Text] [PDF]
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 U. Kammerer, A. O. Eggert, M. Kapp, A. D. McLellan, T. B. H. Geijtenbeek, J. Dietl, Y. van Kooyk, and E. Kampgen Unique Appearance of Proliferating Antigen-Presenting Cells Expressing DC-SIGN (CD209) in the Decidua of Early Human Pregnancy Am. J. Pathol., March 1, 2003; 162(3): 887 - 896. [Abstract] [Full Text] [PDF]
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J. Bayry, S. Lacroix-Desmazes, C. Carbonneil, N. Misra, V. Donkova, A. Pashov, A. Chevailler, L. Mouthon, B. Weill, P. Bruneval, et al. Inhibition of maturation and function of dendritic cells by intravenous immunoglobulin Blood, January 15, 2003; 101(2): 758 - 765. [Abstract] [Full Text] [PDF]
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K. Duperrier, A. Farre, J. Bienvenu, N. Bleyzac, J. Bernaud, L. Gebuhrer, D. Rigal, and A. Eljaafari Cyclosporin A inhibits dendritic cell maturation promoted by TNF-{alpha} or LPS but not by double-stranded RNA or CD40L J. Leukoc. Biol., November 1, 2002; 72(5): 953 - 961. [Abstract] [Full Text] [PDF]
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H. Hackstein, T. Taner, A. J. Logar, and A. W. Thomson Rapamycin inhibits macropinocytosis and mannose receptor-mediated endocytosis by bone marrow-derived dendritic cells Blood, July 18, 2002; 100(3): 1084 - 1087. [Abstract] [Full Text] [PDF]
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A. M. Woltman, C. Massacrier, J. W. de Fijter, C. Caux, and C. van Kooten Corticosteroids Prevent Generation of CD34+-Derived Dermal Dendritic Cells But Do Not Inhibit Langerhans Cell Development J. Immunol., June 15, 2002; 168(12): 6181 - 6188. [Abstract] [Full Text] [PDF]
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M. Relloso, A. Puig-Kroger, O. M. Pello, J. L. Rodriguez-Fernandez, G. de la Rosa, N. Longo, J. Navarro, M. A. Munoz-Fernandez, P. Sanchez-Mateos, and A. L. Corbi DC-SIGN (CD209) Expression Is IL-4 Dependent and Is Negatively Regulated by IFN, TGF-{beta}, and Anti-Inflammatory Agents J. Immunol., March 15, 2002; 168(6): 2634 - 2643. [Abstract] [Full Text] [PDF]
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M. D. Saemann, O. Parolini, G. A. Bohmig, P. Kelemen, P.-M. Krieger, J. Neumuller, K. Knarr, W. Kammlander, W. H. Horl, C. Diakos, et al. Bacterial metabolite interference with maturation of human monocyte-derived dendritic cells J. Leukoc. Biol., February 1, 2002; 71(2): 238 - 246. [Abstract] [Full Text] [PDF]
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H. Hackstein, A. E. Morelli, A. T. Larregina, R. W. Ganster, G. D. Papworth, A. J. Logar, S. C. Watkins, L. D. Falo, and A. W. Thomson Aspirin Inhibits In Vitro Maturation and In Vivo Immunostimulatory Function of Murine Myeloid Dendritic Cells J. Immunol., June 15, 2001; 166(12): 7053 - 7062. [Abstract] [Full Text] [PDF]
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S. Corinti, C. Albanesi, A. la Sala, S. Pastore, and G. Girolomoni Regulatory Activity of Autocrine IL-10 on Dendritic Cell Functions J. Immunol., April 1, 2001; 166(7): 4312 - 4318. [Abstract] [Full Text] [PDF]
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K. D. Kim, Y.-K. Choe, I. S. Choe, and J.-S. Lim Inhibition of glucocorticoid-mediated, caspase-independent dendritic cell death by CD40 activation J. Leukoc. Biol., March 1, 2001; 69(3): 426 - 434. [Abstract] [Full Text]
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 B. Knight, G. C.T. Yeoh, K. L. Husk, T. Ly, L. J. Abraham, C. Yu, J. A. Rhim, and N. Fausto Impaired Preneoplastic Changes and Liver Tumor Formation in Tumor Necrosis Factor Receptor Type 1 Knockout Mice J. Exp. Med., December 18, 2000; 192(12): 1809 - 1818. [Abstract] [Full Text] [PDF]
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K. Shimizu, S.-i. Fujii, K. Fujimoto, K. Kawa, A. Yamada, and F. Kawano Tacrolimus (FK506) treatment of CD34+ hematopoietic progenitor cells promote the development of dendritic cells that drive CD4+ T cells toward Th2 responses J. Leukoc. Biol., November 1, 2000; 68(5): 633 - 640. [Abstract] [Full Text]
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D. Rea, C. van Kooten, K. E. van Meijgaarden, T. H. M. Ottenhoff, C. J. M. Melief, and R. Offringa Glucocorticoids transform CD40-triggering of dendritic cells into an alternative activation pathway resulting in antigen-presenting cells that secrete IL-10 Blood, May 15, 2000; 95(10): 3162 - 3167. [Abstract] [Full Text] [PDF]
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L. Piemonti, P. Monti, M. Sironi, P. Fraticelli, B. E. Leone, E. Dal Cin, P. Allavena, and V. Di Carlo Vitamin D3 Affects Differentiation, Maturation, and Function of Human Monocyte-Derived Dendritic Cells J. Immunol., May 1, 2000; 164(9): 4443 - 4451. [Abstract] [Full Text] [PDF]
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J. Komi and O. Lassila Nonsteroidal anti-estrogens inhibit the functional differentiation of human monocyte-derived dendritic cells Blood, May 1, 2000; 95(9): 2875 - 2882. [Abstract] [Full Text] [PDF]
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G. Penna and L. Adorini 1{alpha},25-Dihydroxyvitamin D3 Inhibits Differentiation, Maturation, Activation, and Survival of Dendritic Cells Leading to Impaired Alloreactive T Cell Activation J. Immunol., March 1, 2000; 164(5): 2405 - 2411. [Abstract] [Full Text] [PDF]
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C. E. Demeure, H. Tanaka, V. Mateo, M. Rubio, G. Delespesse, and M. Sarfati CD47 Engagement Inhibits Cytokine Production and Maturation of Human Dendritic Cells J. Immunol., February 15, 2000; 164(4): 2193 - 2199. [Abstract] [Full Text] [PDF]
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 L. Piemonti, P. Monti, P. Allavena, B. E. Leone, A. Caputo, and V. Di Carlo Glucocorticoids increase the endocytic activity of human dendritic cells Int. Immunol., September 1, 1999; 11(9): 1519 - 1526. [Abstract] [Full Text] [PDF]
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