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Immunobiologie Fondamentale et Clinique, Institut National de la Santé et de la Recherche Médicale U503, Ecole Normale Supérieur Lyon, Lyon, France;
Biologie des Cellules Dendritiques Humaines, Institut National de la Santé et de la Recherche Médicale E 9908, Etablissement de Transfusion Sanguine Strasbourg, Strasbourg, France;
Immunité et infections virales, Faculté de médecine Laennec, IVMC-Centre National de la Recherche Scientifique-Université Claude Bernard Lyon Unité Mixte de Recherche 5537, Lyon, France; and
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Développement Normal et Pathologique du Système Immunitaire, Institut National de la Santé et de la Recherche Médicale U429, Hôpital Necker-Enfants Malades, Paris, France
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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or LPS, respectively. CD40
ligand (CD40L) expressed on activated T cells is shown to induce
terminal differentiation of DCs into mature effector DCs. In contrast,
the CD40L-dependent maturation of DCs is inhibited by MV infection, as
demonstrated by CD25, CD69, CD71, CD40, CD80, CD86, and CD83 expression
down-regulation. Moreover, the CD40L-induced cytokine pattern in DCs is
modified by MV infection with inhibition of IL-12 and IL-1/ß and
induction of IL-10 mRNAs synthesis. Using peripheral blood lymphocytes
from CD40L-deficient patients, we demonstrate that MV infection of DCs
prevents the CD40L-dependent CD8+ T cell proliferation. In
such DC-PBL cocultures, inhibition of CD80 and CD86 expression on DCs
was shown to require both MV replication and CD40 triggering. Finally,
for the first time, MV was shown to inhibit tyrosine-phosphorylation
level induced by CD40 activation in DCs. Our data demonstrate that MV
replication modifies CD40 signaling in DCs, thus leading to impaired
maturation. This phenomenon could play a pivotal role in MV-induced
immunosuppression. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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DCs belong to a family of professional APCs responsible for the
generation of effector CD4+ and
CD8+ T cells (9). They originate
from CD34+ bone marrow progenitors. Immature DCs
form a network within all epithelia, as Langerhans cells (LCs) in the
skin or DCs in the respiratory mucosa. These immature DCs are able to
capture particular Ags via phagocytosis (10) and soluble
Ags via macropinocytosis or receptor-mediated endocytosis
(11). They express low levels of MHC class II (MHC-II)
molecules at their cell surface. To become a potent APC, the immature
DCs need to be activated by stimuli that promote their maturation and
migration to the T cell areas of lymphoid tissues. Living bacteria,
microbial products (LPS), or various cytokines (TNF-, GM-CSF,
IL-1ß) stimulate DC maturation. Upon maturation, MHC-II molecules are
delivered to the plasma membrane (12) and the expression
of costimulatory membrane molecules is increased, thus favoring T cell
activation (13).
When the mature DCs reach secondary lymphoid organs, they interact with T cells, receiving signals which induce their terminal differentiation into mature effector DCs. CD40-CD40 ligand (CD40L) interaction between DCs and T cells is essential for an optimal cytokine production. The best-known consequence of CD40 ligation is the IL-12 production by DCs (14, 15). In human, the X-linked immunodeficiency hyper-IgM syndrome has been attributed to mutations in the CD40L gene (16). Over the past year, it was recognized that the function of CD40 accounts not only for the regulation of T-dependent humoral immune responses, but also for cellular immune responses (17). Several immune dysfunctions observed in CD40L-deficient mice and patients could be explained by a failure properly to activate APCs (18, 19, 20). Recent in vivo studies in mouse demonstrated that CD40 ligation on the DCs can replace CD4+ T cells to prime CD8+ cytotoxic responses (21, 22, 23).
The mechanisms by which MV infection interferes with the functions of DCs remained unknown. In this study, we confirm (24) and further extend that MV replication induces normal maturation of immature monocyte-derived DCs and LCs. But, we show that MV replication leads to an abnormal terminal differentiation of CD40L-activated human DCs. Impairment of CD40/CD40L signaling following MV infection was demonstrated by inhibition of tyrosine-phosphorylation level in MV-infected DCs after CD40 activation. This could explain why DCs display impaired APC functions and may consequently promote MV-induced immunosuppression.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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CD1a-PE (BL6), CD3-PE (UCHT1), CD25-PE (B1.49.9),
CD32-PE (2E1), CD45RO-PE (UCHL1), CD45RA-PE (ALB11), CD69-PE
(TP1.55.3), CD71-FITC (YDJ1.2.2), CD80-FITC (MAB104), CD83-PE
(HB15a), CD86-FITC (HA5.2B7), E-cadherin (67A4), and HLA-DR-FITC
(B8.12.2) Abs were purchased from Immunotech (Marseille, France); MHC-I
(W6/32), CD8-FITC (DK25), and CD4-FITC (MT310) Abs were from DAKO
(Glostrup, Denmark); CD40-PE (LOB7/6), CD86 (BU63), and CD11c-FITC
(3.9) Abs were from Serotec (Oxford, U.K.); and CD80-PE
(L307.4) and MHC-II (L243) were from Becton Dickinson
Immunocytochemistry Systems (San Jose, CA). A FITC-conjugated
IgG1/PE-conjugated IgG2a irrelevant Ab mixture (Immunotech) was used as
isotype controls. Mouse Abs specific for CD46 (20.6; Ref.
25) was produced in our lab. FITC-conjugated,
affinity-isolated F(ab')2 fraction of a sheep
anti-mouse Ig Ab (Silenus, Hawthorn, Victoria, Australia) was used
for indirect immunofluorescence labeling procedures. LPS
(Escherichia coli serotype O127:B8) was purchased from Sigma
(St. Louis, MO). Recombinant human (rh)GM-CSF and IL-4 were generously
provided by the Schering-Plough Laboratory for Immunological Research
(Dardilly, France), whereas rhSCF, rhTNF-, and purified hTGF-ß1
were obtained from R&D Systems (Abingdon, Oxon, U.K.).
Patients
Two patients suffering from X-linked hyper-IgM syndrome were included in this study. Mutations in the CD40L gene were characterized and led to the absence of CD40L expression. Informed consent was obtained from each patient family for this study.
Cells
Immature LCs were generated in vitro from
CD34+ progenitors. Positive selection of
CD34+ cells was performed as previously described
(26). Briefly, PBMC were collected by cytapheresis from
myeloma patients who had received high dose chemotherapy and
hematopoietic growth factors (G-CSF or GM-CSF). Informed consent was
obtained from all patients before cytapheresis. Hematopoietic
progenitors expressing the CD34 Ag were purified (92 ± 2.3%)
using the Ceprate LC34-Biotin Kit (CellPro, Bothell, WA) according to
the cell separation procedure instructions of the manufacturer. A total
of 2 x 104 purified
CD34+ cells/ml were cultured in 75-ml tissue
culture flasks (Falcon 3111, Becton Dickinson Labware, Franklin Lakes,
NJ) in serum-free StemPro-34 complete medium (Life Technologies, Grand
Island, NY) supplemented with L-glutamine (2 mM, Life
Technologies), gentamicin (50 µg/ml, Life Technologies), rhSCF (20
ng/ml), rhTNF- (0.5 ng/ml), rhGM-CSF (200 ng/ml), and hTGF-ß1 (0.5
ng/ml). After 7 days of culture the cell suspensions contained 10%
CD1a+ cells, with 70–98% of these expressing
CD1a and E-cadherin, two markers specifically found on the immature LCs
of the epidermis (13).
Monocyte-derived DCs (Mo-DCs) were generated in vitro, as previously described (4). After 6 days of culture in the presence of 50 ng/ml hrGM-CSF and 500 U/ml hrIL-4, >95% of the cells were DCs as assessed by CD1a labeling. Cultures of the Mo-DCs were performed in 24-well flat-bottom microtiter plates (Falcon), in a total volume of 1 ml, in RPMI 1640 (Life Technologies) supplemented with 10 mM HEPES (Life Technologies), 2 mM L-glutamine (Life Technologies), 40 µg/ml gentamicin (Life Technologies), and 10% FCS (Boehringer Mannheim, Meylan, France).
PBL were activated with a combination of 10 ng/ml PMA (Sigma) and 1 µg/ml ionomycin (Sigma) for 6–12 h. This short-time activation was provided to induce CD40L expression by CD4+ T cells, but no cytokine secretion and a limited background proliferation (5,000 cpm) compared with DC-dependent allogenic proliferation (80,000 cpm) of PBL. After activation, PBL were washed three times. DCs alone were cultured at 106 cells/ml. In PBL cocultures, 0.5 x 106 DCs/ml were cultured together with 0.5 x 106 activated PBL. In the murine fibroblast cocultures, 106 DCs/ml were cultured in the presence of 105 irradiated (7000 rads) fibroblastic CD40L or CD32-transfected L cells (both kindly provided by Schering-Plough Laboratory for Immunological Research).
MV infection and detection
Mo-DCs and LCs were infected, at day 6 and 7, respectively, with 1 PFU/cell of Vero cell-derived MV Hallé (Hallé strain is classified with the vaccine MV strain Edmonston (27)) or pulsed with 1 PFU/cell of MV neutralized by 254 nm UV rays for 30 min (UVMV) or mock-infected. After a 3-h incubation at 37°C, the DCs were washed three times to be free of unattached virus then put in culture. For CD40 stimulation and detection of tyrosine phosphorylation, DCs were previously infected with 4 PFU/cell; all of the DCs were infected and 15% were dead by apoptosis 24 h later when CD40 stimulation was performed. For PFU measurement, virus contents were quantified by limiting dilution from 10 to 10 until 10-10 on confluent Vero cells. A single plaque in the Vero cells confluent culture represents 1 PFU generated by an individual infectious virus. For MV nucleoprotein (NP) staining, after 15 min of permeabilization with 0.33% Saponin (Sigma), cells were stained with anti-NP viral protein mAb (clone 25) kindly provided by F. Wild, followed by incubation with PE-labeled anti-mouse Ig (Immunotech). The apoptosis rate differed between culture conditions according to labeling of fragmented DNA after TUNEL staining: at day 3, 50% of DCs were apoptotic when they are alone or cocultured with L cells, whereas 80% were apoptotic in DC-PBL cocultures (data not shown).
Phenotypic analysis
All immunostaining were performed in 1% BSA and 3% human serum-PBS. Direct immunostainings were performed using 2 µg/ml of FITC-conjugated or PE-conjugated Abs. Indirect immunostainings were performed using 2 µg/ml of the first mouse mAb and revealed with 2 µg/ml of the FITC-conjugated, affinity-isolated F(ab')2 fraction of a sheep anti-mouse Ig Ab. Viable DCs were gated according to negative staining with propidium iodide.
RNase protection assays
RNA was extracted from 107 treated Mo-DCs using RNA NOW-TC reagent (Biogentex, Seabrook, TX). The RNase protection was performed using 4 µg of RNA with the RiboQuant multiprobe RNase assay system (PharMingen, San Diego, CA), following the manufacturer’s specification. In brief, RNA was hybridized overnight with the in vitro-translated 32P-labeled probe (hCK-2 kits, PharMingen). Following hybridization, samples were treated with RNase A+T1 and proteinase K, phenol-chloroform extracted, and ethanol precipitated. The protected fragments were resolved by electrophoresis on a 5% acrylamide/urea gel and exposed on a Phosphor Screen (Molecular Dynamics, Sunnyvale, CA) for 12 h to quantify the intensity of the bands with ImageQuant (Molecular Dynamics).
Cell counts
CD4+ and
CD8+-labeled T cells in DC-PBL cocultures were
numbered by a time-monitored FACS analysis during 2 min at high speed
(1 µl/s). As the CD8 percentages differed in PBL used (34% in
healthy donors, 8% and 20% in CD40L-deficient patients), results of
Fig. 2
were calculated for 5 x 104
CD8+ T cells put in culture at day 0.
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DCs were stimulated with 10 µg/ml of monoclonal anti-CD40 (mAb 89) generously provided by the Schering-Plough Laboratory for Immunological Research or irrelevant IgG1 control Abs (Sigma) for 10 min at 37°C. Stimulation was terminated by lysis in RIPA buffer (150 mM NaCl, 50 mM Tris-HCl (pH 8.0), 1% Nonidet P-40, 0.5% deoxycholate, and 0.1% SDS) containing 5 mM EGTA, 1 mM Na vanadate, and a mixture of protease inhibitors (complete, Boehringer Mannheim) for 15 min at 4°C. Insoluble material was removed by centrifugation at 10,000 x g for 10 min. Proteins from cell lysates were separated by SDS-PAGE under reducing conditions and transferred to Immobilon-P membranes (Millipore, Bedford, MA). Membranes were blocked using 5% nonfat dried milk in TBS-T (20 mM Tris (pH 7.6), 130 mM NaCl, 0.1% Tween 20) and incubated for 1 h with the anti-P-Tyr Ab 4G10 (Upstate Biotechnology, Lake Placid, NY) in TBS-T. Immunoreactive bands were visualized by using secondary horseradish peroxidase-conjugated Abs (Promega, Madison, WI) and chemiluminescence (ECL, Amersham. Little Chalfont, U.K.).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To show that MV replicates in immature DCs, FACS analysis of NP
staining and measurement of infectious virus particles were performed.
NP is the earliest MV protein transcribed during viral cell cycle and
its amount, measured by mean fluorescence intensity (MFI), reflects the
intensity of viral replication in infected cells. Immature LCs or
Mo-DCs were MV-infected. Mo-DCs were cultured for 5 days alone or with
CD40L+-L cells or normal PBL or CD40L-deficient
PBL. At day 3 of culture, 12% of LCs and around 50% of DCs were
NP+, but normal PBL or
CD40L+-L cells enhanced MFI of
NP+ DCs (Table I,
columns 1 and 2). At day 5, PFU were measured in supernatants of
culture. We have previously shown that CD40L+ T
cells enhance viral production by DCs (4). In MV
DC-CD40L-deficient PBL cocultures, the absence of CD40L decreased PFU
measured in supernatant. High MV production was restored by addition of
CD40L+-L cells (Table I, column 3). Thus, high MV replication
in immature DCs correlates with CD40 triggering.
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Physiologically, MV may encounter immature DCs at its entry site
in the respiratory mucosa. As previously described (24),
immature DCs isolated from peripheral blood up-regulated MHC-II, CD83,
and CD86. LCs can be used as a model for epithelial immature DCs
functionally close to the respiratory tract DCs (28).
Immature LCs were MV-infected, then cultured for three days (Table II,
left columns 1–3). At day 3, immature LCs were positive for
E-cadherin, MHC-II, CD1a, and CD80, and negative for CD86 and CD83. In
contrast, after MV replication, E-cadherin and CD1a were
down-regulated, MHC-II and CD80 were up-regulated, whereas CD86 and
CD83 were induced. Mock supernatant and UVMV did not induce phenotypic
maturation (data not shown). Regarding CD1a, CD86, and CD83
expressions, MV replication induced stronger LC maturation than
TNF-.
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left columns 4–6), MV replication down-regulated CD1a, CD11c, and CD32
expression; up-regulated MHC-I, MHC-II, CD80, and CD40 expression; and
induced CD25, CD69, CD71, CD86, and CD83 expression in Mo-DCs. CD46
were also down-regulated as a function of MV production in agreement
with the fact that CD46 is a receptor for MV (29, 30).
Throughout the 14 markers that were studied, a similar maturation
profile was observed after MV infection compared with LPS activation.
Mock supernatant and UVMV did not induce Mo-DC maturation (data not
shown). On the basis of this phenotypic study, we confirm and further
extend to the LCs and Mo-DCs that MV replication induces maturation of
immature DCs. CD40-induced DC maturation is impaired by MV infection
Although MV-induced DC maturation was similar to LPS-induced DC
maturation, MV-infected DCs are deficient in APC functions (4, 5) in contrast to LPS-activated DCs. The CD40L expressed by
activated T cells is a potent signal to induce terminal differentiation
of DCs in mature professional APCs. We compared the consequences of LPS
activation vs MV infection of DCs for the integration of CD40L signal.
Effective CD40 activation of B cells (31) and DCs
(4) were previously shown by using CD40L-transfected mouse
fibroblasts (CD40L+-L cells). Mo-DCs were either
LPS-activated or MV-infected for 3 h, then cocultured with
CD40L+-L cells for 3 days (Table II, right). As
compared with immature DCs (Table II, left), CD40 ligation alone or
CD40 ligation of LPS-activated DCs showed a typical mature phenotype
with high MHC-I, MHC-II, CD25, CD69, CD71, CD40, CD80, CD86, and CD83
expression. In contrast, CD40 ligation of MV-infected DCs inhibited
induction of CD25, CD69, CD71, CD86, and CD83 and up-regulation of CD40
and CD80 expression. MV infection also down-regulated expression of the
CD46/MV receptor. Impaired phenotype was also observed when CD40 was
ligated 48 h before infection, and 24 or 48 h after infection
(data not shown). Thus, CD40-dependent maturation of Mo-DCs is
inhibited by MV replication.
CD40-induced cytokine pattern in DCs is modified by MV infection
We then compared cytokine mRNA productions of uninfected,
LPS-activated, MV-infected, CD40L-activated, and MV-infected +
CD40L-activated DCs. Mo-DCs were treated for 3 h, and then
cultured for 24 h. RNase protection assay was performed by using
specific probes to quantify the level of eight cytokine mRNAs
(Fig. 1). In immature DCs, only low
levels of IL-1ß and IL-1RA mRNAs were detected. LPS activation
induced IL-12p35, IL-12p40, IL-10, IL-1, and IL-6 mRNAs, whereas
IL-1ß and IL-1RA mRNAs were enhanced. MV infection weakly induced
IL-12p35, IL-12p40, IL-1, and IL-6 mRNAs, whereas IL-1ß and
IL-1RA mRNAs were up-regulated. CD40L signal strongly induced IL-12p40;
induced IL-12p35, IL-1, and IL-6 mRNAs; and up-regulated IL-1ß and
IL-1RA mRNAs in immature DCs. MV replication modified the CD40L-induced
cytokine pattern: IL-10 mRNA was induced, whereas neither
CD40L-activation nor MV infection alone induced IL-10 gene
transcription. Furthermore, IL-12p35, IL-12p40, and IL-1/ß mRNAs
were reduced in MV+CD40L condition compared with CD40L activation
alone. Thus, MV replication alters the cytokine pattern induced by
CD40L activation of DCs.
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Our data indicate that MV replication could modify the signal
transduced by CD40L in DCs. To investigate this point, PBL from healthy
donors or from CD40L-deficient patients were used. Normal T lymphocyte
numbers were detected in these patients, but T cell subpopulations were
profoundly affected (Table III), as
patient no. 1 had only 8% CD8+ T cells and 4%
of CD4+/CD45RO+
lymphocytes, whereas patient no. 2 presented normal percentages of
CD4+ and CD8+, but the
number of CD45RO+ T cells was abnormally
low.
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A). In contrast, no
viable CD8+ T cells could be detected at day 5
when DCs were cocultured with CD40L-deficient PBL. Detection of
CD8+ T cells at day 5 was dependent on CD40L
because bystander CD40L activation of the DCs by
CD40L+-L cells permitted us to recover a high
number of CD8+ T cells (x31.6 at day 5) in
CD40L-deficient PBL. Higher proliferation rate obtained with
CD40L+-L cells was attributed to the higher CD40L
expression in transfected L cells than in activated T cells (data not
shown). The absence of CD8+ T cells in the
DC-CD40L-deficient PBL cocultures did not result from an overall
inhibition of T cell proliferation. Indeed, PBL from CD40L-deficient
patients incorporated thymidine and allogeneic
CD4+ T cells were detected in increased number
(x8.9 at day 5; data not shown). Under the culture conditions
performed with a short preactivation of PBL by PMA and ionomycine,
CD8+ T cell proliferation in DC-PBL cocultures
was dependent on CD40-activation of DCs by CD40L+
CD4+-activated T cells.
We then investigated whether MV replication could impair this
CD40L-dependent CD8+ T cell proliferation.
Experiments performed in Fig. 2A were repeated with
MV-infected DCs (Fig. 2B). When the DCs were infected by MV,
CD8+ T cell proliferation was abolished in
allogeneic DC-PBL cocultures. Furthermore, the
CD8+ T cell proliferation obtained by adding
CD40L+-L cells in coculture of DC with
CD40L-deficient PBL was inhibited by MV infection of the DCs.
In conclusion, we have demonstrated that 1) CD40L activation of DCs is required to sustain human CD8+ T cell proliferation, in vitro, and 2) MV infection of DCs prevents this CD40L-dependent CD8+ T cell proliferation.
MV replication impairs CD40 signaling in DCs
To determine whether MV could modify CD40 signaling into the
Mo-DCs, we studied the expression of membrane Ags that were induced or
up-regulated by CD40L activation in DCs. DC-PBL cocultures were
performed using allogeneic PBL either from healthy donors or from
CD40L-deficient patients. In the absence of MV infection (Fig. 3A), DCs cocultured with
normal PBL exhibited a normal mature phenotype
(CD86+, CD80high,
MHC-IIhigh), whereas DCs cocultured with
CD40L-deficient PBL looked like an intermediate stage of maturation
with immature-type expression of CD86. CD86 expression was induced when
CD40L+-L cells were added to the CD40L-deficient
PBL, thus demonstrating that a CD40L signal was required to increase
CD86 expression. By contrast, the CD40L signal, which by itself
enhanced CD80 and MHC-II expression (Table II, right), could be
replaced by other(s) T cell signal(s) that increased CD80 and MHC-II
expressions on DCs (Fig. 3A). When DCs were MV-infected
(Fig. 3B), inhibition of CD86 expression was observed. This
latter event occurred even in the presence of
CD40L+-L cells, suggesting a blockade in CD40
signaling. CD80 expression was inhibited only when MV-infected DCs were
CD40L activated either with CD40L+-PBL or with
CD40L+-L cells. Thus, even in the presence of
other(s) T cell signal(s) able to up-regulate CD80 expression, CD40
triggering of MV-infected DCs did not up-regulate CD80 expression.
Therefore, both MV replication and CD40 triggering of DCs were needed
for inhibition of CD80 and CD86 expression. Although the nature of the
CD40 signaling pathway in DCs has not been elucidated, CD40 signaling
in monocytes and B cells has been shown to involve protein tyrosine
kinase activity (32). Since we demonstrated that MV
infection impairs CD40-induced maturation of DCs, we investigated
whether MV infection influences anti-CD40-induced tyrosine
phosphorylation (Fig. 4). The effect of
CD40 stimulation on overall levels of tyrosine phosphorylation in
mock-treated or MV-infected DCs was examined by Western blot analysis
of total protein using anti-phosphotyrosine Abs. The enhanced
tyrosine phosphorylation was evident in mock-treated DCs after 10 min
of stimulation with anti-CD40. But MV infection strongly inhibited
anti-CD40 enhanced tyrosine phosphorylation.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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/ß mRNA were decreased, whereas
IL-10 mRNA was induced. Using PBL originating from CD40L-deficient
patients and CD40L+-L cells, we demonstrate that
CD40L activation of DCs is required to induce
CD8+ T cell proliferation in vitro and that MV
replication inhibits this CD40L-dependent CD8+ T
cell proliferation. Finally, the inhibition of CD80 and CD86 expression
could be related to the impairment of CD40 signaling, which is
demonstrated by the inhibition of tyrosine-phosphorylation level in
MV-infected CD40-activated DCs. Therefore, the CD40 triggering of
MV-infected DCs could play a pivotal role in MV-induced
immunosuppression. Networks of DCs are found in tissues that come into closest contact with the external environment. According to the studies of Holt et al. (28) in rats, lung wall DCs share some properties of LCs: they can effectively bind inhaled Ags in situ, at the entry site of MV, but require additional maturation/activation signals before they can efficiently present the Ag to T cells. At the present time, we do not have formal proof that DCs are infected in vivo during measles, but five types of immature DCs are susceptible to MV infection and actively suppress T cell proliferation in vitro: Mo-DCs (4), CD34+-derived DCs (5) that contains two subpopulations of immature DCs (13), freshly isolated LCs contrary to surrounding keratinocytes (6), and now CD34+-derived LCs. So far, the best-characterized peripheral DCs are the epidermal LCs. To become potent APCs, these immature DCs must undergo a terminal differentiation step during the migration from the skin to draining lymph nodes that is induced by various stimuli such as LPS, cytokines, or pathogens. Phenotypic DC maturation induced by bacteria (reviewed in Ref. 33) and parasites (34) is documented. Concerning virus-induced DC maturation, poor information is available; blood-derived DCs up-regulate MHC-I, MHC-II, CD38, CD83, and CD86 after infection with influenza virus (35). Our data confirm previous work (24) and further extend to the LCs and Mo-DCs that DC maturation can be obtained with MV. MV induces down-regulation of E-cadherin in LCs; this loss of E-cadherin expression may be of particular relevance for effective migration, because E-cadherin participates in the homophilic adhesion between LCs and keratinocytes in the epidermis (36). Moreover, MV infection of Mo-DCs weakly induces IL-12 and IL-6 mRNAs. Weak IL-12 secretion by MV-infected human blood DC precursors has been already described (24).
When DCs were infected before CD40-mediated signal, MV replication
decreased expression of coactivation (CD40, CD80, CD86), activation
(CD25, CD69, CD71), and maturation (CD83) marker molecules, whereas
MHC-I and MHC-II expressions remained high. Abnormal phenotype was also
observed when Mo-DCs were infected 48 h after CD40 activation
(data not shown). In such culture conditions, DCs showed normal mature
phenotype before they were infected. Thus, MV infection is able to
revert normal CD40-induced mature phenotype in DCs. By contrast, MV
infection did not modified LPS-induced mature phenotype in DCs (data
not shown), suggesting that MV infection specifically blocks
CD40L-mediated DC maturation. Thus, abnormal phenotype induced by MV
infection seems to be dependent on CD40 triggering of DCs rather than
their maturation stage. Functionally, the high CD40L-induced mRNA
synthesis of IL-12 and IL-1/ß were inhibited by MV infection,
whereas IL-10 mRNA synthesis was induced. Thus MV infection inhibits
the CD40-dependent IL-12 secretion by DCs (4) at the
transcriptional level. Consequently, the Th1-type response mediated by
IL-12 secretion of CD40-activated DCs may be inhibited. IL-10 has been
involved in the inhibition of DC maturation and their APC functions
(37). IL-10 mRNA synthesis is also induced by LPS
activation of DCs whether the DCs are CD40 activated or not (data not
shown). Yet, LPS bacterial endotoxin is a potent mediator of DC
maturation. Therefore the presence of IL-10 mRNA, induced by CD40
ligation of MV-infected DCs, may participate to immunosuppression but
cannot account for the various inhibitory effects observed. At a first
sight, DC apoptosis induced by MV infection could explain that
MV-infected DCs are deficient in APC functions, but several features
demonstrate that other mechanisms are involved: 1) cytokine mRNA
synthesis was modified in infected DCs as soon as 24 h after
infection DC, whereas massive apoptosis occurred at day 3 of culture;
2) when DCs were pulsed with UVMV, no apoptosis was observed, yet IL-12
secretion by UVMV-pulsed DCs was 30% decreased and T cell
proliferation was 30% inhibited; 3) though DC apoptosis was delayed 3
days after MV infection, the inhibition of T cell proliferation
occurred at the beginning of the culture (4); and 4)
abnormal DC phenotype in MV-infected CD40-activated DCs was observed on
viable DCs. Thus, MV-induced DC apoptosis certainly reinforces the
inability of DCs to maintain their APC function, but cannot explain the
discrepancies of the APC properties between normal DCs and
MV-infected DCs.
Whatever signal(s) the DCs received to initiate their maturation, the CD40L signal may be needed to achieve their complete maturation and thus generate mature effector DCs. CD40L signaling can be provided by activated T cells in secondary lymphoid organs. Hyper-IgM patients, who are CD40L deficient, display clinical symptoms suggestive of immune deficiency not limited to B cells. In particular, the frequent occurrence of Pneumocystis carenii and Cryptosporidium infections indicate impaired T cell activation (18). As reported in mouse models (21, 22, 23), the lack of CD40 ligation on DCs prevents the generation of CD8+ cytotoxic T cells for certain pathogens. Furthermore, the CD40L signal has been involved in the generation of memory CD8+ CTL (38). The present study demonstrates that 1) the lack of CD40L signal, in vivo, in two hyper-IgM patients, is not strictly correlated with a CD8+ deficiency, but rather with a defective number of CD45RO+ T cells reported as memory T cells. This is confirmed by a recent study where reduced Ag-primed population has been observed in both CD4+ and CD8+ populations, as determined by CD45RO expression (39). 2) CD40L-deficient activated PBL from these patients induce an intermediate stage (CD86- CD80+) of DC phenotypic maturation, in vitro. 3) CD40L activation of DCs is required to activate CD8+ T cells proliferation, in vitro. In the case of MV infection, instead of getting mature effector human DCs able to activate CD8+ T cells proliferation, CD40 triggering of MV-infected DCs prevents CD8+ T cell proliferation.
In DC-PBL cocultures, both CD86 and CD80 expressions were inhibited by the CD40 triggering of MV-infected DCs. CD86 expression was only up-regulated by CD40L+ T cells. In contrast, CD80 up-regulation was also observed using CD40L-deficient T cells. Thus, in the absence of MV infection, other unidentified T cell signals, termed "X," different from CD40L, can up-regulate CD80 expression on DCs. As MV replication abrogated the X pathway when DCs are CD40 triggered, we propose that the negative effect induced by CD40 triggering of MV-infected DCs is dominant and abrogated the X pathway. As a whole, these data suggest that CD40 triggering of MV-infected DCs leads to an inhibitory signal. Indeed, tyrosine-phosphorylation level induced by CD40 activation in DCs is inhibited by MV infection. However, CD40 pathway into DCs have not yet been described. It would be useful to analyze CD40 signaling in DCs and then the relationship between MV infection doses, DC differentiation, DC death, and modification of CD40-signaling in DCs.
The location of DCs identifies them as one of the cell population most likely to have the earliest contact with viruses during infection. DCs have been involved in primary antiviral immune reaction, but also in the propagation of viral infection (reviewed in Ref. 40). Modification of CD40 signaling by MV infection may be the major mechanism which induces MV immunosuppression because: 1) Terminal differentiation of DCs in mature effector DCs is prevented. 2) DCs are used by at least two immunosuppressive viruses as a reservoir that is activated by CD40L and upon interaction with T cells. Indeed, MV replication in DCs is one log enhanced after CD40 activation (4). The same observation was realized for HIV replication (41). Thus the spreading of these immunosuppressive viruses, in vivo, may be linked to CD40 activation of DCs. 3) The requirement of CD40L probably locates the initiation of immunosuppression in T cell area of secondary lymphoid organs where immune response is organized. 4) The impairment of CD40 signaling could also occur in other cell types as macrophages or B cells.
Beyond virus infection, modifying CD40 signaling in DCs could be a powerful tool to modulate immune response.
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Acknowledgments |
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Footnotes |
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e de l’Education Nationale et de la Recherche Technologique, from the Agence Française du Sang (FORTS 96) and by additional support from Association pour la Recherche sur le Cancer (CRC 6108 and SM 9501), Ligue Nationale Contre le Cancer, Programme de Recherche Fondamentale en Microbiologie et Maladies infectieuses et Parasitaires, and Region Rhone-Alpes. 
2 Address correspondence and reprint requests to Prof. Chantal Rabourdin-Combe, Institut National de la Santé et de la Recherche Médicale U503, Immunobiologie Fondamentale et Clinique, Ecole Normale Supérieure de Lyon, 69364 Lyon cedex 07, France. E-mail address:
3 Abbreviations used in this paper: MV, measles virus; CD40L, CD40 ligand; CD40L+-L cells, CD40L-transfected L cells; DC, dendritic cell; LC, Langerhans cell; MHC-I/MHC-II, MHC class I/II; MFI, mean fluorescence intensity; NP, MV nucleoprotein; Mo-DC, monocyte-derived DC; UVMV, UV-inactivated MV; rh, recombinant human.
Received for publication August 9, 1999. Accepted for publication December 2, 1999.
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