By
From the * Laboratory for Immunological Research, Schering-Plough, Dardilly, France; and Immunobiologie Moléculaire, Laboratoire de Biologie Cellulaire et Moléculaire de l'Ecole Normale
Supérieure de Lyon, Lyon, France
Secondary infections due to a marked immunosuppression have long been recognized as a major cause of the high morbidity and mortality rate associated with acute measles. The mechanisms underlying the inhibition of cell-mediated immunity are not clearly understood but dysfunctions of monocytes as antigen-presenting cells (APC) are implicated. In this report, we demonstrate that measles virus (MV) replicates weakly in the resting dendritic cells (DC) as in lipopolysaccharide-activated monocytes, but intensively in CD40-activated DC. The interaction of MV-infected DC with T cells not only induces syncytia formation where MV undergoes massive replication, but also leads to an impairment of DC and T cell function and cell death. CD40-activated DC decrease their capacity to produce interleukin (IL) 12, and T cells are unable to proliferate in response to MV-infected DC stimulation. A massive apoptosis of both DC and T cells is observed in the MV pulsed DC-T cell cocultures. This study suggests that DC represent a major target of MV. The enhanced MV replication during DC-T cell interaction, leading to an IL-12 production decrease and the deletion of DC and T cells, may be the essential mechanism of immunosuppression induced by MV.
In developing countries, measles virus (MV)1 infection
remains a significant cause of mortality accounting for
more than one million deaths per year, especially among
young children. Although malnutrition and an insufficient
supply of medicines may contribute to the severity of measles in the young, it is mainly due to secondary infections
(1). Increased susceptibility to secondary infection correlates with depressed cell-mediated immunity after measles. These immunologic abnormalities were first described by
von Pirquet in 1908 when he noticed a decrease in tuberculin skin reactivity in measles patients (4). The mechanism
of immune suppression is poorly understood but it is widely
assumed that immune suppression is mainly due to MV
replication in leukocytes (5). Infected T cells and monocytes die by apoptosis (6) particularly within syncytia (8).
Syncytia formation is due to the cytopathic effect of MV
(9, 10). The T and B lymphocytes from the PBL of measles patients present a reduced proliferation capacity in response to polyclonal stimulation (11, 12) due to a cell cycle arrest in the end of the G1 phase. In measles, T cells are skewed
from type 1 responses (cell-mediated immunity) towards type
2 responses (antibody-mediated immunity) (13, 14). There
is in vivo and in vitro evidence of a Th2 polarization in cytokine responses during and after measles: production of
IL-4 increases while productions of IL-2 and IFN- This Th2 polarization is suggested to be partly due to the
decreased ability of monocytes to produce TNF- DC are professional APCs, which capture and process
antigens at their immature stage and present antigen to naive T cells to initiate T cell-dependent immune responses
upon their maturation (22, 23). The high capacity of mature
DC to stimulate naive T cells has been attributed to a variety
of factors like the expression of MHC class II molecules,
CD80, CD86, CD40, and diverse adhesion molecules, which
may favor TCR engagement and costimulation (24).
In this study, we used DC, generated by culturing
monocytes in the presence of GM-CSF and IL-4 (28, 29),
to address the following questions: (a) Does MV infect DC
as effectively as monocytes?; (b) Do MV-infected DC
transmit infection to syngenic T cells?; (c) What are the factors which modulate MV replication in the host cells?; and
(d) How does MV infection interfere with the survival and
function of DC and T cells?
Reagents.
Anti-human CD40-ligand (mAb LL2, generated
by Dr. Cees van Kooten at Schering-Plough Laboratory for Immunological Research, Dardilly, France) and anti-CD46 (mAb
20.6, reference 30) were used at 5 and 10 µg/ml final concentrations, respectively. The control antibody mAb 30N (Schering-Plough Laboratory for Immunological Research) was used at 10 µg/ml.
Cell Purifications.
Human peripheral blood was obtained from
the Etablissement de Transfusion Sanguine (Lyon, France).
Mononuclear cells were isolated by density gradient centrifugation using Ficoll/Hypaque, then centrifuged through a 50% Percoll gradient (Pharmacia Fine Chemicals, Uppsala, Sweden) for
20 min at 400 g. The light density fraction from the interface and
the high density fraction from the pellet were recovered and incubated for 10 min at room temperature in 3% human serum-
PBS. Monocytes were purified from the light density fraction by
immunomagnetic depletion (Dynal, Oslo, Norway) using a cocktail of mAbs anti-CD19 (4G7 hybridoma, a gift from Dr. Ron
Levy), CD3 (OKT3; American Type Culture Collection, Rockville, MD), and CD56 (NKH1, Coulter Corp., Miami, FL). The
recovered monocytes were >91% pure as shown by flow cytometry with anti-CD14-FITC (Immunotech, Marseille, France). T
lymphocytes were purified, from the high density fraction, by
immunomagnetic depletion using a cocktail of mAbs anti-CD19,
anti-CD40 (Schering-Plough Research Institute), and anti-CD56
(Coulter), anti-CD14, anti-CD16, anti-HLA-DR, and anti-glycophorin A (Immunotech). After two rounds of depletion, T
lymphocytes were >98% pure as shown by flow cytometry with
anti-CD3-FITC (Immunotech). As assessed by CD1a (Ortho Diagnostic Systems, Inc., Raritan, NJ) staining >95%, DC can be
generated in vitro after 6 d of culture of 5 × 105 monocytes/ml
plus 200 ng/ml GM-CSF and 50 U/ml IL-4 (Schering-Plough Research Institute) in 24-well flat-bottomed microtiter plates (Falcon Labware, Oxnard, CA).
Cell Cultures.
All cultures were performed in 96-well flat-bottomed microtiter plates in a total volume of 200 µl and cultured in RPMI 1640 (Gibco Life Technologies, Inc., Grand
Island, NY) supplemented with 10 mM Hepes (Gibco Life Technologies), 2 mM L-glutamine (Gibco Life Technologies), 40 µg/
ml gentamicin (Schering-Plough, Levallois-Perret, France), and
10% FCS (Gibco Life Technologies). T cells were activated with
a combination of 10 ng/ml PMA (Sigma Chemical Co., St.
Louis, MO) and 1 µg/ml ionomycin (Sigma Chemical Co.) for
6 h and monocytes were activated with 1 µg/ml LPS (Escherichia
coli serotype O127:B8; Sigma Chemical Co.) overnight. After activation, cells were washed three times. Monocytes or DC alone
were cultured at 2 × 105 cells/well. In the T cell cocultures, 105
monocytes or DC per well were cultured together with 105 T
cells/well. In the murine fibroblast cocultures, 2 × 105 monocytes or DC per well were cultured in the presence of 4 × 104 irradiated (7,000 rads) fibroblastic L cells or CD40 ligand-transfected L cells or CD28 ligand-transfected L cells (CD40L+ and
CD28L+ cells; provided by Dr Cees van Kooten, Schering-Plough Laboratory for Immunological Research and Dr. Lewis
Lanier, DNAX Research Institute, Palo Alto, CA, respectively).
10 U/ml M-CSF (Genzyme Corp., Cambridge, MA) were added
to the cultures containing monocytes. 200 ng/ml GM-CSF and
50 U/ml IL-4 were added to the cultures containing DC.
MV Infection and Detection.
Monocytes and DC were infected
with 0.1 PFU/cell of Vero cell-derived MV Hallé (Hallé strain is
classified with the vaccine MV strain Edmonston, see reference
31), either with infectious virus or with neutralized virus by 254 nm ultraviolet rays for 30 min (UV-MV). After a 3-h incubation
at 37°C, the cells were washed four times to be free of unattached
virus. Then infected monocytes or DC were put in culture. Virus
contents of culture supernatants were quantified by limiting dilution from 10 to 10 until 10 IL-12 Titration.
Cell-free culture supernatants were harvested
at various time points. IL-12p70 was measured by a specific
ELISA kit (R&D Systems, Minneapolis, MN), limit of sensitivity
5 pg/ml. Tests were carried out in triplicate and standard deviations are indicated in the figure legends.
Proliferation Assay and Cell and Syncytia Counts.
Cell numbers
were determined with trypan blue dye and syncytia numbers
were counted in situ. DNA synthesis was assessed at different time
points as indicated, after an 8-h pulse with 1 µCi [3H]TdR. Results were expressed as the mean cpm ± SD of triplicate cultures.
FACS® Immunostaining.
After 15 min of permeabilization
with 0.33% Saponin (Sigma Chemical Co.), cells were stained with
anti-nucleoprotein (NP) viral protein provided by F. Wild (Institut Pasteur de Lyon, France; reference 32), followed by incubation with PE-labeled anti-mouse Ig (Immunotech). The ApopTagTM
in situ apoptosis detection kit (S7110-KIT; Oncor Inc., Gaithersburg, MD) was used to detect apoptotic cells by fluorescence detection of digoxigenin-labeled genomic DNA. Anti-CD3, anti-CD14, and anti-DR-FITC-labeled mAbs used to verify the purity
of T, monocytes, and DC, respectively, were from Immunotech.
Immunoenzymatic Staining.
Cytospin preparations of 7 × 105
cells were fixed in acetone for 10 min at 4°C. The slides were
washed in PBS. Double staining was done using mouse IgG1
anti-CD3, anti-CD13, or anti-HLA-DR (Immunotech) and
mouse IgG2a anti-NP. Mouse IgG1 antibodies were revealed by
sheep anti-mouse IgG1 (The Binding Site, Birmingham, UK)
followed by mouse antialkaline phosphatase-alkaline phosphatase complexes (APAAP technique; Dako S.A., Trappes, France). NP
staining was revealed by sheep anti-mouse IgG2a-biotin (The
Binding Site) followed by ExtrAvidin peroxidase (Sigma Chemical Co.). Alkaline phosphatase activity was first developed by Fast
blue substrate; peroxidase activity was developed with AEC
(Sigma Chemical Co.) substrate. Two slides were incubated with
mouse IgG1 and IgG2a isotype control mAbs (Dako S.A.). All of
the slides were washed in tap water before mounting in glycerol
(Dako S.A.).
Both monocytes isolated from PBL and monocyte-derived
DC have been infected by the MV at 0.1 PFU/cell. According to the kinetics of viral particle productions, measured by the number of PFU (Fig. 1), the level of MV replication was equivalent in both populations and reached a
maximum level at day 5. By immunochemistry on cytospins, the detection of MV-NP (which is the earliest viral
protein produced) in both populations confirmed that MV
could replicate in monocytes as well as in DC (Fig. 2, A
and B) in vitro. The wild-type strain of MV, replicated as
well as vaccinal strain (Hallé) (data not shown). We noticed
the presence of syncytia only on the cytospins of MV-infected
DC (Fig. 2 B) but not in MV-infected monocytes. Syncytia
formation in DC cultures depended on virus replication, since no syncytia were observed in DC pulsed with UV-MV (Fig. 2 A).
To analyze how
MV replication was affected during interactions between
monocytes and T cells or DC and T cells, monocytes or
DC were infected with MV, then cocultured for up to 7 d
with syngenic T cells activated with PMA plus ionomycin.
PFU was measured at day 5 of the cocultures (the peak
time point of viral production). In the presence of T cells,
the viral particle productions were greatly increased in both
cultures (Fig. 3 A). While viral production in monocytes
increased fourfold, viral production in DC increased 18-fold in the T cell cocultures.
To identify the signal(s) from activated T cells promoting measles replication in these cocultures, we used an anti-CD40L antibody which blocks CD40-CD40L interactions.
Infected monocytes or DC were cultured with activated T
cells plus anti-CD40L or with an irrelevant antibody (Fig. 3
A). At day 5, anti-CD40L blocked 38 and 45% of PFU
productions within monocyte-T cell and DC-T cell cocultures, respectively. Although increased doses of anti-CD40L antibody failed to completely inhibit the T cell-
dependent increases of viral production in these cocultures,
CD40L+ L cells could replace activated T cells in promoting PFU production in DC or monocyte cocultures (Fig.
3, B and C). This suggested that the anti-CD40L antibody
used might recognize one functional epitope of CD40L.
Alternatively, other T cell-derived signals might be involved. However, CD28+ L cells were not shown to promote
MV production in DC or monocytes (data not shown).
The presence of activated T cells or CD40L+ L cells increased both the size and the number of multinuclear syncytia in MV-infected DC cocultures (Fig. 2, G and H).
The number of syncytia in DC-T cell or DC-CD40L+ L
cell cocultures was 7-15-fold more than in DC cultured
alone. The formation of syncytia in DC-T cell cocultures
was partially inhibited by the anti-CD40L antibody (32-
43% of inhibition, data not shown). All syncytia contained
viral particles as demonstrated by immunohistological staining of NP antigen (Fig. 2, B, D, and F). The infected DC
displayed poor DC morphology and viability (compare Fig.
2, D and F with C and E).
To de-
termine if DC transmit infection to T cells a double staining (anti-NP and anti-CD3) on cytospins of cocultured
APCs-T cells was performed. We observed that in cocultures with DC or monocytes, only a few T cells contained
NP, in contrast to many DC or monocytes (data not
shown). To quantify the percentage of each infected cell
type in these cocultures, double immunofluorescence stainings with anti-NP and anti-HLA-DR or anti-NP and anti-CD3 were performed on DC-T cell cocultures and the results were analyzed by flow cytometry (Fig. 4 A). At day 3 of coculture, 38% (38-44%, n = 5) of DC and 9% (8.5-
10%, n = 5) of T cells expressed NP. Similar observations
were obtained from monocyte-T cell cocultures, where
40% (36-43%, n = 5) of monocytes versus 8% (8-10%,
n = 5) of T cells express NP at day 3 of coculture. At the
end of both cocultures, only 5-10% of the activated T cells
expressed NP, whereas 45% of DC and 40% of monocytes
were shown to express NP at day 5 of coculture (Fig. 4 B).
In MV-infected APC-T cell cocultures, we noticed a
precocious and significant decrease of viable cells, from 106
cells at the beginning of the cocultures to <104 at the end
(only DC used as APCs are shown in Fig. 5 A). By contrast, the UV-MV weakly affected cell viability in both
monocytes or DC-T cell cocultures, with the number of
cells remaining nearly the same as that of uninfected APC-T
cell cocultures.
The decreased cell viability in MV-infected APC-T cell
cocultures correlated with the absence of thymidine incorporation by T cells even at day 1 of coculture, when most
of the cells were still alive (Fig. 5 B). UV-MV partially inhibited T cell proliferation (30%) induced by DC.
To understand mechanisms by which MV decreased viable cell number and thymidine incorporation of T cells, we
looked at apoptosis by the TUNEL technique. In MV-
infected monocytes or DC cultures, ~45% of cells underwent apoptosis at the end of the cultures, whereas in the
cocultures with activated T cells, the percentage of apoptotic APCs increased rapidly to reach 50% of cells at day 1 and 90% at day 7 (only the data using DC as APCs are
shown in Fig. 6 A). This apoptosis was directly linked to
the infection of the cells, as no apoptotic cells were detected in APCs pulsed with UV-MV. Noticeably, T cell
apoptosis reached 65% at day 1 and 95% before the end of
the coculture (only the results in DC-T cell coculture are
shown in Fig. 6 B). This death was largely due to the viral
replication, as the apoptosis background of uninfected activated T cells cultured alone or with UV-inactivated MV
pulsed APCs was <20%. In contrast to the ability of UV-MV
to partially inhibit thymidine incorporation by T cells, UV-MV did not induce apoptosis of either DC or T cells. All
these data suggest that UV-MV can induce cycle arrest of T
cells without inducing apoptosis.
By double staining of NP-expressing cells and of apoptotic cells, we further showed that of 98% of apoptotic T
cells, only 6% had detectable NP expression (Fig. 6 C). In
addition, 34 out of 82% of apoptotic APCs had detectable
NP expression. In contrast, in the absence of T cells all apoptotic DC and apoptotic monocytes expressed NP.
To determine if MV infection of human
DC specifically downregulated IL-12 production as previously described in the case of MV-infected human monocytes (15), the kinetics of IL-12 production was measured
by ELISA in culture supernatants of infected or noninfected DC. IL-12 production was detectable in culture supernatants from DC cocultured with either activated T
cells (Fig. 7 A) or CD40L+ L cells (Fig. 7 B). As anti-CD40L antibody mostly abrogated IL-12-induced production, the essential role of CD40 triggering for IL-12 production by DC was confirmed (33). In these culture conditions,
monocytes did not produce detectable IL-12.
The IL-12 production of DC induced by activated T
cells or CD40L+ L cells was downregulated by adding MV
(77% of the total production at day 3, 65% at day 5, 63% at
day 7) and by adding UV-MV (23% at day 3, 30% at day 5, 30% at day 7) (Fig. 7, A and B). Interestingly, UV-MV,
which did not induce apoptotic cell death, inhibited IL-12 production by 20-30%, suggesting that the inhibition of
IL-12 production was not merely due to cell death.
We have demonstrated the permissivity of DC to MV
infection in vitro. Whereas the level of MV replication in
DC is low, MV replication can be boosted by a DC-T cell
interaction. Thus, DC may capture MV at its site of entry
in mucosa surfaces, then migrate to the T cell areas of lymphoid organs, where DC present immunogenic peptides to
naive T lymphocytes. This DC-T cell interaction allows MV to undergo a massive replication, notably in syncytia.
One of the key molecules which promotes MV replication
was shown to be CD40 ligand, an important molecule for
DC activation and maturation (29, 34, 35). The massive
MV replication mainly occurred in CD40-activated DC,
but not in T cells. However, T cell proliferation function was
profoundly affected. The proliferation of T cells was more
inhibited with the infectious MV than with the UV-MV. Interestingly, our data support the previous observations
demonstrating that the inhibition of T cell proliferation was
not the direct result of MV infection of T cells, but was
due to other cell types being infected by MV within the
blood (36, 37). Our study suggests that DC in the MV-
infected autologous PBL may be responsible for the induction of the arrest of T cell proliferation. T cells as well as DC
were induced to undergo apoptosis by MV directly (by cell
infection) or indirectly (without detectable cell infection). Previous studies have also shown the immunological unresponsiveness and apoptotic cell death of T cells in MV infection (7). This induction of apoptosis may account for the
mechanisms underlying the immunosuppression induced in
MV patients (11, 12, 38, 39), but cannot fully explain the
impairment of T cell proliferation activity, as UV-MV induces an inhibition of T cell proliferation without inducing
apoptosis. It has been proposed that the mechanism of
MV-induced immunosuppression may be due to the interaction of uninfected lymphocytes with MV hemagglutinin
and/or fusion proteins expressed on the surface of an infected APC which then lead APCs to deliver a negative
transmembrane signal to the responding T cells, which
then arrests their proliferation. The ligand(s) on the uninfected T cells are unknown. We propose that members of
the TNF family (for review see reference 40) may be upregulated on DC after MV infection, which may induce a
paracrine-killing of T cells and an autocrine-killing of DC.
Moreover, since apoptosis was observed in all MV-infected
DC, upregulation of apoptosis inducing molecules also occurs in the absence of T cells. A small percentage of DC
and T cells seems to be resistant to MV infection and to
MV-induced apoptosis in vitro. These refractory cells may account for the recovery from measles infection. MV, both
infectious and UV-inactivated, was shown to inhibit IL-12
production by CD40-activated DC. This may explain why
decreases in cellular Th1 responses and increases in humoral Th2 responses in measles patients, often insufficient
to defeat a viral infection, are observed.
Previous studies have shown that monocytes play a key
role in MV infection and in the decrease of cell-mediated
immune responses observed during measles. In this paper,
we confirm the susceptibility of monocytes to MV infection, and demonstrate the higher capacity of MV replication in CD40-activated DC. Syncytia formation could be
detected in MV-infected DC or in MV-infected CD40-
activated DC, but not in MV-infected monocytes or in
MV-infected CD40-activated monocytes. This suggests
that expression of MV hemagglutinin and/or fusion proteins as specific MV-receptor CD46 (41) on monocytes was not sufficient to get fusion between cells. DC might
express protein(s), which is (are) absent from monocytes,
but essential for syncytia formation. During acute infection,
there is evidence that the cytopathic virus induces syncytia
in lung and lymphoid tissues such as thymus, tonsils, lymph
nodes, and spleen (9, 44). Thus, DC in these tissues might
be the privileged target for the cytopathic effects of MV infection. MV infection was shown to downregulate IL-12
production by monocytes induced by LPS and IFN- MV replication in DC-T cell cocultures shared many
features with HIV replication in DC-T cell cocultures: (a)
only a small amount of HIV is needed to infect DC; (b) low
levels of infected- DC can initiate extensive HIV-1 replication in cocultures with memory T cells, or activated T cells
(45); (c) CD40L expressed on activated T cells may
contribute to the DC-dependent HIV replication (47).
However, CD28 on T cells was shown to play a role in the DC-dependent HIV replication, and in our experiments
MV replication by DC was not modulated by CD28 ligation. (d) The conjugation of DC and T cell types leads to
syncytia formation which may facilitate viral replication
(45, 46, 48). More recently, the mechanism by which syncytia promotes HIV replication has been suggested: a syncytium formation allows active NF- The progression to AIDS seems due to the impairment
of the immunological microenvironment to maintain the
capacity for renewal of a balanced, competent Th -cell
population. In this context, it will be extremely important
to understand the molecular and cellular basis of how MV
patients recover from initial immunosuppression occurring
both at the DC and T cell levels.
decrease (14).
and
IL-12 (15, 16). Earlier studies have suggested that the immunodeficiency associated with MV infection could be related in part to the dysfunction of APCs such as monocytes
or B cells (11, 17, 18). Nevertheless, the role played by
dendritic cells (DC), which are the sole professional APCs
able to prime naive T lymphocytes (19), has not yet
been studied in measles.
10 on confluent Vero cells. A single
plaque in the Vero cells confluent culture represents one PFU
generated by an individual infectious virus.
MV Replicates in DC and Induces Syncytia Formation.
Fig. 1.
MV can replicate in DC as well as in monocytes. Kinetics of
viral particle productions by 106 monocytes or by 106 DC pulsed with
UV-MV or infected with MV. Infectious virus produced in the culture
supernatants was quantified by the number of PFU on Vero cell lawn.
Results are representative of six experiments. SD were <10% of variation.
[View Larger Version of this Image (17K GIF file)]
Fig. 2.
Immunochemical and morphological analyses of DC or DC-T cell cocultured with MV or UV-MV (A and B ). At day 3, double stainings on
UV-MV pulsed DC (A) and on MV-infected DC (B) were performed with anti-NP (red) and anti-HLA-DR (blue). NP can be localized in the nuclei,
but is strongly expressed in the cytoplasm where the virus replicates, particularly in syncytia. (C-F). At day 3, anti-NP (red) and anti-HLA-DR (blue)
stainings were performed on UV-MV DC-T cells cocultured (C and E) and on MV DC-T cells cocultured (D and F). (G and H) On CD40L+ L cells,
syncytia were observed only with MV DC (H) and not with UV-MV DC (G). Original magnifications, A and B ×400; C and D ×200; E and F ×1000;
G and H ×100.
[View Larger Version of this Image (91K GIF file)]
Fig. 3.
CD40-CD40L interaction induces a burst of MV
production in MV DC-T cell
cocultures. (A) Kinetics of PFU
of 106 MV monocytes or MV
DC without or with T cells, plus
anti-CD40L mAb or antibody
control. Results are representative of three experiments. SD
was <15%. (B and C) Kinetics of
MV productions by MV monocytes (B) or MV DC (C) in the
presence of CD40L+ L cells,
CD28L+, L cells or T cells. Results are representative of five experiments. SD was <15% of
variation.
[View Larger Version of this Image (32K GIF file)]
Fig. 4.
Percentages of NP+
cells in MV DC-T cell cocultures. (A) At day 3, FACS® analysis of NP+ DC and NP+ T cells
by FITC-anti-HLA-DR/PE-anti-NP and by FITC-anti-CD3/ PE-anti-NP stainings, respectively. Quad limits were positioned on the negative control
which showed a higher autofluorescence in DC than in T cells.
(B) Summary of the FACS®
analysis of the NP+ cells in MV
APC-T cell cocultured up to 5 d.
Results are representative of five
experiments.
[View Larger Version of this Image (38K GIF file)]
Fig. 5.
Number of viable
cells and thymidine incorporation by T cells in DC-T cell
cocultures without or with UV-MV or MV. (A) Kinetic of cell
viability in DC-T cell cocultures. DC have been uninfected, pulsed with UV-MV, or infected
with MV DC. (B) Kinetic of
thymidine incorporation by T
cells cultured alone or with uninfected DC, UV-MV DC, or
MV DC. Results are representative of three experiments. SD
was <10%.
[View Larger Version of this Image (21K GIF file)]
Fig. 6.
FACS® TUNEL
analysis of DC and DC-T cell
cocultures pulsed with UV-MV
or MV. (A) Kinetics of apoptotic
DC from UV-MV DC and MV
DC cultures with or without T
cells. Cells were stained with FITC-antidigoxigenin and DC
were gated according to standard
forward- and side-scatter values.
(B) Kinetics of apoptotic T cells
from DC-T cell cocultures,
pulsed with UV-MV or MV.
Cells were stained with FITC-
antidigoxigenin and T cells were
gated according to standard forward- and side-scatter values.
Results are representative of
three experiments. SD was
<10%. (C) Apoptotic and NP+
cell analysis from MV DC-T cell
cocultures. Cells were stained with anti-NP followed by PE-anti-mouse and FITC-antidigoxigenin. DC and T cells
were gated according to standard
forward- and side-scatter values.
The numbers in each quadrant
represent the percentages of
gated DC or T cells. Quad limits
were positioned on the negative control (not shown). Results are
representative of five experiments.
[View Larger Version of this Image (34K GIF file)]
Fig. 7.
Kinetics of IL-12
production by CD40-activated
DC. IL-12 production of DC
supernatants has been measured
by specific ELISA at various time
points. (A) In cocultures with T
cells, DC were uninfected,
pulsed with UV-MV, or infected
with MV in the presence or absence of mAb anti-CD40L. (B)
Uninfected DC, UV-MV DC,
or MV DC were cultured with
CD40L+ L cells. Results are representative of six experiments.
SD was <10%.
[View Larger Version of this Image (26K GIF file)]
(15).
Here we showed that MV also downregulates IL-12 production by DC induced CD40-ligand. This represents a
key factor contributing to the Th2 polarization observed in
MV-infected patients.
B from the dendritic
cells to conjugate with Sp-1 from the T cells, this combination being possibly involved in the acceleration of viral replication within the heterokaryon (49). It will be interesting
to investigate if similar mechanisms exist in the MV-
induced syncytia. (e) HIV-1-infected monocytes promote
uninfected T cells to undergo apoptosis in the presence of
antigen stimulation (50). (f ) Finally, both HIV and MV
could induce T cell anergy, deletion, and functional polarization to Th2 (for review see reference 51).
Address correspondence to Professor Chantal Rabourdin-Combe, Immunobiologie Moléculaire, Laboratoire de Biologie cellulaire et Moléculaire de l'Ecole Normale Supérieure de Lyon, Unité mixte de Recherche UMR49, C.N.R.S./E.N.S./laboratoire associé I.N.R.A., 69364 Lyon cedex 07, France. Phone: 33-4-72-72-80-16; FAX: 33-4-72-72-86-86; E-mail: Chantal.Rabourdin-Combe{at}ens-lyon.fr
Received for publication 23 April 1997 and in revised form 11 July 1997.
1 Abbreviations used in this paper: DC, dendritic cells; L, ligand; L+, ligand-transfected; MV, measles virus; NP, nucleoprotein; UV-MV, ultraviolet-inactivated measles virus.We thank Dr. J. Chiller for his support; Dr. E. Bates, Dr. F. Fossiez, Dr. C. Arpin, and O. de Bouteiller for critical reading of the manuscript; Mrs. I. Durand for FACS® settings; and Mrs. S. Bonnet-Arnaud and Mrs. M. Vatan for editorial assistance.
This work was supported by Schering-Plough, by institutional grants from the Centre National de la Recherche Scientifique, and from Ministère de l'Education Nationale, de l'Enseignement supérieur et de la Recherche and by additional supports from Association pour la Recherche sur le Cancer (CRC 6108) and Ligue Nationale Contre le Cancer (CRC).
1. | Beckford, A.P., R.O.C. Kaschula, and C. Stephen. 1985. Factors associated with fatal cases of measles: a retrospective autopsy study. S. Afr. Med. J. 68: 858-863 [Medline]. |
2. | Coovadia, H.M., A. Wesley, and P. Brain. 1978. Immunologic events in acute measles influencing outcome. Arch. Dis. Child. 53: 861-867 [Abstract]. |
3. | Tamashiro, V.G., H.H. Perez, and D.E. Griffin. 1987. Prospective study of the magnitude and duration of changes in tuberculin reactivity during complicated and uncomplicated measles. Pediatr. Infect. Dis. J. 6: 451-454 [Medline]. |
4. | Von Pirquet, C.P.. 1908. Das Verhalten der kutanen tuberculin Reaktion Während der Masern. Dtsch. Med. Wochenschr. 34: 1297-1300 . |
5. | Joseph, B.S., P.W. Lampert, and M.B.A. Oldstone. 1975. Replication and persistence of measles virus in defined subpopulations of human leukocytes. J. Virol. 16: 1638-1649 [Medline]. |
6. | Auwaerter, P.G., H. Kaneshima, J.M. McCune, G. Wiegand, and D.E. Griffin. 1996. Measles virus infection of thymic epithelium in the SCID-hu mouse leads to thymocyte apoptosis. J. Virol. 70: 3734-3740 [Abstract]. |
7. | Addae, M.M., Y. Komada, X.L. Zhang, and M. Sakurai. 1995. Immunological unresponsiveness and apoptotic cell death of T cells in measles virus infection. Acta Paediatr. Jpn. 1995: 308-314 . |
8. | Esolen, L.M., S.W. Park, M. Hardwick, and D.E. Griffin. 1995. Apoptosis as a cause of death in measles virus-infected cells. J. Virol. 69: 3955-3957 [Abstract]. |
9. | Enders, J.F., and T.C. Peebles. 1954. Propagation in tissue cultures of cytopathogenic agents from patients with measles. Proc. Soc. Exp. Biol. Med. 86: 277-286 . |
10. | Heneen, W.K., W.W. Nichols, A. Levan, and E. Norrby. 1966. Studies on syncytia formation in a cell line (LU 106) of human origin after treatment with measles virus. Hereditas (Lund.). 57: 369-372 . |
11. | McChesney, M.B., J.H. Kehrl, A.S. Valsamakis, A.S. Fauci, and M.B.A. Oldstone. 1987. Measles virus infection of B lymphocytes permits cellular activation but blocks progression through the cell cycle. J. Virol. 61: 3441-3447 [Medline]. |
12. |
McChesney, M.B.,
A. Altman, and
M.B.A. Oldstone.
1988.
Suppression of T lymphocyte function by measles virus is due
to cell cycle arrest in G1.
J. Immunol.
140:
1269-1273
|
13. | Ward, B.J., R.T. Johnson, A. Vaisberg, A. Jauregui, and D.E. Griggin. 1990. Cytokine production in vitro and the lymphoproliferative defect of natural measles infection. Clin. Immunol. Immunopathol. 55: 315-326 [Medline]. |
14. | Ward, B.J., and D.E. Griffin. 1993. Differential CD4 T-cell activation in measles. J. Infect. Dis. 168: 275-281 [Medline]. |
15. | Karp, C.L., M. Wysocka, L.M. Wahl, J.M. Ahearn, P.J. Cuomo, B. Sherry, G. Trinchieri, and D.E. Griffin. 1996. Mechanism of suppression of cell-mediated immunity by measles virus. Science (Wash. DC). 273: 228-231 [Abstract]. |
16. |
Leopardi, R.,
R. Vainionpää,
M. Hurme,
P. Siljander, and
A.A. Salmi.
1992.
Measles virus infection enhances IL-1 ![]() ![]() |
17. | Esolen, L.M., B.J. Ward, T.R. Moench, and D.E. Griffin. 1993. Infection of monocytes during measles. J. Infect. Dis. 168: 47-52 [Medline]. |
18. | Salonen, R., J. Ilonen, and A. Salmi. 1988. Measles virus infection of unstimulated mononuclear cells in vitro: antigen expression and virus production preferentially in monocytes. Clin. Exp. Immunol. 71: 224-228 [Medline]. |
19. | Flechner, E.R., P.S. Freudenthal, G. Kaplan, and R.M. Steinman. 1988. Antigen-specific T lymphocytes efficiently cluster with dendritic cells in the human primary mixed-leukocyte reaction. Cell. Immunol. 111: 183-195 [Medline]. |
20. | Freudenthal, P.S., and R.M. Steinman. 1990. The distinct surface of human blood dendritic cells, as observed after an improved isolation method. Proc. Natl. Acad. Sci. USA. 87: 7698-7702 [Abstract]. |
21. | Bhardwaj, N., S.M. Friedman, B.C. Cole, and A.J. Nisanian. 1992. Dendritic cells are potent antigen-presenting cells for microbial superantigens. J. Exp. Med. 175: 267-273 [Abstract]. |
22. | Steinman, R.M.. 1991. The dendritic cell system and its role in immunogenicity. Annu. Rev. Immunol. 9: 271-296 [Medline]. |
23. | Austyn, J.M.. 1996. New insights into the mobilization and phagocytic activity of dendritic cells. J. Exp. Med. 183: 1287-1292 [Medline]. |
24. | Inaba, K., M. Witmer-Pack, M. Inaba, K.S. Hathcock, H. Sakuta, M. Azuma, H. Yagita, K. Okumura, P.S. Linsley, S. Ikehara, et al . 1994. The tissue distribution of the B7-2 costimulator in mice: abundant expression on dendritic cells in situ and during maturation in vitro. J. Exp. Med. 180: 1849-1860 [Abstract]. |
25. | Caux, C., and J. Banchereau. 1996. In vitro regulation of dendritic cell development and function. Blood Cell Biochem. 7: 263-301 . |
26. | Cella, M., F. Sallusto, and A. Lanzavecchia. 1997. Origin, maturation and antigen presenting function of dendritic cells. Curr. Opin. Immunol. 9: 10-16 [Medline]. |
27. | Springer, T.A.. 1990. Adhesion receptors of the immune system. Nature (Lond.). 346: 425-434 [Medline]. |
28. | Romani, N., S. Gruner, D. Brang, E. Kampgen, A. Lenz, B. Trockenbacher, G. Konwalinka, P.O. Fritsch, R.M. Steinman, and G. Schuler. 1994. Proliferating dendritic cell progenitors in human blood. J. Exp. Med. 180: 83-93 [Abstract]. |
29. |
Sallusto, F., and
A. Lanzavecchia.
1994.
Efficient presentation
of soluble antigen by cultured human dendritic cells is maintained by granulocyte/macrophage colony-stimulating factor
plus interleukin 4 and downregulated by tumor necrosis factor-![]() |
30. | Naniche, D., T.F. Wild, C. Rabourdin-Combe, and D. Gerlier. 1992. A monoclonal antibody recognizes a human cell surface glycoprotein involved in measles virus binding. J. Gen. Virol. 73: 2617-2624 [Abstract]. |
31. | Rima, B.K., J.A. Earle, R.P. Yeo, L. Herlihy, K. Baczko, V. Ter, Meulen, J. Carabana, M. Caballero, M.L. Celma, and R. Fernandez-Munoz. 1995. Temporal and geographical distribution of measles virus genotypes. J. Gen. Virol. 76: 1173-1180 [Abstract]. |
32. | Giraudon, P., and T.F. Wild. 1981. Monoclonal antibodies against measles virus. J. Gen. Virol. 54: 325-332 [Abstract]. |
33. | Kato, T., R. Hakamada, H. Yamane, and H. Nariuchi. 1996. Induction of IL-12 p40 messenger RNA expression and IL-12 production of macrophages via CD40-CD40 ligand interaction. J. Immunol. 156: 3932-3938 [Abstract]. |
34. | Alderson, M.R., R.J. Armitage, T.W. Tough, L. Strockbine, W.C. Fanslow, and M.K. Spriggs. 1993. CD40 expression by human monocytes: regulation by cytokines and activation of monocytes by the ligand for CD40. J. Exp. Med. 178: 669-674 [Abstract]. |
35. | Van Kooten, C., and J. Banchereau. 1997. Immune regulation by CD40-CD40-L interactions. Front. Biosci. 2: 358-368 . |
36. |
Schlender, J.,
J.J. Schnorr,
P. Spielhofer,
T. Cathomen,
R. Cattaneo,
M.A. Billeter,
V. Ter,
Meulen, and
S. Schneider-Schaulies.
1996.
Interaction of measles virus glycoproteins
with the surface of uninfected peripheral blood lymphocytes
induces immunosuppression in vitro.
Proc. Natl. Acad. Sci.
USA.
93:
13194-13199
|
37. | Sanchez-Lanier, M., P. Guerin, L.C. McLaren, and A.D. Bankhurst. 1988. Measles virus-induced suppression of lymphocyte proliferation. Cell. Immunol. 116: 367-381 [Medline]. |
38. | McChesney, M.B., R.S. Fuginami, N.W. Lerchae, P.A. Marx, and M.B.A. Oldstone. 1989. Virus induced immunosuppression: infection of peripheral blood mononuclear cells and suppression of immunoglobulin synthesis during natural measles virus infection of rhesus monkeys. J. Infect. Dis. 159: 757-760 [Medline]. |
39. | Yanagi, Y., B.A. Cubitt, and M.B.A. Oldstone. 1992. Measles virus inhibits mitogen-induced T cell proliferation but does not directly perturb the T cell activation process inside the cell. Virology. 187: 280-289 [Medline]. |
40. | Nagata, S.. 1997. Apoptosis by death factor. Cell. 88: 355-365 [Medline]. |
41. | Naniche, D.. 1993. Human membrane cofactor protein (CD46) acts as a cellular receptor for measles virus. J. Virol. 67: 6025-6032 [Abstract]. |
42. | Manchester, M., M.K. Liszewski, J.P. Atkinson, and M.B.A. Oldstone. 1994. Multiple isoforms of CD46 (membrane cofactor proteins) serve as receptors for measles virus. Proc. Natl. Acad. Sci. USA. 1994: 2161-2165 . |
43. | Dorig, R., A. Marcel, A. Chopra, and C.D. Richardson. 1993. The human CD46 molecule is a receptor for measles virus (Edmonston strain). Cell. 75: 295-305 [Medline]. |
44. | White, R., and J. Boyd. 1973. The effect of measles on the thymus and other lymphoid tissues. Clin. Exp. Immunol. 13: 343-357 [Medline]. |
45. | Pope, M., S. Gezelter, N. Gallo, L. Hoffman, and R.M. Steinman. 1995. Low levels of HIV-1 infection in cutaneous dendritic cells promote extensive viral replication upon binding to memory CD4+ T cells. J. Exp. Med. 182: 2045-2056 [Abstract]. |
46. | Cameron, P.U., P.S. Freudenthal, J.M. Barker, S. Gezelter, K. Inaba, and R.M. Steinman. 1992. Dendritic cells exposed to human immunodeficiency virus type-1 transmit a vigorous cytopathic infection to CD4+ T cells. Science (Wash. DC). 257: 383-387 [Medline]. |
47. | Pinchuk, L.M., P.S. Polacino, M.B. Agy, S.J. Klaus, and E.A. Clark. 1994. The role of CD40 and CD80 accessory cell molecules in dendritic cell-dependent HIV-1 infection. Immunity. 1: 317-325 [Medline]. |
48. | Frankel, S.S., B.M. Wenig, A.P. Burke, P. Mannan, L.D.R. Thompson, S.L. Abbondanzo, A.M. Nelson, M. Pope, and R.M. Steinman. 1996. Replication of HIV-1 in dendritic cell-derived syncytia at the mucosal surface of the adenoid. Science (Wash. DC). 272: 115-117 [Abstract]. |
49. | Granelli-Piperno, A., M. Pope, K. Inaba, and R.M. Steinman. 1995. Coexpression of REL and SP1 transcription factors in HIV-1 induced, dendritic cell-T cell syncytia. Proc. Natl. Acad. Sci. USA. 92: 10944-10948 [Abstract]. |
50. |
Cottrez, F.,
F. Manca,
A.G. Dalgleish,
F. Arenzana-Seisdedos,
A. Capron, and
H. Groux.
1997.
Priming of human
CD4+ antigen-specific T cells to undergo apoptosis by HIV-infected monocytes.
J. Clin. Invest.
99:
257-266
|
51. | Heeney, J.L.. 1995. AIDS: a disease of impaired Th-cell renewal? Immunol. Today. 16: 515-523 [Medline]. |