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*
Unité de Biologie des Régulations Immunitaires,
Unité d’Histopathologie, and
Unité de Génétique Mycobactérienne, Institut Pasteur, Paris, France
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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) after
administration of live mycobacteria to mice. Experiments were conducted
with Mycobacterium bovis bacillus Calmette-Guerin (BCG)
or a rBCG expressing a reporter Ag. Following infection of mice, DC and
M were purified and the presence of immunogenic peptide/MHC class II
complexes was detected ex vivo on sorted cells, as was the secretion of
IL-12 p40. We show in this study that DC is a host cell for
mycobacteria, and we provide an in vivo detailed picture of the role of
M and DC in the mobilization of immunity during the early stages of
a bacterial infection. Strikingly, BCG bacilli survive but remain
stable in number in the DC leukocyte subset during the first 2 wk of
infection. As Ag presentation by DC is rapidly lost, this suggests that
DC may represent a hidden reservoir for
mycobacteria. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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, to phagocytic cells harboring replicative
intracellular bacteria by increasing their bacteriostatic and
bactericidal functions. In this scheme, macrophages
(M)5 should
represent a privileged APC partner for T cells. Indeed, efficient
uptake of bacteria by M is mediated by various surface molecules
such as FcR, complement receptors, or one of the many receptors for the
glycolipidic components of the bacterial membrane, and is associated
with high bactericidal properties of these cells (1).
Moreover, activated M are a source of IL-12 (2), which
plays a pivotal role in host resistance to bacterial infection by
stimulating IFN- production by NK cells and by promoting Th1
responses (3). However, dendritic cells (DC) represent the
most potent APC for priming naive T cells (4), an
important source of IL-12 following microbial stimuli (5),
and consequently are highly efficient in inducing antiviral and
antitumor immune responses (6). A large body of data
accumulated over the last decade strongly suggests a similar role of DC
in controlling bacterial infection (5, 7, 8, 9, 10, 11), but this
issue is still unresolved. Mycobacteria are facultative intracellular bacteria that can reside and survive in mononuclear phagocytes (12, 13, 14). Interaction of DC with mycobacteria is much less documented, and no information is available in vivo. DC in vitro pulsed with Mycobacterium bovis bacillus Calmette-Guérin (BCG) can efficiently stimulate specific T cells in vivo when injected into mice (7) and also can afford substantial protection against Mycobacterium tuberculosis infection (15). Mycobacteria can signal maturation of DC in vitro, which thus acquire T cell stimulatory activity (16, 17, 18). All this concurs to indicate that DC should play an important role in the elicitation of antibacterial immune responses at early time points post-bacteria delivery, but this question has not yet been directly addressed with live bacteria in vivo.
In the present study, we investigated the infection of DC in vivo
together with APC functions of DC and M during the first 2 wk
following administration of live mycobacteria to mice. Using
Mycobacterium bovis BCG, or a rBCG expressing a reporter Ag,
the Escherichia coli MalE protein (19), DC and
M were purified and up-regulation of APC markers, display of
immunogenic peptides/MHC II complexes, and IL-12 production were
detected ex vivo on DC but not on M. We show that DC is a host cell
for mycobacteria and play an exclusive role in the stimulation of T
cell responses.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Six- to 10-wk-old female BALB/c (H-2d) inbred mice from Janvier (Le Genest Saint-Isle, France) were used.
Bacterial strains and culture conditions
The M. bovis BCG Pasteur vaccine strain 1173 P2 (BCG.wt) was grown in Middlebrook 7H9 (Difco, Paris, France) medium supplemented with 0.1% Tween 80 and albumin dextrose catalase (Difco) or in Middlebrook 7H10 (Difco) solid medium supplemented with oleic acid albumin dextrose catalase (Difco). The E. coli MalE gene was expressed constitutively on a plasmid pIH71 (19). In pIH71, the MalE gene including its signal sequence was fused to the promoter pBlaF*, signal sequence of BlaF. M. bovis BCG Pasteur strain 1173 P2 was transformed with pIH71 by electroporation, and a positive clone was selected and named rBCG(pBlaF*-SSBlaF-SSmalE-malE). In the present study, this rBCG strain was renamed rBCG.MalE. For immunization, BCG strains were first cultivated on Loewenstein-Jensen medium containing 20 µg/ml kanamycin, and then transferred onto Sauton medium.
T cell hybridoma and Ags
BALB/c mice were immunized s.c. with 10 µg of the MalE protein
emulsified in CFA (Sigma-Aldrich, St. Louis, MO). Seven days later,
lymph nodes were removed and a single cell suspension was
prepared and cultured in complete medium (CM) with 10 µg/ml MalE.
Three days later, viable lymphocytes were isolated by fractionation
with Ficoll (Seromed; Biochrom, Berlin, Germany) and fused with BW5147
(
-
-) myeloma cells
as previously described (20). FBU.B11 is specific for
immunodominant p68–82 MalE peptide and MHC II restricted. 45G10 T cell
hybridoma was derived from BALB/c mice and is specific for a poliovirus
(PV) peptide (20). The p68–82 peptide was kindly provided
by J. P. Langeveld (Institute for Animal Science and
Health, Lelystad, The Netherlands); the PV peptide was
synthesized by Neosystem (Strasbourg, France).
mAbs for flow cytometric cell sorting and phenotypic analysis
FITC-, PE-, or allophycocyanin-labeled mAbs specific for
CD11c (HL3), CD11b (M1/70), CD8 (53-6.7), B220 (RA3-6B2), and Gr-1
(RB6-8C5) cell markers were used for FACS sorting. Panels of mAbs were
selected to study the phenotype of APC. We used a combination of
biotinylated or FITC-labeled anti-I-Ad
(AMS-32.1), CD86 (GL-1), CD80 (16-10A1), CD40 (3/23), and
FITC-conjugated streptavidin (Sigma-Aldrich) or PerCP-streptavidin (BD
Biosciences, Mountain View, CA) was used to reveal biotin conjugates.
All labeled Abs were from BD PharMingen (San Diego, CA).
The labeled cells were analyzed on a FACScan or a FACSCalibur for
four-color staining analysis. Files of 50,000–200,000 events were
collected, then subsequently analyzed using CellQuest software (BD
Biosciences).
Cell sorting of spleen cells and ex vivo Ag presentation assay
BALB/c mice were i.v. injected with 0.2–1 x
108 live or heat-killed bacteria and 100 µg of
purified MalE protein in PBS or PBS alone. Spleens were removed and
perfused with collagenase type IV (400 U/ml) containing 50 µg/ml
DNase I (Boehringer Mannheim, Indianapolis, IN). Single spleen cell
suspensions were prepared and separated into low-density cells (LDC)
and high-density cells (HDC) on a dense BSA (Sigma-Aldrich) gradient.
The LDC and HDC populations were further used for staining, sorting,
and FACS analysis or in Ag presentation assays. APC were further
purified from LDC or HDC fractions by sorting on a
FACStarPlus (BD Biosciences) using CD11c, CD8,
CD11b, and B220 cell markers. All APC were 95–99% pure following FACS
analysis. In some experiments, LDC were directly used as an enriched DC
fraction.
Alternatively, DC and M were enriched with an autoMACS (Miltenyi
Biotec, Bergisch Gladbach, Germany). In this case, spleen cells were
first depleted of T cells and neutrophils with anti-Thy1.2 mAb and
anti-Gr-1 mAb (RB6-8C5; kindly given by M.-A. Nahori, Institut
Pasteur, Paris, France) obtained from ascitic fluids and complement
from guinea pig (BioMerieux, Marcy l’Etoile, France). Then
cells were stained with anti-CD11c (N418) Microbeads (Miltenyi
Biotec) before autoMACS separation leading to a positive cell sample of
CD11c+ cells (95–98% pure). The negative
fraction was further incubated with anti-CD11b Microbeads (Miltenyi
Biotec) giving a positive cell fraction containing 80–90%
CD11c-CD11b+ cells.
For ex vivo Ag presentation assay, APC preparations obtained from mice immunized with rBCG.MalE, BCG.wt, or MalE protein were added onto 96-well microplates and serially diluted in CM. A total of 105 T cell hybridomas were added to those APC for 24 h. In some experiments, exogenous protein or peptide was also added to the cell culture containing APC and T hybridoma. Supernatants were harvested, frozen, and tested for IL-2 content by measuring the proliferation of the IL-2-dependent CTLL cell line (results expressed in counts per minute).
Infection assay using labeled fluorescent rBCG.MalE
For labeling, rBCG.MalE was passed through a 25-gauge needle a
few times to break up clumps. Three microliters of component A (Syto-9)
of Live/Dead BacLight Bacterial Viability kit (Molecular Probes,
Eugene, OR) or 5 µl of Calcein AM (Molecular Probes) were added to 1
ml of bacterial suspension to label rBCG.MalE. This suspension was
incubated in the dark at room temperature for 30 min. Bacteria were
then spun down, washed one time, resuspended in PBS, and passed through
a 25-gauge needle to break up clumps of bacteria. Bacteria were checked
under a fluorescent microscope before use. For in vivo infection assay,
mice were i.v. injected with 108 labeled
rBCG.MalE. Then LDC were prepared and stained for FACS analysis of the
infection rate of DC, M, and B cells by labeled rBCG.MalE.
Immunohistochemistry
Mice were injected i.v. with 0.2–1 x 108 CFU of rBCG.MalE or PBS alone, and their spleens were removed and fixed in zinc acetate (0.5%), zinc chloride (0.5%), and calcium acetate (0.05%) in Tris buffer at pH 7 for 48 h. They were then embedded in low-melting point paraffin (37°C) (polyethyleneglycol distearate; Sigma-Aldrich). Paraffin sections (5–6 µm) were deparaffinized in absolute ethanol and air dried. They were incubated in 0.03% H2O2 to neutralize endogen peroxidase activity and, after washing, were incubated in blocking reagent (NEN, Boston, MA) to inhibit nonspecific staining. For immunolabeling of DC, sections were covered for 3 h with a biotinylated anti-CD11c (HL3) in PBS/0.05% saponin. They were incubated with streptavidin-HRP (DAKO, Carpinteria, CA). The reaction was enhanced by incubation in tyramide-biotin (NEN) followed by another incubation in streptavidin-HRP, and peroxidase activity was revealed by amino-ethyl carbazol (Sigma-Aldrich).
Detection of cytokines produced by APC
The different APC were sorted from mice injected i.v. with rBCG.MalE or PBS alone and cultured overnight in CM. The 24-h supernatants were assayed for IL-12 by sandwich ELISA using mAbs C15.6 (p40) or 9A5 (p70) as capture Abs and secondary biotinylated anti-IL-12 mAb C17.8 (BD PharMingen). All assays were standardized with rIL-12 (Genetics Institute, Cambridge, MA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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First, we tracked the presence of BCG bacilli in different
leukocyte subsets of the spleen following i.v. administration to mice.
For this purpose, bacteria were labeled with a green fluorescent dye
(Syto-9) and then used to evaluate by FACS the infection rate of
neutrophils and M
(CD11c-CD11bhigh), DC
(CD11c+), and B cells
(B220+) from the spleen (Fig. 1
A). As could be expected,
2% of the neutrophils/M population was stained with
bacteria-associated dye. However, a similar percentage of the DC
population was also found to be stained by the labeled BCG 4 h
after infection. In contrast, no green-labeled bacteria was found
associated with B cells. These results were confirmed using a rBCG
expressing a green fluorescent protein (our unpublished data).
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B, at day 1,
10% of the viable BCG present in the spleen was found associated with
DC. Interestingly, 12 days after infection, viable BCG was still found
in the DC fraction, but the percentage of CFU found associated with the
DC fraction dramatically dropped. It should be noticed that at this
stage, the number of DC recovered from the spleen of infected mice was
greatly increased (Fig. 1C). The well-documented
hematopoietic effects of BCG (21) probably account for the
abundance of DC at 12 days postinfection. Consequently, the number of
BCG recovered from the total DC population isolated from individual
mice remained unchanged from days 1 to 12. These results suggest that
BCG can survive within the DC pool without apparent growth, although it
cannot be ruled out that they were due to the continuous turnover of
BCG and DC in the spleen. Alternatively, transfer of mycobacteria from
other leukocytes to DC may also occur. In such a case, this may
contribute to a prolonged antimycobacterial immune response.
We next performed immunohistochemical staining of spleen
tissue sections to further characterize the fate of DC.
Compared with control mice, an increased CD11c staining was observed in
the periarteriolar lymphoid sheath (PALS) at 4 and 12 h after
rBCG.MalE infection (Fig. 2, A–C). This result indicates that DC were
redistributed in the T cell zone, suggesting a maturation of DC in
BCG-infected mice. At 12 days postinfection, the central arteriole was
still surrounded by many CD11c+ cells, but their
number was greatly increased in the marginal zone surrounding the
follicle and in the red pulp (Fig. 2D). The presence of BCG
bacilli in DC and M is also shown following purification of
these cells (Fig. 2, E and F). Altogether these
results show that BCG is present in splenic DC as well as in
neutrophils and M during the infection process and that DC
represents a host cell during mycobacterial infection.
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following BCG infection
Because an increased number of DC was observed in the splenic T
cell area following BCG administration, we next analyzed the expression
by these cells of costimulatory molecules required for T cell
activation. Fifteen to 20% of DC showed an up-regulation of CD86 from
4 to 48 h postinfection (Fig. 3A). Up-regulation of CD80 and
CD40 occurred more slowly but was observed on a much larger set of DC
at day 2 postinfection, reaching 50 and 35%, respectively (Fig. 3A). In agreement with the increased number of DC seen in
the PALS (Fig. 2), these phenotypic changes concerned a large set of DC
as compared with the low number of DC infected by the bacilli. This
could indicate that direct interaction with mycobacterial products as
well as bystander effects mediate DC maturation and activation in vivo.
Because DC and M are similarly infected in the spleen, we compared
the phenotypical changes of MHC class II and CD86 (Fig. 3B).
Almost all DC were MHC class II positive (95%) in mice receiving PBS
or BCG, whereas 30 and 25% of M were positive in the PBS and BCG
groups, respectively. CD86 was not detected on M, whereas it was
up-regulated on DC (6 and 21% of positive cells in the PBS and BCG
groups, respectively). These results indicate that DC, but not M,
undergo functional activation/maturation in the early times of
BCG.
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We next investigated the Ag-presenting activity ex vivo by
harvesting spleens from infected mice. Four hours after i.v.
administration of live BCG or rBCG.MalE, we prepared using a BSA
gradient a low density cell fraction enriched in DC from spleens and
monitored their capacity to stimulate the FBU.B11 T hybridoma, specific
for a dominant MalE peptide (Fig. 4A). In these conditions, the
in vivo formation of MalE peptide/MHC complexes on APC from
rBCG.MalE-infected mice, but not from BCG.wt-infected mice, was
detected ex vivo by the T cell hybridoma stimulation. Interestingly,
when mice were i.v. injected with heat-killed rBCG.MalE, direct
stimulation of FBU.B11 was hardly detectable (Fig. 4A),
despite the fact that DC pulsed in vitro with heat-killed rBCG.MalE can
efficiently stimulate FBU.B11 (data not shown). This indicates that
live rBCG.MalE is necessary for efficient Ag presentation by APC in
vivo. We then further investigated the kinetics of Ag-presenting
activity ex vivo by harvesting spleens at various times after i.v.
infection with live rBCG.MalE. The formation of MalE peptide/MHC
complexes was detected at 2, 4, and 12 h after rBCG.MalE infection
but was barely detectable at 48 h (Fig. 4B). When
rBCG.MalE was injected s.c., Ag-presenting activity was detected ex
vivo in draining lymph nodes from days 1 to 2 (data not shown),
confirming the rapid loss of Ag-presenting activity in vivo.
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were found infected with live bacilli, we
assessed the Ag-presenting activity that could be associated with
infection in vivo. Twelve hours after i.v. administration of
rBCG.MalE, DC, M, and B cells were purified by FACS from
spleen and we monitored their capacity to stimulate the FBU.B11 T
hybridoma, specific for a dominant MalE peptide (Fig. 4C).
Ag-presenting activity was only found associated with the DC population
ex vivo. When the different APC were pulsed in vitro with the p68–82
peptide from MalE, all of them were able to stimulate FBUB11 (Fig. 4D). When mice were i.v. injected with soluble MalE protein,
MalE peptide/MHC complexes were detected on DC, M, and B cells (Fig. 4E). In vitro and ex vivo, the same hierarchy of MalE
peptide presentation was observed between the different APC probably
reflecting the level of MHC II expression by these cell types.
Among CD8+ and CD8-
DC, only the CD8- DC subset was suggested to
have phagocytic activity (22). Therefore, we investigated
the capacity of these two subsets to present rBCG.MalE to T cells.
Accordingly, we purified
CD11c+CD8+ and
CD11c+CD8-
subpopulations from rBCG.MalE-infected mice. As shown in Fig. 4F, both DC subsets efficiently stimulated FBU.B11,
demonstrating that MalE peptide/class II complexes were equally formed
in vivo and displayed on these two types of DC. This latter result show
that CD8- as well as
CD8+ DC are able to present
mycobacteria-derived Ags. Altogether, these results show that following
BCG administration, MHC II presentation of mycobacteria-derived
peptides is mainly performed by DC.
Ag presentation by DC is only transient
We then further investigated the kinetics of Ag-presenting
activity during the BCG infection after purification of DC and M.
One week after infection, DC recovered from the spleen were not able to
stimulate FBU.B11, unless peptide was added in vitro, whereas 1 day
following infection DC can directly stimulate FBU.B11 but not the
control poliovirus-specific T hybridoma (Fig. 5A). As hematopoietic events
take place along the infectious process, we checked that recruitment
and production of new DC in the spleen did not dilute the small pool of
Ag-presenting DC. As shown in Fig. 5B, the number of DC
recovered from the spleen of infected mice was stable for the first
48 h, increased 2-fold at 8 days after infection, and increased 5-
to 10-fold 12 days after infection. Therefore, during the first week of
infection, the dilution of the infected DC pool by the arising DC pool
could not account for the loss of in vivo Ag-presenting activity
observed at days 2 and 8 postinfection. Then we asked whether DC became
refractory to presentation for mycobacteria-derived Ag, as DC rapidly
stop MHC II synthesis upon maturation. Mice were first infected
with BCG.wt, and 48 h later they received a second dose
of rBCG.MalE. Four hours after the second injection, the
Ag-presenting activity of DC purified from these mice was compared ex
vivo with DC from mice infected with a single dose of rBCG.MalE and
with DC from mice infected 2 days before with BCG.wt (Fig. 5C). The first round of infection did not shut off the in
vivo Ag-presenting activity of DC, as DC from mice receiving BCG.wt
before rBCG.MalE were even slightly more efficient than DC from mice
having only received rBCG.MalE, clearly showing that the loss of Ag
presentation by DC was not associated with a refractory state.
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-activated M pulsed in vitro
with rBCG.MalE (data not shown), we asked whether M could also
acquire this activity in vivo later during infection. Indeed, IFN-
production by T cells can be already detected a few days after BCG
infection, and a slight increase of MHC II expression was detected on
M 1 wk postinfection (data not shown). After depletion of
contaminating neutrophils, we isolated M by MACS from mice infected
1 and 8 days before and tested their capacity to stimulate FBUB11. No
Ag-presenting activity by these M purified at days 1 and 8
postinfection was detected in this ex vivo assay (Fig. 5A).
Stimulation of FBUB11 was observed when the MalE peptide was added in
vitro, showing that these M were not suppressive for the T cell
readout assay. When M taken from mice infected 8 days before with
rBCG.MalE were titrated with a given dose of PV peptide, they did not
show any increase of stimulatory capacity for the PV-specific T cell
hybridoma (Fig. 5D). In contrast, the same comparison
performed with DC showed a slight increase of MHC II-restricted peptide
presentation. Altogether, these result show that MHC II presentation of
mycobacteria-derived peptides by DC is only transient and does not
exceed the first 2 days. In contrast, MHC II presentation of
mycobacteria-derived peptides by M remains undetectable along the
BCG infection. Characterization of cells producing IL-12 following BCG infection
IL-12 plays a key role both in the control of bacterial infection
and in T cell priming and differentiation. Therefore, we analyzed the
capacity of purified APC from mice infected with rBCG.MalE to produce
IL-12. As shown in Fig. 6A, DC
isolated from the spleen 12 h after in vivo infection produced
IL-12 p40. In contrast, no IL-12 p40 was detected with M and B cells
taken from infected mice (Fig. 6A) or with DC, M, and B
cells isolated from naive mice (data not shown) or from MalE
protein-injected mice. The capacity of DC to produce IL-12 p40 was
further confirmed with DC isolated from draining lymph nodes 24 h
after s.c. administration of rBCG.MalE (our unpublished data).
No bioactive IL-12 p70 (usually produced in amounts 10- to
50-fold less than IL-12 p40 (2)) was detected,
probably reflecting the low number of IL-12-producing cells. Upon
interaction with specific T cells, purified DC from rBCG.MalE-infected
mice produced larger amounts of IL-12 p40 (Fig. 6A). Among
the DC population, both
CD11c+CD8+ and
CD11c+CD8- cells
produced IL-12 in response to rBCG.MalE infection (Fig. 6B).
Interestingly, ex vivo production of IL-12 p40 by spleen DC was only
observed during the first few hours following rBCG.MalE infection but
was not detected later during the course of infection (data
not shown). This latter phenomenon may be due to regulatory events, as
we detected small amounts of IL-10 produced by F4/80 cells 1 wk after
infection (data not shown). This may also correspond to the paralysis
of DC IL-12 production described recently in the Toxoplasma
gondii model (23). Altogether, these results further
confirm that only DC are able to initiate the innate immune response as
well as the primary T cell responses in the early period of BCG
infection.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Effectively, DC are rapidly infected by BCG in vivo and mycobacteria
are still present in the DC population 2 wk after infection, showing
they can survive into these cells. However, it seems that DC do not
permit BCG growth in vivo, suggesting that DC may have bacteriostatic
activity. Because little is known about mycobacteria-DC interaction, in
sharp contrast to what is known about the M host cell, it is not
clear whether the particular properties of the M mycobacterial
phagosome (13, 14) can also take place in DC, allowing
survival of mycobacteria. Recent studies indicate that DC have poor
bactericidal activity leading to M. tuberculosis growth in
human and murine DC in vitro (17, 24). However, the
antimicrobial activity of DC varies depending on their maturity and on
the microenvironment. Thus, mycobacterial growth is inhibited in murine
DC treated with IFN- and LPS (24), and IL-10 can
convert human DC to M with antimycobacterial activity
(16). It remains that these studies show that DC lack
mycobacterial killing activity, which can explain the persistence of
the BCG bacilli within the DC population in vivo. It should be noted
that because Ag presentation by infected DC is rapidly lost, possibly
due to an escape mechanism, DC can also represent a reservoir of
mycobacteria.
Following i.v. administration of live rBCG, DC but not M acquire APC
capabilities for mycobacterial-derived Ags, whereas in vitro studies
showed that both cell types stimulate activated specific T cells
following BCG infection (7). In the spleen, we selected
M expressing CR3, known to mediate phagocytosis of mycobacteria
(25), and we found mycobacteria in this M population
after cell sorting. MHC II expression was low on these cells but
sufficient to detect immunogenic peptide/MHC complexes after i.v.
administration of soluble purified Ag. Although MHC II expression was
slightly increased at late time points post-rBCG inoculation,
presentation of mycobacterial-derived peptides by M was still
undetectable ex vivo. This M population taken from infected mice was
neither suppressive nor altered in its APC functions when fed in vitro
with peptides or proteins. As live mycobacteria interfere with MHC II
presentation in infected M by preventing phagosome acidification
(13) and MHC II transport (26), this most
likely explains the lack of Ag presentation of mycobacteria-derived
peptides by infected M. This phenomenon does not occur in infected
DC, probably because immature DC contain a large intracellular pool of
MHC II molecules (27, 28) ready to be transported at the
membrane once loaded with peptides, which represents a clear advantage
for DC over M in the early times of infection. However, once they
are mature, DC decrease MHC II synthesis (10),
which dampens their capacity to present newly acquired
Ag.
Interestingly, we failed to detect Ag-presenting activity ex vivo on DC
2 days after i.v. infection and along the acute phase of the infection,
suggesting that Ag-specific CD4 T cell stimulation is only transient.
Pancholi et al. (12) showed in vitro that after 2 days of
culture with M. tuberculosis, both M and DC were able to
stimulate CD4 T cells, whereas after 7 days of culture, M, but not
DC, lose this ability. Our data indicate that DC can also be
overwhelmed by mycobacterial infection and that MHC II presentation by
DC is rapidly down-regulated. In vivo, as BCG infection develops, the
number of DC observed in tissue sections is highly increased and they
are widely distributed in all areas of the spleen. Local production and
recruitment of DC from the periphery dilute the infected cell pool,
which remains unchanged in number, and thus may contribute to the loss
of detectable Ag presentation. However, the hematopoietic effects of
BCG are rather limited at 2 days postinfection, a time where MHC II
presentation by DC is almost completely lost. Cell death could explain
the loss of Ag presentation, but in such a case it would be expected
that capture of apoptotic bodies loaded with mycobacteria will lead to
an efficient cross-presentation (29). An alternative will
be that following a first step of infection of immature DC enabling Ag
presentation, DC would die and release mycobacteria that as a second
step will infect mature DC. This possibility is supported by the fact
that in the early phase of infection the number of mature DC is much
larger than the infected DC pool. Because the mature DC have decreased
de novo synthesis of MHC II, presentation of newly acquired
mycobacteria should be much less efficient. However, when we tested
this hypothesis by performing two rounds of infection, the DC were
still efficiently capable to present rBCG.MalE.
The fact that a mycobacteria-derived Ag is efficiently acquired by both
CD8+ and CD8-
subsets of DC for Ag presentation to T cells indicates that there is no
predetermined relationship between one DC subset and intracellular
bacteria. Earlier studies pointed out the poor phagocytic capabilities
of the CD8+ subset (22, 30), but
recent studies seem to indicate that this subset can efficiently
phagocytose (31, 32). Regarding in vivo interaction with
bacteria, our results are consistent with a recent report showing that
bacteria are found in both subsets of DC following infection of mice
with Salmonella dublin (8); however, the two DC
subsets seem to behave differently in terms of cytokine secretion in
Salmonella typhimurium infection (33). Further
studies will determine whether or not the contribution of the different
DC subsets is important for the regulation of the immune response
during mycobacterial infection.
IL-12 plays a pivotal role in the control of mycobacterial
infection (34, 35) because it is involved at the level of
both innate and acquired immunity through IFN- production by NK and
Th1 cells, respectively (2). The capacity of DC to produce
IFN- in vitro in response to IL-12 has also been recently evoked
(36), suggesting that DC may participate directly in the
control of bacterial infections. In many past studies, the production
of IL-12 has been widely attributed to M. However, recent studies
have demonstrated, using in situ detection of IL-12, that DC, but not
M, represent the major source of IL-12, e.g., after injection of
T. gondii extracts (5). Several M.
tuberculosis-derived lipoproteins signal IL-12 production by M
via Toll-like receptor 2 (37), which is also implicated in
M activation by lipoarabinomannans (38, 39). Upon
exposure to BCG, IL-12 production by activated M was shown to be
dependent on IFN-R expression (40), indicating that it
is not an early event of mycobacterial infection. In this study we
demonstrate the superior capacity of DC over M to produce IL-12 p40
following infection of mice with live BCG, but also with heat-killed
BCG (data not shown) during the early phase of infection. In recent
studies it has been pointed out that up-regulation of IL-12 p40 does
not always represents a good indicator for up-regulation of IL-12 p70
in the presence of IL-4 or PGE2 (41, 42). In contrast, up-regulation of IL-12 p70 production was
correlated with IL-12 p40 increase when CD40 was associated with in
vitro soluble tachyzoite Ag stimulation (43). In
the present study, we failed to detect IL-12 p70 in the supernatants of
DC purified from infected animals, probably for sensitivity reasons.
However, IL-12 p70 production by murine and human DC can be induced in
vitro by mycobacteria (17, 24, 44).
In conclusion, we document in this study that DC is the major leukocyte subset involved in the triggering of the immune response to mycobacteria in vivo. In this process, DC activities are only transient and are limited to the early phase of infection, despite the fact that the DC population remains infected for a much longer period of time.
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Acknowledgments |
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Footnotes |
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2 Current address: Laboratory of Immunology, National Institute on Deafness and Other Communication Disorders, National Institutes of Health, Rockville, MD 20850.
3 X.J. and R.L.-M. contributed equally to this work.
4 Address correspondence and reprint requests to Dr. Claude Leclerc, Unité de Biologie des Régulations Immunitaires, Institut Pasteur, 25 rue du Docteur Roux, 75724 Paris Cedex 15, France. E-mail address: cleclerc{at}pasteur.fr
5 Abbreviations used in this paper: M, macrophage; DC, dendritic cell; BCG, bacillus Calmette-Guérin; CM, complete medium; PV, poliovirus; LDC, low-density cell; HDC, high-density cell; PALS, periarteriolar lymphoid sheath.
Received for publication June 8, 2001. Accepted for publication November 27, 2001.
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