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+, and Double-Negative Peyer’s Patch Dendritic Cells1
Immune Cell Interaction Unit, Mucosal Immunity Section, Laboratory of Clinical Investigation, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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- (myeloid) DCs are localized in
the subepithelial dome, CD11b-/CD8+
(lymphoid) DCs in the interfollicular regions, and
CD11b-/CD8- (double-negative; DN) DCs at
both sites. We now describe the presence of a novel population of
intraepithelial DN DCs within the follicle-associated epithelium and
demonstrate a predominance of DN DCs only in mucosal lymphoid tissues.
Furthermore, we demonstrate that all DC subpopulations maintain their
surface phenotype upon maturation in vitro, and secrete a distinct
pattern of cytokines upon exposure to T cell and microbial stimuli.
Only myeloid DCs from the PP produce high levels of IL-10 upon
stimulation with soluble CD40 ligand- trimer, or
Staphylococcus aureus and IFN-
. In contrast, lymphoid
and DN, but not myeloid DCs, produce IL-12p70 following microbial
stimulation, whereas no DC subset produces IL-12p70 in response to CD40
ligand trimer. Finally, we show that myeloid DCs from the PP are
particularly capable of priming naive T cells to secrete high levels of
IL-4 and IL-10, when compared with those from nonmucosal sites, while
lymphoid and DN DCs from all tissues prime for IFN- production.
These findings thus suggest that DC subsets within mucosal tissues have
unique immune inductive capacities. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and
IL-10 (1). Upon Ag-specific triggering, these regulatory T
cells can have Ag nonspecific suppressive effects at systemic sites.
High doses of the same soluble protein Ag given orally can result in
nonresponsiveness due to deletion or anergy of Ag-specific T cells
(2), similar to what occurs following systemic
administration of soluble proteins (3). Not all oral
encounters with Ag result in tolerance, however. Local and systemic T
cell priming can occur when soluble Ags are administered with an
ADP-ribosylating adjuvant, such as cholera toxin, which results in a
Th2-dominant response, or following infection with a mucosal pathogen,
such as Salmonella typhimurium or Toxoplasma
gondii, which results in a Th1-dominant response. Despite recent
progress, the mechanisms by which such disparate T cell immune
responses occur following the administration of orally administered Ags
are still poorly understood. To address this issue, we have focused our
studies on the function of dendritic cells
(DCs)3 of the Peyer’s
patch (PP), because PPs are the primary sites for the induction of
immune responses in the intestinal mucosa and are representative of
lymphoid follicles present in diffuse mucosal tissues.
It has been postulated that the DC system consists of distinct
cellular subsets, which may arise from either a myeloid or lymphoid
precursor (4). In the mouse, these two types of DCs have
been shown to express either high levels of CD11b (myeloid) or
CD8 (lymphoid) molecules, respectively. In addition, we recently
described a third subset of DCs that do not express either of these
markers (double-negative (DN) DCs) (5). By
immunofluorescent staining of tissue sections of PPs, we found that
CD11b+/CD8- DCs are
localized in the subepithelial dome (SED),
CD11b-/CD8+ DCs in the
interfollicular regions, and
CD11b-/CD8- DN DCs at
both sites. Moreover, we have shown that total DC populations, defined
as CD11c+/MHC class
II+/B220-, isolated from
PPs were capable of differentiating naive CD4+ T
cells in vitro into Th cells secreting IFN-, IL-4, and high levels
of IL-10, while total DCs from the spleen primed T cells to secrete
predominantly IFN- (6). This suggested that DCs from
the PP may have a unique intrinsic capacity to induce T cell responses
that are particularly capable of providing help for IgA B cell
differentiation or for regulating systemic immune responses via the
production of IL-4 and IL-10.
In this study, we characterize the phenotype and function of myeloid,
lymphoid, and DN DCs from the PP. First, we describe the presence of a
unique population of intraepithelial DN DCs within the
follicle-associated epithelium (FAE) of the PP. Second, using isolated
cells, we show that the three DC subsets from PP and spleen express
similar levels of MHC class II and costimulatory molecules.
Furthermore, upon overnight in vitro stimulation, all DC subsets
up-regulate expression of DEC-205, MHC class II, CD80, and CD86,
whereas lineage markers are unchanged. Thus, none of the subsets seem
to convert from one type into the other during in vitro maturation.
Third, we demonstrate that upon stimulation in vitro, myeloid DCs from
the PP have a particular capacity to produce IL-10, while lymphoid and
DN DCs from PP produce predominantly IL-12 p70. The ability to secrete
high levels of IL-10 by myeloid PP DC was not an attribute of myeloid
DCs from other lymph nodes examined (mesenteric, axillary, inguinal,
and popliteal). Fourth, we show that only myeloid DCs from the PP
stimulated rapid Ag-specific T cell proliferation and the induction of
cells producing high levels of IL-4 and IL-10. Myeloid DCs from the
spleen induced much lower levels of proliferation, as well as IL-4 and
IL-10 secretion, while all DC subpopulations induced the
differentiation of T cells capable of secreting IFN-.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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PPs were dissected from small intestine of BALB/c, or C57BL/6
mice were frozen in OCT medium (Sakura Finetek, Torrance, CA). Sections
(8 µm) were fixed in cold acetone and stained for DC markers using
the tyramide amplification method (TSA-Direct kit; NEN Life Science
Products, Boston MA), as previously described (5).
Sections were stained with 2 µg/ml anti-CD11c or 10 µg/ml
anti-I-A (M5/114 hybridoma supernatant), anti-CD11b (M1/70
hybridoma supernatant), anti-CD8 (53-6.7; BD PharMingen, San
Diego, CA), anti-CD45R (RA3-6B2; BD PharMingen), or anti-CD3
(CT-CD3; Caltag Laboratories, Burlingame, CA). In Fig. 1
F,
section was stained with anti-CD11c and 2 µg/ml of biotinylated
UEA I-lectin from Ulex europaeus (Sigma-Aldrich, St.
Louis, MO), followed by HRP-conjugated streptavidin and detected with
the Tyramide system, as previously described (5). Slides
were mounted with Fluoromount G (Southern Biotechnology Associates,
Birmingham, AL) and analyzed by confocal microscopy using a Leica
TCS-NT/SP confocal microscope using a x63 objective with oil with zoom
of 2.0x. For Fig. 1A–E, differential interference contrast
(DIC) images were collected simultaneously with the fluorescence images
using the transmitted light detector confocal laser microscope. For
Fig. 1F, the FAE was traced using the Trace Contour filter
in Adobe Photoshop (San Jose, CA).
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Flow cytometric analysis of DC subsets was conducted using
either FITC-, PE-, biotin-, APC-, or CyChrome-conjugated anti-CD11c
(HL3), anti-CD11b (M1/70), anti-CD8 (53-6.7),
anti-I-Ad (AMS-32.1), anti-CD40 (HM40-3),
anti-CD80 (1G10), anti-CD86 (GL1), anti-CD4 (L3T4),
anti-DEC205 (NLDC-145), and anti-CD45R (RA3-6B2). Before
staining, Fc receptors (FcRIII/II) were blocked using anti-mouse
CD16/CD32 (2.4G2). Isotype-matched control Abs used for staining DCs
included rat IgG2a, 
(R35-95); rat IgG2b, (R35-38 or A95-1); and
hamster IgG (G235-2356). The above Abs were purchased from BD
PharMingen, except for NLDC-145, which was purified from hybridoma
supernatant, as described previously (5).
Stimulation of DCs in vitro
Six- to 8-wk-old female BALB/c mice were obtained from National
Cancer Institute (Frederick, MD). DCs were prepared from spleen
(SP) and PP, as previously described (5). Briefly,
isolated tissue was digested with collagenase D and DNase, and
incubated in the presence of EDTA to dislodge lymphoid DCs
(7). Cells were incubated with anti-mouse CD11c-coated
magnetic beads (Miltenyi Biotech, Auburn, CA) and selected on MACS
separation columns. Cells selected on the basis of CD11c expression
were then stained with PE-conjugated anti-CD11c and
CyChrome-conjugated anti-B220 Ab and either FITC-labeled
anti-CD8 Ab (for lymphoid DC) or FITC-labeled anti-CD11b Ab
(for myeloid DC) or both (for DN DLs). Lymphoid, myeloid, and DN DCs
were isolated by flow cytometric sorting performed on a FACStar sorter
(Becton Dickinson, Waltham, MA). Sorted DCs were typically between 95%
and 98% positive for the surface markers of interest and were all MHC
class II+. FACS-purified DC subsets (2 x
105 per well) were incubated overnight in the
presence of either recombinant murine CD40L trimer (10 µg/ml;
Immunex, Seattle, WA) or with Staphylococcus aureus,
Cowan’s (SAC) strain (0.02%; Calbiochem, La Jolla, CA) and IFN-
(100 ng/ml; BD PharMingen) in a total volume of 200 µl per well of
96-well microtiter plate. Supernatants were collected, and IL-10 and
IL-12 p40, p70 levels were measured by ELISA using OptEIA set (BD
PharMingen).
Stimulation of TCR transgenic T cells by DCs
Mice transgenic for a TCR that recognizes OVA323–339 peptide in
the context of I-Ad (DO11.10TCR- transgenic
mice) on a BALB/c background were kindly provided by Dr. Dennis Loh
(Washington University, St. Louis, MO). Spleen T cells from
TCR-transgenic mice were prepared by negative selection on T
cell-enrichment columns (R&D Systems, Minneapolis, MN), followed by
isolation of CD4+/MEL-14+ T
cells by flow cytometric sorting using FITC anti-CD4 and PE
anti-lymphocyte endothelial cell adhesion molecule-1 Abs (BD
PharMingen). Sorted T cell populations were typically 99% pure. In
vitro T cell differentiation assays were performed as previously
described (6). Briefly, primary stimulation cultures were
established by coincubation of purified T cells (5 x
104 cells/well) and sorted DCs from spleen or PPs
(5 x 103 cells/well) pulsed with the
corresponding peptide (3 µM), and 1 ng/ml human rIL-2 (Genzyme,
Cambridge, MA) in a 96-well plate at 200 µl/well for 5-7 days. T
cells were restimulated with anti-CD3 and anti-CD28. Supernatants from
restimulated T cells were collected for detection of IL-4 at 24 h,
and IL-10 and IFN- at 48 h. Cytokine secretion was assayed by a
specific sandwich ELISA for IL-10 and IFN- (BD PharMingen), or IL-4
(Endogen, Boston, MA). Proliferation of T cells was assayed by
incorporation of [3H]thymidine during the final
8 h of a 48-h incubation.
Statistical analysis
Normally distributed continuous variable comparisons were done using the Student t test.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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+
(lymphoid) nor CD11b+ (myeloid) markers (DN DC).
The DN population localizes in the dome as well as in the
interfollicular region of the PP (5). In this study, we
describe the presence of DN DCs within the FAE, a one-cell-thick layer
covering the dome of the PP, depicted by the space between the arrows
in Fig. 1
. The DN DCs
(CD11c+/MHC class II+) are
clearly present within the FAE (Fig. 1a), and they do not
express CD8 or CD11b (green cells in Fig. 1, b and
c). Interestingly, the MHC class II molecules within these
cells appear to be intracellular (Fig. 1A), suggesting they
are immature DCs (8). Myeloid DCs (yellow cells in Fig. 1c) are found to localize in deeper dome areas and not
within the FAE. Furthermore, the DN DCs are not intraepithelial T or B
lymphocytes because they do not express CD3 or CD45R (B270),
respectively (Fig. 1, d and e). The
intraepithelial lymphocytes (red cells in Fig. 1, b,
d, and e) are seen adjacent to the DN DCs (green
cells) and sometimes can be seen to contact each other (Fig. 1d, upper right corner). Occasionally, DN
intraepithelial DCs were found to be associated with M cells within the
M cell pocket (Fig. 1f).
To determine the surface phenotype of the DN DCs, we had enriched for
DC from SP and PP by magnetic selection, and
CD11c+/B220- cells were
analyzed by flow cytometry. For comparison of the surface phenotype of
the three DC subsets, cells were further gated on the
CD11b+ (myeloid), CD8+
(lymphoid), or
CD11b-/CD8- (DN) DC
subsets and analyzed for the expression of various DC markers (Fig. 2A). The DN DCs were found to
express comparable levels of MHC class II, CD40, and costimulatory
molecules (Fig. 2A). DCs expressing CD4 were found within
the myeloid and DN populations in the spleen, but not the PP. Next, the
proportions of the three DC subsets (all
CD11c+/MHC class II+) in
the spleen and the PP, mesenteric lymph nodes (MLN), and peripheral
lymph nodes (combined axillary, popliteal, and inguinal) were assessed
(Fig. 2B). There was a significant DN subset in the PP,
constituting approximately one-third of the entire DC population in
this organ. To our surprise, MLN also contained a similarly substantial
proportion (30%) of the DN subset. In contrast, this DN subset
constituted only a minor and indistinct population in the spleen (9%)
and peripheral lymph nodes (13%).
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and
CD11b on freshly isolated DCs, it was possible that these subsets
represented immature and mature stages of the same lineage of DCs. To
determine whether the DN DC population represents an immature form of
either myeloid or lymphoid DC subsets, we examined the surface
phenotype of DN DCs after in vitro stimulation with
CD40L3. As we have observed previously for
lymphoid and myeloid DC subsets (5), expression of DEC-205
was increased by in vitro activation of DN DCs (Fig. 3). In contrast, neither CD8 nor CD11b
was induced upon DN DC maturation, whereas costimulatory molecules and
MHC class II levels were substantially up-regulated (data not shown).
Thus, the DN DCs, at least during in vitro stimulation, remain DN, and
thus do not likely represent an immature form of either
CD8+ or CD11b+ DC
populations. In addition, as previously reported, neither
CD8+ nor CD11b+ DCs lose
expression of these lineage markers upon in vitro maturation
(5), indicating that the DN subset does not appear to
derive from maturation of these subsets.
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(a microbial stimulus) (Fig. 4). Signaling through the CD40 molecule
led to the production of IL-10 by PP DC, but not by SP DC.
Specifically, only the CD11b+ myeloid DC from the
PP secreted IL-10 upon CD40 cross-linking. The ability to secrete high
levels of IL-10 by myeloid PP DC was not an attribute of DCs from other
lymph nodes because none of the MLN nor peripheral lymph node DC
subsets secreted IL-10 upon CD40L3 stimulation
(data not shown). When DCs were stimulated with SAC and IFN-, the
CD11b+ PP DC secreted even higher levels of
IL-10, and SP CD11b+ DCs secreted low, but
detectable level of IL-10. Myeloid DC subsets from neither PP nor SP
secreted IL-12 p70. In contrast, the lymphoid DCs from PP and SP
secreted high levels of IL-12 p40 and p70 upon microbial stimulation,
but failed to secrete IL-10. The DN PP DC secreted levels of IL-12 p70
comparable with the lymphoid PP DC in response to SAC and IFN-,
whereas the DN population from SP secreted minimal p70. No significant
cytokine secretion was detected from any of the DC subsets in the
absence of CD40 or SAC and IFN- stimulation (data not shown).
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A). T cells primed with
myeloid PP DCs proliferated much more efficiently during secondary
stimulation with anti-CD3 and anti-CD28 (Fig. 5B).
When cytokine secretion from T cells primed with the three DC subsets
from SP and PP was assessed following secondary stimulation, comparable
levels of IFN- secretion were obtained from T cells stimulated with
these DC subsets (Fig. 6). Interestingly,
high levels of the Th2 cytokines (IL-4 and IL-10) were secreted from T
cells primed with PP myeloid DCs compared with those stimulated with
other DC subsets (Fig. 6, B and C). Therefore,
myeloid, but not lymphoid or DN DCs in the PP induce naive T cells to
proliferate rapidly and to secrete high levels of IL-4 and IL-10, and
this function is specific to DCs from the PP.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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or CD11b has been
previously reported in the lymph nodes (combined mesenteric, aortic,
and axillary) (11), upon careful analysis of the
peripheral and mucosal lymphoid tissues, we found that the DN DCs are
abundant in the gut-associated lymphoid organs (MLN and PP), but not in
the peripheral lymph nodes or the spleen. DN DCs constitute almost
one-third of the entire PP DC population. In a prior study, we
demonstrated that DN DCs localized in the SED and IFR of the PP
(5). We now describe the presence of a unique population
of DN DCs within the FAE. The bodies of these cells were located above
the basement membrane with processes appearing to extend to the luminal
surface, and some were found to be associated with M cells within the M
cell pocket. In contrast, such DCs in the villus epithelium of the
small intestine are rare. However, we have not looked extensively for
such cells in the colonic epithelium, a site in which intraepithelial
DCs have been identified in the rat (12). In addition to
their localization, the fact that the FAE DN DCs expressed high levels
of intracellular MHC class II Ags, indicating they are immature DCs,
emphasized the possibility that these cells capture luminal Ags either
directly or early after transport by M cells. Finally, we have been
unsuccessful in demonstrating that DN DCs express CD8 or CD11b, or
that lymphoid or myeloid DCs lose their lineage markers
(5) upon maturation in vitro, suggesting that DN DCs do
not simply represent a different maturation stage of
CD8+ or CD11b+ DCs.
However, it remains a possibility that the DN DCs would require a
specific condition to differentiate into lymphoid or myeloid subset.
Growth factors such as IL-3 and GM-CSF have been shown to induce
differentiation of lymphoid and myeloid DCs in vitro from human DC
precursors, respectively (13).
In our prior study, we determined that total DC purified from the PP
secrete IL-10 upon stimulation via CD40 cross-linking, and induce the
differentiation of naive CD4+ T cells into helper
cells capable of secreting high levels of IL-4 and IL-10 (in addition
to IFN-). Similar total populations from the spleen did not produce
IL-10, nor induce T cells to differentiate into Th2 cells. We now
extend these findings to show that the PP DCs that secrete IL-10 and
induce Th2 differentiation are restricted to the myeloid DC subset.
Consistent with our prior findings, none of the DC subsets from the
spleen secreted IL-10 upon CD40 cross-linking; however, the myeloid DCs
can produce some IL-10 upon stimulation with SAC and IFN-. Neither
PP nor spleen myeloid DCs produced biologically active IL-12 p70 upon
stimulation. In contrast, the lymphoid and DN DCs from the PP secrete
IL-12 p70 and induce the differentiation of Th1 cells. Therefore, DN
DCs from the PP are functionally similar to the lymphoid subset, and
the myeloid subset is responsible for the capacity of DCs from this
organ to induce Th cells producing high levels of IL-4 and IL-10.
In light of these data, one concept that needs to be addressed is
whether different subsets of DCs are more apt to induce T cells to
differentiate into a Th1 or Th2 pathway, so-called DC1 and DC2, or
whether there is significant plasticity in the ability of DCs to
function in this regard. The data presented in this work would support
the concept that lymphoid, and now DN DCs, are more capable of inducing
Th1 differentiation, while myeloid DCs are more capable of inducing Th2
cells (9, 10). In addition, stimulation of DC subsets with
SAC and IFN- resulted in functional IL-12 p70 production only from
lymphoid or DN PP DCs. These data argue that lymphoid and DN DC subsets
are intrinsically functionally different from myeloid DCs in the
setting of either T cell activation to a protein Ag, or in direct
response to a bacterial signal.
That being said, the data presented in this work also support a role
for the tissue microenvironment in modulating these functional
phenotypes. This is because PP myeloid DCs appear to have a particular
ability to produce IL-10 and induce T cells producing IL-4 and IL-10
when compared with myeloid DCs isolated from other organs such as
spleen (Figs. 4 and 5). One possibility is that the microenvironment of
the PP somehow allows myeloid DC precursors to differentiate into high
IL-10-secreting cells. What signals could contribute to such
differentiation is unknown; however, cytokines such as TGF-, IL-10,
and PGE2 may influence the differentiation of
myeloid DCs in the PP, whereas they may not alter the differentiation
or function of PP lymphoid or DN DCs.
A final issue that remains to be addressed is whether the intrinsic phenotype of any PP DC subset can be altered by direct interactions with microbial products or by environmental conditions induced in the PP following infection or administration of mucosal adjuvants, such as cholera toxin. While such alterations are certainly possible, especially given the recent finding that human plasmacytoid DCs can be differentiated in vitro under different conditions to induce either Th1 or Th2 responses (14, 15), to date no in vivo condition has been identified in mice (or in humans) that alters DC function to this extent, i.e., conditions that result in murine myeloid DCs functioning as Th1-inducing, or lymphoid DCs functioning as Th2-inducing cells.
Taken together, we propose the following model of immune induction
within the PP. Intestinal Ags gain entry into the PP via M cells
specialized in transport of certain types of Ags across the FAE. Once
the Ag is transported by the M cells, the first cells to encounter
orally delivered Ag are most likely the myeloid and DN DCs present just
underneath or within the FAE. If the Ag is a soluble protein Ag, the
uptake of the Ag by the SED DCs would not result in the activation or
migration of these cells. Under these conditions, the T cells
interacting with myeloid DCs would become Th2 or Th3 regulatory cells,
whereas those interacting with the DN DCs would secrete IFN- and
most likely be anergized in the absence of activation signals from the
DCs. In support of this possibility, transient IFN- secretion by T
cells after low dose oral feeding has been observed (16).
In addition, Th2/Th3 cells have been argued to be less sensitive than
Th1 cells to anergy induction (17). The end result is the
induction of suppressor T cells following low dose Ag feeding. If
soluble protein Ag is given at high doses, T cell activation occurs in
the PP dome, IFR, as well as in the lamina propria (A. Iwasaki, J.
Chung, and B. L. Kelsall, unpublished observation). Deletion or
anergy of Ag-specific T cells may occur at any of these sites. In the
case of microbial infection with intracellular organism such as
salmonella, the SED DCs would be the first cells to be infected
(18). These DCs, now having been stimulated by the
microbial stimuli such as the LPS on the surface of the bacteria, will
undergo maturation and migrate to the IFR or to the MLN. There, the DN
SED DCs can prime Th1 response either directly, or transfer the
microbial Ag to the IFR-resident lymphoid or DN DCs and cross-prime T
cells. In the current studies, we found that in addition to CD40
cross-linking, SAC and IFN- induced high levels of IL-10 and little
to no IL-12 production by PP myeloid DCs. This implies that during an
intestinal Th1-inducing bacterial infection of the PP, IL-10 will be
produced by myeloid DCs, while the primary source of IL-12 for the
induction of Th1 cells is lymphoid or DN DCs. In this regard, IL-10
produced by myeloid DCs may be acting to control the detrimental
effects of Th1-induced inflammation.
In conclusion, this study clearly demonstrates the presence of three distinct DC subsets in the PP, and shows that myeloid DCs from the PP have the particular ability to secrete IL-10 and to induce naive CD4+ T cells to differentiate into Th2 cells. Moreover, we define a significant population of DN DCs in the PP that is capable of secreting IL-12 p70 upon recognition of microbial stimuli, and is present within the intraepithelial compartment of the FAE.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Brian L. Kelsall, Immune Cell Interaction Unit, Mucosal Immunity Section, Laboratory of Clinical Investigation, National Institutes of Allergy and Infectious Diseases, National Institutes of Health, 10 Center Drive, Room 11N238, Bethesda, MD 20892-1890.
3 Abbreviations used in this paper: DC, dendritic cell; CD40L3, CD40 ligand trimer; DIC, differential interference contrast; DN, double-negative; FAE, follicle-associated epithelium; IFR, interfollicular region; MLN, mesenteric lymph node; PP, Peyer’s patch; SAC, Staphylococcus aureus Cowan’s strain; SED, subepithelial dome; SP, spleen.
Received for publication December 11, 2000. Accepted for publication February 13, 2001.
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