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Laboratoire de Biologie et Thérapeutique des Pathologies Immunitaires, Université Pierre et Marie Curie, Centre National de la Recherche Scientifique (CNRS), Groupe Hospitalier Pitié-Salpêtrière, and
Genopoïétic, Paris, France
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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expression, suggests a lymphoid-related
origin. The sl-DC population is expanded 100-fold after an in vivo flt3
ligand treatment. The sm-DC phenotype suggests a myeloid-related
origin. Interestingly, sm-DCs can acquire i.v. injected macromolecules
in less than 30 min after injection. They may, therefore, play an
important role in the immune response against blood Ags. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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molecule (9).
Granulocyte-macrophage CSF is not required for the development of these
lymphoid-related DCs (10). A common precursor for T cells, B cells, NK
cells, and DCs has also recently been identified in human bone
marrow (11). Langerhans cells may belong to a third DC lineage, and
TGF-ß1 is required for their development (3, 12, 13, 14). In addition to these distinct lineages, DCs differ in their maturation stage and functional properties. "Immature" DCs appear very efficient for macromolecule capture and processing. Inflammatory mediators induce their differentiation into more mature or activated cells that have acquired the capacity to efficiently stimulate T lymphocytes (5, 15, 16, 17). Finally, DCs also appear heterogeneous in terms of turnover (18). Some of them, such as the Langerhans cells (LCs), appear to be renewed very slowly in the absence of inflammation (19, 20). Other DCs have a rapid turnover with a t1/2 < 1 wk (21, 22, 23, 24).
DC heterogeneity has already been described in some lymphoid tissues. In the spleen, two DC populations with distinct phenotypes and localizations have been described: the DCs of the marginal zone and the DCs localized in the periarteriolar T cell region of the white pulp (25, 26, 27, 28), that seem to be of myeloid- and lymphoid-related origin, respectively (9, 29). Similarly, DC heterogeneity has also been described in the Peyer’s patches (30) and tonsils (31).
The characterization of these distinct populations of DCs in various tissues or organs is important because they most probably have different functions in the immune system. Indeed, it has already been suggested that in the spleen the myeloid-related DCs a play major role in triggering immune responses, while the lymphoid-related DCs participate in the regulation of these responses (32, 33). Surprisingly little is known about lymph node (LN) DCs, although these secondary lymphoid organs are major sites of immune response initiation. Until now, a single DC type has been described, which is located in the paracortical zone and referred to as an interdigitating cell (34). It is believed that these cells are derived from immature DCs localized in regional tissues, which migrate to the draining LN via afferent lymph (Refs. 35–37; and reviewed in 38 .
In this work, we present a detailed analysis of LN DCs. We have identified three distinct populations that can be distinguished on the basis of their morphology and phenotype. Interestingly, these cells also differ in their capacity to uptake Ag delivered by different routes and in their dynamics.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Inbred mouse strains DBA/2, C57BL/6, CBA/J, and FVB/N were obtained from IFFA-Credo (L’Arbresle, France). Except where indicated, all experiments were performed with DBA/2 mice. Chimeric mice used to analyze DC turnover were derived from LTR-TK transgenic mice expressing the HSV1-thymidine kinase (TK) in DCs as previously described (24, 39). They were generated by transplantation of 3 to 5 x 106 bone marrow cells from LTR-TK transgenic mice into lethally irradiated (13 Gy) normal recipients with the same DBA/2 genetic background.
Enzymatic cell dissociation
LNs were cut into small fragments and incubated in RPMI 1640 supplemented with 1.6 mg/ml (500 IU/mg) collagenase (type IV, Sigma Chemical Co., Saint Quentin Fallavier, France) and 200 µg/ml DNase I (Boehringer Mannheim, Mannheim, Germany) at 37°C for 30 min. Cells were dissociated by repeated pipetting, reincubated at 37°C for 10 min, and washed. Cell suspensions were then incubated with 200 µg/ml DNase I for 15 min at room temperature and resuspended in staining buffer (PBS, 3% FCS, 0.02% azide) for further flow cytometric analyzes.
MHC class II/CD11c double staining
CD11c expression was analyzed with the unlabeled N418 mAb
(hamster IgG, HB224; American Type Culture Collection (ATCC),
Rockville, MD) revealed by a PE-conjugated F(ab')2
goat anti-hamster IgG (Caltag Laboratories, San Francisco, CA).
Depending on the analysis performed, we used different mAbs against MHC
class II molecules: the M5/114 mAb (ATCC TIB120) was revealed by a
TriColor-conjugated F(ab')2 goat anti-rat IgG (Caltag
Laboratories) (see Figs. 1
, 5, A and B, and 6);
the 14.4.4S mAb (PharMingen, San Diego, CA), either FITC-conjugated or
biotin-conjugated, was revealed by streptavidin TriColor (Caltag
Laboratories) (see Figs. 3, A and B, and 7); the
biotin-conjugated 2G9 mAb (PharMingen) was revealed by streptavidin
TriColor (see Fig. 4); the 28-16-8S mAb (Caltag) was revealed by an
allophycocyanin-conjugated goat anti-mouse IgM (Caltag) (see Figs. 3C and 5C).
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Panels of mAbs were selected to study the phenotype of LN DCs by
three-color flow cytometry analysis. We used a panel of FITC-conjugated
mAbs against B220 (RA3-6B2, Caltag Laboratories), mThy-1.2, mCD4
(CT-CD4, Caltag Laboratories), mCD8 (CT-CD8a, Caltag Laboratories),
Gr-1 (RB6-8C5, Caltag Laboratories), F4/80 (F4/80, Caltag
Laboratories), Mac-1 (M1-70.15, Caltag Laboratories), heat stable Ag
(HSA) (M1/69, PharMingen), MHC class II (M5/114, Boehringer Mannheim),
CD44 (IM7, PharMingen), and CD62L (Mel-14, PharMingen), a panel of
biotin-conjugated mAbs against B7-1 (16.10. A1), B7-2 (GL-1),
H2-Kd (SFI-1.1, PharMingen), and CD54 (KAT-1,
Caltag Laboratories) and a panel of uncoupled rat mAbs against CD2
(AT37, Serotec, Oxford, U.K.), spleen DC (33D1, ATCC TIB 227), DEC 205
(NLDC145 (25)), CD40 (3/23, Serotec), and Fc
II/IIIR (2.4G2,
PharMingen).
To minimize nonspecific binding, cells were preincubated either with 2.4G2 mAb and then analyzed using FITC- or biotin-conjugated third mAbs or with 10% mouse serum and analyzed using mAbs from the panels of uncoupled mAbs. In all stainings, the N418 mAb was revealed by a PE-conjugated F(ab')2 goat anti-hamster IgG (Caltag Laboratories). For MHC class II, the staining differed according to the third mAb used. For FITC-conjugated mAbs, we used a biotin-conjugated anti I-E mAb (14.4.4S) revealed by streptavidin TriColor. For mAbs from the panels of biotin-conjugated or uncoupled rat mAbs, revealed respectively by streptavidin TriColor (Caltag Laboratories) and a TriColor-conjugated F(ab')2 goat anti-rat IgG (Caltag Laboratories), the anti I-E staining was performed with the FITC-conjugated 14.4.4S mAb.
The following isotypic Ig controls were used: FITC-conjugated rat IgG2a (LODDNP-16, Immunotech, Marseille, France), FITC-conjugated rat IgG2b (Cedarlane, Hornby, Ontario, Canada), and biotin-conjugated rat IgG2a, biotin-conjugated hamster IgG, unconjugated rat IgG2a, and unconjugated rat IgG2b (Caltag Laboratories).
Cells were fixed in 1% formaldehyde, and analyses were performed on a FACScan (Becton Dickinson Co., Mountain View, CA).
Flow cytometry cell sorting
After collagenase digestion of inguinal, brachial and axillary LNs from 35 DBA/2 mice, cells were fractionated on a discontinuous gradient of BSA (density = 1.082) as previously described (40). The recovered low-density cells, preincubated with the 2.4G2 mAb to reduce nonspecific binding, were stained with N418 revealed by PE-labeled anti-hamster Ig and FITC-labeled anti-I-E (14-4-4S) mAbs. Cells were sorted at a rate of 3000 events/s on a FACStarPlus (Becton Dickinson) and were kept at 4°C throughout the procedure.
Immunostaining
Cells were cytocentrifuged for 5 min at 300 rpm on slides. Cells were fixed in 1% formaldehyde, 0.2% glutaraldehyde for 5 min at 20°C and conserved at 4°C in PBS until use. Slides were incubated for several min in TBS (50 mM Tris, pH 7.6, 0.9% NaCl), then in TBS supplemented with 0.5% BSA and 0.02% Nonidet P-40 (staining buffer; Sigma Chemical Co.). Slides were immunostained in staining buffer supplemented with saturating levels of M5/114 hybridoma supernatant for 45 min at room temperature, washed twice in TBS, incubated in the staining buffer with peroxidase-conjugated rabbit anti-rat Ig (P 0450, Dako SA, Trappes, France), washed in 50 mM Tris, colored by 3,3'-diaminobenzidine hydrochloride (DAB tablets, Sigma Fast, Sigma Chemical Co.) for 2 to 3 min, rinsed 5 min in H2O, counterstained with hematoxylin, rinsed 5 min in H2O, fixed in 70% ethanol then 100% ethanol, and mounted.
Detection of CD4 transcription by RT-PCR
LN cells were lysed in an RNA extraction solution (RNA-BTM; Bioprobe, Montreuil, France). Cellular RNA was then reverse transcribed at 42°C for 1 h in a 20-µl reaction containing: 1 mM of all four deoxynucleotide triphosphates, 0.04 U of random primer P(dN)6, 40 U of RNase inhibitor (Pharmacia LKB Biotechnology, Uppsala, Sweden), and 200 U of MMLV reverse transcriptase (Life Technologies, Gaithersburg, MD). Five microliters of the reverse transcription product were used for amplification in a 50-µl reaction containing: 1 µM of murine CD4 or ß-actin primers, 200 µM of all four deoxynucleotide triphosphates, 1.5 mM MgCl2, and 1 U Taq DNA polymerase (Goldstar DNA polymerase; Eurogentec, Seraing, Belgium). The murine CD4 primers were: sense primer, 5'-TGTGGCAGTGTCTGCTGAGTGA-3', in the D4 domain; antisense primer, 5'-TGGCAGGTCTTCTTCTCACTGA-3', in the cytoplasmic region. There are two introns present between the genomic position of these primers, and thus they can only amplify CD4 cDNA. Reactions were performed in a DNA thermal cycler (Hybaid Ltd., Teddington, U.K.) as follow: an initial denaturation cycle lasting 10 min at 94°C, followed by 35 cycles of amplification each comprising denaturation for 30 s at 94°C, annealing for 30 s at 62°C, and extension for 30 s at 72°C. The last cycle was followed by an extension cycle lasting 10 min at 70°C.
Allogeneic MLR
Nylon wool-passed T cells from mesenteric, brachial, axillary, and inguinal LN cells and graded numbers of FACS-sorted stimulator cells irradiated at 20 Gy were cocultured in RPMI 1640 supplemented with 10% FCS (Flobio, Asnieres, France), 50 µM 2-ME, and antibiotics in U-bottom 96-well microplates. After 108 h of culture, cells were pulsed with 1 µCi/well of [3H]TdR for an additional 12 h before harvesting and scintillation counting.
FITC skin painting
Mice were anesthetized with avertin (2.5% tribromoethanol) and the skin painted at the level of the triceps muscle or on the abdomen at the level of inguinal LN with 25 µl of 0.8% FITC (isomer 1, Sigma Chemical Co.) dissolved in a 1:1 mixture of acetone:dibutylphthalate just before application.
FITC-dextran
Mice were anesthetized with avertin and injected i.v. in the retro-orbital sinus with 100 µl of 50 to 100 mg/ml FITC-dextran, m.w. 40,000 (FD-40, Sigma Chemical Co.) or 150,000 (FD-150, Sigma Chemical Co.). Potential small FITC contaminants were removed by dialysis through a dialysis membrane (membra-cell 24037, Polylabo France) with a m.w. cut-off of under 12,000 to 16,000. Dialysis was performed for 24 to 48 h at 4°C in PBS in the dark.
Ganciclovir and Flt3 ligand administration
Ganciclovir (GCV) was continuously administered at a dose of 50 to 55 mg/kg/day for 7 days by using miniosmotic pumps (model 2001, Alza Corp., Palo Alto, CA) implanted s.c. as previously described (24). Flt3 ligand (Immunex, Seattle, WA) was injected i.p. once a day for 7 days at a dose of 10 µg/day.
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Results |
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Double staining of LN cells, performed using the N418 mAb, which
recognizes the murine CD11c (41), and a mAb against MHC class II,
allowed us to distinguish four cell populations. Besides
double-negative and MHC class II+/CD11c-
cells, which are mainly T and B cells, respectively, two distinct
populations of double-positive cells were detected (Fig. 1
A). One expressed high
CD11c levels and cell surface MHC class II at a level comparable to
that of B cells (s-cells, Fig. 1A). The other
expressed 5 to 10 times more cell surface MHC class II and lower levels
of CD11c (l-cells, Fig. 1A). These cells represented
approximately 0.5 and 1.5% of all LN cells, respectively. Size
analysis of gated cells showed that l-cells were significantly larger
than s-cells, hence the label s (small) and l (large) (Fig. 1B). Two MHC class II+/CD11c+
populations were identified in all of the LNs (brachial, inguinal,
axillary, para-aortic, or mesenteric) and mouse strains (DBA/2,
C57Bl/6, CBA/J, FVB) analyzed (not shown). However, in mesenteric LNs,
l-cells expressed lower levels of cell surface MHC class II compared
with peripheral LNs.
To analyze their T cell stimulatory capacity, s- and l-cells were sorted by flow cytometry and used as stimulators in mixed lymphoid reaction. Both s- and l-cells had a strong T cell MLR-stimulating activity, similar to that of splenic DCs used as control (data not shown).
Morphology and phenotype delineate three distinct LN DC populations
To observe the morphology of s- and l-cells, flow cytometry cell
sorting was performed. Cytospins of sorted cells revealed for both
types of cells the typical dendritic cellular processes that
characterize DCs (1). However, some differences could be observed:
l-cells were larger and had longer and more numerous dendritic
processes; s-cells contained intracellular clusters of MHC class
II-rich compartments (Fig. 2).
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). The two populations did not express
the classical markers of B cells (B220), T cells (Thy-1), or
granulocytes (Gr-1). They did express a set of molecules associated
with the Ag-presenting function of DCs such as MHC molecules and
costimulatory and adhesion molecules ICAM-1, B7-1, B7-2, and CD40.
l-Cells expressed intermediate to high levels of these molecules, while
expression levels were lower in s-cells.
l-Cells did not express CD4, CD8, the myeloid marker F4/80, and the
33D1 marker of splenic DC; they expressed low levels of the
interdigitating cell marker DEC-205 and heterogeneous levels of HSA and
Mac-1. In contrast, these molecules clearly defined two distinct
populations of s-cells. One expressed CD4, F4/80, Mac-1, 33D1, and low
levels of HSA and did not express DEC-205, while the other population
expressed the lymphoid DC marker CD8 and DEC-205, high levels of
HSA, and low to zero levels of CD4, F4/80, Mac-1, and 33D1 (Fig. 3 and
data not shown). To verify that the detected CD4 expression was due to
endogenous production, we analyzed CD4 transcription by RT-PCR on
sorted cells. CD4 transcripts were indeed detected in s-DC using
primers that encompass an intron and thus cannot amplify genomic DNA
(data not shown). Altogether, based on their phenotype, sm- and sl-DCs
appear to be myeloid or lymphoid related, respectively (9, 29).
Therefore, based on morphology, phenotype, and T cell-stimulatory capacity, s- and l-cells can be considered as typical DCs (1) and will subsequently be referred to as s-DC and l-DC, according to their small and large size, respectively. Based on their putative lymphoid- or myeloid-related origin, the two s-DC populations will subsequently be referred to as sl-DC and sm-DC, respectively.
l-DC acquire skin-painted Ags
The migration of DCs from extravascular compartments of
nonlymphoid tissues to draining LNs via afferent lymph has been well
documented (36, 37, 38). In this respect, 24 h after skin painting
with FITC diluted in organic solvents, this tracer was detected in LN
DCs, suggesting that epidermal LCs migrate to the draining LN (42, 43).
We analyzed FITC staining of s-DC and l-DC at different time points
after FITC skin painting. After 24 h, up to 60% of l-DCs showed
high levels of FITC fluorescence in the draining brachial LN, whereas
no staining was observed in the contralateral one. FITC staining of
l-DCs became detectable 12 h after application and peaked at
24 h (Fig. 4). Interestingly, in
preliminary experiments, when we applied a 10-fold higher volume of
FITC, we observed a significant increase in l-DCs proportion,
suggesting that new cells immigrated into the LN (data not shown).
However, this procedure was not suitable for additional experiments due
to staining of the contralateral LN DCs. The proportion of FITC-stained
cells then decreased, with only a few stained cells remaining at day 5
(Fig. 4); this could be due to either their migration outside the LN,
their death in situ, or FITC degradation. During the 5-day follow-up,
no significant FITC staining could be detected in CD11c-negative cells,
nor in either s-DC population (Fig. 4).
sm-DC rapidly and efficiently acquire blood macromolecules
We similarly tested the capacity of the three populations of LN
DCs to acquire blood Ag using i.v. injection of FITC-dextran. High m.w.
dextrans are known to remain in the plasmatic compartment (44). They
have also been used as a marker to quantify the endocytic capacity of
DCs (17). Cells of peripheral LN were analyzed at different time points
after i.v. injection of FITC-dextran. At 30 min, 17% of s-DCs were
highly fluorescent in both brachial and inguinal LNs. In contrast, only
3% of l-DCs were FITC+, and moreover, the
fluorescence of these stained cells was weak (Fig. 5). The proportion of FITC-stained s-DCs
increased rapidly during the first 30 min after injection, and then
more slowly during the next 12 h to reach 30% of the cells. This
percentage then remained stable for 5 days. The percentage of
fluorescent l-DCs increased slowly, reaching the same proportion after
12 h as that observed at 30 min for s-DCs. At day 5, the
percentages of fluorescent s- and l-DCs were similar. During this time
period, only traces of CD11c- cells were FITC+
(Fig. 5). Similar results were obtained using FITC-dextran of 40,000 or
150,000 m.w. or after dialysis to remove potential small m.w. FITC
contaminants (data not shown).
To analyze which of the s-DC acquire FITC-dextran, we performed
four-color flow cytometry analysis. 30 min after FITC-dextran i.v.
injection, 17% of the sl-DCs were stained, while >50% of the sm-DCs
were FITC+ (Fig. 5C). However, sl- and
sm-DCs populations overlap based on CD4 staining (Fig. 3A), and it should be noticed that the FITC-stained
sl-DCs are positioned close to the bar separating sm- and sl-DC on the
CD4 staining axis. Therefore, it can be assumed that most if not all
stained cells are sm-DCs. Similar results were obtained at 12 h
after FITC-dextran injection (data not shown).
Careful analysis of FITC staining in s- and l-DCs revealed two
interesting features. First, there was a progressive increase in the
cell surface expression of MHC class II in most FITC+
cells, which resulted in a shift from the s- to the l-DC population
(Fig. 6A). Furthermore,
while CD11c expression was heterogeneous among l-DCs, the l-DCs that
became FITC+ had high CD11c expression levels, comparable
to that of s-DCs. Second, the intensity of FITC staining increased with
time among s-DCs and also, but in a delayed manner, in l-DCs (Fig. 6B).
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We next analyzed the in vivo turnover of l- and s-DCs. We used a
model of transgenic mice allowing the conditional ablation of dividing
DC precursors expressing an HSV1-TK gene upon GCV treatment (24, 39).
This enzyme allows the conversion of the nontoxic GCV into GCV
triphosphate, which can be incorporated into elongating DNA, inducing
elongation termination and ultimately cell death. Because only those
cells that express HSV1-TK and that are dividing can be killed, size
variation of a cell population during a GCV treatment reflects the
turnover of these cells if they divide or have a dividing precursor.
However, this system will not discriminate between the rapid turnover
of a resident cell population that divides in situ or of a circulating
cell population with a dividing precursor. Using this model, we
previously showed the rapid turnover of spleen DCs, amounting to 10 to
15% renewal per day, in agreement with previous measurements obtained
with other techniques (21). Here, we found that a 7-day GCV treatment
led to the almost complete disappearance of both populations of s-DCs,
while the l-DC population was only slightly reduced (Table I). These results reveal further
differences between the three populations of LN DCs, sm- and sl-DC
having a rapid turnover, l-DC a slow turnover.
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The in vivo administration of Flt3 ligand, a stimulator of
hemopoietic progenitor cells, has recently been described to induce
hypertrophy of lymphoid tissues and to dramatically increase the
proportion of spleen DC (45, 46). The effects of Flt3 ligand on the
distinct subpopulations of LN DCs were analyzed by three-color flow
cytometry after a 7-day treatment. The l-DCs did not appear to be
affected by Flt3 ligand. In contrast, the proportion of s-DCs reached
30% of total LN cells vs 0.5% in controls (Fig. 7A). The expanded
population did not express CD4, F4/80, or Mac-1, but expressed low
levels of CD8 and high levels of HSA, indicating that Flt3 ligand
treatment affected mostly if not exclusively the sl-DC subpopulation
(Fig. 7B). A moderate increase in the other DC
populations could be masked by the dramatic increase of the sl-DCs.
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Discussion |
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In mouse lymphoid tissues, N418 is the only mAb that strongly reacts with DCs but not with freshly isolated macrophages or lymphoid cells (29, 41). We looked for a possible heterogeneity of DCs in LN within CD11c+, as already described in the spleen (27, 29, 32, 47) and in Peyer’s patches (30). We used double-staining analysis of whole LN cells without any purification procedure or culture, which could have modified the cell characteristics or resulted in the loss of a DC subpopulation. Three distinct CD11c+/MHC class II+ populations were clearly disclosed by four-color flow cytometry analysis. Given their overall heterogeneity, it is admitted that identification of DCs is based on a combination of characteristics including: 1) their morphology, with the existence of dendrites; 2) their phenotype, showing the presence of molecules involved in T cell activation; and 3) their T cell stimulatory capacity (1). The three CD11c+/MHC class II+ cell populations fulfill all of these criteria and can thus be considered as typical DCs.
Due to their high level cell surface expression of MHC class II and costimulatory molecules, their size, and their dendritic morphology with numerous and long cell processes, l-DCs resemble mature DCs. On the other hand, due to the mainly intracellular location of MHC class II molecules, their smaller size, and their smaller dendrites, both populations of s-DCs resemble more immature DCs (15, 17, 48). Despite differences in MHC class II expression, all of these cells strongly stimulate in vitro allogeneic responses to similar levels. This is probably due to a rapid in vitro maturation of s-DCs during the culture, as suggested by preliminary experiments (data not shown).
Expression of the Mac-1 and F4/80 myeloid markers on the sm-DC
population supports a myeloid origin of these cells. CD4 expression
does not argue against such a hypothesis, since CD4 is not considered a
marker of lymphoid-related DCs (49). Nonetheless, sm-DC are the only
identified DCs that clearly express CD4 at levels similar to that of
CD4 T cells. In addition, based on 33D1 expression and low HSA
expression levels, sm-DCs resemble marginal zone splenic DCs (26, 29),
a subpopulation considered to be myeloid related (9, 50). In contrast,
sl-DCs expressed CD8, a marker considered to define lymphoid-related
DCs (9). sl-DCs also express DEC-205 and high levels of HSA, thus
resembling splenic DCs of the T cell zone that have been considered
lymphoid related (9, 26). Finally, the observation that Flt3 ligand
treatment induced the expansion of only sl-DCs reinforces the
hypothesis that the two s-DCs populations have distinct lineages.
Different turnover of the lymph node DCs
In addition to these phenotypic differences, we showed that these DC subpopulations have a different turnover by using an animal model of conditional ablation of HSV1-TK-expressing DC upon GCV treatment. Using this model, we previously showed that a 7-day GCV treatment led to complete disappearance of splenic DCs of the white pulp marginal zone, while LCs that also expressed HSV1-TK were minimally affected (24, 39). Since GCV kills only HSV1-TK-expressing cells that are dividing (51), these experiments indicate that the splenic DCs have a very rapid turnover, whereas LC have a slow turnover, in agreement with previous observations (19, 20, 21). Using the same experimental system, we show here that sm- and sl-DC populations have a rapid turnover, since a 7-day GCV treatment resulted in the disappearance of almost all s-DCs. This rapid turnover could be due to either a high division rate of the s-DC themselves or to a rapid transit into LN of cells that have a rapidly dividing precursor. Additional experiments will be needed to address this issue. On an other hand, GCV treatment induced only a slight depletion of l-DCs (1.2 vs 1.6% in control mice). This result is compatible with the persistence of the LC-derived l-DC and the disappearance of the sm-DC-derived l-DC (see below).
Origin of the l-DCs
The majority of l-DCs come from the peripheral territories drained by the regional LN. Indeed, at 3, 12, 24, and 120 h after FITC skin painting, l-DCs but not s-DCs were FITC stained. Because there was a progressive increase in the proportion of l-DC-stained cells from 12 to 24 h after skin painting, without any detectable stained s-DCs throughout this time period, it is unlikely that we could have missed a transition from stained s-DCs to l-DCs. This suggests that neither sm- nor sl-DCs represent an intermediate differentiation stage from LCs to l-DCs and that LCs have completed their phenotypic maturation when they arrive in the draining LN. These l-DCs were thus derived from LCs, which themselves are believed to derive from the CFU-DC progenitors yielding pure DC colonies in semisolid medium (3, 12). A maximum of 60% of all l-DCs was stained after skin painting. This may indicate that not all the territory drained by the analyzed LN was painted and/or that some l-DCs have a different origin. In this respect, there is some evidence that a fraction of the l-DC population is derived from sm-DCs. First, sm-DCs were the first to be stained after i.v. injection of FITC-dextran, followed by a delayed staining in l-DCs, which appeared with kinetics compatible with a maturation of sm-DC to l-DC. Second, although CD11c expression in l-DCs was quite heterogeneous, those l-DCs stained by the plasmatic tracer had a CD11c expression that was more homogeneous and was similar to that of s-DCs. Finally, preliminary experiments suggest that some s-DCs can evolve toward an l-DC-like phenotype upon culture (data not shown).
Altogether, these results suggest that LNs contain DCs from the three DC lineages that have been proposed so far, originating from: 1) a pure DC progenitor (for most of l-DC) (3, 12); 2) a common myeloid/DC precursor (for sm-DC) (3, 4); and 3) a common lymphoid/DC precursor (for sl-DC) (10).
Capture of blood Ag by sm-DCs
Intravenous injection of FITC-dextran led to a remarkably rapid staining of sm-DCs. This staining was observed with both 40,000 and 150,000 m.w. FITC-dextran. These molecules are known to remain in the blood circulation and are actually used in humans to replenish the plasma compartment. Therefore, it is unlikely that they diffused directly from the blood to the LN parenchyma. On the contrary, the staining was specific in many ways. First, only sm-DCs, representing 0.2% of all LN cells, were stained by FITC-dextran 30 min after i.v. injection, whereas all of the LN cells were stained when incubated with FITC-dextran in vitro (data not shown). Second, contamination with plasma molecules present in the LN blood vessels and occurring during preparation of LN cells was ruled out; no staining could be observed when LN cells from an untreated mouse were prepared in the presence of the plasma of a FITC-dextran treated mouse (data not shown). Finally, stained s-DCs were only detected in peripheral but not in mesenteric LN. These observations also indicate that the staining of sm-DCs was not due to passive diffusion of FITC-dextran by blood vessel leakage. Therefore, these results suggest that FITC-dextran was acquired within the blood compartment by sm-DCs; this could be due to either the existence of DCs within the blood vessel endothelium layer of peripheral LNs or to a translocation of blood DCs to LNs. In this respect, we observed that the intensity of FITC staining in sm-DCs increased with time, suggesting that some sm-DCs were exposed several times to the plasmatic tracer. These cells may therefore be extremely mobile, migrating from the blood to LNs and back to the blood. Such a recirculation has not been observed for spleen or lymph-borne DCs (52, 53, 54). Altogether, our results suggest a possible circulation of DC from blood to LN. Further careful analysis will be required to clarify this important point, which conflicts with the current paradigm for DC circulation (38).
Until now, Ags delivered through the blood supply have been assumed to be trapped mainly by APCs in the spleen, as well as in the liver for particulate Ags (38, 55). Our results demonstrate that regardless of the mechanism, previously unidentified LN DCs can specifically uptake a plasmatic molecule. These cells, therefore, might play an important role in the control of blood pathogens. In this line, this cell population seems to play a role in the transport of HIV from the blood to the LNs (our manuscript in preparation).
In conclusion, we show for the first time the existence of a DC heterogeneity in the LN. The three DC populations identified appear to belong to distinct lineages and to differ in their capacity to uptake Ags administrated by different routes. These results, together with further analysis of the functional properties of these DCs, may have important implications for a better understanding of the relationship between the route of Ag introduction and the nature of the subsequent immune response.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. David Klatzmann, CERVI, Groupe Hospitalier Pitié-Salpêtrière, 83 blvd. de l’Hôpital, 75651 Paris Cedex 13, France. E-mail address:
3 Abbreviations used in this paper: DC, dendritic cell; LN, lymph node; PE, phycoerythrin; HSA, heat-stable antigen; GCV, ganciclovir; s- and l-cells, small and large cells; sl-, small lymphoid; sm-, small myeloid; HSV, herpesvirus; TK, thymidine kinase.
Received for publication July 18, 1997. Accepted for publication October 2, 1997.
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