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HEMATOPOIESIS
From E00-13 Institut National de la Recherche
Scientifique (INSERM)- Université Paris 6, service
d'Immunologie Biologique and laboratoire d'Immunologie Cellulaire et
Immunopathologie de l'Ecole Pratique des Hautes Etudes; Laboratoire
d'Immunologie Cellulaire, UMR7627 Centre National de la Recherche
Scientifique; INSERM U494; hôpital de la
Pitié-Salpêtrière, Paris; INSERM U346, hôpital
Edouard Herriot, Lyon, France.
To better characterize human dendritic cells (DCs) that originate
from lymphoid progenitors, the authors examined the DC differentiation pathways from a novel CD7+CD45RA+ progenitor
population found among cord blood CD34+ cells. Unlike
CD7 Dendritic cells/Langerhans cells (DCs/LCs) are
essential antigen-presenting cells for initiating and maintaining the
adaptative immune response.1 Recently, the DC system has
become increasingly complex, with reports of discrete populations in
humans and mice, the significance of which remains
enigmatic.2-10 For example, human DCs can differentiate
from CD34+ hematopoietic progenitor cells (HPCs) in the
cord blood,11 the adult bone marrow or
blood,5,12,13 or the thymus,14 and also from
monocytes,15-17
ILT3+/ILT1 Classically, distinct DC subsets are obtained from human
CD34+ HPCs along 2 major pathways leading to either
CD1a+ precursor-derived LC-like DCs or CD14+
precursor-derived DCs related to monocyte-derived
DCs,20-24 a view that has been confirmed in
mice.25 However, this dual LC versus "myeloid" DC
model has been challenged by data that HPCs with lymphoid potential
also generate DCs. The concept of lymphoid DCs arose from the
observation that DCs and T lymphocytes can both develop from murine
Sca+CD4lo thymic precursors26:
Such DCs are now recognized as an original population regarding their
CD8 Isolation of CD34+ cells from the cord blood
and thymus
Thymic progenitors were purified as described34 from
normal thymus of children undergoing cardiac surgery. Thymocytes were obtained by Ficoll-Paque centrifugation, CD34+ cells were
purified with CD34 mAb 561-coated M-450 Dynabeads, labeled with CD34-PE
(clone 581) and CD1a-FITC (clone BB5, Diaclone, Besançon, France)
mAbs, and CD34+CD1a Culture of natural killer cells
Culture of dendritic cells HPCs (1-2 × 104/mL) were cultured as reported22,23,37,38 in 6-well plates (ATGC, Noisy le Grand, France) in RPMI 1640, 10% FCS, 1% glutamine, 1% antibiotics, with the following cytokines: 20 ng/mL granulocyte-macrophage colony-stimulating factor (GM-CSF) (specific activity 1.125 × 107 U/mg; gift of Schering Plough, Kenilworth, NJ), 50 ng/mL SCF (gift of Amgen, Thousand Oaks, CA), 50 ng/mL Flt3 ligand (FL; gift of Immunex, Seattle, WA), and 50 U/mL tumor necrosis factor (TNF)- (Genzyme, Cambridge, MA). When used, IL-4 (50 ng/mL,
gift of Schering Plough; specific activity 2.41 × 107
U/mg) and trimeric CD40 ligand (CD40LT; 0.5 µg/mL; gift of Immunex) were added from culture day 5, whereas macrophage (M)-CSF (250 ng/mL;
specific activity 1.5 × 105 U/µg; R&D Systems,
Minneapolis, MN) or transforming growth factor (TGF)- 1 (10 ng/mL;
specific activity 5 × 105 U/mg; Genzyme) was added at
culture initiation or on day 5. Medium and cytokines were renewed every
3 days. Thymic CD34+CD1a progenitors were
cultured under the same conditions, except that IL-7 was added during
the first 3 days of culture.
Limiting dilution analysis and single-cell cultures Limiting dilution analysis was performed as reported.35 CD34+CD7+CD45RA+ and CD34+CD7CD45RA+ HPCs were seeded
into 96-well plates (ATGC) at 300, 100, 30, 10, 3, and 1 cell per 200 µL/well. Cultures supplemented with SCF, FL, GM-CSF, and TNF- (DC
condition) or with SCF, IL-2, IL-7, and IL-15 (NK condition) were
maintained with half medium change and fresh cytokines added every 6 to
7 days. Plates were examined weekly, and cell-containing wells were
scored under the microscope. At the end of 3 weeks, DCs were identified
by FACS, which was based on CD1a expression, or morphology after
48-hour culture in the presence of CD40LT (CD40LT; 0.5 µg/mL) when
less than 500 cells per well were available. NK cells were detected by
FACS after labeling with CD56-PE (clone Leu-19, Becton Dickinson) or under the microscope after 20 minutes incubation with CD56 mAb (clone
Leu 19)-coated magnetic beads (Dynal).35 The maximum likelihood estimate of DC or NK progenitors was calculated according to
the single hit Poisson model.35
For single-cell suspension cultures,
CD34+CD7+CD45RA+ HPCs were plated
in 96-well plates (0.3 cell per 200µL/well), and cultured as
previously described, but in the simultaneous presence of SCF, FL,
GM-CSF, TNF- Flow cytometry cell surface marker analysis Cells were incubated for 30 minutes at 4°C with mAbs (1:100 final unless specified) in PBS, 2% FCS, with or without 2% AB serum (Site Transfusionnel Pitié-Salpêtrière, Paris, France), washed, and analyzed with a FACscalibur (Becton Dickinson). HPCs were labeled with CD34-PE-Cy5 and the following mAbs: CD2-PE (clone 39C1.5), CD3-FITC (clone SK7), CD7-FITC (clone 8H8.1), CD38-FITC (clone T16), and CD44-FITC (clone J173) from Immunotech; CD7-PE (clone SK7), CD10-FITC (clone W8E7), CD13-PE (clone L138), CD19-PE (clone 4G7), and CD33-PE (clone P67-6) from Becton Dickinson; CD45RA-PE (clone HI100), CD45RO-FITC (clone UCHL1), CD64-FITC (clone 10.1), CD86-PE (clone FUN-1), and CD90/Thy1-FITC (clone 5E10) from Pharmingen; and rat anti-CLA mAb HECA-452 (gift from L. J. Picker and P. R. Bergstresser, Dallas, TX). In addition to CD1a-FITC, DCs were examined with the following mAbs: CD11b-PE (clone bear 1), CD11c-PE (clone BU15), CD83-PE (clone HB15A), CD116-PE (GM-CSFR; clone SCO6), and CD127-PE (IL-7R; clone R34.34, diluted 1:5) from Immunotech; CD14-PE (clone LeuM3) and CD14-allophycocyanin (clone M P9), CD123-PE (IL-3R;
clone 9F5, diluted 1:5) from Becton Dickinson; anti-E-cadherin (clone
HECD-1; R&D Systems). When anti-E-cadherin or anti-CLA unconjugated
mAbs were used, labeling was developed by goat antimouse
F(ab')2-PE or antirat F(ab')2-FITC (anti-CLA) secondary antibodies. Isotype-matched FITC-, PE-, and
allophycocyanin-conjugated irrelevant control mAbs were from Immunotech
and Becton Dickinson; unconjugated mouse and rat isotype controls were
from Immunotech and Caltag (San Francisco, CA), respectively.
Secondary cell sorting Culture day 5 or day 8 cells (1-5 × 106/mL) were incubated at 4°C for 30 minutes with CD14-PE and CD1a-FITC mAb (1:50), resuspended in PBS, 2% FCS, and sorted with a FACStar Plus, based on CD1a or CD14 expression. This resulted in 99% ± 1% pure populations (n = 12).Confocal microscopy Cells were washed with PBS and cytospun onto glass slides, which were dried before 10-minute fixation in PBS, 3% paraformaldehyde at 20°C, and permeabilized with PBS, 0.05% saponin, 0.2% BSA, 0.5% AB serum. Cells were stained with 1:100 diluted unconjugated mouse anti-Lag mAb39 (a gift from F. Furukawa, Handacho, Hamamastu; K. Yoneda and S. Imamura, Kyoto; Japan) or with rabbit anti-S100 antibodies (Dako, Glostrup, Denmark) diluted 1:500. Staining was developed with a tetramethyl-rhodamin-isothiocyanate-conjugated (TRITC) swine antirabbit antibody (Dako) diluted 1:40, or with biotinylated swine antigoat, -mouse, and -rabbit multilink antibodies (Dako) diluted 1:50, followed by tetramethyl-rhodamin-isothiocyanate-conjugated streptavidin (Immunotech) diluted 1:200, and mounted in fluorescent-mounting medium (Dako). Normal rabbit serum (Dako) or mouse IgG1 (Immunotech) were used as isotype-matched controls. Confocal laser scanning microscopy and fluorescence analysis were performed using a Sarastro CLSM1000 confocal microscope (Molecular Dynamics, Sunnyvale, CA). Excitation was obtained by an argon laser filtered at 514 nm, which ensures low background light and TRITC emission. The selected dichroic filter was DF 530 nm; to avoid excitation noise at emission, fluorescence acquisition was performed using a 550-nm high-pass filter in 256 × 256 or 512 × 512 pixel matrices, and a 50-µm pinhole size. Laser power was set to 25 mW and the PM detector was set at values ranging from 778 to 1078 V (sensitivity adjustments). Images at 0.25 µm pixel size were obtained at × 40 magnification, 1.0 numerical aperture, and analyzed as reported.40 Also, visualization of images in pseudocolor provided a supplementary tool for interpretation when the signal was low. The blue representation of corresponding values emphasized legibility of images.Electron microscopy Culture day 8 cells were washed and fixed for 18 hours with 2% glutaraldehyde in cacodylate buffer, then with 1% osmium tetroxyde, and embedded in epoxy medium after dehydration through a graded series of ethanols. Ultrathin sections were poststained with uranyl acetate and lead citrate and examined on a JEOL 1200EX electron microscope (CMEABG, Université de Lyon, France). A minimum of 100 to 150 cells of each population were analyzed for their morphology and the presence of Birbeck granules (BGs).Mixed leukocyte reaction assay Responder allogeneic adult T lymphocytes enriched to more than or equal to 85% as described41 (105 cells per well in 96-well U-bottomed culture microplates; Costar) were cultured for 6 days with 0.1 to 5 × 103 stimulating cells in RPMI 1640, 10% heat-inactivated AB serum, 1% glutamine, 1% antibiotics. Stimulating cells were day 10 mature DCs from sorted CD45RA+CD7,
CD45RA+CD7+, or bulk CD34+ HPC
cultures. Incorporation of [3H]thymidine (0.037 MBq [1
µCi] per well; Amersham, Amersham, UK) was assessed by
18-hour pulse. Results are mean counts per minute (cpm) of triplicates.
Detection of cytokines in culture supernatants CD1a+CD14and
CD1aCD14+ precursors were sorted on day 5 from cultures of CD34+CD7+CD45RA+
or CD34+CD7CD45RA+ HPCs and
cultured for another 48 hours under the standard condition plus IL-4.
Immature CD1a+83 DCs (0.5 to
1 × 105 cells/mL) were then recovered, washed, and
cultured for 72 hours with CD40LT (0.5 µg/mL, Immunex) before
supernatants were collected. Because only limited DC numbers were
available, cytokine levels were normalized according to total viable
cell counts, and expressed as picograms per 105 cells.
Cytokines were also assayed in day 3, 5, and 7 supernatants of mixed leukocyte reaction (MLR) (ratio: 5 × 103 DCs per 105 T lymphocytes). Alternatively, MLR day 7 T lymphocytes (2.5 × 105 cell/mL) were restimulated for 48 hours with immobilized CD3 and CD28 mAbs (Immunotech) before supernatants were collected. Supernatants were kept at Statistics Results are shown as means ± SD of data. Statistical analyses were performed with the paired Student t test or by analysis of covariance.
Phenotypic characterization of cord blood CD34+ hematopoietic progenitor cells We first examined markers that distinguish myeloid from lymphoid HPCs (CD13, CD33, CD44, CD45RA, CD64, CD86 vs CD2, CD3, CD7, CD10, CD19),14,33,42-48 or immaturity markers (CD38, CD90/Thy1): 17% ± 4% of CD34+ HPCs (n = 17) expressed CD7, 12% ± 4% and 27% ± 4% were CD10+ or CD45RA+, respectively, whereas less than 1% were CD2+, CD3+, or CD19+. Colabeling for CD34, CD38, and CD7 or CD10 showed that CD34+CD7+ HPCs expressed intermediate to high CD38 levels, whereas CD34+CD10+ HPCs were mainly CD38loCD90/Thy1+ and thus corresponded to more immature cells (data not shown). According to CD45RA and CD7 expression, 3 CD34+ HPCs populations were distinguished (Figure 1): these CD7+CD45RA,
CD7+CD45RA+, and
CD7CD45RA+ cells represented 11% ± 3%,
5% ± 2%, and 22% ± 4% of HPCs, respectively, with no
difference being noted whether they were analyzed in enriched CD34+ cells or among bulk mononuclear cells (data not
shown). Of note, none of these populations expressed membrane or
cytoplasmic CD3 (data not shown). CD13, CD33, and CD44 were expressed
by more than or equal to 95% of CD34+ HPCs independent of
CD7 or CD45RA, but CD45RO, CD64, and CD86 were undetectable or barely
detectable. Finally, about 75% of CD34+ HPCs were
CLA+,8 but there was no correlation with CD7
or CD45RA expression.
Natural killer cells differentiate only from CD34+CD7+CD45RA+ hematopoietic progenitor cells We first examined the 3 HPC populations for the capacity to generate NK cells as a marker of their lymphoid potential. Bulk HPCs and sorted subsets were cultured with SCF, IL-2, IL-7, and IL-15, which has proved as efficient for the generation of NK cells from the cord blood HPCs.49 Growth of cells with the NK phenotype was monitored on the basis of CD56 expression.50 Under the conditions used here, only CD7+CD45RA+ HPCs had NK cell potential: On culture days 14 to 21, they yielded 59% ± 6% CD56+CD8cells (n = 7) with NK cytotoxic
activity (data not shown) relative to 1% ± 0.5% for
CD7CD45RA+ and 2% ± 0.1% for bulk HPCs
(Figure 2). Surprisingly, no NK cells were detected in CD7+CD45RA HPC cultures,
even after up to 60 days (data not shown). The CD56 cells
in the cultures were almost exclusively myeloid CD14+/
CD1a+/ cells. CD3 and CD19 labeling was undetectable,
which indicates the lack of T or B lymphocyte differentiation in this
system. At variance with thymic CD34+CD1a
HPCs,34 differentiation of NK cells from
CD34+CD7+CD45RA+ HPCs was dependent
on IL-15 (data not shown).
Differentiation of dendritic cells from
CD34+CD7 , referred to as the
standard condition.37,38,51 This was followed based on
CD1a expression. CD34+CD7CD45RA+
cells (referred to thereafter as single positive [SP]) expanded at
similar levels than bulk HPCs, whereas growth of
CD34+CD7+CD45RA+ (double positive
[DP]) cells was more limited (Figure
3): Starting from 1 × 105
cells on day 0, day 12 cell recovery was
3.7 ± 1.0 × 106 cells for DP cells versus
15.5 ± 5.2 × 106 for SP cells. Nonetheless, the
highest CD1a+ DC percentages were found in DP cell
cultures. There were already 47% ± 6% DCs as early as culture day
5, 78% ± 8% on day 8, and 73% ± 22% on day 12, relative to
33% ± 10%, 37% ± 15%, and 45% ± 17% DCs for SP HPCs
(P  .02; day 8 and day 12). Also, at variance with
thymic CD34+CD1a HPC,34 DP HPC
survival, growth, and differentiation did not require exogenous IL-7,
which was in agreement with the lack of CD127 expression (data not
shown). Finally, because of limited expansion, DP HPCs generated lower
DC numbers than SP HPCs (2.6 ± 1.1 × 106 vs
7.7 ± 4.1 × 106; P = .04; n = 6). Of
note, comparable DC percentages and absolute numbers were found in
cultures of SP and bulk HPCs.
CD34+CD7+CD45RA Altogether, these data indicate that, as with respect to NK cell differentiation, DP HPCs also display the highest capacity to generate DCs under the conditions used here. Dendritic cells and natural killer cells differentiate from common double positive progenitors We thus examined whether DP HPCs comprised common DC and NK cell progenitors. To this end, cells were plated at limiting dilutions and cultured for 3 weeks under the DC or the NK condition before scoring positive wells. DCs and NK cells were then assessed by FACS, based on CD1a or CD56 expression. When cell numbers were too low, DCs were identified according to their morphology after a 48-hour culture in the presence of CD40LT, and NK cells by the binding of CD56 mAb-coated beads. In 2 independent experiments, clonogenicity was estimated as 1:3 and 1:4 under the DC condition, relative to 1:22 and 1:42 under the NK condition, DCs being found in all positive wells cultured under either condition; NK cell progenitor frequency were estimated as 1:23 and 1:76 under the NK condition. As to SP HPCs, almost all wells still scored positive for DCs at the 1 cell per well dilution under the DC condition, and no NK cells were found under the NK condition.Because these data suggested that DP HPCs might contain bipotent DC/NK progenitors, cells were seeded under clonal conditions (0.3 cell per well) and cultured for 3 weeks with cytokines of both the DC and NK conditions to allow for the simultaneous differentiation of both cell types. Again, in 2 independent experiments, typical DCs were found in all cell-containing wells, whereas only 5% and 30% of the clones also yielded NK cells that bound CD56 mAb-coated beads (data not shown). Thus, at least a subset of DP HPCs has the capacity to generate both DCs and NK cells under the conditions used here. Phenotypic characterization of dendritic cells differentiated from single positive and double positive hematopoietic progenitor cells We then examined whether DCs from SP or DP HPC origin differed as to expression of cytokine receptors CD116/GM-CSFR, CD123/IL-3R, CD127/IL-7R, as well as of CD11b, CD11c, CD83, and E-cadherin. Irrespective of origin, DCs were homogeneously CD11b+CD11c+ (data not shown); DCs from SP or bulk HPC cultures were CD123+, expressed intermediate CD116 and CD127 and low to undetectable E-cadherin levels, whereas DCs from DP HPC cultures displayed a more complex pattern: Most CD1ahi DCs were CD123, about 50% being also
CD116lo/, whereas CD1aint/lo DCs were
CD116+CD83+ (Figure
4; data not shown), indicating that these
corresponded to immature and mature DCs, respectively. Of note,
36% ± 13% of DP HPC-derived DCs expressed E-cadherin (E-cadh)
irrespective of CD1a level, relative to 13% ± 9% and 16% ± 9%
(P .03; n = 6) for DCs from SP or bulk HPC cultures.
Because such discrepancies could relate to differences in DCs
differentiation/maturation kinetics as well as origin, IL-4, TGF-1
or CD40LT was added from day 5 onward to SP and DP HPC cultures with
SCF/FL/GM-CSF/TNF-. On day 8, 3 days later, immature
CD1ahi DCs were all
CD83CD123lo/CD127E-cadh+
in cultures with IL-4 or TGF-1, whereas CD40LT induced homogeneous mature
CD1alo/CD83+CD123+CD127+E-cadh+
DCs (data not shown). That no difference was noted between SP-, DP- or
bulk HPC-derived DCs indicates that CD123, CD116, or E-cadherin expression relates to the DC differentiation/maturation stage rather
than origin.
Double positive hematopoietic progenitor cells do not require
exogenous tumor growth factor , with only 10%
to 25% being S100+. In contrast, DCs originating from DP
HPCs were S100+ and approximately 30% were
Laghi. Interestingly, most DCs differentiated from
CD34+CD1a thymic HPCs were also
S100+, and 30% to 40% coexpressed Lag (Figure
5). Because these data argued for an
association between the lymphoid and LC differentiation potential of
the HPCs, we examined whether TGF-1 affected Lag expression by the
DC progeny of DP and SP HPCs. In cultures conducted for 8 days in the
presence of TGF-1, the DCs obtained expressed similar CD1a,
E-cadherin, CD83 and CD116 levels, and 60% to 80% became
Laghi irrespective of their origin (data not
shown).
On electron microscopy (EM) examination, no BGs were observed in DCs
differentiated under the standard condition (data not shown). As to DP
HPC-derived DCs, this discrepancy with Lag labeling was not unexpected
because these are transient structures found only in less than or equal
to 20% of CD34+ HPC-derived Lag+
LCs.52 In the presence of TGF-
Thus, DP and SP HPCs can both generate BG+ LCs, but only SP
HPCs require exogenous TGF- Analysis of the CD1a+CD14 thymic HPCs have shown the
predominance of CD1a+CD14over
CD1aCD14+ precursor-derived
DCs,34 we examined these pathways in SP and DP HPC
cultures by sequential CD1a and CD14 double labeling of cells (Figure
7). Total day 5 CD1a+ cell
percentages were greater among the progeny of DP than of SP HPCs
(47% ± 6% vs 33% ± 10%; P = .07; n = 4), which
was primarily due to the occurrence of
CD1a+CD14+ cells and resulted paradoxically in
2-fold higher total CD14+ cell percentages (51% ± 9%
vs 25% ± 9%; P = .07). However, on day 8, percentages
of CD1a+CD14 (46% ± 8% vs
20% ± 11%; P = .002; n = 8) as well as of
CD1a+CD14+ DCs (32% ± 5% vs 18% ± 6%;
P = .004) were greater in DP HPC cultures, indicating that
most day 5 CD14+ cells had then acquired CD1a. Accordingly,
total CD14+ cell percentages in DP HPC cultures decreased
from about 50% to 15% from day 5 to day 12, whereas they remained
around 25% in SP HPC cultures. Thus, although DP HPCs do not
preferentially differentiate into DCs via
CD1a+CD14 precursors, their
CD1aCD14+ progeny readily differentiate into
DCs under the standard condition. Of note, SP HPC-derived
CD14+ cells failed to down-regulate CD14 even when cultured
for up to 15 to 20 days, unless it was with 10-fold more TNF- (500 instead of 50 U/mL), which indicates that this phenomenon does not
depend on the kinetics of SP and DP HPC growth and differentiation
(data not shown).
We next assessed the effect of M-CSF on SP and DP HPC expansion and
differentiation, based on the assumption that it would up-regulate CD14
expression by c-fms/CD115+ precursor-derived
DCs.16,20 When M-CSF was added to the other cytokines,
only SP HPCs expanded from 1 × 105 day 0 cells to
29 ± 15 × 106 on day 12 relative to
19 ± 11 × 106 without M-CSF (P = .006;
n = 6), whereas DP HPC cultures yielded only
4 ± 2 × 106 cells whether M-CSF was present or not.
In parallel, relative CD1a
These data indicate that most DC precursors generated from SP HPCs are
c-fms/CD115+ whereas DP HPCs yield
CD1a+CD14 Functional characterization of dendritic cells differentiated from single-positive and double-positive hematopoietic progenitor cells We then examined whether DC function could depend on the HPC they derive from by comparing the allogeneic MLR stimulating capacity of mature DCs derived from either SP or DP HPCs. Immature CD1a+ DCs sorted from day 8 standard condition cultures, to which IL-4 was added from day 5, were cultured for 3 more days in the presence of CD40LT, yielding homogeneously mature CD83+ DCs (data not shown). When these were added to allogeneic adult T lymphocytes, the same MLR stimulation level could be reached with about 2-fold less DP HPC-derived than SP HPC-derived DCs (P = .12; NS) (Figure 9A). Because we previously showed that, overall, CD1a+CD14 precursor-derived DCs elicit
stronger MLR than those from CD1aCD14+
precursors,22 we also compared the MLR-stimulating
capacity of mature CD1a+CD14 and
CD1aCD14+ precursor-derived DCs from SP and
DP HPC cultures. Day 5 sorted precursors were cultured as above for 5 more days under the standard condition plus IL-4, and CD40LT for the
last 72 hours. On average, CD1a+CD14
precursor-derived DCs from DP HPC cultures stimulated the MLR 3-fold
more efficiently than their CD1aCD14+
counterparts, both eliciting 3- to 6-fold stronger reactivity than
either type of DC from SP HPC cultures (Figure 9B).
This led us to examine the cytokine profile of allogeneic T lymphocytes
cocultured with the different types of DCs. Only IFN- When we assessed cytokine production in culture supernatants of the DCs
(Figure 10), both
CD1a+CD14
Thus, DP HPC-derived DCs display limitedly, albeit significantly, higher T-cell stimulatory and cytokine production capacities than their SP HPC-derived counterparts.
Analysis of in vitro DC differentiation pathways from discrete HPC
populations with defined developmental characteristics, eg, lymphoid or
myeloid potential, represents an attractive strategy to compare DCs
from different origins and, hence, to approach characterization of
so-called lymphoid DCs. Because cord blood is rich in HPCs spanning
from pluripotent to committed lineage-restricted populations, we
examined CD34+ HPCs for markers of lymphoid or myeloid
potential. Only CD45RA and CD7 appeared relevant in this respect in
that they permitted to individualize 3 CD34+ HPC
populations, defined as CD7+CD45RA Also, at variance with the thymic HPCs, cord blood DP HPCs
differentiate into DCs via both CD1a+CD14 In the last part of this report, we examined whether DCs that
differentiate from DP or SP HPCs could differ regarding their phenotype
and function. DCs from both DP and SP HPC cultures actually coexpressed
CD11b and CD11c myeloid markers, whereas they differed regarding
markers such as E-cadherin (which was low or undetectable in DCs from
SP cultures), CD116, CD127, or CD123, but none of these was really
discriminant. Confocal microscopy examination of cells cultured with
IL-4 showed that the immature DC progeny of SP and DP HPCs differed as
to expression of LC markers S100 and Lag; whereas DCs that originated
from DP or from thymic CD34+CD1a Altogether, our results provide evidence that the DCs that differentiate from cord blood DP CD7+CD45RA+ HPCs represent an original population regarding their developmental pathways and function. It may thus be possible that these DCs are rather dedicated to adaptative immunity, whereas DCs of myeloid origin would be more involved as an interface between the innate and adaptative immune systems.16,30,59
We gratefully acknowledge the help of the following colleagues and companies: Pr. J. Milliez, service de Gynécologie-Obstétrique, hôpital Saint-Antoine (Paris, France) for the gift of cord blood samples; Dr E. Thomas (Immunex, Seattle, WA), for the gift of recombinant Flt3 ligand and CD40LT; Schering Plough (Kenilworth, NJ), for the gift of recombinant GM-CSF and IL-4; Amgen (Thousand Oaks, CA), for the gift of recombinant SCF; Drs L. J. Picker and P. R. Bergstresser (Southwestern Medical Center, Dallas, TX), for the gift of anti-CLA mAb HECA-452; Drs F. Furukawa (Hamamastu University School of Medicine, Handacho, Hamamastu, Japan), K. Yoneda, and S. Imamura (Faculty of Medicine, Kyoto, Japan), for the gift of the anti-Lag mAb; and Catherine Durieu (ESA 7087, hôpital Pitié-Salpêtrière, Paris, France) for cell sorting.
Submitted February 25, 2000; accepted August 3, 2000.
Supported by the Agence Nationale de Recherche contre le SIDA, the Association de Recherche contre le Cancer, the Comité de Paris de la Ligue Nationale contre le Cancer, and the Association pour la Recherche sur les Déficits Immunitaires Viro-Induits (Paris, France).
B.C. and S.C. contributed equally to this work.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Bruno Canque, Laboratoire d'Immunologie, CERVI, hôpital de la Pitié-Salpêtrière, 83 Blvd de l'Hôpital, 75651 Paris Cedex 13, France; e-mail: b_canque{at}club-internet.fr.
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R. Haddad, P. Guardiola, B. Izac, C. Thibault, J. Radich, A.-L. Delezoide, C. Baillou, F. M. Lemoine, J. C. Gluckman, F. Pflumio, et al. Molecular characterization of early human T/NK and B-lymphoid progenitor cells in umbilical cord blood Blood, December 15, 2004; 104(13): 3918 - 3926. [Abstract] [Full Text] [PDF]
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K. Talvensaari, E. Clave, C. Douay, C. Rabian, L. Garderet, M. Busson, F. Garnier, D. Douek, E. Gluckman, D. Charron, et al. A broad T-cell repertoire diversity and an efficient thymic function indicate a favorable long-term immune reconstitution after cord blood stem cell transplantation Blood, February 15, 2002; 99(4): 1458 - 1464. [Abstract] [Full Text] [PDF]
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Top
Abstract
Introduction
2 × 106 U/mg), and 20 ng/mL IL-15 (specific activity greater than or equal to
2 × 106 U/mg) (all from Valbiotech, Paris, France).
Cultures were maintained for up to 2 months with half change and
cytokine addition every 3 to 4 days. CD56+CD8
20°C until used. IL-1
, and
TNF-
): DCs differentiated from CD1a+CD14
): DCs differentiated
from CD1a