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*
First Department of Internal Medicine and
First Department of Pathology, Kansai Medical University, Moriguchi, Osaka, Japan;
Department of Zoology, Kyoto University, Sakyo, Kyoto, Japan;
Cellular Technology Institute, Otsuka Pharmaceutical Co., Tokushima, Japan; and
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Schering-Plough, Laboratory for Immunological Research, Dardilly, France
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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, but not GM-CSF
receptors. After culture with IL-3, fraction 3 acquired the
characteristics of mature DCs; however, the expression of CD62L (lymph
node-homing molecules) remained unchanged, indicating that fraction 3
can be a precursor pool for previously described plasmacytoid T cells
in lymphoid organs. Strikingly, the
CD1a+/CD11c+ DCs (fraction 1) quickly acquired
LC characteristics when cultured in the presence of GM-CSF + IL-4 +
TGF-
1. Thus, E-cadherin, Langerin, and Lag Ag were expressed within
1 day of culture, and typical Birbeck granules were observed. In
contrast, neither CD1a-/CD11c+ (fraction 2)
nor CD1a-/CD11c- (fraction 3) cells had the
capacity to differentiate into LCs. Furthermore, CD14+
monocytes only expressed E-cadherin, but lacked the other LC markers
after culture in these cytokines. Therefore,
CD1a+/CD11c+ DCs are the direct precursors of
LCs in peripheral blood. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Hemopoietic progenitors that differentiate into various types of DCs
are present in the cord blood (15, 16), bone marrow
(17), and peripheral blood (18, 19). At least
two separate developmental pathways of DCs from
CD34+ progenitors have been shown (16, 19): one pathway involves
CD1a+/CD14- cells with the
features of epidermal LCs. The other pathway includes a
CD1a-/CD14+ intermediate
that can differentiate into monocyte-derived or dermal DCs that lack LC
markers. GM-CSF and TNF- are required in both pathways, but TGF-1
is found to be important for the formation of Birbeck granules and Lag
Ag of LCs (20, 21, 22). It has also been found that peripheral
blood monocytes cultured with a combination of GM-CSF, IL-4, and
TNF- differentiate into mature DCs (23), and that LCs
may develop if TGF-1 is added (24). The identification
of DC intermediates in the peripheral blood is, therefore, important,
because it is thought that most blood DCs are either en route from the
bone marrow to peripheral tissues, or from nonlymphoid tissues to the
regional lymph nodes and spleen.
Previously, DCs were purified from the peripheral blood after a 1- to 2-day culture period. These DCs were morphologically, phenotypically, and functionally different from the DCs present in the circulation (25, 26). Recently, DCs have been isolated from the peripheral blood by immunoselection methods without culture. DCs thus prepared are not the morphologically typical DCs, and, interestingly, there are two subsets with distinct phenotype (27, 28, 29). The nature, origin, and commitment of these DCs require further analysis. In the present study, we have isolated fractions of blood DCs using magnetic beads and a multicolor sorting system. We have distinguished three distinct fractions by extensive surface marker analyses, and have examined their functions and developmental capacities. During these investigations, we have identified a distinct subset of DCs that quickly differentiates into LCs.
Strunk et al. have reported that, in the peripheral blood, only
CD34+ progenitors expressing the skin-homing
receptor differentiate into LCs (after 10- to 18-day culture)
(19). It has recently been reported that, in the presence
of GM-CSF, IL-4, and TGF-1 for 6 days, human peripheral blood
monocytes (CD14+ cells) can differentiate into
LCs characterized by the expression of CD1a, E-cadherin, Lag Ag, and
Birbeck granules (24). We show in this study, however,
that the most direct precursors of LCs in human blood are a distinctive
subset of
CD1a+/CD11c+/CD14-
DCs.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The following culture medium was used throughout experiments:
RPMI 1640 supplemented with 2 mM L-glutamine, 100 U/ml
penicillin, 100 µg/ml streptomycin, and heat-inactivated 10% human
AB serum (ICN Biomedicals, Aurora, OH). Monocyte-conditioned medium was
prepared according to Reddy et al. (30). Briefly, blood
mononuclear cells were layered onto human 
-globulin-coated
plates for 1 h, and nonadherent cells were washed away. The
dish-adherent cells were incubated in fresh medium for 24 h. The
supernatant was harvested and frozen at -20°C until use. The
following recombinant human cytokines were purchased from Boehringer
Mannheim (Indianapolis, IN): GM-CSF (used at a concentration of 100
ng/ml), TNF- (2.5 ng/ml), TGF-1 (1 ng/ml), IL-3 (10 ng/ml),
and IL-4 (50 ng/ml). The sources of mAb used in our studies are listed
in Table I
. FITC-, PE-, PerCP-, PE Cy5-,
or biotin-labeled isotype-matched controls were purchased from Becton
Dickinson (Sunnyvale, CA) or PharMingen (San Diego, CA). When unlabeled
mAb or biotinylated mAbs were used as the primary reagents, PE-labeled
goat anti-mouse IgG F(ab')2 (Jackson
ImmunoResearch, West Grove, PA), FITC-labeled goat anti-mouse IgG
F(ab')2 (Becton Dickinson), FITC-labeled
polyclonal anti-rat Ig (PharMingen), or tri-color-labeled
streptavidin (streptavidin-RED670; Life Technologies, Glasgow, U.K.)
were employed as secondary reagents. The mAbs against Langerin (DCGM4)
(31) and Lag-1 (32) were used to determine
the differentiation into LCs.
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Enrichment and analyses of peripheral blood DCs
PBMC were isolated by Lymphoprep (Nycomed Pharma, Oslo, Norway) gradient centrifugation of heparinized blood (50–60 ml) obtained from each healthy volunteer. PBMC were incubated with anti-CD3 (HIT3a) and anti-CD14 (M5E2) mAbs (both from PharMingen) for 30 min on ice, and cells binding these mAb were removed using sheep anti-mouse Ig-coated magnetic beads (M-450; Dynal, Oslo, Norway). CD3-/CD14- cells were further incubated with CD4-conjugated microbeads (Miltenyi Biotec., Bergisch Gladbach, Germany), and the CD4+ cells were then enriched through Mini MACS magnetic separation column (Miltenyi Biotec.). Using this protocol, the percentage of DCs (<1% of total PBMC) increased to 20–50%, depending on the individuals (n > 30). The resultant DC-enriched population (CD4+/CD3-/CD14- cells) was stained with PE-labeled anti-CD11c (LeuM5), FITC-labeled anti-CD1a (BB-5), PerCP-labeled HLA-DR (L243), and a mixture of biotinylated mAbs against lineage markers (CD3; M2AB, binding to a different determinant from that recognized by the previous anti-CD3 mAb, CD7; G34, CD14; UCHM1, binding to a different determinant from that recognized by the previous anti-CD14 mAb, CD16; 3G8 and CD19; HIB19), followed by RED613-streptavidin (Life Technologies). The stained cells were analyzed (or sorted for cytologic assay, T cell proliferation assay, and analyses of endocytosis) by an EPICS ELITE flow cytometer (Coulter, Hialeah, FL). Consequently, three phenotypically distinct fractions of DCs were found: CD1a+/CD11c+/lin-/DR+ (fraction 1), CD1a-/CD11c+/lin-/DR+ (fraction 2), and CD1a-/CD11c-/lin-/DR+ cells (fraction 3).
Purification and characterization of DC fractions
The DC fractions were obtained also by a two-color FACStar cell
sorter (Becton Dickinson), as follows. 1) Purification of
CD1a+/CD11c+ DCs (fraction
1): after staining the DC-enriched population with PE (or FITC)-labeled
mAbs against lineage markers plus FITC (or PE)-labeled anti-CD1a
mAb, CD1a+/CD11c+ DCs were
sorted as CD1a+/lin- cells
(note that all CD1a+/lin-
DCs are quite comparable with
CD1a+/CD11c+/lin-
DCs when analyzed after staining with anti-CD11c mAb). 2)
Purification of
CD1a-/CD11c+ (fraction 2)
and CD1a-/CD11c-
(fraction 3) DCs: the DC-enriched population was stained with
FITC-labeled mAbs against lineage markers and CD1a plus PE-labeled
anti-CD11c (LeuM5), and the
lin-/CD1a-/CD11c+
and
lin-/CD1a-/CD11c-
fractions were then collected using a FACStar as fractions 2 and 3,
respectively. These fractions collected using the FACStar were
phenotyped. Purity of the sorted cells was always greater than 96% by
reanalysis using a FACScan (Becton Dickinson). The sorted fractions,
thus prepared, were then stained with PerCP-labeled HLA-DR. The cells
bearing HLA-DR (DR+) were gated as DCs, and
further analysis was conducted using a FACScan after staining with a
panel of mAbs (listed in Table I
) conjugated with FITC or PE. Before
staining, the cells were incubated with an excess amount of unlabeled
polyclonal human or mouse Ig to block nonspecific binding of
labeled mAbs.
Culture of DCs
The sorted DCs were cultured in 96-well flat-bottom tissue
culture plates at 1 x 105 cells/well in
medium supplemented with monocyte-conditioned medium (final
concentration 50% v/v) for 1–6 days. In the culture of
CD1a-/CD11c- DCs
(fraction 3), medium supplemented with IL-3 was used. In some
experiments, a mixture of cytokines such as GM-CSF + TGF-1, GM-CSF +
IL-4, GM-CSF + TNF-, GM-CSF + TNF- + TGF-1, or GM-CSF + IL-4 +
TGF-1 was used. Half of the medium was removed and replenished every
2 days.
Analysis of endocytosis
To assess endocytosis of DC fractions, FITC-dextran (Polysciences, Warrington, PA) was used according to the method described previously (33). Briefly, the cells were incubated with 0.1 mg/ml FITC-dextran at 37°C for 15 or 45 min. The results were displayed as mean fluorescence intensity after subtracting the background in which cells were incubated with FITC-dextran at 4°C.
Cytologic assays
Cells were cytocentrifuged onto a slide and stained with May-Giemsa solution, or by mAb to CD68 or Lag, then visualized using the ABC method using DAKO LSAB (labeled streptavidin biotin) kit plus hematoxylin and diaminobenzidine.
For electron-transmission microscopy, cells were fixed with 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) and postfixed with 1% OsO4. After dehydration with graded ethanol, they were embedded in Epon. Ultrathin sections were stained with lead citrate and uranyl acetate, and studied using a Hitachi H-600a electron microscope (Hitachi, Ibaragi, Japan).
Mixed leukocyte reaction
Freshly prepared blood DCs, cultured DCs (in the presence of
GM-CSF and IL-3), or blood monocytes (purified by magnetic beads as
CD14+ cells) were irradiated at 15 Gy, and
graded doses were then added to 2 x 105
allogeneic T cells (collected by magnetic beads as
CD3+ cells) in 96-well flat-bottom culture plates
for 6 days. The cells were pulsed with 1 µCi of
[3H]TdR during the last 16 h of the
culture period.
Cell cycle and viability assays
For cell cycle analyses, 105 cells were suspended in 70% ethanol at 4°C for 1 h, followed by resuspension in 500 µl PBS, 250 µl RNase (1 mg/ml; Sigma, St. Louis, MO), and 250 µl propidium iodide (PI) (100 µg/ml; Molecular Probes, Eugene, OR). Cell viability was determined using Annexin V-FITC Apoptosis Detection Kit (Genzyme, Cambridge, MA). The stained cells were analyzed with a FACScan.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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DC-enriched populations prepared from the peripheral blood
(CD4+/CD3-/CD14-
cells) were four-color analyzed after staining with PE-labeled
anti-CD11c, FITC-labeled anti-CD1a, PerCP-labeled HLA-DR, and a
mixture of biotinylated mAbs against lineage markers, followed by
RED613-streptavidin.
lin-/HLA-DR+ cells (DCs)
(Fig. 1A) were of medium cell
size (between lymphocytes and monocytes) by light scatter profiles. Two
distinct populations of
lin-/DR+ were observed
with respect to the expression of CD11c (Fig. 1B).
Furthermore, CD1a (detected by mAb clone BB-5)-positive cells were
newly observed as a blood DC fraction (Fig. 1B). Therefore,
three fractions of DCs were identified with the phenotypes of
CD1a+/CD11c+/lin-/DR+
(fraction 1),
CD1a-/CD11c+/lin-/DR+
(fraction 2), and
CD1a-/CD11c-/lin-/DR+
cells (fraction 3). CD1a has been detected only on epidermal LCs
(8, 34) and dermal DCs (10) in vivo (or on
DCs derived from CD34+ progenitors (15, 16) or monocyte-derived DCs (23) in the presence of
cytokines in vitro), but not on freshly isolated blood DCs (27, 28). CD1a+ DCs were also observed when a
different anti-CD1a mAb (BL-6) was used (data not shown). The
overall intensity of CD1a expression, however, was lower than that on
CD34+ progenitor-derived DCs (data not shown)
when determined by either BB-5 or BL-6 Ab.
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C shows the sorting gate of fraction 1 as
lin-/CD1a+ cells (note
that all CD1a+ cells are
CD11c+, as shown in Fig. 1B), and R2
and R3 in Fig. 1D represent the sorting gates of
lin-/CD1a-/CD11c+
cells (fraction 2) and
lin-/CD1a-/CD11c-
cells (fraction 3), respectively. Morphology of peripheral blood DC fractions
Freshly prepared or cultured (for 3 days in monocyte-conditioned
medium or IL-3-supplemented medium) DC fractions were examined by light
and electron microscopes (EM). Freshly sorted
CD1a+/CD11c+ DCs (fraction
1) (Fig. 2, A and
B) and
CD1a-/CD11c+ DCs (fraction
2) (Fig. 2, D and E) were found to be monocyte
like, characterized by short cell processes, indented nucleus, high
nuclear-cytoplasmic ratio, and few cytoplasmic dense bodies and
vesicles, in contrast to typical DCs. After 3-day culture, both
fractions showed typical DC features such as long cell processes, large
cell size (Fig. 2, C and F), and a prominent
tubular-vesicular system (data not shown).
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, G and
H). These cells correspond to plasmacytoid T cells
within the T cell zone of lymphoid organs, as recently characterized by
Grouard et al. (14). After culture with IL-3, they showed
a typical DC morphology (Fig. 2I) with marked decreases in
rough endoplasmic reticulum (data not shown), and this process was
dependent on the presence of IL-3. No Birbeck granules were found in
either of the three presently identified fractions of DCs before or
after culture with monocyte-conditioned medium or IL-3 (data not
shown). Phenotype of peripheral blood DC fractions
The fluorometrical characterization of the three fractions of DCs
was next conducted. The surface phenotype of the three DC fractions
before and after culture is highlighted in Figs. 3 and 4.
The common features of these three fractions are as follows: none of
the DC fractions expressed lineage markers for T cells (CD8, TCR,
Thy-1/CD90), B cells (CD10, CD20), NK cells (CD56), monocytes (CD14),
granulocytes (CD15, CD35), or erythrocytes (glycophorin A) before or
after culture. In addition, they possessed neither CD27 and CD30 (TNF
receptor families) nor CD70 (ligand for CD27). IL-1R type I (CD121a),
G-CSFR (CD114), IL-2R-chain (CD122), and IL-6R (CD130) were
negative on all freshly prepared DC fractions (data not shown). All
fractions clearly expressed MHC class II, CD4, CD9, CD36, CD38, CD45,
and adhesion molecules (CD11a, CD15s, CD18, CD29, CD44, CD49d, CD50,
and CD54). Costimulatory molecules (CD80, CD86, and CD40) were dimly
expressed on all fractions (fraction 3 little expressed CD40), and were
up-regulated after culture. Activation molecules CD25 and the
DC-associated marker CD83 were also detected on all fractions after
overnight culture, but not on freshly isolated cells. In addition, CLA
(a skin-homing molecule (35), which is constitutively
expressed on LCs (36)), and L-selectin (CD62L, known to be
a lymph node-homing molecule (37)) were present on all
fresh DC fractions. The characteristic features of each fraction are as
follows.
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receptors), and cytokine receptors (GM-CSFR (CD116) and IL-3R
(moderately positive)). Furthermore, they dimly expressed CD5 and
CD11b. After culture, CD11b was up-regulated, and CD32/64 was
down-regulated. CD62L became undetectable during culture in contrast to
freshly prepared cells. On the basis of the expressions of CD2, CD9,
CD11b, CD11c, CD13, CD32, CD33, CD64, and GM-CSFR, fraction 1 is
thought to be closely associated with the monocyte lineage. Actually,
the phenotype of fraction 1 was quite similar to that of blood
monocyte-derived DCs and that of CD34+
progenitor-derived DCs (but not LC type) (16).
Fraction 3.
In contrast to fraction 1,
CD1a-/CD11c- DCs
expressed dimly or little CD33, CD13, CD1c, CD2, CD49e, CD45RO, CD32,
CD64, but were significantly positive for CD45RA. In comparison with
the other two fractions, they were highly positive for CD4, but
moderately positive for HLA-DR/DQ. They also possessed IL-3R
(brightly positive), but not GM-CSFR. Fraction 3 brightly expressed
intracytoplasmic CD68, but the other two fractions were only weakly
positive (data not shown). Therefore, fraction 3 is closely related to
previously reported plasmacytoid T cells, not only morphologically but
also according to phenotypical features. Furthermore, freshly prepared
fraction 3 had no CD95. However, CD95 was induced during 3-day culture
with IL-3, whereas the expression of CD62L remained unchanged. When
compared with culture in medium alone, this fraction exhibited
increased levels of CD83 and HLA-DR/DQ in the presence of IL-3 (data
not shown).
Fraction 2. CD1a-/CD11c+
DCs were a minor population, and uniformly expressed CD11c and CD33,
like fraction 1. However, they were found to be heterogeneous in the
expression of some surface molecules such as CD2, CD13, CD45RA, CD45RO,
IL-3R, and GM-CSFR, although they were morphologically homogeneous.
Fraction 2, in contrast to fraction 1, had no CD1a, CD1c, CD11b, or
CD64. However, they had CD11c, CD33, CD13, and GM-CSFR, indicating that
they were monocyte-lineage cells. Based on the expression of the CD2
molecule, fraction 2 could be divided into two minor subfractions:
CD2high and CD2low DCs.
After purifying these two populations and staining them with a panel of
mAbs, we found that the former were
CD13low/CD45RA+/CD45RO-/GM-CSFR+
(bright)/IL-3R+ (bright). On the other hand,
CD2low DCs were
CD13high/CD45RA+
(dim)/CD45RO+/GM-CSFR+
(dim)/IL-3R+ (dim). Therefore, both
subpopulations in fraction 2 seem to be related (but not completely
matched) to fractions 1 and 3 on the basis of the expression of
surface molecules. The physiological features of the heterogeneous
fraction are unknown at present.
Finally, CD1a molecules were not induced on the
CD1a- cells even when both fraction 2 and
fraction 3 were cultured with monocyte-conditioned medium or
IL-3-supplemented medium, respectively (Fig. 4), or with any
combinations of cytokines examined (GM-CSF, IL-3, TNF-, and
TGF-1) (data not shown).
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The stimulatory activity for allogeneic T cells was compared among
the three DC fractions, as shown in Fig. 5. The order in the stimulatory activity
of freshly prepared fractions was as follows: fraction 1 =
fraction 2 > fraction 3, being dependent on the order of the
expression of MHC class II molecules. The stimulatory activities
became stronger after culture for 3 days with GM-CSF plus IL-3
(note that fractions 1 and 2 are dependent on GM-CSF, and fraction
3 is dependent on IL-3), indicating that they become mature
DCs.
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,
freshly isolated fractions 1 and 2 had a potent ability to endocytose
FITC-dextran during a 45-min incubation compared with
CD34+ progenitor-derived mature DCs, although
this capacity was less than monocytes. The endocytic activity
of fractions 1 and 2 decreased after culture for 3 days with
monocyte-conditioned medium (data not shown). Fraction 3, which had
been freshly isolated or cultured (with IL-3), showed little
FITC-dextran uptake.
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It should be noted that no Birbeck granules were found in
CD1a+/CD11c+ DCs (fraction
1) before or after culture with monocyte-conditioned medium. To further
characterize fraction 1, we compared the effects of combinations of
cytokines (GM-CSF + TGF-1, GM-CSF + IL-4, GM-CSF + TNF-, GM-CSF +
IL-4 + TGF-1, and GM-CSF + TNF- + TGF-1) on their maturation
and differentiation. The expression of E-cadherin, the LC-restricted
Langerin (detected by mAb DCGM-4), and Lag Ag was kinetically examined.
In the presence of GM-CSF + IL-4 + TGF-1, E-cadherin and Langerin
were detected (Fig. 7A)
already after 1 day of culture, and intracellular Lag was also slightly
positive both in FACS (data not shown) and cytochemical (Fig. 7B) examinations. Expression of the LC-specific molecules
increased at 3 days, as shown in Fig. 7. Finally, fraction 1 showed DC
morphology, and typical Birbeck granules were observed 6 days after
culture (Fig. 8). In addition, the
intensity of CD1a expression increased concomitantly (Fig. 7A).
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1, but not with any cytokine
alone nor any combinations tested. Of importance, neither fraction 2
nor 3 had the capacity to differentiate into LCs when cultured with
GM-CSF + IL-4 + TGF-1 or other cytokines tested (GM-CSF, IL-3,
TNF-, and TGF-1) (data not shown). Furthermore, induction of
LC-specific molecules was not observed in CD14+
monocytes during culture with the same cytokine combination (GM-CSF +
IL-4 + TGF-1). On day 6, E-cadherin and CD1a became positive, as has
been reported by Geissmann et al. (24); however, the more
sensitive Lag and Langerin markers were not detected in our culture
condition (Fig. 7). Furthermore, when we examined the frequency of
dividing cells (evaluated by PI staining) and viable cells (evaluated
by PI and annexin V staining) before and after the culture, the
percentages of dividing cells were less than 1% (before and after 1-,
3-, and 6-day culture), and the percentages of viable cells were more
than 90% throughout the culture period (data not shown). In addition,
15 Gy irradiation did not alter LC differentiation from fraction 1
(data not shown). These findings clearly demonstrate that 1) fraction 1
has the capacity to differentiate into LCs without cell proliferation,
and 2) fraction 1 is the direct/immediate precursor of LCs, compared
with monocytes.
As shown in Fig. 9, 3 days after culture
in the presence of GM-CSF, IL-4, and TGF-1, fraction 1 DCs expressed
higher intensities of cutaneous lymphocyte-associated Ag and CD1a, with
lower HLA-DR, CD54, CD40/CD80/CD86, and CD83 (mostly negative, a few
positive) than those cultured with monocyte-conditioned medium.
Activation-associated markers, CD25 and CD71, were not induced, and CD2
was down-regulated in contrast to the cells before culture.
Furthermore, CD23 (Fc
RII) was induced on fraction 1 under this
condition (but not with monocyte-conditioned medium), as reported by
Schmitt et al. (38) on LCs. These findings suggest that
the combination of IL-4, GM-CSF, and TGF-1 is essential for
differentiation of fraction 1 into LCs rather than terminal maturation
and/or activation.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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As previously reported, less mature APCs in the blood, with little
expression of CD11c and CD33, have been speculated to be on the way
from the bone marrow to nonlymphoid tissues such as the skin, and more
mature potent APCs bearing CD11c and CD33 are thought to be en route to
the lymph nodes from the skin (27). When freshly isolated,
these DCs display immature dendritic morphology and express CD4
(5, 27), but not CD83 (26, 39), in agreement
with our results (Figs. 2 and 3). Notably, DCs in the blood have been
found within a CD1a-negative fraction (26, 27, 28, 39).
However, we have identified one fraction of DCs as CD1a low-positive
cells (fraction 1). The difference in detection of CD1a between our
present results and previous studies could be due to the following: 1)
in previous studies, T cells were depleted by SRBC-rosette formation,
and this caused the depletion of some DCs bearing CD1a because this DC
population coexpresses CD2 (binding site of SRBC) (5, 26, 27); 2) part of the CD1a+ DC fraction
coexpressing CD11b might be depleted along with myelomonocyte-lineage
cells when anti-CD11b mAb was used in immunoselection (29, 39, 40); 3) the level of CD1a expression on this fraction is
much lower than that on the CD34+
progenitor-derived DCs or monocyte-derived DCs; therefore,
CD1alow+ DCs were classified as CD1a-negative
cells; in this context, Brown et al. reported that CD1a was weakly
expressed on a small amount of blood DCs (41); and 4)
different anti-CD1a mAbs employed in detection, this being
considered to be a main reason. In our study, a difference in the
intensity of CD1a was actually observed when two anti-CD1a mAbs
(clones BB-5 and BL-6) were compared. The
CD11c+/CD33+ DC subset in
the blood that O’Doherty et al. (27) and Thomas et al.
(28) previously identified might contain fraction 2
(CD1a-/CD11c+ DCs) and a
part of fraction 1
(CD1a+/CD11c+ DCs). In
their studies, CD1alow+ DCs might be classified
as CD1a-negative cells.
Fraction 1 is morphologically similar to monocytes and, after culture,
the cells resemble monocyte-derived DCs. Furthermore, fraction 1 has
myelomonocyte-associated Ags, and has the capacity to uptake
FITC-dextran. These findings indicate that fraction 1 is a homogeneous
subset, and most likely a CD14-negative monocyte-associated or
monocyte-derived DC. Strikingly, cells in fraction 1, but not fraction
2 or 3, rapidly became cells with typical LC characteristics in the
presence of GM-CSF, IL-4, and TGF-1. E-cadherin (Fig. 7A)
and Lag (Fig. 7B) were detected after short-term culture,
and the newly described LC-specific Ag, Langerin (31), was
concomitantly induced (Fig. 7A). Furthermore, the
differentiation of LCs from fraction 1 was confirmed by the appearance
of Birbeck granules (Fig. 8). DCs bearing CD1a are induced from
CD14+ blood monocytes after culture with GM-CSF
and IL-4 (23, 33). Geissmann et al. (24) have
recently reported that blood monocytes differentiate into LCs after a
6-day culture in the additional presence of TGF-1. We did not
observe acquisition of a complete LC phenotype from
CD14+ monocytes in our culture condition (Fig. 7). Furthermore, the differentiation of LCs from fraction 1 occurred
during a very short-term culture. In addition, the possibility that a
dividing or a distinct subpopulation in fraction 1 selectively
differentiates into LCs was ruled out by the following: there was <1%
of dividing cells, there was a low cell mortality rate, and a low rate
of apoptosis throughout the short-term culture. Therefore, fraction 1
is a distinct direct/immediate precursor of LCs in the peripheral
blood, and might be an intermediate stage between monocytes and
LCs.
LCs are myeloid-lineage DCs that originate from the bone marrow.
However, little is known about their migration/differentiation pathway.
Our results might help elucidate this question, as fraction 1 is the
direct precursor of LCs (in response to GM-CSF + IL-4 + TGF-1) and a
bipotent cell that can be induced to differentiate into non-LCs such as
dermal DCs (in response to monocyte-conditioned medium). This indicates
that common precursors for the LC lineage and the monocyte-associated
DC lineage are present in the peripheral blood. It can be speculated
that part of this fraction migrates to the epidermis to differentiate
into LCs, and that the other migrates to nonlymphoid tissues such as
the dermis and differentiate into dermal DCs. This process might be
dependent upon cytokines produced by regional microenvironments.
Fraction 3 (CD1a-/CD11c-
DCs) is another homogeneous subset in the peripheral blood, and
distinct from fraction 1, based on its morphology, the expression of
lineage-associated Ags, lack of ability to uptake FITC-dextran, and
dependency on IL-3. From our detailed studies on blood DCs, the
features of fraction 3 correspond completely to those of plasmacytoid T
cells in the T zone of the lymphoid organ (found to be close to or
within the high endothelial venule) that could be derived from
CD4+/CD11c- blood DCs
(14). Fraction 3 possesses high levels of the lymph
node-homing molecule; CD62L, and its expression, remains unchanged
during culture, although it is down-regulated on fraction 1 (Figs. 3 and 4). These findings support the possibility that fraction 3 in the
blood may be a precursor pool for plasmacytoid T cells that directly
migrate to the T zone in the lymphoid tissues via the high endothelial
venule. Furthermore, fraction 3 requires IL-3, but not GM-CSF, for cell
maturation and development, whereas typical myeloid-lineage DCs are
known to be dependent on GM-CSF, indicating that fraction 3 is not in
the monocyte-associated DC lineage. Grouard et al. have pointed out
that plasmacytoid T cells in the lymphoid organs are lymphoid-derived
DCs for this reason (14). However, fraction 3 also
expresses several myelo-monocytic markers (CD9; Fig. 3, CD36 and CD68;
data not shown); therefore, it is not evident at the moment whether
they indeed are lymphoid-derived DCs. This issue will require further
studies on these cells. Our results clearly indicate that fraction 3 is
a different lineage from that previously examined in in vitro
differentiation studies of CD34+ progenitors to
LCs or dermal DCs.
Although examination of the minor fraction, fraction 2 (CD1a-/CD11c+ DCs), has been hampered by the limitation of cells collected from the peripheral blood, this fraction has similar features to fraction 1 in their morphology, allostimulatory activity, and the expression of several myelo-monocytic Ags, suggesting that cells in this fraction are monocyte-associated or monocyte-derived DCs, or that they are in an intermediate stage preceding fraction 1. Furthermore, fraction 2 is phenotypically similar to recently reported DCs located in the germinal center (GCDC) (13); therefore, the possibility exists that fraction 2 becomes GCDCs, which have been presumed to be derived from blood CD4+/CD11c+ DCs.
In summary, we have characterized three fractions of DCs in the human peripheral blood using a panel of mAbs. It is noted that fraction 1 has the capacity to become LCs. These results suggest that blood DC fractions (subsets) are different not only in their maturational stages, but also in their lineage or differentiation pathways. Our studies of the DC fractions were conducted using healthy volunteers. Therefore, the findings in this study also offer basic data for the diagnosis of some immune-related diseases. We are now proceeding with experiments related to these points.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Susumu Ikehara, First Department of Pathology, Kansai Medical University, 10-15 Fumizono-cho, Moriguchi City, Osaka 570-8506, Japan. E-mail address:
3 Abbreviations used in this paper: DC, dendritic cell; EM, electron microscope; LC, Langerhans cell; PerCP, peridini-chlorophyll protein; PI, propidium iodide.
Received for publication January 11, 1999. Accepted for publication May 20, 1999.
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), fraction 3
(
), and monocytes (
). Vertical bars represent SDs of triplicate
cultures. Representative data of three experiments are shown.
). Results shown are
representative of three experiments.
20–25% of cultured DCs) by EM
examinations. Original magnification: left, x6,000;
right, x12,000; inset, x20,000.