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Divisions of
*
Immunology, Allergy, and Infectious Diseases and
General Dermatology, Department of Dermatology, University of Vienna Medical School, Vienna, Austria
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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-stimulated CD34+ stem cells. The
CD14+CD1a- progenitors, on the other hand, can
give rise to a separate DC type characterized by its
CD1a+CD11b+CD36+FXIIIa+E-cad-BG-
phenotype (non-LC DC). Although GM-CSF/TNF- are important for both
LC and non-LC DC differentiation, TGF-ß1 is thought to preferentially
promote LC development in vitro and in vivo. However, the hemopoietic
biology of this process and the nature of TGF-ß1-responsive LC
precursors (LCp) are not well understood. Here we show that
CD14+ precursors in the presence, but not in the absence,
of TGF-ß1 give rise to a progeny that fulfills all major criteria of
LC. In contrast, LC development from CD1a+ progenitors was
TGF-ß1 independent. Further studies revealed that CD14+
precursors contain a CD11b+ and a CD11b-
subpopulation. When either subset was stimulated with GM-CSF/TNF-
and TGF-ß1, only CD14+CD11b- cells
differentiated into LC. The CD11b+ cells, on the other
hand, acquired non-LC DC features only. The higher doubling rates of
cells entering the CD14+ LCp rather than the
CD1a+ LCp pathway add to the importance of
TGF-ß1 for LC development. Because
CD14+CD11b- precursors are multipotent cells
that can enter LC or macrophage differentiation, it is suggested that
these cells, if present at the tissue level, endow a given organ with
the property to generate diverse cell types in response to the local
cytokine milieu. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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gives rise to a progeny of
CD1a+, E-cadherin+
(E-cad+), Birbeck granule (BG)-containing cells
with immunostimulatory properties strikingly similar to those of LC
isolated from human skin (3, 4). Subsequent studies have
tried to delineate the phenotype of LC progenitors at their various
states of maturation/differentiation. Around day 6 of in vitro culture
of CD34+ HPC in GM-CSF/TNF--supplemented
medium, CD1a+ cells appear that, upon
prolongation of the culture until days 12–14, develop into typical DC
displaying all the features found in and on epidermal LC (5, 6). Besides this CD1a+ LC precursor,
CD14+CD1a- cells emerge
early during the culture. Apparently, their lineage commitment differs
from that of the CD1a+ cells, as they give rise
to a monocyte/macrophage phenotype when exposed to M-CSF while
differentiating into non-LC DC in the continued presence of GM-CSF and
TNF- (5). Phenotypically, these non-LC DC are
characterized by the abundant expression of factor XIIIa (FXIIIa),
CD1a, CD11b, and CD36 and the virtual absence of E-cad and BG and,
thus, display striking similarities to dermal DC (1, 5, 6, 7, 8).
Although certain stimuli, i.e., stem cell factor and Flt3 ligand
(Flt3-L), amplify the DC differentiation pathways initiated by GM-CSF
and TNF- without any apparent selectivity for LC or non-LC DC
development (9, 10), TGF-ß1 seems to be of unique
importance in LC ontogeny. This is best evidenced by the lack of LC in
TGF-ß1-/- mice
(11) and by the preferential development of
CD1a+BG+ cells in
GM-CSF/TNF--containing, TGF-ß1-supplemented serum-free stem cell
cultures (12).
Although recent studies showed that TGF-ß1 prevents apoptosis of and,
together with GM-CSF/TNF-, acts as a growth factor for myeloid
precursor cells (13), the exact role of TGF-ß1 for the
selective development of LC from cytokine-stimulated
CD34+ HPC remains unknown. In this study we
isolated various well-defined DC precursor populations that emerge
during the culture of GM-CSF/TNF--stimulated
CD34+ stem cells and sought to determine which
cellular phenotype requires the presence of TGF-ß1 for entering the
LC differentiation pathway.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The FITC-conjugated mouse mAbs (clones) used were anti-CD1a (OKT6; Ortho Diagnostic Systems, Raritan, NJ), anti-CD11b (BEARI; Immunotech, Marseilles, France), anti-CD14 (Leu M3; Becton Dickinson, Mountain View, CA), anti-CD36 (OKM5; Ortho Diagnostic Systems), and anti-CD86 (FUN-1; PharMingen, San Diego, CA). PE-conjugated mAbs included anti-CD1a (HI149; PharMingen), anti-CD11b (D12), anti-CD11c (S-HCL-3), anti-CD14 (Leu M3; all from Becton Dickinson), and anti-CD86 (FUN-1; PharMingen). Peridinin chlorophyll protein (PerCP)-labeled anti-HLA-DR (clone L243) was purchased from Becton Dickinson. Purified anti-E-cadherin mAbs were obtained from Transduction Laboratories (mAb clone 36; Lexington, KY) and Immunotech (clone 67A4). Biotinylation of mAbs was performed according to standard protocols. The anti-CD45 mAb (clone MEM-28) and the mAb Lag were provided by Dr. Horejsi (Praha, Czech Republic) and Dr. Imamura (Kyoto, Japan), respectively. Rabbit anti-FXIIIa was obtained from Behring Diagnostics (Marburg, Germany). Appropriate fluorochrome- and biotin-labeled mouse control mAbs were purchased from Becton Dickinson. Nonlabeled IgG1 control mAbs (MOPC-21) were obtained from Sigma (St. Louis, MO). FITC-labeled goat F(ab')2 anti-mouse IgG + IgM was purchased from An der Grub (Kaumberg, Austria). Streptavidin-conjugated to PE (SA-PE) and PE-Cy5 (SA-Cy5) were obtained from Becton Dickinson and Dako (Glostrup, Denmark), respectively. mAbs used in immunodepletion experiments included anti-CD3, anti-CD16, anti-CD19, anti-CD34, anti-CD41, and anti-CD56 (all from Immunotech); anti-CD8 and anti-HLA-DR (both from Becton Dickinson); and anti-CD45RO (from Dako).
Growth factors
Recombinant human growth factors used in this study were GM-CSF
(6 x 106 U/mg; provided by Novartis, Basel,
Switzerland), TNF- (1 x 107 U/mg) and
IL-4 (1 x 107 U/mg; both from Genzyme,
Cambridge, U.K.), Flt3-L (Serotec, Oxford, U.K.), IL-6 (1 x
107 U/mg; Life Technologies, Gaithersburg, MD),
and M-CSF (1.5 x 108 IU/mg; R&D Systems,
Minneapolis, MN). Purified human platelet-derived TGF-ß1 (3.2 x
107 U/mg) and neutralizing anti-TGF-ß mAbs
were obtained from R&D Systems.
Isolation of CD34+ hemopoietic stem cells
Cord blood (CB) was obtained according to institutional guidelines. MNC were prepared by discontinuous density gradient centrifugation (Ficoll-Hypaque; Pharmacia, Uppsala, Sweden) of heparinized CB. CD34+ cells were separated from CB-MNC by a positive immunoselection procedure (CD34 multisort kit; Miltenyi Biotec, Bergisch Gladbach, Germany). Briefly, CB-MNC (2–3 x 108) were incubated with anti-CD34 mAb-coated paramagnetic microbeads for 30 min at 4°C. After several washings, bead-bound CD34+ HPC were isolated on MiniMACS separation columns using a magnet (MiniMACS, Miltenyi Biotec). CD34+ cells (0.5–1 x 106) were recovered at a purity of >95% as determined by immunostaining with a PE-labeled anti-CD34 mAb (HPCA-2; Becton Dickinson) recognizing a CD34 epitope distinct from that bound by the mAb used for immunoselection.
In vitro culture of CD34+ CB-MNC
Purified CD34+ CB-MNC were seeded at a
density of 1–3 x 104 cells/ml in RPMI 1640
containing 2 mM L-glutamine, 125 U/ml penicillin, 125
µg/ml streptomycin, 50 mM ME, and 10% heat-inactivated FCS (all from
Life Technologies). This culture medium was further supplemented with
200 U/ml GM-CSF, 50 U/ml TNF-, and 50 ng/ml Flt3-L. Cells were
cultured in 25-cm2 tissue culture flasks (Costar,
Acton, MA) at 37°C in a humidified atmosphere in the presence of 5%
CO2. On day 6, cells were collected and labeled
either with anti-CD1a FITC/anti-CD14 PE or with anti-CD14
FITC/anti-CD11b PE, and stained cells were resuspended in PBS/5 mM
EDTA to prevent aggregation.
CD1a+CD14-,
CD1a-CD14+,
CD11b+CD14+, and
CD11b-CD14+ cell
populations were sorted using a FACStarPlus flow
cytometer (Becton Dickinson). The purity of the sorted fractions was
always >98%. Sorted cells were resuspended in serum-free X-VIVO 15
medium (BioWhittaker, Walkersville, MD) containing GM-CSF (200
U/ml)/TNF- (50 U/ml) or M-CSF (2500 IU/ml)/IL-6 (200 U/ml). Where
indicated, GM-CSF/TNF--containing cultures were additionally
supplemented with TGF-ß1 (1 ng/ml) or neutralizing anti-TGF-ß
mAbs (20 µg/ml). On day 10, half the culture supernatant was removed
and replaced by fresh medium supplemented with the same cytokines/mAbs
and cytokine/mAb concentrations that had been added to the cultures on
day 6. On day 12 or 14, cells were subjected to further analyses.
Generation of monocyte-derived DC
Monocyte-derived DC were prepared according to protocols described previously (14, 15). Briefly, heparinized peripheral blood of adult donors was subjected to Ficoll-Hypaque density gradient centrifugation, and the resulting PBMC fraction was incubated with mAbs recognizing CD3, CD16, CD19, CD34, CD41, and CD56. After two washings, mAb-bound PBMC were allowed to bind to rat anti-mouse IgG1-coated immunomagnetic beads (MACS, Miltenyi Biotec) and were depleted using a magnetic separator (MiniMACS) according to the manufacturer’s instructions. This procedure routinely resulted in the recovery of CD14+ PBMC at a purity of >90%. Cells thus isolated were cultured in X-VIVO 15 medium containing GM-CSF (800 U/ml) and IL-4 (1000 U/ml) for 7 days. Where indicated, TGF-ß1 was added to the culture medium at a final concentration of 1 ng/ml.
Flow cytometry analyses
Cells (2 x 105) were incubated with the appropriate concentrations of the indicated biotin- and/or fluorochrome-conjugated mAbs for 30 min at 4°C. The binding of biotinylated mAbs was revealed by a consecutive incubation step with SA-PE or SA-Cy5. For the detection of intracellular Ags, cells were fixed and permeabilized before immunostaining using the Fix&Perm reagent (An der Grub). Briefly, 5 x 105 cells were exposed to the formaldehyde-based fixative for 15 min at room temperature and then incubated with mAb Lag- or IgG1 control mAb-supplemented (each 1 µg/ml) permeabilization solution for 15 min at room temperature. The binding of the first step mAbs was revealed by exposing the cells to 3 µg/ml of FITC-labeled goat anti-mouse IgG+IgM F(ab')2. After two rounds of washings, free cell-bound anti-mouse binding sites were blocked by saturating concentrations of an irrelevant IgG1 control mAb, and cells were exposed to anti-CD1a PE or PE-labeled mouse IgG1 control mAbs. Cellular fluorescence and light scatter parameters of individual cells were recorded on a FACScan flow cytometer (Becton Dickinson).
Immunohistochemistry
Cells (3–5 x 104) were placed onto individual fields of adhesion slides (Bio-Rad, Richmond, CA) and fixed in acetone for 10 min at 4°C. After rinsing the slides in PBS/1% H2O2, cells were exposed to mAb Lag or mouse IgG1 (each 2 µg/ml) in PBS/1% normal horse serum (Vector, Burlingame, CA). Cell-bound mAbs were conjugated with biotinylated horse anti-mouse IgG (Vector) and visualized using avidin/biotinylated peroxidase complexes (ABC Elite Vectastain Kit, Vector) and 3-amino-9-ethylcarbazole (Sigma). Cells were counterstained with Harris’ hematoxylin (Merck, Darmstadt, Germany) and, finally, were embedded in Glycergel (Dakopatts, Glostrup, Denmark).
Western blotting
Cells were solubilized in Tris lysis buffer (20 mM, pH 8.2) containing 1% Nonidet P-40 (Sigma), 140 mM NaCl, 2 mM EDTA, 1 mM iodoacetamide, 1 mM PMSF, and aprotinin and leupeptin (both at 10 µg/ml; all from Sigma). Lysates were kept on ice for 30 min, reacted with SDS sample buffer and analyzed by immunoblotting as described previously (16). Briefly, aliquots of lysates were submitted to 8% SDS-PAGE and blotted onto Immobilon-P nitrocellulose membranes (Amersham, Aylesbury, U.K.). Membranes were blocked with 5% nonfat dry milk/0.1% Tween-20 (Sigma)/PBS for at least 6 h. Thereafter, they were exposed to mouse mAbs against CD45 (clone MEM-28; 2.4 µg/ml) and E-cad (clone 36; 0.25 µg/ml), or to rabbit anti-FXIIIa (1:200). Primary Ab binding was revealed by an incubation step with peroxidase-conjugated sheep anti-mouse IgG (1/10,000) or sheep anti-rabbit IgG (1/10,000; both from Bio-Rad) and were visualized by chemiluminescence (ECL Western blotting detection kit, Amersham).
Transmission electron microscopy
Sample preparation for transmission electron microscopy and examination was performed as described previously (4).
Allogeneic MLR
Graded numbers of differentially stimulated CD34+ HPC-derived cells were dispensed in individual wells of 96-well round-bottom microtiter plates (Costar, Cambridge, MA) and were cocultured with 1 x 105 allogeneic naive CD4+ T cells for 6 days in RPMI 1640/10% AB serum (PAA Laboratories, Linz, Austria). Naive CD4+ T cells were purified by anti-CD8/CD16/CD19/CD45RO/CD56/HLA-DR-based immunomagnetic depletion of PBMC as described previously (17). During the last 16 h of the APC-T cell coculture, cells were pulsed with 0.5 µCi [3H]thymidine (Amersham), and the incorporated radioactivity was measured by ß-scintillation spectroscopy (1205 Betaplate, Wallac Instruments, Meriden, CT). Results are expressed as mean counts per minute of duplicate cultures. Background counts of controls (responder T cells or stimulator cells alone) were always <100 cpm.
Analysis of precursor frequencies and cumulative cell divisions
Due to its amphophilic properties, 5- (and 6)-carboxyfluorescein
diacetate succinimidyl ester (CFDA; Molecular Probes, Eugene, OR)
permeates into the cytosol of cells, where its acetate groups are
removed by cellular esterases (18, 19). This process
yields a brightly fluorescent, amine-reactive fluorophore that binds to
proteins in a covalent fashion. Freshly isolated
CD34+ HPC (1 x 106)
were incubated in 1 µM CFDA/PBS for 20 min at 37°C. After several
washings to remove noninternalized/metabolized CFDA, the
CD34+ HPC-containing cell population was divided
into two equally sized aliquots, one of which was irradiated with 3000
rad. CFDA-labeled nonirradiated and irradiated
CD34+ HPC were cultured in RPMI 1640/10% FCS
supplemented with GM-CSF, TNF-, and Flt3-L. On day 6 of the culture,
cells were recovered, stained with anti-CD1a PE, anti-CD14 PE,
or both mAbs (to define the double negative population) or were
simultaneously exposed to anti-CD14 PE and biotinylated
anti-CD11b followed by SA-Cy5 and were subjected to multicolor FACS
analysis.
Irradiated cells that served as an internal standard for nondividing
cells produced a single-peaked histogram in the high CFDA fluorescence
intensity range confirming dye stability over the culture period. CFDA
segregates equally between daughter cells upon cell division, resulting
in the sequential halving of cellular fluorescence intensity with each
successive generation. When analyzed by flow cytometry, this sequential
halving of fluorescence is visualized as distinct peaks or populations
of cells. Thus, the simultaneous multicolor analysis of CFDA content
and Ag expression profile allows tracking cell divisions in mAb-defined
(sub)populations of proliferating cells. To estimate the "size" of
the proliferative burst, we relied on the fact that the size of the
(mAb-defined) daughter cell populations on day 6 and the size of their
CD34+ precursor pools on day 0 are related by a
function of 2n, where n is the number
of division cycles achieved during clonal expansion. Because the number
of events under individual CFDA fluorescence peaks (E)
equals the number of (mAb-defined) day 6 cells that have undergone
n division cycles, the number of CD34+
precursor cells that must have divided n times to generate
them can be extrapolated by dividing En by
2n. As previously described (20, 21), the sum of the extrapolated precursor numbers for each
division cycle gives the size of the precursor sample pool
(P) sufficient to have generated the cell sample
(E) with the given division pattern (Equation 1
):
 | (1) |
):
 | (2) |
En), and
this quotient was multiplied by n; second, the sum of all
these values directly reveals Da (Equation 3):
 | (3) |
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Suggestive evidence exists that TGF-ß1, both in vitro
(12) and in vivo (11), is of essential
importance for the development of LC from bone marrow-derived HPC. In
an attempt to characterize the to date unknown TGF-ß-responsive LC
precursor(s), we identified and isolated the well-defined
CD14+CD1a- and the
CD1a+CD14- DC precursor
populations arising around day 6 from
GM-CSF/TNF-/Flt3-L-stimulated
CD34+ HPC (Fig. 1)
(5, 6). In initial experiments, the conventional serum-containing and,
thus, latent TGF-ß-containing culture model and an entirely
TGF-ß-free culture system (i.e., serum-free X-VIVO 15 medium) were
used in parallel for this primary culture period. Although
CD14+CD1a- and
CD1a+CD14- cells appeared
on day 6 under both conditions, the frequency of their occurrence (21
vs 35% CD14+ and 12 vs 22%
CD1a+ cells in X-VIVO and in FCS/RPMI,
respectively; mean values; n = 5) as well as the
overall cell expansion (i.e., 5- to 10-fold less in serum-free compared
with FCS-conditioned medium) were drastically lower under serum-free
conditions (data not shown). This poor cell recovery/expansion under
serum-free conditions hampered a solid search for TGF-ß1-responsive
DC precursors before day 6. Importantly, however, cells that had been
expanded for 6 days in FCS-conditioned medium and then were further
propagated in the X-VIVO- and FCS-based culture system yielded
comparably sized progenies on day 12. This allowed us to study in
detail the effect of exogenous TGF-ß1 on the differentiation of
CD1a+ and CD14+ DC
precursors. Consequently,
CD1a+CD14- and
CD14+CD1a- cells were
isolated on day 6 and propagated in TGF-ß1-conditioned or
nonconditioned X-VIVO medium until day 12, and cellular progenies were
subjected to immunostaining with mAbs recognizing differentiation Ags
that are either DC subtype restricted (e.g., E-cad, CD1a, CD11b, and
CD36) or occur in virtually all members of the DC family (e.g., HLA-DR
and B7-2) (1, 2). As shown in Fig. 1, CD14+
cells, when cultured in the presence of TGF-ß1, gave rise to a
cellular progeny that expressed high levels of CD1a, E-cad, and HLA-DR.
In striking contrast, when the same precur- sors were cultured
in the absence of this cytokine, cells emerged that hardly displayed
these LC-associated features, but expressed HLA-DR in quantities
comparable to those on the TGF-ß1-stimulated cells (Fig. 1).
Interestingly enough, when cells were cultured in the presence of
TGF-ß1, but not in its absence, a small subpopulation appeared that
displayed CD11b, an Ag expressed by monocyte-related non-LC DC such as
dermal DC and monocyte-derived DC (Fig. 1). Although the emergence of a
CD1a+E-cad+ progeny from
CD14+ cells strictly depended on the presence of
TGF-ß1, CD1a+ progenitors gave rise to
CD1a+E-cad+HLA-DR+CD11b-
cells in both the presence and the absence of this cytokine (Fig. 1).
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The TGF-ß1 dependency of E-cad synthesis by
CD14+ progenitor-derived cells was further
investigated by protein immunochemistry. In accordance with the surface
immunostaining results, we observed, by Western blotting, that the
expression of 120-kDa anti-E-cad immunoreactive moieties in
CD14+ precursor-derived cells occurs in a
TGF-ß1-dependent fashion (Fig. 2A). In contrast,
CD1a+ intermediates acquire E-cad expression
regardless of whether the cultures were supplemented with TGF-ß1
(Fig. 2A). As previously reported, we found that
GM-CSF/IL-4-elicited monocyte-derived DC are induced to express E-cad
when generated in the presence of TGF-ß1 (22, 23) (Fig. 2, A and B). Fig. 2B also shows that
TGF-ß1 mediates enhanced CD1a and decreased HLA-DR expression. Note,
however, that regardless of exposure to TGF-ß1, monocyte-derived DC
maintain expression of factor XIIIa (FXIIIa; Fig. 2A) and
the myeloid marker CD11b (Fig. 2B), two Ags reportedly
expressed by dermal type, non-LC DC. This is in contrast to most
CD14+ HPC-derived LC-type DC that do not express
these two epitopes after TGF-ß1-dependent culture and differentiation
(1, 5, 6). Thus, it appears that the majority of
CD14+ HPC-derived cells can acquire the exact
immunophenotype of LC in a TGF-ß1-dependent manner, while
CD14+ peripheral blood monocyte-derived DC do not
undergo complete LC differentiation. We further investigated whether
quantitative differences in TGF-ß production by
CD14+ and CD1a+
progenitor-derived cells could explain their mutually exclusive E-cad
expression pattern. Although we found that CD1a+
progenitor-derived cells secreted somewhat higher amounts of this
cytokine than did the progeny of the CD14+ cells
(90 vs 32 pg/105 cells, mean values obtained in
six independent experiments), the neutralization of TGF-ß could
neither prevent nor reduce E-cad production and surface expression by
these cells (Fig. 2A and data not shown). Thus, it appears
unlikely that the occurrence of a cell type-restricted autocrine
TGF-ß circuit is responsible for the mutually exclusive expression of
LC-associated molecules by the progeny of the two precursor
subsets.
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To determine whether TGF-ß1 stimulation of
CD14+ progenitors gives rise to cells that
fulfill the cardinal phenotypic features of LC, we first analyzed
CD1a+ cells arising from TGF-ß1-stimulated
CD14+ HPC-derived precursors for the expression
of the cytoplasmic, BG-associated protein Lag. Using either cytoplasmic
immunolabeling and FACS analysis or immunohistochemical staining, we
observed that the majority of the TGF-ß1-stimulated cells were
Lag+ (Fig. 3,
A and C), whereas cells that had been cultured in
the absence of this cytokine were entirely devoid of this
immunoreactivity (Fig. 3, A and B). In the former
cells, the Lag Ag was distributed in a granular pattern,
indistinguishable from that seen in LC generated from
CD1a+ precursors (Fig. 3E). In keeping
with our previous results, CD1a+ progenitors gave
rise to Lag+ LC regardless of whether
TGF-ß1 was added to the cultures (Fig. 3, D and
E). By EM, we found that the cells that emerged from
TGF-ß1-stimulated CD14+ progenitors displayed
pronounced DC morphology (Fig. 4B) and, thus, resembled LC
generated from CD1a+ precursors (Fig. 4C). Both cell populations exhibited abundant dendritic
membrane protrusions and contained many multilamellar organelles and
indented or lobulated nuclei in a slightly off-center position. As the
most revealing feature, both DC populations contained BG, cytoplasmic
organelles characterized by their typical double-membrane junctions and
their rod-shaped structure (Fig. 4, E and F).
Numeric evaluations revealed that BG are encountered in comparable
frequencies in ultrathin-sectioned cell profiles (i.e., 10–20%) of
TGF-ß1-stimulated CD14+ precursor-derived and
CD1a+ precursor-derived DC. In the absence of
TGF-ß1, CD14+ precursors gave rise to cells
with a roundish, monocyte-like morphology (Fig. 4A).
Although these cells displayed a nuclear architecture similar to that
of CD14+ and CD1a+
precursor-derived LC, their cytoplasm contained many multivesicular,
rather than multilamellar, organelles and was entirely devoid of
BG.
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Data obtained to date demonstrated that
CD34+ stem cell-derived
CD14+ precursors can develop into LC in a
TGF-ß-dependent fashion. This is best evidenced by the observations
that TGF-ß1 up-regulates CD1a, E-cad, and Lag expression and induces
BG and DC morphology in the progeny of those cells. It was interesting
to note, however, that this LC-promoting potency did not uniformly
occur with all CD14+ hemopoietic precursors
identified and isolated. In fact, small populations of
CD11b+ and weakly E-cad+,
or even E-cad-, cells could be recovered from
TGF-ß1-stimulated CD14+ precursor cell cultures
(Fig. 1
). To further characterize these cells, we performed
anti-E-cad and anti-CD11b double-immunolabeling experiments,
which revealed that in the presence of TGF-ß1, two mutually exclusive
cell populations emerged from CD14+ progenitors.
The major population (>75%) expresses E-cad but not CD11b and, thus,
corresponds to the CD14+ cell-derived LC
(Fig. 5). Most of the remaining cells are
E-cad-/low but CD11b+
(Fig. 5). It is noteworthy that in the absence of TGF-ß1, the
appearance of the CD11b+ subset is only
incomplete.
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). To analyze
the LC/DC differentiation potential and lineage commitment of the
CD11b+ and
CD11b-/lowCD14+ cells, the
subsets were sorted and propagated with TGF-ß1 or M-CSF. In the
presence of TGF-ß1, the CD11b- cells gave rise
to a typical LC progeny with homogeneous, high level expression of
CD1a, E-cad, HLA-DR, and B7-2, but no anti-CD14 or anti-CD11b
and only negligible anti-CD36 immunoreactivity (Fig. 6).
Conversely, when the same cell population was exposed to M-CSF,
macrophage-like cells with high CD14, CD11b, and CD36; weak B7-2; and
no CD1a and E-cad expression developed (Fig. 6). Thus, it appears that,
unlike the CD1a+ LC precursor, which dies in the
presence of M-CSF (5), the
CD11b-CD14+ population is
at least bipotent, giving rise to LC or macrophages upon appropriate
cytokine stimulation. When we subjected the
CD11b+CD14+ precursor to
the same isolation, cell culture, and immunophenotyping protocol, we
found that TGF-ß1 stimulation led to the emergence of DC that express
CD1a, HLA-DR, and B7-2, but, unlike LC, display only limited E-cad
expression and are homogeneously CD11b+ and
CD36+ (Fig. 6). Furthermore, these cells did not
express CD14 (Fig. 6) and, thus, by immunophenotype strikingly resemble
dermal DC, a non-LC DC population constitutively present in human skin
(5, 7). As also shown in Fig. 6, M-CSF stimulation of
CD11b+CD14+ cells resulted
in the appearance of a typical macrophage progeny.
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A comparative assessment of the MLR-stimulating potency of both
subpopulations generated in the presence of TGF-ß1 revealed that LC
generated via either pathway were effective stimulators of naive T
cells and elicited T cell proliferation at low APC:T cell ratios (Fig. 7A). When
CD11b-CD14+ cells were
stimulated with TGF-ß1 or M-CSF and their cellular progeny was tested
for its MLR-stimulating capacity, we observed that, at the per cell
level the LC emerging in the presence of TGF-ß1 were at least
100-fold more potent than the M-CSF-dependent macrophages (Fig. 7B). Macrophages generated from M-CSF-stimulated
CD11b+CD14+ cells also
displayed only poor accessory cell function (Fig. 7B).
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To investigate whether TGF-ß1-responsive
CD14+ LC precursors originate from a
CD34+ HPC pool differing in size and
proliferation potential from that giving rise to conventional
CD1a+ LC-committed progenitors, we performed
fluorescence-based precursor frequency analyses. Purified
CD34+ HPC were labeled with CFDA, a brightly
green fluorescent and stable cell tracker that is equally distributed
to daughter cells during mitoses and thus allows the quantification of
cells that have undergone a defined number of divisions (18, 19). Labeled CD34+ HPC and, as a control,
irradiated, and thus proliferation incompetent, CFDA-labeled
CD34+ HPC were stimulated with
GM-CSF/TNF-/Flt3-L and subjected to multicolor flow cytometric
analyses on day 6 of the culture. Irradiated cells produced a
single-peak histogram in the high CFDA fluorescence intensity range
that serves as the internal standard for nondividing cells (Fig. 8 and data not shown). The CFDA labeling
procedure did not adversely affect HPC proliferation and LC
differentiation, because it neither altered the yields and percentages
of CD1a+ and CD14+ cells on
day 6 nor negatively influenced the TGF-ß1-dependent and -independent
development of E-cad+CD1a+
DC from CD14+ and CD1a+
precursors, respectively (data not shown). The comparative analysis of
the fluorescence profiles of the CD14+ and
CD1a+ progenitors as well as of the remaining
double-negative cells directly shows that CD14+
cells have undergone the most and
CD14-CD1a- cells the
fewest numbers of divisions (Fig. 8). When we calculated, on the basis
of cell counts per given cell division number, LC precursor frequencies
at the CD34+ HPC stage, it appeared that the
CD11b-CD14+ and the
CD1a+ LC progenitors originate from a similarly
sized CD34+ stem cell pool (Table I). Interestingly, the
CD11b-CD14+ progenitors
had undergone one cell division more than the
CD1a+ LC precursors until day 6 (Table I). Thus,
the more frequent occurrence of CD14+ than of
CD1a+ LC precursors on day 6 (Table I) is due to
a higher proliferative potential and not due to a larger
CD34+ stem cell pool of the CD14 pathway of LC
development. These calculations also revealed that the
CD11b+CD14+ subpopulation
emerges from a CD34+ HPC pool that is only half
the size of that giving rise to their CD11b-
counterparts (Table I). When CD14+ and
CD1a+ LC progenitors were sorted, subjected to
CFDA labeling, and cultured in the presence or the absence of TGF-ß1,
we observed that LC generated from either pathway underwent an average
of only 0.5 cell divisions between days 6 and 12 (data not shown).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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and TGF-ß1 originate not only
from CD1a+ progenitors but also from a pool of
CD14+CD1a- precursors; and
2) that these two progenitor cells differ in their TGF-ß1
requirements. It was shown previously that TGF-ß1 can amplify LC
development in vitro (12) and, based on experimental data
in gene-targeted TGF-ß1-/- mice, in vivo
(11). However, the biology of TGF-ß1-mediated
amplification of LC development is poorly understood to date. The data
available suggest that TGF-ß1 inhibits the apoptotic death of LC
precursors (13). We now have evidence that another main
function of this cytokine is to redirect a large population of
monocytic precursors into the differentiation pathway(s) of
cutaneous DC.
Upon TGF-ß1 signaling,
CD11b-CD14+ cells
homogeneously develop into LC (Fig. 6). However, not all
CD14+ cells enter the same route of
differentiation.
CD11b+CD14+ cells give rise
to cells with the immunophenotypic characteristics of dermal DC (DDC)
rather than of LC. We have evidence that TGF-ß1 is also needed for
the development of these
E-cad-CD11b+ DDC-like
cells from CD14+ precursors (Figs. 5 and 6).
At first glance, the latter results contradict the previous observations showing that CD14+ precursors, even in the absence of exogenous TGF-ß, can give rise to non-LC DC (5, 6). Because bovine serum-containing and, thus, latent TGF-ß-containing culture media were used in these studies, we also analyzed the role of TGF-ß1 for LC and non-LC DC differentiation of CD14+ precursors during culture in FCS-conditioned medium. In the absence of exogenous bioactive TGF-ß1 and in accordance to previous observations (5, 6), no E-cad+CD11b- LC could be recovered from CD14+ precursors, and the cells rather homogeneously acquired a DDC-like phenotype (data not shown). In striking contrast, TGF-ß1 supplementation of these cultures resulted in preferential LC development, as assessed by immunophenotypic and ultrastructural criteria (data not shown). Thus, it appears that 1) CD14+ precursors cannot sufficiently activate and/or use xenogeneic latent TGF-ß1 to undergo LC transformation; and 2) FCS-derived factors other than TGF-ß1 can promote non-LC DC development from CD14+ precursors.
In accordance with earlier observations (22, 23) we found
that DC generated from blood monocytes in the presence of GM-CSF/IL-4
could display E-cad in a TGF-ß1-dependent fashion (Fig. 2, A and B). However, TGF-ß1 does not allow these
DC to undergo unequivocal LC differentiation, as it did not abolish the
expression of the monocyte/macrophage-related Ag FXIIIa (Fig. 2A) and the myeloid Ag CD11b (Fig. 2B), two
markers never detected on classical LC. These data strengthen the
concept that not all CD14+ DC precursor subsets
can undergo full TGF-ß-dependent LC differentiation and, furthermore,
suggest that the differentiational stage of the
CD11b-CD14+ intermediate
is likely to be the last checkpoint during myelopoiesis from which
TGF-ß-dependent, unambiguous LC differentiation can occur.
It should not be forgotten that the
CD11b-CD14+ cells are
versatile precursors that, depending on the cytokine signals delivered,
can enter various lineages of myeloid differentiation. Unlike
CD1a+ LC precursors, which do not express
receptors for M-CSF and die in the sole presence of this cytokine
(5),
CD11b-CD14+ precursors
differentiate into macrophages in the presence of M-CSF (Fig. 6). We
have preliminary evidence that in the combined presence of TGF-ß1 and
M-CSF, CD11b-CD14+
precursors develop into weakly E-cad- and CD1a-positive DDC-like cells.
Thus, it appears that the
CD11b-CD14+ subpopulation
contains multilineage myeloid precursors with LC, non-LC DC, and
monocyte/macrophage differentiation potential. It is also conceivable
that the early receipt of either TGF-ß1 or M-CSF signals by a
certain, poorly differentiated common progenitor cell type results in
the emergence of either CD1a+ LC precursors or
CD11b+CD14+ non-LC DC
precursors on day 6 of the stem cell cultures. The latter argument is
supported by our observation that the supplementation of FCS- as well
as serum-free CD34+ stem cell cultures with
neutralizing anti-TGF-ß mAbs can reduce the frequency of
CD1a+ LC precursors on day 6 (data not shown).
However, the inhibition of CD1a+ LC precursor
development was never complete. Thus, it is possible that 1) in certain
CD1a+ LC precursors, TGF-ß delivery may occur
in a compartmentalized autocrine or perhaps even intracrine fashion not
accessible to mAb-based inhibition; or 2) stem cell pools exist that
allow for TGF-ß-independent LC development. In fact, our studies of
the precursor frequencies and proliferation potentials of the various
CD34+ HPC-derived subpopulations showed that
CD11b-CD14+ and
CD1a+ precursors emerge from stem cell pools that
differ in their proliferative potential (Fig. 8 and Table I). Due to
their similar size, however, it cannot be decided whether the stem cell
pools of these precursor subsets are identical, partially overlapping,
or mutually exclusive. Unfortunately, our attempts to identify and
purify cord blood-derived CD34+ HPC subsets with
mutually exclusive CD1a or CD14 differentiation potentials have
remained unsuccessful. Thus, it appears that
CD1a+ and CD14+ precursors
originate from phenotypically similar stem cell pools, but enter
divergent pathways early after the start of the culture. This argument
finds support in the observation that LC derived from
CD1a+ precursors homogeneously express the
maturation-related surface Ag CD83 by day 10, while
CD14+ precursor-derived LC start to up-regulate
this molecule only after day 13 of the culture (data not shown).
At the present time it is not clear which cell type serves as the main TGF-ß1 source in vivo and, perhaps more important, as the biologically relevant LC/DDC precursor. Although TGF-ß1 is constitutively produced by keratinocytes (1), cell transfer studies in mice showed that radiation-resistant host cells other than keratinocytes could be important in this regard (24). As it is well established that TGF-ß is secreted as a latent complex (25, 26), the activity of TGF-ß-liberating factors rather than the mere presence of this cytokine may be of importance for the regulation of LC development. Concerning the nature of the LC/DDC progenitor, it remains to be seen whether certain CD1a+BG- cells or even CD1a- cutaneous lymphocyte Ag (CLA)+ cells (27) reside within the dermis and can develop into typical LC. The involvement of the CD14 pathway of LC/DDC development can be deduced from our recent identification of a CD45+HLA-DR+CD14+ but CD11b- cell population within human dermis that by immunophenotype corresponds to the TGF-ß1-responsive CD14+ LC precursor seen in CD34+ HPC-derived cultures (data not shown). Should this cell type give rise to LC in the presence of TGF-ß1 but develop into macrophages in the presence of M-CSF it would provide the skin with the unique option to rapidly generate functionally diverse cell types in response to the local cytokine milieu.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Dieter Maurer, Division of Immunology, Allergy and Infectious Diseases, Department of Dermatology, University of Vienna Medical School, Waehringer Guertel 18-20, A-1090 Vienna, Austria. E-mail address:
3 Abbreviations used in this paper: LC, Langerhans cells; ACC, accessory cells; BG, Birbeck granules; CB, cord blood; CFDA, carboxyfluorescein diacetate; DC, dendritic cells; DDC, dermal dendritic cells; EM, electron microscopy; HPC, hemopoietic progenitor cells; E-cad, E-cadherin; Flt3-L, Flt3 ligand; FXIIIa, factor XIIIa; ML, multilamellar organelles; MV, multivesicular organelles; PerCP, peridinin chlorophyll protein; SA, streptavidin; SA-PE, PE-conjugated streptavidin.
Received for publication March 29, 1999. Accepted for publication August 16, 1999.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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