By
From the * Division of Immunology, Allergy and Infectious Diseases, Department of Dermatology, Institute of Transfusion Medicine, University of Vienna Medical School, A-1090 Vienna, Austria;
and § Contrat jeune formation Institut National de la Santé et de la Recherche Médicale,
94-03, Etablissement de Transfusion Sanguine, F-67065 Strasbourg, France
We have recently described a system for the generation of dendritic cells (DC) and Langerhans
cells (LC) from defined CD34+ precursors purified from peripheral blood of healthy adult volunteers (1). This study has now been extended by the characterization of two distinct subpopulations of CD34+ cells in normal human peripheral blood as defined by the expression of the
skin homing receptor cutaneous lymphocyte-associated antigen (CLA). CD34+/CLA+ cells
from normal peripheral blood were found to be CD71LOW/CD11a+/CD11b+/CD49d+/
CD45RA+ whereas CD34+/CLA cells displayed the CD71+/CD11aLOW/CD11bLOW/CD49d(+)/
CD45RALOW phenotype. To determine the differentiation pathways of these two cell populations, CD34+ cells were sorted into CLA+ and CLA
fractions, stimulated with GM-CSF and
TNF-
in vitro, and then were cultured for 10 to 18 d. Similar to unfractionated CD34+ cells,
the progeny of both cell populations contained sizable numbers (12-22%) of dendritically
shaped, CD1a+/HLA-DR+++ cells. In addition to differences in their motility, the two dendritic cell populations generated differed from each other by the expression of LC-specific
structures. Only the precursors expressing the skin homing receptor were found to differentiate into LC as evidenced by the presence of Birbeck granules. In contrast, CLA
precursor cells
generated a CD1a+ DC population devoid of Birbeck granule-containing LC. Provided that
comparable mechanisms as found in this study are also operative in vivo, we postulate that the
topographic organization of the DC system is already determined, at least in part, at the progenitor level.
Dendritic cells (DC) are bone marrow-derived leukocytes with potent immunostimulatory properties. They
occur in small numbers in most non-lymphoid tissues and
are well equipped to take up various types of antigen. After an
antigenic challenge, these cells undergo phenotypic changes
that allow them to leave their residence, to migrate to the
regional lymphoid organs, and, upon arrival in the T-dependent zones, to activate resting T cells (2). This is best exemplified by the antigen-induced metamorphosis of epidermal Langerhans cells (LC; CD45+/CD1a+/CD32+/
E-cadherin+/Birbeck granule+/ATPase+/MHC class I+
and II+) into interdigitating reticulum cells (CD45+/CD1a Together with the findings that the HECA-452-defined
E-selectin ligand cutaneous lymphocyte-associated antigen
(CLA) functions as a skin homing molecule for certain
memory T cells (5), that LC within the skin are CLA+ (6),
and that intravenously injected LC specifically home to the
skin (7), our recent observation that approximately half of
the CD34+ hematopoietic progenitor cells circulating in
the peripheral blood of healthy adults (PBPC) react with
the mAb HECA-452 (1) led us to the hypothesis that CLA
might also be involved in the migration and/or the maturation of LC precursors. Here, we provide evidence that circulating CD34+/CLA+ but not CD34+/CLA Purification of CD34+ PBPC from Healthy Volunteers.
CD34 + PBPC were isolated essentially as described (1). Following this
procedure, 2 × 106 CD34+ PBPC with a purity of >95% and a
viability of >96% can be reproducibly procured from one leukapheresis product (1.5-3 × 109 PBMC) (1).
Three-Color Flow Cytometry of PBPC.
Freshly isolated PBPC
were adjusted to 5 × 105 cells/ml in PBS containing 1% FCS and
0.1% NaN3 and quenched with normal sheep serum (10% vol/
vol, 30 min on ice) to reduce nonspecific reactivity. Cells were
reacted for 30 min on ice with a panel of mouse mAbs against human leukocyte differentiation antigens including CD45RA (clone
2H4; Coulter, Hialeah, FL), CD71 and HLA-DR (clones LO1.1 and L243; Becton Dickinson Immunocytometry Systems, BD-IS,
San Jose, CA), CD11a, CD11b, CD18, CD49d, CD49e, CD29
(clones 25.3, Bear 1, BL5, HP2.1, SAM1, K20; all Immunotech,
Marseille, France) as well as two different anti-CD1a mAb, which
also served as isotype controls (clone OKT6, mouse IgG1; Ortho,
Raritan, NJ, and B17.20.9, mouse IgG2a; Immunotech). With
washings after each incubation, cells were consecutively exposed to
biotinylated F(ab Suspension Culture of PBPC Subpopulations.
To separate CLA+/
CD34+ from CLA/
CD32
/E-cadherin
/Birbeck granule
/ATPase
/MHC
class I++ and II+++) (3, 4). In contrast to the substantial information about the factors governing DC trafficking from
the periphery to the lymphoid organs, relatively little is
known about the reverse process, e.g., about the mechanisms underlying the immigration of circulating LC precursors in their ultimate tissue of residence, i.e., the skin
and epidermis.
PBPC can
differentiate into LC in vitro.
)2 fragments of a polyclonal sheep anti-mouse
Ig (10 min, on ice; Amersham Inc., Arlington Heights, IL), normal mouse serum (10% vol/vol, 20 min, on ice) and, finally,
streptavidin-PerCP (BD-IS) simultaneously with mouse anti-human
CD34-PE (8G12-PE; BD-IS) and HECA-452-FITC (5) (kindly
provided by Dr. L. Picker, University of Texas Southwestern
Medical Center, Dallas, TX) or the respective isotype control mAbs
(30 min, on ice). For data analysis, CLA+/CD34+ and CLA
/
CD34+ events were gated and reactivity with the third mAb was
measured as mean fluorescence intensity using a FACScan® flow
cytometer (BD-IS) equipped with LysysTM II (BD-IS) software.
/CD34+ cells, 24-well cell culture plates (Costar, Cambridge, MA) were coated with purified HECA-452 mAb
(10 µg/ml, overnight, 4°C) and then extensively washed with PBS.
Freshly isolated CD34+ PBPC were incubated in HECA-452-
coated wells for 60 min at 37°C (105 cells/well). Thereafter,
plates were rinsed 3× with 1 ml HBSS to collect the nonadherent cell population.
(Genzyme, Cambridge, MA). Alternatively, aliquots of unfractionated freshly isolated PBPC were cultured at 5-7 × 104 cells/ml in the same medium in 24-well culture plates, which were either coated with the
HECA-452 mAb as described above or mock (HBSS)-coated.
Cultures were maintained for at least 3 wk (37°C, humidified atmosphere, 5% CO2) by replacing one-fourth of the medium with
fresh cytokine-supplemented complete medium every 4-5 d.
Phenotypic Analysis of PBPC-derived Cells by Immunocytochemistry, Flow Cytometry, and Electron Microscopy.
Cells derived from
unfractionated PBPC or the HECA-452-sorted subpopulations
were harvested after defined periods of culture with GM-CSF/
TNF- (10, 14, 16, 18 d) and subjected to immunohistochemistry, flow cytometry, and immuno-electron microscopy.
Allogeneic Mixed Leukocyte Reaction.
Allogeneic mixed leukocyte reactions (MLR) were performed to test for the presence of
potent accessory cells within the populations of PBPC-derived cells.
Various numbers of mitomycin C (50 µg/ml, 30 min, 37°C; Sigma
Chem. Co., St. Louis, MO)-treated PBPC-derived cells and, for
control purposes, plastic-adherent monocytes from the same individual were cocultured with 105 purified allogeneic T cells in
complete medium supplemented with 10% plasma of PBPC donor origin in 96-well round-bottomed culture plates (Costar).
MLRs were maintained for 4, 5, or 6 d (37°C, humidified atmosphere, 5% CO2) before [3H]thymidine (1 µCi/well; Amersham)
was added for additional 16 h. Incorporation of the radionucleotide
was measured with a -scintillation spectroscope (Packard Instruments, Meriden, CT). Results are expressed as mean cpm SD of
triplicate cultures. Proliferation of 105 T cells or mitomycin
C-treated stimulators alone resulted in an incorporation rate of
<500 cpm.
In a first
series of experiments, we used three-color flow cytometry
to determine the immunophenotype of HECA-452-reactive and HECA-nonreactive CD34+ PBPC. We found that
both populations were CD1a, expressed equal levels of
HLA-DR and common CD45 molecules, but exhibited pronounced quantitative and qualitative differences in their distribution of various leukocyte differentiation antigens. Quantitatively, CLA+/CD34+ cells showed a three- to fivefold
higher expression of the
2-integrins LFA-1 (CD11a/CD18)
and Mac-1/C3bi (CD11b/CD18), and a 1.5-fold higher
expression of the
1-integrins VLA-4 (CD49d/CD29) and
VLA-5 (CD49e/CD29) as compared with CLA
/CD34+
cells (Fig. 1). It will be interesting to determine whether
the higher level of LFA-1/VLA-4 expression on CLA+/
CD34+ PBPC also allows for a more efficient binding to
VCAM-1+/E-selectin+ endothelia (5, 9).
Qualitatively, the major phenotypic differences between
the two progenitor cell populations are the almost selective
anti-CD45RA reactivity of CLA+ PBPC and the eightfold
higher transferrin receptor (CD71) expression on CLA/
CD45RA
PBPC (Fig. 1). Since CD45RALOW/
/CD34+
cells in bone marrow, cord blood, and growth factor-mobilized blood represent the more immature progenitor cell subset when compared with CD45RA+/CD34+ cells (10),
CLA+/CD45RA+ PBPC probably constitute an already
committed cell population of only limited proliferative potential (see below). Their low expression of transferrin receptors supports this assumption since the anti-CD71 reactivity of a given cell usually correlates with its state of
proliferation (11).
To determine whether both
CLA+ and CLA PBPC can differentiate into LC, we established a subtractive culture system that is based on the
panning of one subpopulation of PBPC (i.e., CLA+ cells)
directly in the culture well and on the transfer and subsequent culture of the nonadherent subtracted (i.e., CLA
)
cells. This procedure allows for excellent cell recovery in both the panned and nonpanned (<0.1% contamination
with CLA+ cells) fractions. When the two PBPC subsets
were cultured for 2 wk with GM-CSF/TNF-
, the total
number of nucleated cells recovered from the CLA
fraction was three to five times higher than that collected from the CLA+ subset. These differences in the multiplication
rate reflect the phenotypic and maturational differences between the precursor cells (see above).
To test the hypothesis that CLA expression is involved
in the generation of LC, immunohistochemistry was used
to search for the presence of CD1a+ and Lag-reactive cells
(8, 12). After 10 d of stimulation with GM-CSF/TNF-,
CD1a+ dendritically shaped cells were easily detectable in
cultures from either CLA+ or CLA
precursors. The absolute
numbers of CD1a+ cells were always higher in the cell fraction originating from CLA
precursors because these cells
showed an approximately three- to fivefold higher proliferation compared to the CLA+ fraction (data not shown).
With regard to the progeny of CLA+/CD34+ PBPC, the percentage of CD1a+ cells ranged from 12% (day 10) to 22.5% (day 18) and that of Lag+ cells from 5% (day 10) to 20% (day 18; Table 1, Fig. 2 A). Double labeling on day 18 revealed that Lag reactivity of a given cell was restricted to the CD1a+ population (data not shown). Whereas on day 10 the percentage of Lag+ among CD1a+ cells ranged between 35-40%, ~90% of CD1a+ cells gave positive Lag immunostaining on day 18 of culture. The majority of CD1a+/Lag+ cells tended to form clusters (Fig. 2 A) and were dendritic in shape. Occasionally, we observed isolated cells with a distinctive dendritic morphology devoid of Lag reactivity (data not shown).
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In sharp contrast to these findings, cells derived from
CLA/CD34+ PBPC, while containing various numbers
of CD1a+ dendritic cells (Table 1), proved to be completely Lag
at all time points investigated (Fig. 2 B, Table 1).
These data were confirmed at the ultrastructural level. Although Birbeck granules were easily detected in one third
of CD1a+ LC (5/15 cells investigated) derived from CLA+
progenitors (Fig. 3), the search for these organelles in the CD1a+ progeny of CLA
cells yielded negative results (data
not shown). Concerning other phenotypic features of the
CD1a+ progeny of CLA+ and CLA
progenitors, we found
that, on day 18 of culture, CD1a+ cells derived from either
CLA+ or CLA
progenitors uniformly expressed MHC
class II and CD80 (data not shown). CD14 was detected on
a minor subpopulation of CLA+ PBPC-derived (15%) and
CLA
PBPC-derived (6%) CD1a+ cells.
Other differences emerged when we investigated the functional properties of the two populations. Although cells derived from both CLA+ and CLA progenitors induced vigorous proliferation of allogeneic T cells at all time points
tested and, in this capacity, were 10-50 times more potent
than syngeneic monocytes (Fig. 4), they clearly differed from
each other in their migratory properties. During the routine daily inspection of the PBPC progeny, we noticed that
cultures derived from either CLA+/CD34+ or unfractionated CD34+ cells, but not from CLA-depleted CD34+ cells,
contain a population of dendritically shaped cells which, in
contrast to their symbionts, moved continuously in the culture well. Microphotography revealed that these cells, per
minute, cover distances which were multiples of their own
diameter (Fig. 5). Since LC, upon receipt of an antigenic
(danger? [13]) signal, leave their epidermal residence and
migrate to the regional lymphoid organs (2), the migrating
DC observed in our cultures (assuming that they are CD1a+
LC) might reflect this in vivo situation and may therefore
serve as a paradigm of any peripheral non-lymphoid DC in
its transition to a terminally differentiated stimulator cell of
primary immune responses.
We were concerned that Birbeck granule formation in
the progeny of CLA+ PBPC, instead of being an intrinsic
property of this cellular subset, may have resulted from a
HECA-452-induced cross-linking of CLA epitopes. In fact,
the occurrence of a similar event has been described for
murine dendritic epidermal T cells upon stimulation with a
mitogenic anti-Thy-1 mAb (14). We consider this possibility highly unlikely for several reasons: (a) we (1) and others (15) have shown that unfractionated CD34+ cells can
differentiate into Birbeck granule+ LC without engaging the
CLA molecule; (b) comparable absolute numbers of CD1a+/
Lag+ LC were generated from unfractionated or CLAsorted starting cell populations containing equivalent numbers of CLA+ cells; (c) freshly isolated, unfractionated PBPC,
when cultured in the presence or absence of the mAb
HECA-452, exhibited similar proliferation rates and Lag
frequencies upon stimulation with GM-CSF/TNF- (data
not shown). The latter observation implies that the differences in the multiplication rate between CLA
and CLA+/
CD34+ PBPC are not due to an antiproliferative signal resulting from the HECA-452-induced occupancy of CLA
epitopes but rather reflect differences in the maturation
state of the two cell populations.
At the present time, it is not known whether the CLA
PBPC can develop/mature into CLA+ ones and, if so,
which factor(s) is (are) needed for this to occur. Also, it remains to be determined whether the expression on PBPC
of HECA-452-reactive CLA and of other molecules necessary for adhesion to and transmigration through microvascular endothelial cells (e.g., CD49d/CD29) allows for
the homing of progenitors to the skin. The mutually nonexclusive possibility exists that CLA expression merely reflects a distinct maturational stage of the progenitor cell that
renders it susceptible to stimuli favoring LC development.
In this context, recent attention has focused on TGF-
1
whose presence is apparently needed for the development of LC in vitro (16) and in vivo (17). We are currently investigating whether the CLA+ cell population differs from
its CLA
counterpart in the responsiveness to TGF-
1 and,
conversely, whether factors promoting the development of
lymphoid DC (e.g., relB) (18) are selectively/predominantly expressed in CLA
PBPC and their progeny. Should
this be the case, we would conclude that the topographic
organization of the DC system is already determined at the
progenitor level.
Received for publication 25 November 1996 and in revised form 16 January 1997.
This study was supported, in part, by grant S06702-MED from the Austrian Science Foundation (Vienna, Austria) and by grant FORTS 96 from the Agence Française du Sang (Paris, France).The authors are grateful to H. Bausinger, Dr. A. Bohbot, and Dr. D. Spehner for their help with electron microscopy. We wish to thank Dr. S. Imamura (Kyoto, Japan) for providing us with the mAb Lag.
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