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,§
*
Department of Cell Biology and Ludwig Institute for Cancer Research,
Department of Laboratory Medicine,
Molecular Cardiobiology Program, and
§
Section of Immunobiology, Yale University School of Medicine, New Haven, CT 06520
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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, LPS, or CD40
ligand, the LCs could be synchronously induced to mature. Depending on
the agent used, distinct types of LCs emerged differing in their
capacity for T cell stimulation, IL-12 production, intracellular
localization of MHC products, and overall morphology. Most
interestingly, the expression of different sets of Toll family
receptors is induced or down-regulated according to the maturation
stimulus provided. These results strongly suggest that different
proinflammatory stimuli might drive distinct developmental
events. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In vitro, DC maturation can be triggered by a variety of cytokines
(e.g., TNF-) and bacterial products (e.g., LPS) (6, 7).
Maturation in vivo has best been described for epidermal DCs
(Langerhans cells, LCs), an immunologically important DC population.
LCs exist as immature cells in the skin that migrate into the afferent
lymphatics and then to lymphoid tissue. Migration is enhanced by the
presence of maturational stimuli, as would occur concomitant with
infection. However, movement of LCs to lymph nodes also occurs
constitutively, with maturation subsequently induced in lymphoid tissue
upon binding of CD40 on LCs to CD40 ligand (CD40L) on T cells (8, 9). There is increasing evidence that the control of LC
maturation may help determine the induction of tolerogenic vs
immunogenic T cell responses (10). However, progress in
understanding LC maturation at the molecular level has been limited by
the inability to produce immature LCs in culture. Although the
maturation of human monocyte- and mouse bone marrow-derived DCs have
begun to be evaluated in some detail (3, 11), LCs
have proved difficult to isolate in any quantity, purity, or defined
maturational stage starting from bone marrow aspirates, leukapheresis
products, or umbilical cord blood (12, 13, 14, 15, 16). Thus, little
is known regarding the maturation of this important DC population.
By using CD34+ cells from G-CSF-mobilized patients, we have been able to produce large numbers of LCs that can be maintained in an immature state and induced to mature synchronously by the addition of inflammatory mediators. Remarkably, the type of activation stimulus used controls the type of mature DC that emerges, both in terms of their overall cellular organization, cytokine production, and their expression of receptors that play critical roles in innate immunity.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cancer patients who had undergone chemotherapy followed by G-CSF
treatment in preparation for autologous stem cell transplantation (or
healthy donors mobilized with G-CSF alone) were selected on the basis
of how efficiently CD34+ stem cells were
mobilized into their peripheral blood. Typically, patients with 
1%
CD34+ PBMCs were asked to donate either a part of
a standard (60 ml) leukapheresis or to undergo a separate apheresis
procedure for subsequent donation. Informed consent was obtained from
all patients, and the protocol was approved by the Yale University
School of Medicine Human Investigational Studies Committee.
CD34+ stem cells were immunomagnetically purified
from leukapheresis products with either a midi-MACS system (CD34
progenitor cell isolation kit or multisort kit) following the protocol
of the manufacturer (Miltenyi Biotec, Auburn, CA) or a Baxter Isolex
device (Baxter, Deerfield, IL). Yields of CD34+
cells varied from 1 x 107 to 2.5 x
108/ml leukapheresis; purity ranged from 85 to
95% CD34+ after a single selection.
CD34+ cells were frozen in aliquots of 2.5
x 106 in PBS/20% human albumin/10% DMSO and
stored in liquid nitrogen.
Culture conditions
Cells were thawed and cultured at 1 x
104/ml/well in 24-well plates in media prepared
exactly as per Strobl et al. (16). Specifically, cells
were grown in X-VIVO 15 containing 100 ng/ml GM-CSF (5.6 IU/mg), 20
ng/ml stem cell factor (5 x 104 U/mg), 2.5
ng/ml TNF- (2 x 107 U/mg), 0.5 ng/ml
TGF-ß1 (2 x 107 U/mg), and 100 ng/ml Flt3
ligand (Flt3L). All cytokines were purchased from PeproTech (Rocky
Hill, NJ), except for GM-CSF and Flt3L, which were obtained from
Immunex (Seattle, WA). Cultures were incubated at 37°C with 5%
CO2 in a humidified environment for 7–10 days
without feeding or replating; by this time total cell number had
increased by 50- to 100-fold. Clusters were purified by gently
harvesting cells with a pipette and layering on top of 6 ml of 7.5%
BSA (Sigma, St. Louis, MO) in 15-ml tubes: up to eight wells were
loaded per column. No adherent cells remained in the wells upon
harvesting. After 30 min on ice, single cells in suspension were
removed by aspirating the BSA columns until 3.5 ml remained. Clusters
were concentrated by centrifugation at 300 x g,
resuspended in growth media, and cultured at 5 x
105 cells/ml/well in 24-well plates for an
additional 2 days. For maturation, X-VIVO + 1x cytokines was
supplemented with additional 10 ng/ml of TNF- or with 250 ng/ml of
LPS (Sigma). Gel-purified platelets were prepared, activated, and fixed
exactly as per Tuszynski et al. (17). A total of 1.2
x 108 platelets/well were added to 24-well
plates, spun at 1200 x g, and washed two times with
PBS. Cluster-purified DCs were then plated on top of these platelet
lawns.
Antibodies
Abs to the following proteins were used: CD1a-PE (clone BL6), CD83-PE (Immunotech, Marseille, France); CD1a-FITC (clone HI149), CD14-FITC, CD66b-FITC, CD86-PE, CD107a, HLA-DR-FITC, HLA-DR-CyChrome, (PharMingen, San Diego, CA); HLA-A,B,C-FITC (Clone W6-32, Biodesign International, Kennebunkport, ME); CLA 1 (American Type Culture Collection clone HECA-452; Manassas, VA). LFA-3 Abs were obtained from mouse ascites. We would like to thank the following people for kind gifts of Abs: Kayo Inaba for mAb anti-Lag (18); Peter Cresswell for the polyclonal Ab anti-HLA-DR (19). Fluorescence microscopy was performed as described in Pierre et al. (3). Flow cytometry was performed by standard procedure. When detection of intracellular protein was required, cells were previously fixed and premeabilized as decribed in Inaba et al. (20).
Electron microscopy
For conventional plastic sections, cells were fixed with 2.5% glutaraldehyde in 100 mM cacodylate, pH 7.4, for 1 h at room temperature, washed once with 100 mM cacodylate, treated with 2% OsO4 for 1 h, treated with 1% uranyl acetate in 50 mM maleate buffer, pH 5.2, for 1 h, dehydrated using a graded ethanol series and acetone, and pelleted before embedding in Epon and sectioning. For protein A gold labeling of Lag-stained cryosections, cells were fixed in 4% paraformaldehyde (PFA)/100 mM HEPES, pH 7.4, for 1 h at room temperature and then processed exactly as per Sodeik et al. (21).
T cell isolation
PBMCs were obtained by leukapheresis from adult volunteer donors and further purified by centrifugation over Lymphocyte Separation Medium (Organon Teknika, Durham NC) according to the manufacturer’s instructions. Isolated PBMCs were washed three times in HBSS (Mg2+ and Ca2+ free) and either used immediately or suspended in 10% DMSO and 90% heat-inactivated FCS and cryopreserved in liquid nitrogen. No differences were seen in the responses of cells recovered from cryopreservation compared with freshly isolated cells. CD4+ and CD8+ T cells were isolated as described respectively in Ma and Pober (22) and Biedermann and Pober (23).
Alloreaction
Alloreactions were set up in round-bottom 96-well plates in
triplicate. Where not otherwise indicated, 300,000 purified
CD4+ T cells or 150,000 purified
CD8+ T cells were added to 1,000 fixed DCs.
During the last 18–24 h of coculture on the indicated days, 1 µCi
[3H]thymidine (NEN Life Science Products,
Boston, MA) was added to each well, and proliferation was assessed by
[3H]thymidine incorporation. The plates were
harvested with a 96-well harvester (Tomtec, Orange, CT) and counted on
a Microbeta scintillation counter (Wallac, Gaithersburg, MD). For IL-2
detection in alloreaction, Ab to IL-2 receptor 
-chain (anti-TAC,
IgG1, used at 20 µg/ml, a gift from Dr. T. Waldmann) was added to
prevent cytokine utilization. Supernatant was collected on day 1 or day
3 and assayed for IL-2 by ELISA.
RT-PCR assay
Total RNA was isolated from differently treated DC using TRIzol (Life Technologies, Gaithersburg, MD), and 0.5 µg was used for reverse transcription using avain myeloblastosis virus reverse transcriptase (Life Technologies). Primer sequences were as follows: TLR1, 5'-CCTGGCAAGAGCATTGTGGAA and 3'-TGTAATCTATTTCTTTGCTTGCTCTGTCAG; TLR2, 5'-GTGAAGAGTGAGTGGTGCAAGTAT and 3'-CATAAAGATCCCAACTAGACAAAGACTGG; TLR3, 5'-TGGGTCTGGGAACATTTCTCTTCA and 3'-ATAAATTTAATGTACAGAGTTTTTGGATCC; TLR4, 5'-CTGAGCAGTCGTGCTGGTATCATC and 3'-ACCCAGCTGGGCAAGAAATGCCTCAGGAGG; and TLR5, 5'-GCCCAGGGCAGGTGCTTATCT and 3'-GATAACTTGGTGCAAATACAAAGTGAAGA.
IL-12 assay
Cell-culture supernatants were assayed for IL-12 content by ELISA for p70 Ag using a kit from Endogen (Woburn, MA).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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,
TGF-ß, stem cell factor, and Flt3L. Cells were grown for 7–10 days
without further manipulation.
By day 7, total cell number had increased 50- to 100-fold with 35–60%
of the cells being positive for CD1a. A representative profile of
cell-surface expression for various markers at day 8 is shown in Fig. 1
A. Interestingly, and in
contrast with the majority of other methods for producing DCs from
CD34+ precursors, most of the cells in culture
were HLA-DRdim as expected for immature DCs. No
monocyte- or granulocyte-related cells were detected in culture as
indicated by the absence of CD14+ and
CD66b+ cells; these markers were also not
detected at earlier times of culture (days 2–6, data not shown),
suggesting that there was not even transient expression of early
granulocyte markers. The majority of the cells were positive for CD13,
suggesting that they were myeloid in origin. Interestingly, most of the
cells were CLA+, a feature consistent with their
being LC-committed precursors (25).
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B, CD1a expression did not decrease with time in culture
when day 10 cells were replated either in serum-free X-VIVO or in RPMI
1640 containing 5% FCS. However, the serum-fed cells exhibited a
marked increase in surface HLA-DR (data not shown), suggesting that
maturation had been induced by a cytokine or LPS contaminant in the
serum. In any event, it is clear that the presumptive LCs could be
maintained, and apparently in an immature state, for days if grown in a
defined serum-free medium (see below).
We next performed a more detailed analysis of DCs generated from the
mobilized CD34+ precursors. Specifically, we were
interested in determining whether homogenous populations of presumptive
LC precursors could be enriched from the cultures, maintained in an
immature DC phenotype and then triggered to mature in a synchronous
fashion. From day 4 on, loosely adherent aggregates of cells appeared
in culture, which increased in size and number with time. At day 7,
presumptive LCs were found only in multicellular clusters, as indicated
by staining with CD1a and with the unique LC marker Lag
(18) (Fig. 2,
A–D). The cluster cells also exhibited most of their MHC
class II intracellularly with very little on the plasma membrane,
typical of immature DCs (3). These features were highly
reminiscent of the DCs produced from mouse bone marrow in which
proliferating and differentiating DCs accumulate as immature DCs in
analogous cell clusters (26).
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E). The LC marker Lag was expressed by >70% of the
cluster-purified cells. Electron microscopy of the purified cells
revealed the abundant presence of pathognomonic LC-specific organelles,
Birbeck granules (Fig. 2F, arrows). The granules persisted
for several days in immature LC in culture (data not shown). Birbeck
granules were found by protein A-gold immunocytochemistry of
cryosections to be specifically labeled by anti-Lag Abs (Fig. 2G) but negative for MHC class II or Lamp 1 (not shown).
If left undisturbed, cells in the proliferating clusters exhibited this
immature LC phenotype for up to 12 days. However, upon disruption of
the clusters by vigorous repeated pipetting, the cells appeared to
mature as indicated by increased surface expression of MHC class II and
other markers of mature DCs (CD86, CD40, CD80) (Fig. 3B). Yet, when clusters were
purified and gently replated in fresh growth medium (X-VIVO plus
cytokines), the immature phenotype was maintained and spontaneous
maturation was not observed, as indicated by the low level of HLA-DR
and costimulatory molecules expressed on the cell surface (Fig. 3A). Previous attempts to cultivate human DCs from
CD34+ precursors have yielded only mature cells
with high levels of cell-surface MHC class II products and
costimulatory molecules (14, 16). Conceivably, this
difference may reflect the sensitivity of DCs to physical manipulation,
which might disrupt and activate cell clusters.
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, LPS, or
CD40L. Fixed, activated human platelets were used as a source of
membrane-bound CD40L (27). After 2 days, FACS analysis
demonstrated that these agents greatly enhanced the expression of
HLA-DR, costimulatory molecules, and CD83 on the plasma membrane (Fig. 3, C and D and Fig. 5E).
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, LPS, or CD40L induced
a terminal DC differentiation from an immature to a mature phenotype.
Although each of these treatments induced qualitatively similar
differences when assayed by FACS, analysis by confocal microscopy
revealed significant differences among the differently treated cells,
in terms of both morphology and maturation stage. Cells from purified
clusters, replated in regular growth medium, exhibited the phenotype
typical of early DCs: class II is intracellularly distributed and
perfectly colocalized with lysosomal markers (Fig. 4, top row) (3).
Following TNF- treatment (12.5 ng/ml), nearly all of the cells
exhibited the "intermediate" phenotype previously observed only in
rat and mouse DCs. The intermediate DCs were characterized by the
accumulation of MHC class II on the surface as well as in peripheral
intracellular vesicles (class II vesicles or "CIIV") devoid of
lysosomal markers (Fig. 4, second row). The lysosomes were
largely depleted of MHC class II and clustered in the perinuclear
cytoplasm. The accumulation of cells in this intermediate phenotype
suggests a role for TNF- in the early but not the late events of DC
or LC activation. Thus, TNF- may be unable to drive the maturation
process to completion.
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-treated cells. The
maturation driven by LPS seemed to be more advanced; the cells appeared
smaller, but with abundant phyllopodia and more of the MHC class II at
the plasma membrane than in the TNF--treated cells (Fig. 4, third row). Therefore, the LPS-treated cells were
reminiscent of the mature or "late" stage identified in the murine
system, although the structure of the lysosomal compartment in
LPS-treated human DCs was much less compact, and indeed more similar to
what was observed in intermediate cells.
Activated platelets, a source of CD40L, induced the most dramatic
change in cell morphology (Fig. 4, bottom row). Again,
virtually all of the MHC class II was found at the plasma membrane,
which was now organized into long dendrites. Class II-negative
lysosomes were tightly clustered in the perinuclear region.
Importantly, these mature phenotypes were stable, with the LCs expressing high levels of surface HLA-DR and maintaining CD1a for days (data not shown). Indeed, simply plating clusters in RPMI 1640 supplemented only with FCS, as opposed to cytokine-supplemented X-VIVO, supported LC maturation. Together, these findings indicate that LCs prepared in this way are irreversibly committed to a DC phenotype.
We then asked if the observed differences in morphology and cell
organization corresponded to differences in function. We tested the T
cell stimulatory capacity of the differently treated DCs in an
alloreaction using CD8+ T cells. DCs were cluster
purified at day 7 and replated in regular growth media or in the
presence of LPS or TNF-; on day 9 DCs were mixed with
CD8+ T cells. After 2–6 days, cells were
harvested, washed extensively, and proliferation was evaluated. As
shown in Fig. 5A, LPS-treated
DCs were the most efficient in stimulating CD8+ T
cell proliferation when compared with control or TNF--treated DCs,
although the differences were not very pronounced. In fact, even
control DCs (i.e., cells not treated with TNF- or LPS before assay)
were able to support a T cell response.
Interaction with T cells alone is almost certainly capable of inducing
DC maturation due to the presence of CD40L on T cells. Thus, it was
likely that maturation of even the untreated control DCs occurred
during the course of the proliferation assay, obscuring any possible
differences in stimulatory capacity. To circumvent this problem, we
decided to use for the assay previously fixed DCs to prevent further
maturation due to T cell contact. As before, control, TNF--, and
LPS-treated DC were harvested, but fixed with 0.5% PFA and washed
extensively, before coculture with freshly isolated
CD4+ and CD8+ T cells.
Although the magnitude of the T cell proliferative response was
markedly lower using fixed DCs, the results clearly indicated that both
CD4+ and CD8+ responses
were completely dependent on prior exposure of the DCs to a
maturational stimulus with LPS-treated DCs being the most efficient
(Fig. 5B). Similar results were obtained when IL-2
production by CD4+ cells was assayed (Fig. 5C). Despite the differences in stimulatory capacity between
TNF-- and LPS-treated DCs, both treatments caused equivalent
up-regulation of MHC class I, MHC class II, LFA-3 (CD58), and the
DC-maturation marker CD83 (Fig. 5E). Although just 10% of
the total T cell stimulatory activity was left upon DC fixation, the
magnitude of these responses was still clearly dependent on the number
of DCs added to the assay (Fig. 5D). The T cell stimulatory
activity of human DC seems to be much more affected by PFA fixation
than what has previously been shown for mouse DCs (3).
While this difference may reflect differences between the mouse and
human DCs used, it should also be noted that the murine DC studies were
conducted using T cell hybridomas as a read out, as opposed to the
primary T cells used here.
One interesting feature of our culture system is that the cells are
sensitive to LPS, although they do not express CD14, generally thought
to be an important LPS receptor. Because the cells were grown in
serum-free media, another potential mediator of LPS responsiveness, LPS
binding protein, was also unlikely to play a role (28).
Recently, a new family of molecules involved in the response to LPS,
the Toll-like receptors (TLR1 through TLR5), has been cloned and
partially characterized (29). Therefore, we investigated
which, if any, TLRs were expressed by our in vitro-derived LCs and if
their levels of expression level was affected by maturation. Using
specific oligonucleotide primers for each of the human TLR molecules,
RT-PCR was performed on cDNA obtained from cluster-purified cells
treated for 48 h with or without TNF- or LPS. As shown in Fig. 6A, we observed distinct
expression patterns for the five receptors depending on the type of
inflammatory agent used. While the expression of TLR1 mRNA was almost
unchanged regardless of whether TNF- or LPS were added to the
cultures, LPS but not TNF- induced a dramatic down-regulation of
TLR2. Conversely, expression of TLR3 and TLR4 were up-regulated to
varying extents by both mediators, but more so by LPS than by TNF-.
Thus, TNF- and LPS exert markedly different effects on the
expression of individual members of the TLR family, further emphasizing
that different mediators can elicit qualitatively different types of DC
maturation.
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-matured, DCs produced significant levels
of IL-12 (Fig. 6B). |
Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Maturation was induced by three different mediators, TNF-, LPS, and
CD40L. Each of these agents generated mature cells that were similar on
the basis of their surface marker profiles as determined by flow
cytometry. However, several striking differences were uncovered by more
detailed cell biological and functional analysis. By confocal
microscopy, each agent was found to produce cells exhibiting markedly
different morphologies. Perhaps most important was the finding that
TNF- treatment drove the cells no further than the
"intermediate" DC phenotype, characterized by an accumulation of
nonlysosomal intracellular vesicles that, in mouse bone marrow-derived
DCs, carry peptide-loaded MHC molecules to the plasma membrane
(3).5
Thus, TNF- may be capable of inducing only the earliest steps in DC
maturation. This is in contrast to LPS and CD40L that can convert
immature cells, which typically localize the bulk of the MHC class II
products in late endosomes and lysosomes, to mature cells that express
little if any intracellular class II (4, 11, 31).
The phenotypic differences observed were reflected, at least in part,
at a functional level. Using an allogeneic response as an assay,
diminished T cell stimulatory activity was consistently observed when
TFN--treated rather then LPS-treated DCs were used as APCs. We also
observed that the important Th1-driving cytokine, IL-12, was produced
only by LPS-treated DCs and not by TNF--treated cells.
We also found that the cells were markedly different in another feature
of likely functional importance, namely the expression of TLR family
members (32, 33). Recent evidence strongly suggests that
TLRs play critical roles in innate immunity by providing a means
whereby cells of the innate immune system distinguish between self and
invading microorganisms (34). LPS is thought to bind and
activate at least two members of this family, TLR2 and TLR4 (35, 36). Thus, the expression of these two receptors and their
relatives can be expected to play an important role in DC function.
Interestingly, we found that LPS and TNF- elicited markedly
different patterns of TLR gene expression following induction of DC
maturation. In particular, LPS induced the complete down-regulation of
mRNA encoding one of its presumptive receptors TLR2, while TNF- had
no effect on TLR2 expression. On the contrary, TLR4 is completely
absent in immature cells, and its expression is differentially induced
upon maturation. That LPS induced higher levels of TLR3 and TLR4 than
did TNF- implies a further qualitative difference in the effects of
these mediators, but also may reflect the ability of LPS to drive
maturation to "completion" or at least beyond the intermediate
phenotype elicited by TNF-.
The functional consequences of TLR expression on DC or LC function are not yet known, although it does seem likely that these alterations will control one or more important activities. Cella et al. (37) have recently found that influenza virus infection or double-stranded RNA can cause the maturation of human monocyte-derived DCs in a fashion that favors their ability to selectively trigger the development of Th1 T lymphocytes. This observation also clearly illustrates that different DC activators can generate mature DCs with important functional differences.
The advent of a cell culture system that allows the differentiation and controlled maturation of DCs from uncommitted precursors itself has created some interesting opportunities. First, because the cells produced under the conditions used in our experiments appear identical with epidermal LCs, it should now be possible to apply the same cell biological and functional techniques to the study of LCs as have previously been used to study monocyte-derived or bone marrow-derived DCs. Cell fractionation has already enabled us to isolate highly purified fractions of Birbeck granules, which can now be analyzed biochemically in vitro (M.A.V., M.W.E., and I.M., unpublished observations).
In addition, our system for LC production from mobilized CD34+ cells may have good potential for clinical applications. Not only can large numbers of autologous cells be produced with reasonable ease, but the fact that the LCs generated remain immature in culture will allow them to remain in the state best suited for Ag accumulation. Also relevant to any potential clinical applications was the fact that once differentiated, the LCs we have produced are phenotypically stable. To be maximally effective, DCs must maintain their differentiated features upon reintroduction into the bloodstream or tissues, environments in which the cytokine composition is markedly different from the medium in which they had been cultured. DCs differentiated from peripheral blood monocytes, for example, may rapidly lose their DC features upon removal of IL-4 or GM-CSF, suggesting that they may not resemble DCs for long after in vivo reintroduction. While more work will be required to document the in vivo behavior of any reintroduced DC population, the in vitro stability of the CD34-derived LCs described here suggest that they may have critical advantages as potential immunotherapeutic vehicles.
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Acknowledgments |
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Footnotes |
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2 E.G. and M.A.V. contributed equally to this work.
3 Address correspondence and reprint requests to Dr. Ira Mellman, Department of Cell Biology, Yale University School of Medicine, 333 Cedar Street, P.O. Box 208002, New Haven, Connecticut 06520-8002. E-mail address:
4 Abbreviations used in this paper: DC, dendritic cell; LC, Lanerhans cell; CD40L, CD40 ligand; Flt3L, Flt3 ligand; CLA, cutaneous lymphocyte-associated antigen; TLR, Toll-like receptor; PFA, paraformaldehyde.
5 S. Turley, K. Inaha, W. Garrett, M. Ebersold, R. M. Steiman, and I. Mellman. Selective transport of MHC class II-peptide complexes and costimulatory molecules to the surface of developing dendritic cells. Submitted for publication.
Received for publication July 26, 1999. Accepted for publication January 14, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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cooperate in the generation of dendritic Langerhans cells. Nature 360:258.[Medline]
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L. Hertel, V. G. Lacaille, H. Strobl, E. D. Mellins, and E. S. Mocarski Susceptibility of Immature and Mature Langerhans Cell-Type Dendritic Cells to Infection and Immunomodulation by Human Cytomegalovirus J. Virol., July 1, 2003; 77(13): 7563 - 7574. [Abstract] [Full Text] [PDF]
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 A. Camporeale, A. Boni, G. Iezzi, E. Degl'Innocenti, M. Grioni, A. Mondino, and M. Bellone Critical Impact of the Kinetics of Dendritic Cells Activation on the in Vivo Induction of Tumor-specific T Lymphocytes Cancer Res., July 1, 2003; 63(13): 3688 - 3694. [Abstract] [Full Text] [PDF]
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 M. Lu, M. Zhang, R. L. Kitchens, S. Fosmire, A. Takashima, and R. S. Munford Stimulus-dependent Deacylation of Bacterial Lipopolysaccharide by Dendritic Cells J. Exp. Med., June 16, 2003; 197(12): 1745 - 1754. [Abstract] [Full Text] [PDF]
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A. L. Ackerman and P. Cresswell Regulation of MHC Class I Transport in Human Dendritic Cells and the Dendritic-Like Cell Line KG-1 J. Immunol., April 15, 2003; 170(8): 4178 - 4188. [Abstract] [Full Text] [PDF]
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G. Ratzinger, J. L. Reagan, G. Heller, K. J. Busam, and J. W. Young Differential CD52 expression by distinct myeloid dendritic cell subsets: implications for alemtuzumab activity at the level of antigen presentation in allogeneic graft-host interactions in transplantation Blood, February 15, 2003; 101(4): 1422 - 1429. [Abstract] [Full Text] [PDF]
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S. Beaulieu, D. F. Robbiani, X. Du, E. Rodrigues, R. Ignatius, Y. Wei, P. Ponath, J. W. Young, M. Pope, R. M. Steinman, et al. Expression of a Functional Eotaxin (CC Chemokine Ligand 11) Receptor CCR3 by Human Dendritic Cells J. Immunol., September 15, 2002; 169(6): 2925 - 2936. [Abstract] [Full Text] [PDF]
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F. Geissmann, M.C. Dieu-Nosjean, C. Dezutter, J. Valladeau, S. Kayal, M. Leborgne, N. Brousse, S. Saeland, and J. Davoust Accumulation of Immature Langerhans Cells in Human Lymph Nodes Draining Chronically Inflamed Skin J. Exp. Med., August 19, 2002; 196(4): 417 - 430. [Abstract] [Full Text] [PDF]
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 Y. Nagata, S. Ono, M. Matsuo, S. Gnjatic, D. Valmori, G. Ritter, W. Garrett, L. J. Old, and I. Mellman Differential presentation of a soluble exogenous tumor antigen, NY-ESO-1, by distinct human dendritic cell populations PNAS, August 6, 2002; 99(16): 10629 - 10634. [Abstract] [Full Text] [PDF]
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N. Hilf, H. Singh-Jasuja, P. Schwarzmaier, C. Gouttefangeas, H.-G. Rammensee, and H. Schild Human platelets express heat shock protein receptors and regulate dendritic cell maturation Blood, May 15, 2002; 99(10): 3676 - 3682. [Abstract] [Full Text] [PDF]
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 A. R. Hayman, P. Macary, P. J. Lehner, and T. M. Cox Tartrate-resistant Acid Phosphatase (Acp 5): Identification in Diverse Human Tissues and Dendritic Cells J. Histochem. Cytochem., June 1, 2001; 49(6): 675 - 684. [Abstract] [Full Text]
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P. A. MacAry, M. Lindsay, M. A. Scott, J. I. O. Craig, J. P. Luzio, and P. J. Lehner Mobilization of MHC class I molecules from late endosomes to the cell surface following activation of CD34-derived human Langerhans cells PNAS, March 27, 2001; 98(7): 3982 - 3987. [Abstract] [Full Text] [PDF]
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 E. Cario and D. K. Podolsky Differential Alteration in Intestinal Epithelial Cell Expression of Toll-Like Receptor 3 (TLR3) and TLR4 in Inflammatory Bowel Disease Infect. Immun., December 1, 2000; 68(12): 7010 - 7017. [Abstract] [Full Text] [PDF]
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E. J. Soilleux, R. Barten, and J. Trowsdale Cutting Edge: DC-SIGN; a Related Gene, DC-SIGNR; and CD23 Form a Cluster on 19p13 J. Immunol., September 15, 2000; 165(6): 2937 - 2942. [Abstract] [Full Text] [PDF]
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 M. Brenner, C. Rossig, U. Sili, J. W. Young, and E. Goulmy Transfusion Medicine: New Clinical Applications of Cellular Immunotherapy Hematology, January 1, 2000; 2000(1): 356 - 375. [Abstract] [Full Text] [PDF]
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90%) of
cells after cluster purification and reculture are also
Lag+. F, Electron microscopy analysis of LC
in culture reveal the presence of abundant Birbeck granules (indicated
by arrowheads). G, Only BGs (and not other structures
such as multilamellar lysosomes, indicated by asterisks) were heavily
decorated by anti-Lag Abs. However, the multilamellar structures
did label with Abs to MHC class II molecules (not shown).
