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Blood, Vol. 96 No. 3 (August 1), 2000:
pp. 878-884
CLINICAL OBSERVATIONS, INTERVENTIONS, AND THERAPEUTIC TRIALS
In vivo generation of human dendritic cell subsets by Flt3 ligand
Eugene Maraskovsky,
Elizabeth Daro,
Eileen Roux,
Mark Teepe,
Charlie R. Maliszewski,
Jeannie Hoek,
Dania Caron,
Mel E. Lebsack, and
Hilary J. McKenna
From Immunex Corporation, Seattle, WA.
 |
Abstract |
Dendritic cells (DCs) represent a family of ontogenically distinct
leukocytes involved in immune response regulation. The ability of DCs
to stimulate T-cell immunity has led to their use as vectors for
immunotherapy vaccines. However, it is unclear whether and to what
degree in vitro-generated DCs are representative of DCs that develop
in vivo. Treatment of mice with human Flt3 ligand (FL) dramatically
increases the number of DCs. We report here that administration of FL
to healthy human volunteers increased the number of circulating
CD11c+ IL-3R low DC (mean 44-fold) and
CD11c IL-3Rhigh DC precursors (mean
12-fold). Moreover, the CD11c+ DCs were efficient
stimulators of T cells in vitro. Thus, FL can expand the number of
circulating, functionally competent human DCs in vivo.
(Blood. 2000;96:878-884)
© 2000 by The American Society of Hematology.
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Introduction |
Dendritic cells (DCs) are rare, bone marrow
(BM)-derived antigen-presenting cells (APCs) that are involved in
immune surveillance, antigen (Ag) capture, and Ag
presentation.1 DCs are uniquely able to present Ag to naive
T cells and induce their activation and proliferation.1 DCs
can be generated from myeloid-committed precursors, including
CD14+ and CD14 BM-derived precursors,
Langerhans cells of the skin (Lc-DCs),1,2 peripheral blood
(PB) monocytes (monoDCs),3,4 and immature neutrophils.5 Alternatively, DCs can be generated from
lymphoid-committed precursors, which also give rise to B cells, natural
killer (NK) cells, and T cells.6-9 The myeloid growth
factor granulocyte-macrophage colony-stimulating factor (GM-CSF) is
obligate for the development of myeloid-related DCs in vitro but does
not appear to be required for DC development from
lymphoid-committed precursors.10 The seeming
diversity in DC subsets and their distinct anatomic locations suggest
that DCs may be composed of a system of several cell types, each of
which may express overlapping but distinct functions.
In the PB of healthy individuals, DCs represent a minor fraction
(< 1%) of mononuclear cells and can be distinguished from other
mature cell lineages by their characteristic dendritic morphology; their lack of cell surface expression of CD3, CD14, CD19, and CD56;
their high level of expression of CD1b/c, CD11c, CD33, and HLA-DR; and
their expression of CD4. In addition, a DC precursor subset lacking
CD11c expression but expressing interleukin (IL)-3R , and previously referred to as the plasmacytoid T cell, has been identified in lymphoid tissue and PB.11-14
The efficient capacity of DCs to initiate and regulate
lymphocyte-mediated immunity has led to the study of DCs as cellular vaccine adjuvants for the immunotherapy of cancer.15,16
Numerous studies in mice have demonstrated the efficacy of DC-based
immunization protocols for the generation of antitumor
responses.15 However, despite these promising results, the
feasibility of using DCs as cellular vectors for immunotherapy is
limited by the extremely small number of DCs that can be isolated
from human PB. Moreover, the use of only 1 type of DC precursor
(ie, monocytes) or the preferential expansion of only myeloid-related
DCs in vitro for use in clinical studies may not necessarily
provide the appropriate APC-derived signals that would
result in the generation of optimal T-cell immunity.17 The
use of cytokines that increase the number of DCs in vivo without
altering subset diversity may obviate many of the potential problems
surrounding current DC-based immunotherapy strategies.
Flt3 ligand (FL) is a hematopoietic growth factor that induces the
proliferation and survival of primitive hematopoietic progenitor and
stem cells.18 When administered to mice, FL has the unique ability to expand the number of both myeloid-related and
lymphoid-related DC subsets.19,20 The expansion of DCs in
mice has resulted in successful vaccination with a soluble Ag in
the absence of chemical adjuvants21 as well as the
generation of protective immune responses to established
tumors.22,23 A phase I clinical study was therefore
performed in healthy human volunteers, and the ability of FL to expand
DCs in these subjects was examined. Our findings indicate that FL can
be administered to humans to expand significantly the number of
circulating CD11c+ and CD11c DC subsets
and DC precursors.
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Patients, materials, and methods |
Administration of FL to healthy human volunteers
PB samples were obtained from a randomized, placebo-controlled,
phase I dose-escalation study performed in healthy human volunteers. The study was conducted at the Phase I clinic of Pharmaco International (Austin, TX), a commercial contract research organization. The protocol
was approved by Pharmaco's IRB, Research Consultants' Review
Committee, and informed consent was obtained from all individuals. FL
was produced by recombinant DNA technology in a Chinese hamster ovary
(CHO) cell line. FL was supplied as a sterile lyophilized preparation
of 5 mg of FL, with 40 mg mannitol, 10 mg sucrose, and 25 mmol/L of
tromethamine (Tris) per vial. It was reconstituted before
administration in bacteriostatic water for injection. Twenty volunteers
received once-daily subcutaneous injections of 10, 25, 50, 75, or 100 µg · kg1 · d1 of human
FL (n = 3 per group) or matching placebo (n = 1 per group) for 14 consecutive days. PB samples (20 mL) were obtained on
days 1 (before the first injection), 3, 5, 7, 9, 11, 13, 15, 17, 19, and 21 of FL or placebo administration. Safety monitoring included
clinical laboratory evaluation and assessment of adverse events.
Treated individuals were monitored on site for signs of adverse events.
Differential white blood cell (WBC) count and serum chemistries were
performed. Samples were analyzed by flow cytometry for changes in the
distribution of various leukocyte populations.
Cell preparation and flow cytometric isolation of
peripheral blood mononuclear cell (PBMC) populations
PBMCs were isolated after centrifugation over a discontinuous
density gradient using Ficoll (1.077 g/mL, Accu-Prep; Accurate Chemicals, Westbury, NY), followed by centrifugation in
phosphate-buffered saline (PBS) containing 5% fetal bovine serum (FBS)
(Intergen, New York, NY). Cells were then incubated with directly
conjugated antibodies, which included CD1a, CD1b/c, CD2, CD3, CD4, CD5,
CD8, CD11b, CD11c, CD14, CD19, CD33, CD40, CD56, CD80, CD86, CD123w (IL-3R), CD116 (GM-CSFR), HLA-DR, macrophage mannose receptor (MMR) (Pharmingen, San Diego, CA), and CD83 (Immunex
Corp, Seattle, WA), for 30 minutes at 4°C in PBS containing 0.01%
NaN3 supplemented with 10% goat serum and 10% rabbit
serum to block Fc receptors. Propidium iodide (PI) (1 µg/mL) was
included in the final wash to allow exclusion of dead cells.
Isotype-matched antibodies were used as negative controls.
Multiparameter flow cytometric analysis and sorting of PBMCs were
performed on a FACStar Plus (Becton Dickinson, San Jose, CA). Cytospins
of sorted PBMC populations were performed by centrifugation of
5 × 104 cells onto slides (500 rpm), followed by
staining with Wright-Giemsa according to the manufacturer's
instructions (Fisher Diagnostics, Pittsburgh, PA).
Generation of CD1a+ DCs from
CD34+ BM cells
CD1a+ DCs were derived by culturing CD34+ BM
progenitor cells. BM cells were obtained from healthy donors (kindly
provided by Dr M. A. Caligiuri, Roswell Park Cancer Institute, Buffalo,
NY). The mononuclear cells were isolated after centrifugation over a
discontinuous Ficoll density gradient, followed by centrifugation in
PBS containing 5% FBS. CD34+ BM cells were isolated using
an anti-CD34 immunoaffinity column according to the manufacturer's
instructions (Ceprate; Cell Pro, Bothell, WA). Briefly, BM mononuclear
cells were incubated with anti-CD34-biotin and passed over a
streptavidin-conjugated Ceprate gel matrix column. The column was
washed extensively, and bound CD34+ BM cells were dislodged
from the Ceprate column matrix by mechanical disruption and collected
in PBS containing 5% FBS. Fluorescence-activated cell sorter (FACS)
analysis revealed a purity of 85% to 92% CD34+ cells. DCs
were generated by culturing CD34+ cells with recombinant
human (rhu)GM-CSF (20 ng/mL), rhuIL-4 (20 ng/mL), rhu
tumor necrosis factor (TNF)- (20 ng/mL), plus rhuFL (100 ng/mL) in
modified McCoy's culture medium for 14 days. DCs were purified by cell
sorting on the basis of CD1a, CD86, and HLA-DR expression. All
cytokines used in these cultures were produced and purified at Immunex.
Mixed leukocyte culture (MLC) and
Ag-specific presentation assays
T cells were isolated from PBMCs of healthy donors using opsonized
sheep red blood cells. Purified CD4+ T cells (90% to 95%)
were isolated using the MACS CD4 T-cell isolation kit (Miltenyi Biotec,
Sunnyvale, CA). MLC or Ag presentation assays were performed in 96-well
round-bottom culture plates (Nunc, Naperville, IL). Autologous T cells
were obtained from cryopreserved PBMCs obtained from healthy volunteers
before FL treatment. Allogeneic CD4+ T cells
(1 × 105) were incubated with varying numbers of
irradiated (2000 rad), cell-sorted PBMC populations from FL-treated
individuals or control CD1a+ DCs in 0.2 mL modified
McCoy's medium containing 10% FBS and 104
2-mercaptoethanol (culture medium) for 5 days in humidified 10% CO-2 in air. Autologous CD4+ T cells
(1 × 105) were incubated with irradiated (2000 rad)
CD11c+ CD14 DCs or CD11c+
CD14+ monocytes from FL-treated individuals in 96-well
round-bottom plates in AIM-V serum-free medium (Life Technologies,
Gaithersburg, MD) with or without 25 µg/mL each of either keyhole
limpet hemocyanin (KLH; Calbiochem Corp, La Jolla, CA), tetanus toxoid
(TT; Connaught, Swiftwater, PA), ovalbumin (Ova; Pierce Chemical Co.,
Rockford, IL), or hepatitis B surface Ag peptide (HBsAg peptide; Sigma, St Louis, MO) for 6 days at 37°C in 10% CO2 in
air. The cultures were then pulsed with 0.5 µCi
3H-thymidine for 24 hours, and the cells were harvested
onto glass fiber sheets for counting on a gas-phase  counter. Values
represent the mean ± SEM of 3 replicate cultures.
Fluid-phase uptake of fluorescein isothiocyanate
(FITC)-ovalbumin
Fluid-phase uptake was assessed by incubation of PBMCs with 2 mg/mL
FITC-Ova (Molecular Probes, Eugene, OR) at either 0°C or 37°C
for 30 minutes. The 0°C samples were first metabolically fixed by
preincubation in 0.1% sodium azide at 37°C and rapidly chilled
before incubation with FITC-Ova. These metabolically fixed, 0°C
samples were used to assess nonspecific cell-surface staining. The
cells were then incubated with anti-CD14-APC and
anti-CD11c-PE (Pharmingen), and PI was used to exclude
dead and dying cells. CD11c+ CD14 DCs,
CD11c+ CD14+ monocytes, and
CD11c CD14 lymphocytes were gated
electronically and examined for FITC-Ova uptake as a function of the
mean fluorescence intensity (MFI) in FITC staining. The data are
presented as the MFI ± SEM of 3 FL-treated individuals, and similar
results were obtained in 3 separate experiments. Examination of cells
by fluorescence microscopy demonstrated that FITC-Ova was internalized.
| |
Results |
FL increases circulating CD11c+ DCs in vivo
FL was well tolerated by all subjects at all doses tested compared
with the placebo controls, with only 2 adverse events considered to
be related to FL administration: enlarged lymph nodes and
injection-site reactions. FL did not cause allergic-type reactions. FL
increased the number of WBCs, PBMCs, and CD14+ monocytes
but did not significantly alter the number of circulating lymphocytes.24
Flow cytometric analyses of the PBMC fraction from a
representative FL-treated volunteer (75 µg/kg) and a placebo-treated volunteer are presented in Figure 1A. PBMCs
that lacked the mature lineage markers CD3, CD14, CD19, and CD56
(Lin) and expressed CD33 or HLA-DR were rare in the
blood of placebo-treated individuals (Figure 1A) and also in
pretreatment samples (data not shown). However, after 14 days of FL
treatment, approximately 14% of PBMCs were Lin and
CD33+. Similarly, approximately 14% of PBMCs expressed
CD11c but not the monocyte marker CD14 (Figure 1B). Four-color flow
cytometry revealed that all Lin CD33+
cells expressed CD11c. In contrast, approximately 19% of PBMCs were
Lin HLA-DR+ (Figure 1A), suggesting that
FL treatment generated a population of approximately 4% of
Lin HLA-DR+ cells that were
CD33 CD11c and that were rare in
the blood of placebo-treated individuals (discussed later). Finally, FL
treatment increased the proportion of the CD11c+
CD14+ monocyte fraction (Figure 1B).
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| Fig 1.
Flow cytometric analysis of PBMCs from healthy volunteers
treated with either placebo or FL for 14 consecutive days.
(A) Correlation of lineage-specific marker expression (CD3, CD14, CD19,
and CD56) with CD33 or HLA-DR expression. (B) Correlation of CD14 with
CD11c expression. (C) Expression of cell surface molecules on
CD11c+ CD14 PBMCs from FL-treated
individuals. CD1a, CD1b/c, CD2, CD4, CD5, CD8, CD11b, CD40, CD80,
CD83, CD86, HLA-DR, GM-CSFR, IL-3R, and MMR expression are
presented as the shaded histograms. Unshaded histograms represent
CD11c+ CD14 cells incubated with
isotype-matched controls. (D) Expression of CD5 and MMR on gated
CD11c+ CD14 PBMCs. Data are presented
from a representative individual treated with FL at 75 µg · kg1 · d1. Similar
profiles were observed with treatment at 10, 25, 50, and 100 µg · kg1 · d1.
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Further phenotypic analysis of the FL-generated
CD11c+ CD14 PBMCs revealed that they
were CD1a, CD1b/c+, CD2+,
CD4+, CD8, CD11b+,
CD40, CD80,
CD83, CD86+, and
HLA-DRbright (Figure 1C), a phenotype consistent with PB
DCs (hereafter referred to as CD11c+ DCs). Analysis
revealed that the gates used for CD11c+ DCs contained less
than 1% CD16+ cells, indicating that the
CD14low CD16+ monocytes were not represented
within this gate (25,26 and data not shown). The expression
of CD11b and the low to undetectable levels of CD40, CD80, and CD83
suggested that the circulating CD11c+ DCs were not fully
mature.27-31 However, DCs isolated from oncology patients
treated with FL up-regulate costimulatory molecules in short-term
cultures (15 hours) supplemented with GM-CSF plus IL-4 (manuscript in preparation), indicating that FL-generated DCs can be
further matured. The CD11c+ DCs expressed detectable levels
of GM-CSFR but low to negligible levels of IL-3R
(Figure 1C). A subset of the CD11c+ DCs expressed the
lymphoid marker CD5 (Figure 1C). Similarly, a subset of
CD11c+ DCs expressed MMR, which is involved in
receptor-mediated endocytosis.28,32 Interestingly,
expression of CD5 and MMR was mutually exclusive, and there were 3 distinct populations of CD11c+ DCs: CD5+
MMR, CD5 MMR+, and
CD5 MMR (Figure 1D).
Wright-Giemsa staining of cytospins of the sorted PBMC populations
indicated that the FL-generated CD11c+ DCs displayed a
distinctive multilobulated nuclear morphology and expressed veiled and
dendritic processes, typical of PB DCs (Figure
2A). Unlike monocytes from the
placebo-treated individuals (Figure 2D), the CD14+
monocytes from FL-treated individuals (Figure 2B) also contained multilobulated nuclei but did not express dendritic processes, suggesting that FL treatment had altered the morphology of monocytes within the CD11c+ CD14+ fraction. Finally, the
CD11cdull CD14dull cells from FL-treated
individuals (Figure 1B) were identified as neutrophils based on their
polymorphonuclear morphology and size (Figure 2C).
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| Fig 2.
Morphology of sorted PBMCs from FL- or placebo-treated
volunteers.
(A) CD11c+ CD14 DC fraction from an
FL-treated individual. (B) CD11c+ CD14+
monocyte fraction from an FL-treated individual. (C)
CD11cdull CD14dull neutrophil fraction from an
FL-treated individual. (D) CD11c+ CD14+
monocyte fraction from a placebo-treated individual. Cytospins were
prepared from the sorted PBMC populations depicted in Figure 1B and
stained with Wright-Giemsa stain.
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FL-generated CD11c+ PB DCs stimulate
Ag-specific T-cell proliferation
Various PBMC subsets were examined for their capacity to stimulate
the proliferation of alloreactive T cells in vitro in MLC.First, total PBMCs taken before and after FL treatment from the same donor
were compared. PBMCs containing maximal numbers of circulating DCs (day
15 of FL treatment) contained significantly higher MLC stimulatory
capacity than PBMCs derived from the pretreatment sample (day 0 of FL
treatment) (Figure 3). This suggests that the increased number of circulating DCs after FL treatment resulted in
enhanced MLC-stimulating capacity of the PBMCs. To demonstrate definitively that this enhancement was due to the increased numbers of
DCs in FL-treated volunteers, we sorted DCs on the basis of CD11c,
HLA-DR, and lack of CD14 expression. These sorted DCs were compared
with sorted CD1a+ DCs generated by culturing
CD34+ BM cells in GM-CSF, IL-4, FL, and TNF- for 14 days
for their capacity to stimulate alloreactive T-cell proliferation in
vitro. The CD11c+ DCs from an FL-treated individual were
30- and 300-fold more efficient at stimulating alloreactive T cells to
proliferate than the autologous CD11c+ CD14+
monocyte fraction or the CD11clo CD14lo
neutrophil fraction, respectively (Figure
4A). Interestingly, the CD11c+
DCs from FL-treated donors were 3-fold less efficient than
CD1a+ BM-derived DCs. Although the DCs generated from the
BM and the PB were not derived from the same donors, and therefore
direct comparison of APC potency is difficult, the trend in a number of
experiments was always for the BM-derived DCs to stimulate higher
levels of proliferation in the MLC experiments. In addition, CD11c+ DCs from an FL-treated individual were more
efficient than CD11c+ CD14+ monocytes at
stimulating the proliferation of autologous CD4+ T cells to
recall Ag (TT) and were the only cells capable of inducing T-cell
proliferation to the nominal Ags Ova, KLH, or HBsAg (Figure 4B). These
data suggest that FL-generated PB DCs can capture, process, and present
protein or peptide Ag to naive and memory autologous T cells and induce
their expansion in vitro.
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| Fig 3.
Function of in vivo FL-generated PBMCs.
Alloreactive T-cell-stimulating capacities of PBMCs from day-0 ( )
and day-15 PB ( ) samples from the same donor were
compared. Data represent the mean ± SEM of
triplicate wells. Results from a representative FL-treated donor and
the corresponding placebo-treated individual from the 25-µg/kg cohort
are presented. Similar results were seen with PBMCs from other
FL-treated donors.
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| Fig 4.
Function of in vivo FL-generated DCs.
(A) Comparison of alloreactive T-cell-stimulating capacity
of the various PBMC populations depicted in Figure 1B. Sorted
CD11c+ CD14+ monocytes (),
CD11c+ CD14 DCs ( ), and
CD11clo CD14lo neutrophils
( ) from day-15 PB samples were compared with control
CD1a+ DCs generated in vitro from CD34+ BM
progenitors ( ). Data represent the mean ± SEM of triplicate
wells. Results are representative of 4 separate experiments. (B)
Comparison of Ag-induced T-cell proliferation by purified
CD11c+ CD14 DCs and CD11c+
CD14+ monocytes from FL-treated individuals.
CD11c+ CD14 DCs or CD11c+
CD14+ monocytes were isolated by flow cytometry and
cultured with purified autologous T cells derived from cryopreserved PB
samples taken before commencement of the study, in the presence of
recall Ag (TT) or nominal Ags (Ova, KLH, or HBsAg peptide). Data
represent the mean ± SEM of triplicate wells. Results are
representative of 3 separate experiments. (C) Comparison of Ag uptake
by PBMCs from FL-treated individuals. PBMCs were cultured with FITC-Ova
at either 0°C or 37°C for 30 minutes and then incubated with
anti-CD14-APC and anti-CD11c-PE. CD11c+
CD14 DCs, CD11c+ CD14+
monocytes, and the enriched lymphocyte fraction
(CD11c CD14) were examined by
flow cytometry for internalized FITC-Ova. Cells incubated at 0°C
were used to discriminate between nonspecific cell surface staining and
active uptake at 37°C. The data are presented as the mean
fluorescence intensity (MFI) ± SEM from 3 FL-treated individuals.
Results are representative of 3 separate experiments.
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FL-generated DCs exhibit low fluid-phase uptake
To assess the capacity of FL-generated DCs to capture soluble Ag by
fluid-phase endocytosis and macropinocytosis, we incubated the cells
with 2 mg/mL FITC-Ova for 30 minutes. Examination of the various PBMC
subsets revealed that the FL-generated CD11c+ DCs were
capable of internalizing soluble Ova, albeit less efficiently than
circulating CD14+ monocytes (Figure 4C). As expected, the
lymphocyte-enriched fraction (CD11c
CD14) was inefficient at uptake of soluble FITC-Ova
(Figure 4C). Because immature DCs are highly efficient at uptake of
small solutes28,32-34 (eg, Ova), these data suggest that
FL-generated CD11c+ DCs are not immature and may represent
a more intermediate phenotype.
Kinetics of in vivo expansion of CD11c+ DCs with FL
FL-mediated expansion of DCs in mice has been shown to be transient
and reversible, with increased DC numbers detectable 3 days after
initiating FL treatment and a return to basal levels within 7 days of
the last FL injection.19 An increase in circulating human
CD11c+ DCs was detected by day 5 of FL treatment, peaking
at an average 44-fold above basal or placebo control levels by day 9 (Figure 5A). Seven days after the last FL
injection, CD11c+ DC numbers remained well above placebo
levels.
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| Fig 5.
Expansion of CD11c+ DCs in the PB of
FL-treated individuals.
(A) Increase in total numbers of CD11c+ DCs over time in
FL-treated (, , ) and placebo-treated () individuals
from the cohort receiving a dose of 100 µg · kg1 · d1. Similar
results were obtained in the cohorts receiving 10, 25, 50, and 75 µg/kg. (B) Number of CD11c+ DCs detected at day 15 in
each FL-treated or placebo-treated (*) individual from each dose
cohort.
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Interestingly, there was no apparent dose response in the number of DCs
attained after 14 days of FL treatment at a dose ranging from 10 to 100 µg · kg1 · d1,
suggesting that even the lowest dose (10 µg/kg) was capable of
generating maximal numbers of circulating CD11c+ DCs
(Figure 5B). In the 15 FL-treated volunteers, there was a mean 44-fold
increase in the absolute number of circulating CD11c+ DCs,
with DCs constituting approximately 15% of PBMCs or
8.8 × 105 DCs per milliliter of blood (Table
1). Over the same period, the proportion or
absolute number of circulating DCs did not change in the 5 placebo-treated individuals, in whom DCs constituted approximately
1.0% of PBMCs or 1.8 × 104 DCs per milliliter of
blood (Table 1).
FL increases circulating CD11c DC precursors in vivo
A recent report described the isolation of CD4+,
CD11c, CD45RA+ cells, termed
plasmacytoid T cells,11 from human tonsils. These cells are
also in the PB of normal individuals.12-14 They lack surface expression of mature lineage markers (CD3, CD14, CD19, and
CD56) and CD33 but express high levels of IL-3R and
HLA-DR.11,12 When cultured in IL-3 and CD40L, these cells
develop into functional DCs. Furthermore, once matured, these
CD11c DCs appear to differentially regulate the
cytokine repertoire of naive T cells, preferentially generating
IL-4-, IL-5-, and IL-10-secreting T cells as
compared with CD11c+ DCs, which induce interferon
(IFN)- -secreting T cells.14 We examined whether the 4%
of PBMCs in FL-treated individuals that were Lin
CD11c CD33 HLA-DR+
(Figure 1A) expressed IL-3R. The CD11c PBMC
fraction in FL-treated individuals contained an increased proportion of
large cells (based on forward light scatter) lacking mature lineage
markers and expressing high levels of IL-3R (Figure 6A). These cells were increased
approximately 3-fold as a proportion of PBMCs compared with
placebo-treated individuals (or baseline levels). These
CD11c cells were CD4+,
CD40low, CD45RA+, CD86low, and
HLA-DR+ and lacked CD1b/c, CD2, and CD80 expression (Figure
6B). Thus, the Lin CD11c
CD33 HLA-DR+ PBMCs detected in
Figure 1A represent the CD11c IL-3R+
DC precursor subset. The lineage derivation of these
CD11c DC precursors remains unclear. A lymphoid
relationship has been proposed because of their responsiveness to IL-3
but not to GM-CSF.11,13,14 However, others have suggested
that this subpopulation may be of myeloid origin, distinct from that of
the Lc-DC or monoDC pathways.12
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| Fig 6.
Detection of CD11c IL-3R+
DCs in FL-treated but not placebo-treated individuals.
(A) PBMCs from FL-treated (100 µg · kg1 · d1) or
placebo-treated individuals (day 15) were analyzed by flow cytometry
for large (FSChigh) CD11c cells (shown
gated) or cells that lacked the mature lineage markers CD3, CD14, CD19,
and CD56 but expressed high levels of IL-3R. (B) Expression of
CD1b/c, CD2, CD4, CD40, CD45RA, CD80, CD86, and HLA-DR on
CD11c IL-3R+ PBMCs from an FL-treated
individual. Results are representative of 4 separate experiments.
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| |
Discussion |
In this phase I clinical study, increasing doses of FL were
administered subcutaneously to healthy volunteers over 14 consecutive days to evaluate the safety of FL and its effect on the number of
circulating DCs. FL administration was well tolerated and resulted in a
dramatic increase in the numbers of at least 5 types of DCs or DC
precursors. These included: a 44-fold increase in CD11c+ PB
DCs (which can be subdivided into CD5+
MMR, CD5 MMR+, and
CD5 MMR subsets) (Figure 1A, B,
D; Table 1), a 12-fold increase in CD11c
IL-3R+ DC precursors (Figure 6A; Table 1), and a 10-fold
increase in CD14+ monocytes (Figure 1B). Although CD5 and
MMR expression has been detected on blood DCs and on in
vitro-generated monocyte-derived DCs,28,32,35 it is
unclear whether expression of CD5 or MMR defines the maturational
status or ontogenic derivation of the FL-generated CD11c+
DCs. However, the conspicuous absence of a CD5+
MMR+ intermediate subset suggests that the CD5+
MMR and CD5 MMR+ DCs
may represent distinct DC subpopulations rather than DCs progressing along a maturational continuum. The lack of CD16
expression by the CD11c+ DCs argues against the
MMR+ subpopulation representing contaminating
CD14low CD16+ MMR+
monocytes,25,26 as these were excluded by the stringency of gating. Howard et al36 have identified CD5+
MMR and CD5 MMR+ DC
subsets in the afferent lymph of cattle. Although these 2 DC subsets
are equivalent in their capacity to stimulate Ag-specific CD4+ T cells, the CD5 MMR+
DCs are more efficient at priming naive respiratory syncytial virus-specific T cells. These 2 DC subsets may therefore play distinct
roles in the induction of primary immune responses.36
Interestingly, cells in the CD14+ monocyte fraction
from FL-treated individuals were morphologically distinct from
monocytes isolated before treatment and those from placebo control
individuals (Figure 1B, D). The appearance of these
morphologically distinct cells within the CD14+ PBMC
fraction may represent de novo-generated CD14+ monocytes
or CD14+ myelomonocytic precursors that accumulate in the
PB of FL-treated individuals. This would be consistent with the
previously reported myelopoietic effects of FL treatment in
mice.37
It is generally believed that immature DCs are specialized in Ag uptake
and processing, and mature DCs are specialized in T-cell priming and
stimulation.32,34 FL-generated CD11c+ DCs were
capable of fluid-phase Ag uptake, but not to the degree that the
CD14+ monocytes were (Figure 4C). The CD11c+
DCs were also able to stimulate alloreactive T cells (although less so
than the mature BM-derived DCs) and could stimulate proliferation in
response to a recall Ag (TT) and to the nominal Ags KLH, Ova, and HBsAg
peptide. These data suggest that on the basis of their low uptake of
fluid-phase Ag, the FL-generated CD11c+ DCs are relatively
mature compared with immature monoDCs, which are very efficient at Ag
uptake.28,32-34 However, although the levels of Ag capture
exhibited by FL-derived DCs are low, they were sufficient to drive
T-cell proliferative responses, perhaps indicating that these DCs have
other specialized Ag presentation features. It is possible that
FL-generated DCs could have matured during the in vitro culture period
with T cells (4 days), acquiring the capacity to efficiently stimulate
T-cell proliferation. In this regard, recent experiments using DCs
generated in FL-treated cancer patients have indicated that these blood
DCs mature rapidly (15 hours) upon in vitro culture, up-regulating
CD80, CD83, CD86, and HLA-DR expression (manuscript in preparation).
Spontaneous maturation of blood DCs in vitro has also been
reported.38 Although this could explain the apparent
discrepancy between immature surface phenotype and an intermediate to
mature functional capacity, it is also possible that FL-generated blood
DCs may be more related to in vitro-generated Lc-DCs, which are less
efficient at Ag uptake as compared with in vitro-generated
monoDCs.32 FL-generated DCs are, therefore, not completely
immature on the basis of their low Ag capture activity, high cell
surface major histocompatibility complex class II expression, and
efficient Ag presentation, but they are also not fully mature cells
because they express low to negligible levels of CD40, CD80, and
CD8327-31,33 and do not present alloantigen as efficiently
as mature CD1a+, in vitro-generated, BM-derived DCs. Taken
together, the data suggest that FL-generated PB DCs are either
intermediate in their maturation status or are not functionally
equivalent to in vitro-generated monoDCs.
Clinical trials currently examining the safety and efficacy of
using DCs to deliver tumor Ag vaccines in vivo are targeting the
monocyte-derived DC subset.16 Functional distinctions
between Lc-DCs and monoDCs have been reported and suggest that these DC subsets are not equivalent in their capacity to stimulate T- and B-cell
responses.17,32 In addition, unlike Lc-DCs or monoDCs, the
CD11c DCs fail to secrete IL-12 upon stimulation and
secrete high levels of IFN- upon viral infection.14,39
The CD11c DCs may also differentially influence the
types of cytokines that T cells are induced to secrete.14
These studies emphasize how understanding the functional heterogeneity
of DC subsets is essential for their appropriate use in immunotherapy.
In this respect, the most successful immunotherapy strategies will
likely be those that maintain the diversity of DC subsets or those that can specifically target their distinct functional characteristics in vivo.
The ability of FL to expand the number of DCs in vivo for immunotherapy
offers an alternative strategy to the use of in vitro-generated DCs.
FL treatment of healthy volunteers was safe and well tolerated. Large
numbers of functionally competent DCs with various phenotypes were
generated in vivo. The ability to expand a diverse repertoire of DC
subsets may obviate many of the issues regarding which type of in
vitro-generated DC population (monoDCs, Lc-DCs, or lymphoid-related DCs) is most appropriate for the generation of clinically effective immunity in vivo.
FL has been shown to generate effective T-cell and NK cell-mediated
antitumor responses in tumor-bearing mice.22,23 Clinical studies using FL to expand DC populations in cancer patients are currently underway. It will be of great interest to determine whether
the increase in DC numbers will result in increased immunity to
endogenous tumor Ags. As an alternative approach, it may be possible to expand DC subsets by FL treatment, isolate the DCs for transient manipulation with tumor vaccines, and reinfuse them into
cancer patients. Thus, FL may be an important cytokine for augmenting
antitumor immune responses in vivo.
| |
Acknowledgments |
We thank S. Braddy, D. Hirschstein, and A. Alpert for assistance with
flow cytometry and Dr K. Shortman from the Walter and Eliza Hall
Institute, Melbourne, Australia; and Dr M. B. Widmer, Dr D. H. Lynch,
Dr D. E. Williams, and A. Aumell from Immunex Corporation for critical
advice and assistance.
| |
Footnotes |
Submitted July 16, 1999; accepted March 31, 2000.
Reprints: E. Maraskovsky, The Ludwig Oncology Unit, Ludwig
Institute for Cancer Research, 6th Floor, Harold Stokes Building,
Austin and Repatriation Medical Center, Studley Rd, Heidelberg VIC 3084 Australia; e-mail: eugene.maraskovsky{at}ludwig.edu.au.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
| |
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Abstract
Introduction