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Department of Medicine and The Vanderbilt Cancer Center, Vanderbilt University Medical Center, Nashville, TN 37232;
Cardinal Bernardin Cancer Center, Loyola University Medical Center, Maywood, IL 60153; and
Imperial College School of Medicine, Northwick Park Institute for Medical Research, Harrow, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In recent years several groups have described defective function of DC in tumor-bearing mice and in cancer patients (3, 4, 5, 6). It is now considered one of the important mechanisms of tumor escape from immune system control. The major findings in these studies were the lack of expression of costimulatory molecules in tumor-associated DC consistent with the phenotype of immature DC. A population of DC isolated from the peripheral blood of patients with breast and head and neck cancer demonstrated significantly reduced ability to cluster and stimulate allogeneic and Ag-specific T cell responses (7, 8). These cells have a substantially lower level of expression of MHC class II (HLA-DR) and costimulatory molecules than DC isolated from control donors. In agreement with these reports DC isolated from tumor-bearing mice also had a decreased expression of B7-2 and MHC class II as well as some adhesion molecules. These cells were unable to induce effective peptide-specific and antitumor cytotoxic immune responses and were ineffective as a tumor vaccine (9).
Thus, DC in tumor-bearing hosts are functionally defective. Previous studies have shown that several tumor-derived factors (vascular endothelial growth factor, IL-6, M-CSF, and gangliosides) affect DC maturation from hemopoietic progenitor cells (HPC) in vitro. However, mature DC were functionally competent (10). This was consistent with the fact that functionally competent DC can be generated in the absence of tumor-derived factors from bone marrow progenitor cells of tumor-bearing mice and from peripheral blood progenitors of cancer patients (8, 11). Thus, there is now enough evidence that tumor-derived factors affect DC differentiation. This may result in a substantial decrease in the number of mature DC in cancer patients. This hypothesis has been directly confirmed in two recent studies in which the number of DC was significantly decreased in cancer patients (12, 13). The decrease in the presence of DC in peripheral blood from cancer patients closely correlated with the stage and duration of the disease. We have previously demonstrated that this decrease was also closely associated with appearance of a large number of cells lacking markers of mature lymphoid and myeloid cells in peripheral blood (13). We termed those cells immature cells (ImC). We hypothesized that those ImC might have an impact on immune response in cancer. In this study we characterized the nature of ImC, demonstrated that these cells actively suppressed Ag-specific T cell responses, and identified factors able to differentiate these cells in vitro. This may suggest a new approach to improve immune response in cancer.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Forty-four patients, 32–79 years of age, with histologically confirmed cancer were enrolled in the study. Of these 44 patients, 13 had squamous cell carcinoma of the head and neck (HNSCC), 21 had non-small cell lung carcinoma, and 10 had breast cancer. The vast majority of patients were newly diagnosed, but a few had recurrent disease with no prior therapy for at least 1 year before the study. All patients had advanced diseases (stages III–IV) in accordance with the American Joint Committee on Cancer Criteria. Seven healthy volunteers served as controls.
Reagents
Complete culture medium (CCM) included RPMI 1640 (Life
Technologies, Gaithersburg, MD) supplemented with 10% FCS and
antibiotics. Ficoll-Paque was obtained from Pharmacia-Biotech (Uppsala,
Sweden). PE-, FITC-, or Quantum Red-conjugated anti-human MHC class
I (W6/32), anti-HLA-DR, CD3, CD13, CD14, CD19, CD57, and CD15 Abs
as well as isotype-matched mouse Ig were purchased from Sigma (St.
Louis, MO). FITC-conjugated anti-CD1a, anti-CD86, and
anti-CD40 Abs were obtained from PharMingen (San Diego, CA),
FITC-conjugated anti-CD34 Abs were purchased from Becton Dickinson
(Franklin Lakes, NJ) and Coulter (Hialeah, FL), and FITC-conjugated
anti-CD33 Ab was obtained from Serotec (Raleigh, NC). SRBC were
obtained from Cocalico (Reamstown, PA), and metrizamide was purchased
from Nyegaard (Oslo, Norway) and Sigma. Recombinant human GM-CSF, IL-4,
G-CSF, M-CSF, and CD40 ligand were purchased from RDI (Flanders, NJ).
Recombinant human TNF-
, TGF-
, and Flt-3 ligand were purchased
from R&D Systems (Minneapolis, MN). All-trans-retinoic acid
(ATRA) was obtained from Sigma. A stock solution of ATRA in absolute
ethanol (10-3 M) was stored at -30°C.
NG-monomethyl-L-arginine
(LMMA) was obtained from Calbiochem (San Diego, CA).
Peptide GILGFVFTL derived from the matrix of the influenza virus was synthesized by SynPep (Dublin, CA). This peptide has a high affinity with HLA-A2.
Cell isolation
DC and T cells were isolated from peripheral blood as previously described (14) with some modifications. Briefly, mononuclear cells obtained after centrifugation of peripheral blood over a Ficoll-Paque gradient were incubated with 2-aminoethylisothiouronium bromide (Sigma)-treated SRBC. Cells that adhered to the red cells (R+) and those that did not (R-) were separated on a Ficoll-Paque gradient. RBCs were then incubated for 18 h in CCM. Nonadherent cells were centrifuged over a metrizamide gradient (7.25 g of metrizamide in 50 ml of CCM) to obtained enriched fraction of DC. These cells were used in additional experiments.
In some experiments R- mononuclear cells were cultured for 2 h in CCM. After that time nonadherent cells were removed, and adherent cells were cultured in CCM supplemented with 30 ng/ml GM-CSF and 10 ng/ml IL-4. The same amount of cytokines in 0.5 ml of CCM was added on day 3. After 5–6 days of culture DC were collected, washed, and used in further studies.
R+ cells were further processed by osmotic lysis of red cells to obtain an enriched T cell fraction followed by overnight incubation in CCM at 37°C. More than 90% of nonadherent cells were T cells, as estimated by flow cytometry.
MLR and Ag-specific T cell proliferation
The ability of DC to stimulate allogenic T cells was tested in MLR. Fifty thousand T cells were plated in each well of 96-well round-bottom plates, and DC and T cells were cultured at ratios of 1:20, 1:40, 1:80, and 1:160 for 5 days. One microcurie of [3H]thymidine (sp. act., 25Ci/mmol) was added to each well 18 h before harvesting the cells. [3H]thymidine uptake was counted in a liquid scintillation counter (Beckman, Palo Alto, CA).
Ag-specific T cell response was measured using tetanus toxoid (TT). DC were cultured with autologous T cells in the presence of 1.0 µg/ml TT. [3H]thymidine was added after 4 days of culture, and uptake was counted 18 h later in a liquid scintillation counter. Background levels of T cell proliferation (without TT) were subtracted.
Generation of peptide-specific CTLs
Flu peptide-specific CTLs were generated from the peripheral blood of an HLA-A2-positive donor. DC (2 x 105 cells) isolated as described above were pulsed for 2 h with the peptide (10 µM), washed, and incubated in CCM with 2 x 106 T cells in 24-well plates in the presence of IL-7 (25 ng/ml). IL-2 (1.5 ng/ml) was added 2 days later. T cells were restimulated with peptide-pulsed DC on days 7, 14, and 21. IL-7 (25 ng/ml) was added immediately after restimulation, and IL-2 (1.5 ng/ml) was added 2 days later. CTLs were harvested on day 21 or day 28 and either used immediately or cryopreserved in liquid nitrogen until being tested in an enzyme-linked immunospot (ELISPOT) assay.
The peptide-specific CTLs were tested by cytolysis of the
peptide-loaded target T2 cells. Briefly, T2 cells (1.6 x
105/well) were loaded with 10 µg/ml of
2-microglobulin (Sigma) and 100 µM of FLU
peptide. After overnight incubation cells were washed, labeled with
51Cr, and used as targets in a standard 6-h
chromium release assay. As a control, T2 cells were incubated with
2-microglobulin alone.
IFN-
ELISPOT assay
The 96-well multiscreen filtration plates (Millipore, Bedford,
MA) were coated with 50 µl of mouse anti-human IFN- mAb
(MAB285, R&D Systems; final concentration, 12.5 µg/ml). After
overnight incubation at 4°C, wells were washed four times with PBS.
The remaining protein binding sites were blocked by incubating plates
for 2 h at 37°C with 200 µl/well RPMI 1640 supplemented with
10% human serum. T cells (105 cells/well) were
incubated in final volume of 200 µl with 104
sorted DC or ImC in the presence of 10 µg/ml specific peptide and 2
ng/ml IL-2. After 24-h incubation at 37°C wells, were washed six
times with PBS containing 0.05% Tween 20 (PBS-T). Wells were then
incubated overnight at 4°C with 100 µl (5 µg/ml) of biotinylated
goat anti-human IFN- Ab (BAF285; R&D Systems), washed six times
with PBS-T, and incubated for 2 h at room temperature with 50 µl
(1.25 µg/ml) of avidin-alkaline phosphatase (Sigma). Wells were
washed three times with PBS-T and three times with PBS and then
incubated with 50 µl of substrate
(5-bromo-4-chloro-3-indolyl-phosphate/nitro blue tetrazolium, Sigma)
for 10–15 min. The reactions were stopped by discarding the substrate
and washing the plates under tap water. The plates were then air-dried,
and colored spots were counted using a stereomicroscope.
Flow cytometry
Cells were labeled with PE-, FITC-, or Quantum Red-conjugated Abs by incubation on ice for 30 min followed by washing with PBS. Data acquisition and analysis were performed on a FACScalibur flow cytometer (Becton Dickinson, Mountain View, CA) using CellQuest software. For intracellular labeling cells were fixed for 60 min with 2% paraformaldehyde, permeabilized for 20 min with 0.2% Tween 20, and stained with FITC-conjugated anti-HLA DR Ab. Nonspecific binding was measured using FITC-conjugated isotype-matched mouse Ig. Cell sorting was performed on FACStar cell sorter (Becton Dickinson)
Assessment of colony formation
Colony formation by HPC was measured using semisolid 1% methylcellulose medium supplemented with recombinant cytokines (erythropoietin, stem cell factor, GM-CSF, G-CSF, IL-6, and IL-3) supporting the optimal growth of erythyrocyte bone-forming unit, GM-CFU, M-CFU, G-CFU, and GEMM-CFU colonies (Methocult H4436; Stem Cell Technologies, Vancouver, Canada). Sorted peripheral blood cells were seeded at 15,000 cells/plate. Colonies were scored on days 12–13.
Electron microscopy
The cells were fixed in 3% glutaraldehyde in 0.1 M sodium phosphate buffer (pH 7.4), washed in buffer, and embedded as a pellet in 1.5% low gelling temperature agarose (Sigma). The cells were given two additional washes with the sodium phosphate buffer, postfixed in 1% osmium tetroxide in 0.1 M phosphate buffer (pH 7.4) for 1 h, washed with water, block stained in 2% aranyl acetate for 2–4 h, washed again, and dehydrated using an acetone gradient. They were gradually infiltrated with Araldite resin over 8–24 h, embedded in the resin, and cured for 18 h at 65°C. The blocks were sectioned using a Reichert-Jung Ultracut E ultramicrotome (Vienna, Austria), and ultrathin (100 nm) sections on copper grids were stained with Reynold’s lead citrate, carbon coated, and viewed using a JEOL JEM-1200 Ex electron microscope (Peabody, MA). The cells present were identified, and at least 100 cells were counted for each sample. The cell diameters (37–45 cells/sample) were measured at their widest points from electron micrograph prints, and the mean and SEM were calculated.
Statistical analysis
Statistical analysis was performed using parametric and nonparametric methods and JMP statistical software (SAS Institute, Cary, NC).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Phenotype of ImC in peripheral blood of cancer patients
First, we tested the hypothesis that ImC were HPC. Enriched DC fractions from healthy volunteers and cancer patients were labeled with PE-conjugated lineage (Lin)-specific Abs (anti-CD3, -CD14, -CD19, and -CD57), Quantum Red-conjugated anti-HLA-DR Ab, and FITC-conjugated anti-CD34 Ab (two different clones obtained from Becton Dickinson and Coulter were used). CD34 molecules are expressed on HPC. The presence of CD34+ cells was analyzed in the Lin-HLA-DR- ImC population. In four experiments CD34+ cells represented from 0.9–1.8% of this cell population.
To investigate the presence of colony-forming HPC Lin-HLA-DR- ImC and Lin-HLA-DR+ DC from three cancer patients and three control donors were sorted on a cell sorter (FACStar, Becton Dickinson). Cells were then cultured in semisolid methylcellulose medium supplemented with growth factors supporting the growth of myeloid and erythroid colonies (stem cells). Lin-HLA-DR+ obtained from cancer patients and control donors gave rise to 195.4 ± 25.9 colonies/100,000 cells, whereas Lin-HLA-DR- gave rise to 1,750.5 ± 250.6 colonies/100,000 cells (p < 0.05). Most of these colonies were erythyrocyte bone-forming units and GM-CFU. No differences in the total number of colonies and the types of the colonies were found between control donors and cancer patients (data not shown). Thus, the Lin-HLA-DR- population of ImC was enriched for progenitor cells. However, the proportion of colony-forming cells among ImC was only about 1.7%, which was consistent with the number of CD34+ cells. Thus, these data demonstrate that HPC represent only a minor fraction of ImC.
To investigate the nature of ImC further, DC fractions isolated from
cancer patients were labeled with PE-conjugated lineage-specific Abs,
Quantum Red-conjugated anti-HLA-DR Ab, and FITC-conjugated Abs
specific for different cell lineages. At least four experiments have
been performed for each marker. More than 99% of
Lin-HLA-DR+ DC and >95%
of Lin-HLA DR- ImC
expressed myeloid cell marker CD33 (Fig. 1
A). Only 0.8 ± 0.4% of
Lin- HLA DR+ DC and
0.9 ± 0.3% of
Lin-HLA-DR- ImC expressed
granulocyte marker CD15 (p > 0.1; Fig. 1A). At the same time, 8.3 ± 1.0% of
Lin-HLA-DR+ DC and
32.4 ± 8.7% of
Lin-HLA-DR- ImC expressed
M-CSF receptor (CD115) specific for mature and ImC of the
monocyte/macrophage cell lineage (p < 0.05;
Fig. 1B). About two-thirds of all ImC expressed myeloid cell
marker CD13, and more than one-third of these cells expressed CD11c
marker specific for macrophages/DC (Fig. 1C). In three
independent experiments almost all ImC
(Lin-HLA-DR-) expressed
MHC class I molecule (Fig. 1C).
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B). Thus, together these data demonstrate that with the exception of small proportion of HPC, ImC were represented by MHC class I-positive myeloid cells. About one-third of the ImC were relatively immature monocytes/macrophages and DC. The remaining cells probably represented earlier stages of cell differentiation.
Ultrastructure of immature myeloid cells
Two populations of cells, ImC
(Lin-HLA-DR-) and DC
(Lin-HLA-DR+) cells, were
sorted on a FACStar cell sorter and analyzed by electron microscopy.
Two separate experiments were performed. These two populations of cells
did not differ with regard to the presence of apoptotic or necrotic
cells; the total numbers of dying cells in the two experiments were,
respectively, 11 and 4% for
Lin-HLA-DR- and 10 and
3% for Lin-HLA-DR+ cells.
Relatively small ImC represented a significant proportion of the
Lin-HLA-DR- cells (Fig. 2, A and B). In the
two experiments, the mean diameter at the widest points of all cells in
this Lin-HLA- DR-
population were 6.1 ± 0.2 and 6.4 ± 0.2 µM compared with
7.8 ± 0.2 and 7.8 ± 0.3 µM in the
Lin-HLA-DR+ population
(p < 0.05). In the two samples, 25 and 29% of
the total Lin-HLA-DR-
cells were identifiable as morphologically immature monocytes and DC
(Fig. 2, C–E). These cells are small, only 4–6 µm in
diameter. Many vacuoles are present. Although DC can also express
vacuoles, this is generally a monocyte/macrophage characteristic,
particularly if different sizes of vacuoles are present and abundant.
Another characteristic is the horseshoe-shaped nucleus that may be
developing in this cell (Fig. 2C). The volume of the
cytoplasm compared with that of the nucleus is small and somewhat
similar to that in lymphocytes. However, the cell surface is
irregularly shaped and is not like that in lymphocytes or lymphoblasts,
but is in the form of developing veils (Fig. 2D). As the DC
develop further the volume of the cytoplasm in relation to the nucleus
increases, and distinct veiled projections can be seen. The cytoplasm
has fewer inclusions than macrophages. Despite this veiled morphology,
the immaturity of these cells is evident from the size and the still
heterochromatic nucleus (Fig. 2E). Such ImC were rarely
found in the population of
Lin-HLA-DR+ that contained
cells of a more mature appearance. These cells are larger, with larger
veiled projections. The chromatin is becoming more euchromatic,
suggesting that maturation is occurring. The cytoplasm is paler than
that in macrophages with fewer organelles and occasional large apparent
vacuoles, probably produced from the invaginations of the veiled
surface morphology (Fig. 2F). Thus, these data are in
agreement with the results of surface marker expression and suggest
that ImC belong to the myeloid cell lineage.
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We asked whether these ImC could be differentiated into mature
cells in vitro. T cell-depleted mononuclear cells obtained from
patients with advanced disease were cultured overnight, and nonadherent
cells were collected and either analyzed immediately or cultured in the
presence of various cytokines. Half the medium was replenished, and
fresh cytokines were added on day 3 in all experiments. After 5–6 days
of culture cells were collected, washed, and labeled with a cocktail of
Abs as described above. Less than 10% of the cells survived 5–6 days
of incubation in medium alone. In preliminary experiments we tested two
cytokines important for the survival of different populations of
myeloid cells: GM-CSF and IL-3. All cells survived a 5- to 6-day
incubation with 20–30 ng/ml GM-CSF or 20–50 ng/ml IL-3 (data not
shown). However, neither of these cytokines significantly changed the
proportion of ImC. The culture of ImC for 5–6 days with IL-3 (1–20
ng/ml) or with a combination of GM-CSF and IL-3
also did not affect the proportion of ImC (Fig. 3). We used GM-CSF in all subsequent
experiments to maintain cell viability. Several combinations of factors
were used: GM-CSF and IL-4 (10 ng/ml), TNF- (1 ng/ml), TGF- (10
ng/ml), FLT-3 ligand (100 ng/ml), LPS (1 µg/ml), and CD40L (200
ng/ml). At least three experiments with each combination were
performed. Only the combination of GM-CSF and IL-4
significantly increased the proportion of DC (Fig. 3). The proportion
of DC was further increased (almost to control levels) when a
combination of three cytokines (GM-CSF, IL-4, and CD40L) or (GM-CSF,
IL-4, and TNF-) was used (data not shown). However, previous studies
have shown that the combination of GM-CSF and IL-4 induced generation
of DC from plastic adherent progenitors in peripheral blood (16, 17). These cells actively proliferate during the first couple of
days in culture. A substantial proportion of these
cells might have been present in the culture, and DC generated from
those progenitors might then significantly decrease the proportion of
ImC. To eliminate this possibility
Lin-HLA-DR- cells were
first sorted and then incubated with GM-CSF (control), GM-CSF and IL-4,
or GM-CSF, IL-4, and TNF- for 6 days as described above. In three
experiments the combination of these cytokines decreased the presence
of ImC (Lin-HLA-DR-,
B7-2-, or CD40-) by only
15–20% (data not shown). These data indicate that the considerable
decrease in the proportion of ImC observed in the presence of these
cytokines in previous experiments was due to accumulation of DC
generated from the progenitors but not from ImC.
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Functional significance of immature myeloid cells
To investigate the possible functional role of ImC,
Lin-HLA-DR- and
Lin-HLA-DR+ cells were
isolated from the peripheral blood of patients with advanced cancer
using cell sorting. Cells were then cultured with allogeneic T cells
isolated from healthy volunteers. DC isolated from healthy individuals
were used as controls. As reported earlier, the cells in the DC
fraction from patients with advanced cancer demonstrated a profound
defect in their ability to stimulate allogeneic T cells (8, 13). Here, in four experiments
Lin-HLA-DR- cells were
not able to stimulate allogeneic T cells (Fig. 4A). At the same time,
Lin-HLA DR+ DC
demonstrated control levels of T cell stimulation (Fig. 4A).
Thus, it appears that removal of ImC completely restored the ability of
the DC population to stimulate allogeneic T cells.
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B). At an ImC/DC ratio of 1:1 (usually seen in cancer
patients (13)) the T cell response was reduced >3-fold
(Fig. 4B). No effect of ImC on T cell proliferation in
absence of TT was found (data not shown). It was possible that in these
experiments ImC might compete with T cells for nutrients in the medium.
To exclude this possibility we used sorted autologous
CD14+ cells instead of ImC. The addition of as
many as 5000 CD14+ monocytes to the culture with
DC did not significantly affect TT-specific T cell proliferation (data
not shown).
For analysis of the MHC class I-restricted response, a CTL line was
generated from HLA-A2-positive healthy volunteer by stimulation of T
cells with influenza virus-derived peptide with high affinity to HLA-A2
molecule. The peptide specificity of the CTLs was confirmed in
cytotoxicity assay with peptide-loaded T2 cells (data not shown). ImC
and DC were isolated from peripheral blood of HLA-A2-positive cancer
patients and were incubated with the specific CTLs with or without
peptide. Production of IFN- by T cells was analyzed 24 h later
using an ELISPOT assay. DC from a cancer patient increased the number
of peptide-specific IFN- producing cells >3-fold. However, the
presence of ImC in the mixture at an ImC/DC ratio of 1:1 almost
completely abrogated that effect (Fig. 4C). These data
demonstrated that ImC were able to inhibit Ag-specific T cell
responses.
What could be a mechanism of the effects of ImC? These cells are
comprised of immature macrophages and myeloid cells known to produce
NO. NO is a well-described factor that inhibits T cell function
(18). To investigate the role of NO in the observed
effects of ImC we used a competitive inhibitor of NO synthase, LMMA
(19). LMMA did not significantly affect the inhibitory
effect of ImC on TT-dependent T cell proliferation or on FLU
peptide-specific IFN--producing cells (Fig. 5). To study the role of soluble factors
released by ImC, these cells and DC were cultured for 24 h in
U-bottom 96-well plates at a concentration of 5 x
105 cells/ml. Cell supernatants were collected
and immediately used in experiments with TT-dependent proliferation and
FLU peptide-specific response. In these experiments DC and T cells
isolated from HLA-A2-positive healthy volunteer were used. The presence
of supernatants at a final concentration as high as 20% (v/v) did not
significantly affect the T cell response (data not shown).
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ATRA is a natural oxidative metabolite of vitamin A and is known
to be a regulator of cell differentiation (20, 21). ATRA
induces terminal differentiation of promyelocytes into mature
neutrophils in patients with M3 (acute promyelocytic) leukemia
(22). Because the majority of ImC were represented by
immature myeloid cells at early stages of differentiation, we
hypothesized that these cells can be differentiated into mature cells
by ATRA. To test this hypothesis cells from DC fractions of cancer
patients were cultured with 30 ng/ml of GM-CSF and different
concentrations of ATRA (5 nM to 10 µM). After 5 days the total number
of cells and cell viability were measured. ATRA at concentrations of 5
and 10 µM was toxic for the cells. Cell viability returned to a
control level at 1 µM and remained at the same level at all other
tested concentrations (Fig. 6A). The total number of
recovered cells remained at the control (no ATRA) level at
concentrations from 10-6 to 5 x
10-8 M and increased at lower concentrations of
ATRA (Fig. 6B). In all subsequent experiments we used ATRA
at concentrations of 1 µM and lower. Five-day incubation of cells
with ATRA dramatically reduced the presence of ImC and increased the
presence of DC. In four experiments the proportion of
Lin-HLA-DR+ DC increased
from 30.4 ± 4.8% (in the presence of GM-CSF alone) to 70.2
± 5.6% (at an ATRA concentration of 1 µM) and was slightly lower at
an ATRA concentration of 0.5 µM. A further decrease in the ATRA
concentration cancelled the effect (Fig. 7A). Similar dramatic changes
were observed in the presence of
Lin-B7-2+ DC (Fig. 7A). The proportion of CD83+ (marker
specific for mature DC) cells in the DC fraction increased in the
presence of 1 µM ATRA by almost 4-fold. To confirm that these cells
were indeed DC we used an allogeneic MLR (a specific function of
relatively mature DC). In five independently performed experiments ATRA
dramatically increased the ability of these cells to stimulate control
allogeneic T cells (Fig. 7B). These data indicate that ATRA
was able to differentiate the majority of ImC to DC.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Effective function of the DD is an important element of antitumor immunity. The presence of functionally competent DC is critical for effective antitumor control and for the success of cancer immunotherapy. There is ample evidence of inadequate function of these cells in tumor-bearing hosts. At the same time, mature DC remaining in tissues demonstrate normal levels of functional activity and functionally potent DC can be generated from progenitors in patients even with advanced stages of cancer (3, 4, 5, 6, 8, 9, 10). Tumor-derived factors dramatically affect DC differentiation in vitro (23, 24, 25, 26, 27). Decreased DC production was associated with accumulation of monocytes/macrophages and immature myeloid cells. In addition, a significant decrease in the proportion and absolute numbers of DC in peripheral blood from cancer patients has been recently reported (12, 13). All these data support the idea that abnormal differentiation of DC forms a basis for defects of DC in cancer. We have previously demonstrated that the decreased presence of DC in cancer patients was closely associated with the accumulation in peripheral blood of cells lacking markers specific for mature cells of lymphoid and myeloid lineages (13). These cells were termed ImC. The presence of these cells dramatically increased in patients with early stages of cancer. In patients with advanced disease the number of ImC was significantly higher than the number of DC (13). The presence of ImC dropped considerably within 3–4 wk after surgical removal of the tumor (13). This drop is consistent with the hypothesis that the generation of these cells was due to the production of soluble tumor-derived factors. Thus, the appearance of ImC in peripheral blood of cancer patients was clinically relevant.
DC belong to the myeloid cell lineage, and DC and macrophages share a common progenitor. Therefore, it was not surprising that inhibition of DC differentiation in vitro results in the appearance of immature myeloid cells and monocytes (23, 24). Here we investigated the nature of the ImC generated in cancer patients. First, we tested the hypothesis that these cells could be CD34+ HPC. Increased production of these cells in patients with head and neck cancer has been previously reported (28). The population of ImC indeed contained an increased proportion of CD34+ HPC. However, the total percentage of these cells was <2%. The rest of the cells were MHC class I-positive myeloid cells. ImC did not express granulocyte markers. However, about 30% of these cells expressed M-CSF receptor (CD115), specific for cells of monocyte/macrophages lineage. The same proportion of cells expressed another marker of this cell lineage, CD11c. About 60% of ImC were CD13 positive. This molecule is expressed ona portion of immature myeloid cells, monocyte/macrophage, and DC. About 20% of the cells expressed intracellular HLA-DR, which might be considered characteristic of immature DC. Similar data were obtained by electron microscopy. This was confirmed by in vitro maturation experiments with growth factors. About one-third of the ImC became CD14+, plastic-adherent macrophages in the presence of M-CSF. About 20% of cells became Lin-HLA-DR+ DC in the presence of GM-CSF and IL-4. The presence of G-CSF or GM-CSF did not lead to differentiation of these cells into granulocytes. Taken together these data indicate that about one-third of ImC are represented by ImC of the macrophage/DC cell lineages. The remaining two-thirds of ImC are probably the cells at earlier stages of myeloid differentiation.
The exact mechanism of increased production of immature myeloid cells in cancer patients is not clear. However, it is known that tumor cells may produce several growth factors and cytokines able to stimulate myelopoiesis (GM-CSF, M-CSF, and IL-6). In addition, vascular endothelial growth factor produced by many tumors is able to affect myelopoiesis (29). It is possible that increased production of these growth factors may affect the normal pathway of cell differentiation resulting in the accumulation of immature myeloid cells.
We also asked whether these immature myeloid cells might affect immune
function. Elimination of ImC using cell sorting completely restored the
functional potency of the DC fraction. This is consistent with
previously reported observation that DC sorted based on phenotypic
characteristics were functionally competent in tumor-bearing mice
(10). These data indicate that the appearance of ImC may
be responsible for the decreased function of DC in cancer patients
observed in previous studies. To investigate the effect of ImC on the
Ag-specific T cell response we used two experimental systems. In both
these systems (MHC class II-associated TT-specific T cell proliferation
and MHC class I-restricted IFN- production) ImC actively inhibited
the T cell response in the presence of functionally competent DC. This
effect was seen at an ImC/DC ratio of 1:1. This or higher ratios were
observed in almost all patients with advanced stages of cancer and in
some patients with early stages of the disease. These data demonstrate
that ImC may actively suppress immune responses and thus contribute to
tumor nonresponsiveness. Several mechanisms may be responsible for the
observed effects of ImC. Our preliminary data ruled out a direct role
of apoptosis; ImC did not induce apoptosis of T cells (unpublished
observations). Supernatants from ImC also did not significantly affect
either T cell proliferation in response to TT or IFN- production in
response to FLU peptide. The experiments with the NO inhibitor LMMA
suggest that NO was not actively involved in the observed effects of
ImC. Thus, it is more likely that ImC exert their effects via direct
cell-cell contact. The fact that almost all ImC express MHC class I
molecule suggests that ImC might induce T cell anergy by engaging the
TCR complex on T cells in the absence of costimulatory signals. At this
time we are investigating this possibility.
Thus, it appears that ImC can be an important factor in immunosuppression in cancer. Hyperproduction of these cells might impede any attempt to induce a strong Ag-specific T cell response in cancer patients. It could be one of the factors that make cancer vaccines ineffective in patients with advanced stages of the disease. Therefore, it will be beneficial to find ways to eliminate these cells. One of the approaches would be to differentiate ImC into relatively mature cells. We have used several combinations of growth factors that promote differentiation of DC, macrophages, and granulocytes. They demonstrate only a minor effect on the population of ImC. The precise mechanism of this nonresponsiveness is not clear. It could be explained by the fact that the majority of ImC were represented by immature myeloid cells at early stages of differentiation. It is possible that these cells may have an altered differentiation program. This may be due to the lack of expression of the growth factors’ receptors or to affected molecular mechanisms responsible for cell differentiation. These mechanisms are under investigation at this time. Because of the nature of ImC, we have used ATRA in an attempt to differentiate these cells into granulocytes. A naturally occurring isomer of retinoic acid, ATRA is a well-known factor capable of induction of differentiation of the human leukemia cell line HL-60 and freshly isolated acute promyelocytic leukemia cells (30, 31). It is successfully used in differentiation induction therapy in patients with acute promyelocytic leukemia (32, 33). ATRA may also affect the growth of normal hemopoietic progenitors and blast progenitors in acute myelogeneous leukemia. However, these effects of ATRA depend on culture conditions (34, 35, 36). To our surprise, in the presence of GM-CSF and ATRA the majority of ImC were differentiated into relatively mature DC. The ability of GM-CSF to enhance the effect of ATRA on differentiation of human myeloblastic leukemia ML-1 cells into granulocytes has been previously described (37). No information is available about the ability of ATRA, in combination with any other growth factor, to induce differentiation of the DC. However, early studies in maturing splenic DC from mice showed that exposure to low doses of retinoid resulted in DC with an enhanced capacity to stimulate T cell proliferation; higher amounts resulted in a lower stimulatory capacity (37). It is likely that this effect depends on the nature of ImC. More studies are underway to identify the mechanism of the effects of ATRA on DC differentiation.
In conclusion, here we have demonstrated a significant accumulation of immature myeloid cells in peripheral blood of cancer patients. These cells actively suppressed the Ag-specific T cell response in cancer patients and thus could be involved in immunosuppression in cancer. ATRA was able to differentiate the majority of these cells into relatively mature DC. This observation may suggest a new approach to treatment of solid tumors and may be useful in the immunotherapy of cancer, especially in patients with advanced disease.
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Acknowledgments |
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Footnotes |
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2 Current address: H. Lee Moffitt Cancer Center, University of South Florida, Tampa, FL 33612.
3 Address correspondence and reprint requests to Dr. Dmitry Gabrilovich, H. Lee Moffitt Cancer Center, University of South Florida, MRC-2E, Room 2067, 12902 Magnolia Drive, Tampa, FL 33612.
4 Abbreviations used in this paper: DC, dendritic cell(s); ImC, immature cell(s); HNSCC, squamous cell carcinoma of the head and neck; CCM, complete culture medium; ATRA, all-trans-retinoic acid; LMMA, NG-monomethyl-L-arginine; TT, tetanus toxoid; ELISPOT, enzyme-linked immunospot; PBS-T, PBS containing 0.05% Tween 20; HPC, hemopoietic progenitor cells; Lin, lineage.
Received for publication July 27, 2000.
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