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Department of Medicine and Vanderbilt Cancer Center, Vanderbilt University School of Medicine, Nashville, TN 37232
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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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Here, we demonstrate a systemic defect in these LC functions in tumor-bearing mice. We show that those defects are closely associated with length of time that the animal was exposed to the tumor and most likely due to the effect of tumor-derived factors on the process of LC maturation. We also demonstrate that these defects can be partially reversed by inhibiting one of these tumor-derived factors by the administration of anti-VEGF Ab.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Female BALB/c and A/J mice, 6 to 8 wk old, were purchased from Harlan (Indianapolis, IN) and were housed in specific pathogen-free units of the Division of Animal Care at Vanderbilt University Medical Center.
Cell lines and Abs
Two tumor cell lines have been used. D459 tumor cells were constructed by transfection of BALB/c 3T3 cells with EJ ras and mutant human p53 open reading frame. Details of this cell line were described elsewhere (10, 11). MethA sarcoma cells was obtained from Dr. L. J. Old. This is a transplantable 3-methylcholanthrene-induced sarcoma of BALB/c origin passaged as an ascitic tumor (12). The following Ab-producing hybridomas were obtained from the American Type Culture Collection (Manassas, VA) and used as culture supernatants: anti-CD4 (L3T4, TIB-207), anti-CD8 (Lyt-2.2, TIB-210), anti-B cells (HB-146), anti-MHC class II (I-Ad,k, HB-120). FITC- and PE-labeled Abs used in flow cytometry were purchased from PharMingen (San Diego, CA): anti-CD11c (N418), anti-CD-86 (B7-2), anti-CD40, and anti-I-Ad FITC- and PE-conjugated IgG were used in controls. Biotinylated I-Ad and I-Ak Abs (PharMingen) were used in LC analysis. Neutralizing goat anti-mouse VEGF Ab was purchased from R&D Systems (Minneapolis, MN). Control goat Ig was obtained from Sigma (St. Louis, MO).
Cell preparation
Cells were prepared as described earlier (3). Briefly, a single-cell suspension was prepared from inguinal, axillary, and brachial lymph nodes by pressing the tissues through a wire mesh. Cells were then washed and layered onto a metrizamide (Nygaard, Oslo, Norway) gradient (14.5 g plus 100 ml of RPMI 1640 medium) and centrifuged for 10 min at 600 x g (metrizamide can be also obtained from Sigma). Cells at the interface were washed once and resuspended in complete culture medium (RPMI 1640 (Life Technologies, Gaithersburg, MD) with 100 IU/ml penicillin, 0.1 mg/ml streptomycin, 1 x 10-5 M 2-mercaptoethanol, and 10% FCS (HyClone, Logan, UT)). DC were identified by their distinctive morphology and by labeling with N418 Ab. In some experiments, FITC-positive cells from lymph nodes were sorted using a FACStar flow cell sorter (Becton Dickinson, Mountain View, CA). Pelleted cells from the lymph node suspension were passed through nylon wool columns to obtain >90% pure T cells.
Tumor induction and assay of LC migration
Two hundred thousand D459 cells or 6 x 105 MethA sarcoma cells were injected s.c. into the shaved back of mice. LC migration was measured at different times after tumor inoculation as described earlier (13). Briefly, mice were painted on the shaved back or abdomen with 50 µl of 1% FITC (isomer 1; Sigma) dissolved in a 50:50 (v/v) acetone-dibutylphthalate mixture. Twenty-four hours after painting with FITC or vehicle, mice were killed and single-cell suspensions were prepared from inguinal, axillary, and brachial lymph nodes. The DC fraction was enriched by separation on a metrizamide gradient. Cells were labeled with PE-conjugated anti-I-Ad or B7-2 Abs and analyzed with the gate set around the cluster of large cells using a FACSCalibur flow cytometer (Becton Dickinson, Mountain View, CA).
Epidermal sheet preparation and LC identification
Epidermal sheets were prepared from abdominal skin using the EDTA separation procedure described earlier (13, 14, 15). The ventral trunk skin was shaved, and the mice killed by cervical dislocation. The remaining hair was removed by chemical depilation with a thioglycolate-based commercial depilatory cream. The keratin layer was then removed by two or three applications of cellophane tape, and the skin was surgically excised while firmly attached to fresh cellophane tape. The skin was incubated for 2.5 h at 37°C in PBS containing 20 mM EDTA (pH 7.3) and 0.001% trypsin. The cellophane tape, with the epidermis attached, was separated from the dermis, washed, and incubated overnight at 4°C with biotinylated anti-I-Ad or anti-I-Ak Abs. After that time, these sheets were washed and incubated with streptavidin-peroxidase (PharMingen) for 2 h at room temperature. I-A+ cells were visualized in 0.7% mg/ml, 3,3'-diaminobenzidine containing 2 mg/ml H2O2 (Sigma FAST DAB tablet set; Sigma) for 5 min at room temperature. The sheets were then washed, lightly dried, and mounted on slides. I-A+ cells were counted in 1 mm2 by counting 10 separate, randomly selected fields.
Assessment of LC development using radiation bone marrow chimeras
Control and tumor-bearing BALB/c mice (Dd, Kd, Ld, I-Ad, I-Ed) were irradiated with two fractions of radiation to a total dose of 1080 cGy (two doses of 540 cGy with a 3-h interval). Mice were then immediately injected with 3 to 4 x 105 bone marrow cells obtained from A/J mice (Dd, Kk, Ld, I-Ak, I-Ek) after it was depleted for T-, B-, and MHC class II-positive cells using mAbs and complement (Low-Tox-M Guinea Pig Complement; Cedarlane, Hornby, Ontario, Canada). From 3 to 4 wk later, LCs in skin were analyzed as described above using a biotinylated I-Ak Ab. For control of experimental conditions bone marrow also was injected into nonirradiated BALB/c mice.
T cell proliferation assay
DC were obtained from lymph nodes of FITC-painted mice as described above. Cells were irradiated (2000 cGy) and added in triplicate to 5 x 104 T cells obtained from healthy BALB/c mice at different DC:T cell ratios. After a 3-day incubation in 96-well U-bottom plates, the cultures were pulsed with 1 µCi of [3H]thymidine (Amersham, Arlington Heights, IL). [3H]Thymidine uptake was counted using a liquid scintillation counter.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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To study the effect of tumor on LC migration and function, BALB/c
mice were inoculated with 2 x 105 D459 cells. Six
weeks later, when tumors reached 15 to 17 mm in diameter (220–280
mm2), mice were painted with FITC. In an initial series of
experiments, we compared the effect of painting the skin around the
tumor in the dorsal trunk ("local site") and painting the skin at a
distant site (the ventral trunk). The number of DC in lymph nodes was
counted using morphologic criteria and then confirmed by analysis of
cell surface molecule expression. Since there is no single marker that
can identify all lymph node DC, we used several different markers. The
percentage of cells positive for each marker was counted. We found no
statistically significant differences between the percentage of
CD11c+ and B7-2 MDS-2+ cells in all samples.
These two markers were used in most of the experiments. The changes in
FITC-painted tumor-bearing mice were also confirmed by staining with
MHC class II Abs. Figure 1
A
demonstrates the results of a typical experiment looking at the
fraction of CD11c-positive cells in the lymph nodes of FITC-painted
mice.
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B).
The number of DC in lymph nodes obtained from control animals was also
increased more than twofold. However, almost no increase was detected
in tumor-bearing mice. No differences were found between mice painted
with FITC near the tumor or at a distant site. The percentage of
FITC-positive DC in lymph nodes was analyzed using flow cytometry.
Lymph node DC were labeled with PE conjugated anti-B7-2 or
anti-I-Ad. Double-positive (FITC and PE) cells
represent LC that had taken up FITC and migrated to the lymph nodes.
Tumor-bearing mice had a 2.5-fold reduced percentage of FITC-positive
DC compared with control animals (p < 0.01,
Fig. 2). This was observed in mice
painted both near the tumor and at a distant site (data not shown). We
then asked whether the observed decrease in the number of FITC-positive
cells was also associated with a decreased ability of DC to stimulate
primary FITC-specific T cell responses. DC were isolated from the lymph
nodes of painted tumor-bearing or control mice and irradiated, and
their number was adjusted to equal levels using morphologic criteria
confirmed by staining with anti-CD11c Ab. These DC were then
incubated with T cells from control untreated BALB/c mice. DC from
control FITC-painted mice stimulated significant T cell proliferation,
whereas very low responses were detected when DC were isolated from
tumor-bearing mice (Fig. 3).
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Effect of tumor on function and number of LC in skin
Next, we attempted to identify the possible mechanism of the
observed defective LC migration to lymph nodes. We have previously
shown in an animal model that defective function of DC in cancer was
associated with a reduced expression of MHC class II and the
costimulatory molecule B7 on the surface of DC (3, 4). B7 expression is
also low in tumor-associated DC in patients with cancer (7). Here, we
investigated the expression of MHC class II and B7-2 molecules on DC
from FITC-painted mice. Control and tumor-bearing mice were painted
with FITC (distant site), and 24 h later DC from lymph nodes were
labeled with PE-conjugated anti-I-Ad or anti-B7-2.
FITC-negative and FITC-positive cells were analyzed separately. In four
independent experiments, expression of B7-2 and I-Ad in
FITC-negative DC was significantly lower in tumor-bearing mice than in
controls (42.1 ± 4.2% decrease, p = 0.008). Two
typical experiments are shown in Figure 4. The level of B7-2 expression in
FITC-positive DC was also decreased in tumor-bearing mice. However,
that decrease was less pronounced (28.6 ± 3.4%;
p = 0.02). The same changes were detected in the level
of MHC class II (data not shown).
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). Treatment with FITC decreased
the number of LC by 
30% both in control and in tumor-bearing mice
(data not shown). These data suggest that a major reason for defective
LC migration to nodes in tumor-bearing mice may be a decreased LC
density in the skin of tumor-bearing hosts.
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, despite a more than threefold
decrease in the number of FITC-positive LC in tumor-bearing mice,
sorted FITC-positive cells were fully competent in stimulating primary
T cell responses. This suggests that if a LC can take up an Ag and
transport it to the lymph node, then it is capable of stimulating a
primary response. Tumor-bearing animals thus have a decreased number of
LCs capable of taking up FITC and transporting it to the lymph nodes,
and this results in the observed decreased stimulatory capacity of
nodal DC.
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One of the possible explanations for the decreased numbers of LC in the skin of tumor-bearing mice is that LCs migrate toward the tumor because of the effect of tumor-derived chemotactic factors. Because these stimuli are continuously present in tumor-bearing mice, this may result in fewer LC in skin. To test this possibility, we inoculated greater numbers of tumor cells (10 times higher than in previous experiments) to shorten the length of time required for the tumor to reach a given size. At this dose, tumors reached 220 to 280 mm2 within 2 to 2.5 wk, twice as fast as in the previous experiments. If the described effects were solely the result of migration of LC out of skin toward the tumor, we would expect to see the same level of decrease in the number of FITC-positive DC in lymph nodes as in the previous set of experiments. However, when we evaluated the number of FITC-positive DC in draining lymph nodes from these animals, only a minimal decrease in the percentage of FITC-positive DC was detected in lymph nodes of tumor-bearing mice (data not shown). We then studied how further increased numbers of inoculated tumor cells affected LC number in skin during shorter exposures to tumor (12 days). Even a 50-fold increase in the number of injected tumor cells did not change the number of LC in the skin in both experimental models (data not shown). It is important that in all these experiments, the tumor was injected into the dorsal trunk and the skin was obtained from ventral trunk. These experiments demonstrated that decreased numbers of LCs in the skin and the decreased responses of these cells to additional stimulation with Ag require relatively long exposure to tumor. This suggests that the tumor may affect the process of LC production and maturation from precursors rather than directly affecting mature LC. Since the half-life of LC in skin is relatively long, the observed effect may require the prolonged presence of tumor to be observed.
LC maturation in tumor-bearing bone marrow chimeric mice
To distinguish newly produced LC from preexisting ones, bone
marrow chimera experiments were performed. Tumor-bearing BALB/c mice
(haplotype d/d; tumor size, 50–70 mm2) were lethally
irradiated, and bone marrow from A/J mice (haplotype d/k) was used for
reconstitution. The bone marrow from the donor was depleted for
lymphocytes, macrophages, and DC. After 3 weeks, mice were still free
from visible signs of graft-vs-host disease. When tumors reached
120–150 mm2, mice were killed, and the number of
donor-derived LC (I-Ak+ cells) were counted in the skin as
described in Materials and Methods. The same experiments
were performed in parallel in tumor-free mice. As a control, bone
marrow cells were also injected into nonirradiated control mice. As
shown in Figure 7, large numbers of
I-Ak+ cells were detected in the skin of control irradiated
mice (Fig. 7B). No I-Ak-positive cells were
detected in the skin of nonirradiated mice (data not shown). The number
of LC in the skin of tumor-bearing mice was significantly lower than
those observed in control animals (Fig. 7A).
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The data shown above indicate that tumor cells may affect the
process of maturation from bone marrow progenitors to mature LC in
skin. Several factors could potentially contribute to this process.
Recently, we have shown that VEGF, which is produced by most tumors can
directly affect DC maturation in vitro (16). We thus hypothesized that
blockade of VEGF with a neutralizing Ab may improve the function of LC
in tumor-bearing hosts. D459 cells produce VEGF in vitro (our
unpublished observations) and in mice bearing 220- to
280-mm2 tumor, the serum level of VEGF was 120 to 180 pg/ml
as was determined by ELISA (R&D Systems). Since direct antitumor
effects of VEGF blockade are possible and tumor regression by any means
may improve LC function, we tested whether anti-VEGF Ab was able to
affect the growth of established, poorly immunogenic D459 tumors. Mice
were injected with D459 cells, and when tumors reached 5 to 6 mm in
diameter, treatment with anti-VEGF Ab was initiated. We tested
three doses of Ab (2, 5, and 10 µg/mouse i.p. twice a week during 4
wk). These doses were chosen on the basis of the in vitro neutralizing
activity of the Ab. As controls, mice were treated with the same
concentration of goat Ig. In six independently performed experiments,
anti-VEGF Ab did not affect tumor growth (4 mice/group in each
experiment, data not shown). No differences were found between mice
treated with goat Ig and untreated animals. Then, we asked whether
anti-VEGF Abs had any effect on the ability of LC to migrate to
draining lymph nodes. Tumor-bearing mice were treated with 10
µg/mouse of anti-VEGF Ab for 4 wk. After that time, they were
painted with FITC as described above. Figure 8A shows the result of one of
these experiments demonstrating that treatment with anti-VEGF Ab
increased the percentage of FITC-positive DC in LN. This was associated
with an improved ability of these cells to stimulate primary
FITC-specific T cell responses (Fig. 8B). Treatment with
anti-VEGF Ab also reversed the tumor-associated decrease in the
number of LC in the skin (Fig. 8C).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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However, it was not clear whether the described defects were localized
or systemic. Also not clear was whether LC in tumor-bearing animals had
a decreased ability to take up Ag and migrate to the site of lymphoid
stimulation (mostly lymph nodes), the major function of LC. These
questions are important not only for our understanding of the
immunopathogenesis of cancer but also for devising better strategies
for the immunotherapy of cancer and cannot be addressed with in vitro
models. Therefore, we adopted an in vivo model of LC migration in
response to the topical application of FITC. As has been shown before,
after application of FITC, LCs migrate to draining lymph nodes and
stimulate primary FITC-specific T cell responses (8, 9). Transfer of
these cells into naive mice led to development of FITC hypersensitivity
(22). We use this model to investigate the possible effect of tumors on
LC function. The two tumor models used in our study have been described
elsewhere (10, 23, 24). In most experiments, we used the poorly
immunogenic D459 tumor. This tumor reaches 15 mm in diameter (220–280
mm2) 6 wk after the s.c. injection of 2 x
105 cells. After that time, tumors quickly exceed the 2 cm
in diameter often associated with ulceration of the skin above the
tumor. Therefore, we chose 6 wk as an end point in this study. Painting
of the skin with FITC resulted in significant increases in the number
of lymphocytes and DC in the lymph nodes of control animals, a usual
finding after the application of contact sensitizers. However, the
total number of DC and the number of FITC-positive DC in lymph nodes
was significantly lower in tumor-bearing mice. To our surprise, the
same results were obtained when mice were painted at the tumor site
(dorsal trunk) or at a site distant from the tumor (ventral trunk).
These data indicate that there is a systemic defect in LC function in
mice bearing tumors. The mechanism of this effect is most likely via
soluble tumor-derived factors. We first tested the hypothesis that
these factors affected the ability of LC to migrate to lymph nodes and
stimulate T cells via the down-regulation of LC surface receptors known
to play a major role in DC function. We asked whether tumor cells
induce down-regulation of those molecules in FITC-positive DC in
draining lymph nodes and whether the total number of DC in the nodes of
tumor-bearing mice was decreased. Our data demonstrated only a slight
decrease in the number of FITC-negative DC in lymph nodes from
tumor-bearing mice compared with control animals (15%), whereas the
number of FITC-positive DC was decreased almost threefold. This implies
defective Ag uptake and transport in these animals. On the surface of
DC from tumor-bearing mice, we found a reduced expression of MHC class
II and B7-2 molecules compared with DC from control animals. This
decrease was seen on the surface of FITC-negative DC and, to a lesser
extent, on FITC-positive cells. However, in tumor-bearing mice sorted
FITC-positive LC that migrated from the skin to the lymph nodes had the
same ability to stimulate naive T cells as these cells from control
mice. This indicates that defects we observed in the ability of
FITC-positive cells from tumor-bearing mice to stimulate primary T cell
responses were not due to decreased expression of the surface molecules
on DC, but rather due to reduced numbers of LC taking up FITC and
migrating from the skin. We then examined the number of LC in skin.
Significantly lower numbers of LC were found in tumor-bearing mice than
in control animals. The same decrease was observed in mice bearing
another syngeneic tumor (MethA sarcoma). These data indicate that the
substantial decrease in the number of FITC-positive cells in lymph
nodes was not due to defects in the function of LC capable of conveying
Ag to the nodes, but rather a consequence of a decreased pool of LC in
the skin.
There are two possible explanations for a generalized decrease in the number of LC in the skin of tumor-bearing animals. First, tumor-derived factors may work as chemokines to attract LC to the site of the tumor, depleting LC from the skin in spite of normal rates of LC production. Increased numbers of LCs at the site of some (but not all) types of human tumors have been described and are correlated with a good prognosis (reviewed in 25 , consistent with this hypothesis. The other possible explanation is that tumor-derived factors may also prevent replenishment of LC in the skin by interfering with their production and maturation from precursors. The half-life of LC has been reported to be around several weeks (26, 27), so it is possible that a 6-wk exposure to a tumor is sufficient to cause decreased numbers in the skin due to decreased generation of LC from progenitors. The histologic analyses of tumors so far reported in the literature cannot distinguish between these hypotheses.
To address these questions, we established tumors using increased numbers of tumor cells, thus shortening the time the tumor takes to achieve a given size. We determined the dose that resulted in the same tumor size used before (220–280 mm2) but achieved this size in half the time (2–2.5 wk). If the migration of LC toward the tumor was the only reason for the decreased LC number in the skin and their decreased migration to lymph nodes after FITC painting, we should see roughly the same effect after this short exposure to the tumor. However, these effects were much smaller in mice exposed to tumor for 2 to 2.5 wk than in mice exposed to the same size tumor for 6 wk. A 12-day exposure to a 50-fold increased number of tumor cells did not affect the number of LC in the skin distant from the tumor at all. These data strongly suggest that defective replenishment of LCs in the skin after naturally occurring LC turnover or migration induced by tumor-derived factors is the major cause for defective LC function in tumor-bearing animals. These findings were also supported by our experiments with radiation chimeras, as donor bone marrow cells transplanted into tumor-bearing mice showed considerably reduced generation of donor LC in the skin of tumor-bearing mice.
The observed systemic effects of tumors can best be explained by soluble factors, perhaps directly produced by tumors. Tumors are known to produce factors that may affect maturation of LC from progenitors. For example, numerous studies of the inhibition of LC function by IL-10 have been described (28, 29, 30, 31, 32, 33, 34, 35). Elevated levels of IL-10 in the sera of cancer patients have been reported by several groups (36, 37). Expression of IL-10 mRNA was detected in renal carcinoma cells (38). It is possible that IL-10 or other cytokines or tumor-derived factors may affect the maturation of LC in vivo. However, it is important that IL-10 affects the latest stages of in vitro maturation of LC from a relatively immature state (freshly isolated LC) to a fully mature state. Since even these relatively immature LCs still express MHC class II, they would be detected immunohistochemically by the methods used in this study. Therefore, decreases in the number of LC observed in our study are more likely to be due to effects of tumor-derived factors on earlier stages of LC maturation.
Previously, we have reported on the possible involvement of VEGF in
defective DC maturation in vitro (16). This effect could be mediated by
the blockade of TNF-
-inducible activation of the transcription
factor NF-
B in hemopoietic progenitor cells (39). VEGF is produced
by almost all tumors and was found in the sera from the tumor-bearing
mice in this study. We thus asked whether blockade of VEGF would
improve LC function in this system. This assessment is complicated by
the fact that VEGF directly promotes tumor neovascularization (reviewed
in 40 , and anti-human VEGF inhibits the growth of human
tumors in nude mice (41, 42, 43, 44). The effect of anti-VEGF Ab is most
pronounced in the inhibition of the growth of small tumors and
micrometastases, so we tested whether anti-mouse VEGF Ab affected
the growth of the established tumors used in our study. In six
independent experiments (four mice per group per experiment) we did not
find any effect of anti-VEGF Abs at doses as high as 10 µg per
mouse treated twice a week for 4 wk. We did not escalate this dose
further. It is possible that antitumor effects could be seen at higher
doses of the Ab than were tested here. It is important to emphasize
that we used anti-mouse VEGF Ab, not the anti-human one used in
previous studies. Despite the lack of antitumor effects at these doses,
we detected a marked improvement in LC function after 4 wk of treatment
with anti-VEGF Ab in tumor-bearing mice. The number of LC in the
skin was also increased compared with tumor-bearing mice treated with
goat Ig. Thus, anti-VEGF Ab was able to improve LC function in
tumor-bearing mice, consistent with our previously reported data that
VEGF might be one of the factors responsible for defective Ag
presentation function in cancer. Despite the effects of this Ab on LC
function, we did not observe significant spontaneous induction of
antitumor immunity. D459 cells are very poorly immunogenic and have not
been observed to induce spontaneous immune responses or spontaneous
rejection under any experimental conditions. Apparently, low
expression of tumor-specific Ags does not induce an immune response
even when DC function is improved. However, when these mice are
immunized with tumor-specific Ag, anti-VEGF Ab markedly increases
the effectiveness of immunotherapy (manuscript in preparation).
In conclusion, we describe for the first time that tumors, via soluble tumor-derived factors (including VEGF), considerably affect the function of LC even at sites distant from the tumor. This effect requires relatively prolonged exposure to the tumor and probably works through inhibition of LC differentiation. This phenomenon may represent one of the possible mechanisms of tumor escape from immune system control. Anti-VEGF Ab may provide a new tool for the improvement of DC function in cancer and therefore could be potentially used in immunotherapy of cancer.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Dmitry Gabrilovich, Vanderbilt Cancer Center, Vanderbilt University School of Medicine, 649 MRB II, Nashville, TN, 37232-6838. E-mail address:
3 Abbreviations used in this paper: DC, dendritic cells; LC, Langerhans cells; VEGF, vascular endothelial growth factor.
Received for publication February 20, 1998. Accepted for publication June 23, 1998.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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 K. Mashino, N. Sadanaga, F. Tanaka, M. Ohta, H. Yamaguchi, and M. Mori Effective Strategy of Dendritic Cell-based Immunotherapy for Advanced Tumor-bearing Hosts: the Critical Role of Th1-dominant Immunity Mol. Cancer Ther., August 1, 2002; 1(10): 785 - 794. [Abstract] [Full Text] [PDF]
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M. L. Disis, K. Rinn, K. L. Knutson, D. Davis, D. Caron, C. dela Rosa, and K. Schiffman Flt3 ligand as a vaccine adjuvant in association with HER-2/neu peptide-based vaccines in patients with HER-2/neu-overexpressing cancers Blood, April 15, 2002; 99(8): 2845 - 2850. [Abstract] [Full Text] [PDF]
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G. M. Halliday and S. Le Transforming growth factor-{beta} produced by progressor tumors inhibits, while IL-10 produced by regressor tumors enhances, Langerhans cell migration from skin Int. Immunol., September 1, 2001; 13(9): 1147 - 1154. [Abstract] [Full Text] [PDF]
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D. J. Nelson, S. Mukherjee, C. Bundell, S. Fisher, D. van Hagen, and B. Robinson Tumor Progression Despite Efficient Tumor Antigen Cross-Presentation and Effective "Arming" of Tumor Antigen-Specific CTL J. Immunol., May 1, 2001; 166(9): 5557 - 5566. [Abstract] [Full Text] [PDF]
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B. Almand, J. I. Clark, E. Nikitina, J. van Beynen, N. R. English, S. C. Knight, D. P. Carbone, and D. I. Gabrilovich Increased Production of Immature Myeloid Cells in Cancer Patients: A Mechanism of Immunosuppression in Cancer J. Immunol., January 1, 2001; 166(1): 678 - 689. [Abstract] [Full Text] [PDF]
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B. Almand, J. R. Resser, B. Lindman, S. Nadaf, J. I. Clark, E. D. Kwon, D. P. Carbone, and D. I. Gabrilovich Clinical Significance of Defective Dendritic Cell Differentiation in Cancer Clin. Cancer Res., May 1, 2000; 6(5): 1755 - 1766. [Abstract] [Full Text] [PDF]
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D. I. Gabrilovich, T. Ishida, S. Nadaf, J. E. Ohm, and D. P. Carbone Antibodies to Vascular Endothelial Growth Factor Enhance the Efficacy of Cancer Immunotherapy by Improving Endogenous Dendritic Cell Function Clin. Cancer Res., October 1, 1999; 5(10): 2963 - 2970. [Abstract] [Full Text] [PDF]
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S. Corinti, D. Medaglini, A. Cavani, M. Rescigno, G. Pozzi, P. Ricciardi-Castagnoli, and G. Girolomoni Human Dendritic Cells Very Efficiently Present a Heterologous Antigen Expressed on the Surface of Recombinant Gram-Positive Bacteria to CD4+ T Lymphocytes J. Immunol., September 15, 1999; 163(6): 3029 - 3036. [Abstract] [Full Text] [PDF]
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J. E. Ohm, M. R. Shurin, C. Esche, M. T. Lotze, D. P. Carbone, and D. I. Gabrilovich Effect of Vascular Endothelial Growth Factor and FLT3 Ligand on Dendritic Cell Generation In Vivo J. Immunol., September 15, 1999; 163(6): 3260 - 3268. [Abstract] [Full Text] [PDF]
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 S. Rosic-Kablar, K. Chan, M. D. Reis, I. D. Dube, and M. R. Hough Induction of tolerance to immunogenic tumor antigens associated with lymphomagenesis in HOX11 transgenic mice PNAS, November 21, 2000; 97(24): 13300 - 13305. [Abstract] [Full Text] [PDF]
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