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Immunology |
Divisions of Pulmonary and Critical Care Medicine [M. D. R., S. M. K., A. N. P., M. M.] and Hematology/Oncology [B. J. G., N. M., R. A. F.] and the Jonsson Comprehensive Cancer Center [M. D. R., R. A. F.], University of California at Los Angeles School of Medicine, Los Angeles, California 90095
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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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Whereas little is known about the in vivo origins and
trafficking of human DCs, in vitro studies have identified
two cell populations that can act as DC precursors: (a)
CD34+ stem cells originating from bone marrow
(16
, 17)
; and (b) circulating
CD14+ monocytes (18, 19, 20)
.
CD34+ stem cells mature into DCs in response to
GM-CSF and tumor necrosis factor 
, an effect that can be enhanced by
a variety of other factors including IL-4, flt3 ligand, and CD40 ligand
(21
, 22)
. CD14+ monocytes mature
into DCs primarily in response to the combination of GM-CSF and IL-4
(19
, 23) , although similar effects have been reported when
GM-CSF is combined with IFN-, CD40 ligand or IL-13
(24, 25, 26)
. Based on evidence that viable DC progenitors
exist in cancer patients (27)
and can respond to cytokine
stimulation, we set out to determine whether systemic therapy with the
combination of GM-CSF and IL-4 would work in vivo as it does
in vitro to increase the number and/or function of
circulating APCs. A total of 21 patients with metastatic solid tumors
were enrolled in a Phase I, dose-escalating, multicohort study to
evaluate the effects of daily therapy with GM-CSF alone
versus the combination of GM-CSF and IL-4 on the number,
phenotype, and function of circulating APCs.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Clinical Protocol.
Patients were recruited into a total of six successive treatment
cohorts (cohorts AI, AII, B, C, D, and E) in a serial manner. Patients
were treated with daily s.c. injections of sterile human recombinant
GM-CSF (specific activity = 1.125 x 107 units/mg) and IL-4 (specific
activity = 2.414 x 107 units/mg) provided by Schering-Plough
Research Institute (Kenilworth, NJ). For cohorts AI–D, the 28-day
protocol included 14 continuous days of therapy followed by 14 days of
observation. For cohort E, the 28-day protocol alternated 7-day periods
of cytokine therapy with 7-day periods of observation. Injections were
self-administered s.c. to the skin of the thigh and/or abdomen as a
home-based therapy. Patients in cohort AI received a fixed dose of
GM-CSF (2.5 µg/kg/day), whereas patients in subsequent cohorts
received the same dose of GM-CSF in combination with IL-4 at 0.5
(cohort AII), 1.0 (cohort B), 2.0 (cohort C), 4.0 (cohort D), or 6.0
(cohort E) µg/kg/day. All patients successfully completing the AI
cohort were enrolled as subjects in the AII cohort, but new subjects
were enrolled into all subsequent groups (cohorts B–E). Patients
achieving objective clinical responses or stable disease were eligible
to continue therapy without further determination of biological
activity. Patients were monitored twice weekly for evidence of toxicity
according to the National Cancer Institute Common Toxicity Criteria.
Blood was drawn on days 0, 7, 14, and 21 for in vitro
testing to assess the biological effects of therapy. To assess
antitumor efficacy, radiological imaging of measurable and evaluable
disease and tumor marker serologies (when applicable) were obtained
serially for as long as patients remained on therapy. Standard
Southwest Oncology Group response criteria were used to determine
objective responses.
Laboratory Reagents and Antibodies.
CM was composed of RPMI 1640 supplemented with glutamine (Irvine
Scientific, Irvine, CA), 0.01 M HEPES buffer,
antibiotic-antimycotic mixture (Life Technologies, Inc., Grand Island,
NY), and 10% heat-inactivated human AB serum (Gemini Bioproducts,
Inc., Calabasas, CA). Dulbecco’s PBS (Life Technologies, Inc.) was
supplemented with 2% human AB serum. M450 magnetic beads coated with
goat antimouse IgG were obtained from Dynal Inc. (Lake Success, NY).
Fluorescein-labeled dextran (Mr
40,000) was purchased from Molecular Probes (Eugene, OR).
Fluorochrome-conjugated mAbs used for FACS analysis included anti-CD3,
anti-CD13, anti-CD14, anti-CD15, anti-CD20, anti-CD40, and anti-CD57
from Caltag Laboratories (South San Francisco, CA); anti-HLA-DR and
anti-CD80 (B7-1) from Becton Dickinson (San Jose, CA); anti-CD11c and
anti-CD86 (B7-2) from PharMingen (San Diego, CA); anti-CD1a from
Serotec (Raleigh, NC); and anti-CD83 (clone HB15a) from Beckman Coulter
(Fullerton, CA).
FACS Analysis.
Peripheral blood samples from study days 0, 7, 14, and 21 were
collected in heparinized tubes, and leukocytes were isolated by
differential centrifugation over Ficoll-Paque gradients
(Pharmacia Biotech, Inc., Alameda, CA). Leukocytes were analyzed for
their expression of cell surface markers by three-color FACS analysis
using a single FACScan II flow cytometer that was calibrated daily with
CaliBRITE beads and AutoCOMP software (all from Becton Dickinson).
Between 5,000 and 300,000 events (depending on the cell population and
marker) were acquired for each sample using CellQuest software (Becton
Dickinson) that simultaneously acquired data for forward scatter, side
scatter, FL1 (FITC label), FL2 (PE label), and FL3 (either
tricolor or PerCP label). The settings for all of these
parameters were optimized at the initiation of the study and maintained
constant during all subsequent analyses to allow direct comparison
between samples analyzed on different days. For cohort E, a new lot of
anti-CD14 mAb resulted in lower expression as compared to prior
cohorts. The relative expression of a given marker was expressed by the
LFI. The number of circulating CD14+ or
CD83+ cells per milliliter of blood was
determined by multiplying the percentage of leukocytes staining with a
given marker (as determined by FACS analysis) by the number of
leukocytes recovered per milliliter of blood.
Endocytosis Assay.
The temperature-dependent uptake of FITC-labeled dextran was used to
measure endocytotic function according to a modification of the
procedure described by Sallusto et al. (28)
.
Fresh leukocytes from study patients in the D cohort were resuspended
in 0.5 ml of CM (containing 25 mM HEPES) and
cultured with FITC-labeled dextran (1 mg/ml) for 60 min at either 0°C
or 37°C. Cells expressing CD14 were stained by adding anti-CD14-PE
during the last 5 min of the assay, and the reaction was terminated by
adding 3 ml of ice-cold PBS containing 0.1% azide. Cell pellets were
washed four times, and the CD14+ population was
analyzed immediately for the intracellular accumulation of the FITC
label by FACS analysis. The degree of endocytosis was determined by
comparing the intracellular uptake at 37°C with the nonspecific
binding that occurred at 0°C.
Allogeneic MLR.
APCs collected on different days of therapy were evaluated for their
ability to stimulate the proliferation of allogeneic T cells using a
one-way MLR. This approach, as compared to a soluble antigen
presentation assay, was specifically chosen to prevent any treatment-
or cancer-related effects on the patient’s own T cells from
interfering with the measurement of APC function (29
, 30)
.
APCs were prepared either by depleting contaminating neutrophils from
PBMCs with endotoxin-free anti-CD24 mAb (2
µg/106 cells; PharMingen) and immunomagnetic
beads (bead:cell ratio, 4:1) or by preparing purified
CD14+ or
CD14-/CD83+ cells by
fluorescence-activated cell sorting analysis using a
FACStarplus flow cytometer and Lysys II software
(Becton Dickinson) as described previously (19)
. APCs were
irradiated with 30 Gy (cesium source) and cocultured with 1 x 105 responder T cells for 6 days at APC:T
cell ratios of 1:5 to 1:100. Wells were pulsed with 1 µCi of
[3H]thymidine (DuPont New England Nuclear,
Boston, MA) and harvested 18 h later using an automated cell
harvester (PhD cell harvester; Cambridge Technology, Cambridge, MA).
cpm were determined by liquid scintillation counting, and each data
point represented the average ± SD of three to six
wells.
Data Analysis.
Data from replicate measurements of a single assay are expressed as the
mean value ± 1 SD, and data representing entire cohorts
are represented as the mean value for all of the subjects in a
cohort ± 1 SE. The hypothesis that treatment either
increased or decreased an outcome variable within a given cohort or
between different cohorts was evaluated using either a paired
Student’s t test or an unpaired Student’s t
test, respectively. A two-way ANOVA with correction for multiple
comparisons was used to compare MLR results for the same subject when
measured on different days of the treatment. A P of 
0.05
was considered significant.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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. Reversible, cytokine-induced leukocytosis occurred in all patients
with maximal day 14 peripheral blood counts averaging 22,300 ± 10,070 in cohort A and 30,300 ± 7,238 in cohort
D (no significant differences). Maximal day 7 counts in the E cohort
averaged 15,100. Side effects consisted primarily of grade 1 and 2
constitutional symptoms: fever; bone pain; local injection site
erythema; nausea and vomiting; thrombocytopenia; or transaminitis.
Grade 3 hepatic toxicity (elevated alkaline phosphatase) occurred in
one subject each in cohorts C and D, and grade 3 headache occurred in
one subject in cohort E, all of which resolved spontaneously without
interruption in therapy. No dose-limiting toxicity was observed, but
one patient in cohort AI receiving GM-CSF alone voluntarily withdrew
from the study after developing a rash at the injection site.
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4.0 µg/kg/day IL-4, there was a
further increase in the number of circulating
CD14+ cells beyond that observed with GM-CSF
alone (Fig. 1
). On day 7, peripheral blood CD14+ counts
averaged 5-fold higher in cohort D (P < 0.05) and 2.2-fold higher in cohort E as compared with the increase
that resulted from GM-CSF alone. The number of
CD14+ cells returned to baseline within 7 days of
completing cytokine therapy in all cohorts.
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. This effect was detectable on day 7 in the C cohort and was
statistically significant at both days 7 and 14 in the D cohort.
Similar effects were observed by day 7 in the E cohort, where CD14
expression was decreased by 65% compared to the day 0 value.
Concurrent with the decrease in CD14, there was an IL-4-dependent
increase in the expression of HLA-DR in the same cells (Fig. 2
). By day 7 of treatment, HLA-DR expression increased to 229 ± 87% of day 0 levels in the D cohort (P < 0.05) and 308 ± 83% of day 0 levels in the E
cohort (P < 0.05). Significant increases in
the expression of HLA-DR were not observed in patients receiving GM-CSF
alone. A similar trend was observed for the up-regulation of CD11c
(Fig. 2
).
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) and expression of CD86 was not increased at all on circulating
CD14+ cells from treated patients. To evaluate
this further, peripheral blood leukocytes were isolated from cohort E
on day 7 of in vivo therapy and cultured for 24 h
in vitro in the absence of any exogenous cytokines. This
short-term culture resulted in a dramatic up-regulation of CD80, CD86,
CD11c, and HLA-DR on the CD14dim+ cells,
producing a pattern of expression similar to that observed when
monocytes are cultured in vitro with GM-CSF and IL-4 for a
full 7 days (19)
. Overnight culture also resulted in
expression of CD83, a marker usually expressed only on functionally
mature DCs.
|
).
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and 5
). The administration of GM-CSF, alone or in combination with 0.5 or 1.0
µg/kg/day IL-4, did not significantly increase the percentage of
CD83+/HLA-DR+ cells.
However, there was a dose-dependent increase in the percentage of
CD83+/HLA-DR+ cells at
higher IL-4 doses, with the maximal effect observed in the D cohort
(2.35 ± 1.8% of PBMCs in the D cohort
versus 0.09 ± 0.09% of PBMCs in the AI
cohort, P < 0.05). In terms of the total number
of circulating
CD83+/HLA-DR+ cells
recovered per milliliter of blood, there was an average 130-fold
increase by day 7 in patients treated with 4 µg/kg/day IL-4 (range,
26–399-fold increase; Fig. 1B
). This
CD83+/HLA-DR+ population
was comprised of two distinct subsets, one containing smaller cells
with minimal autofluorescence, and the other containing larger cells
with a relatively high autofluorescence (Fig. 6
). Both subsets expressed HLA-DR and CD1a in the absence of CD14, a
pattern consistent with DCs of stem cell origin.
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|
). In cohorts D and E, the CD14+ populations were
purified by cell sorting and compared for their allostimulatory
activity. A treatment-related increase in activity was observed in two
of the four patients in cohort D and in four of the four patients in
cohort E (Fig. 7
). In one patient (patient D4),
CD83+ cells were also prepared by cell sorting
and found to stimulate greater T-cell proliferation than either PBMCs
or sorted CD14+ cells collected from day 0.
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Whereas the ex vivo generation of cytokine-derived DCs offers an important new approach to cancer therapy, it requires considerable resources and expertise. A leukopharesis is required to obtain precursors, recovered cells need to be purified and differentiated in vitro under sterile conditions, and mature DCs need to be loaded with antigen and returned to patients. As an alternative to ex vivo therapy, we hypothesized that the combination of GM-CSF and IL-4 might be administered directly to cancer patients to generate cytokine-derived APCs in situ. This approach has several potential advantages in terms of the large number of CD14+ and CD34+ precursors that exist in vivo; the capacity to activate APCs directly within lymphoid organs, tissues, and tumor sites; and the potential for a simplified home-based therapy. Patients in this study quickly learned how to administer their own injections at home. The expansion of APCs in this manner could also be combined with standard vaccine approaches to target responses against specific antigens. However, even in the absence of antigen targeting, the placement of cytokine-derived APCs directly within murine tumors has been shown to induce tumor regression (41) . Similarly, the in vivo expansion of APCs with systemic flt3 ligand induces antitumor immunity in a variety of tumor models (42, 43, 44) . These animal models, in combination with human studies that find a correlation between the number of tumor-associated APCs and clinical outcome (8, 9, 10, 11) , support the hypothesis that increasing the number of APCs in vivo might activate antitumor immunity.
The feasibility of combining GM-CSF and IL-4 as an in vivo
therapy was based on past experience with these cytokines. As others
have reported (45
, 46)
, we found that a 2-week course of
GM-CSF was well tolerated and induced a 5–20-fold increase in the
number of circulating CD14+ cells. Although there
is less experience with IL-4, a prior study observed 5–6 µg/kg/day
to be the maximum tolerable dose when given alone as a daily injection
for 2 weeks (47)
. Pharmacokinetic analysis documented
blood levels in the range of 1–12 ng/ml after a 4 µg/kg/day
injection of IL-4 and suggested that peak levels might be sustained for
up to 8 h (48)
. Preliminary in vitro
studies (data not shown) confirmed that as little as 1.25 ng/ml IL-4
was sufficient, when combined with GM-CSF, to promote human monocytes
to mature into APCs, with higher levels producing more rapid and
complete DC maturation. The fact that neither GM-CSF nor IL-4 has
produced marked antitumor responses in the past when used as single
agents is not surprising (45
, 49
, 50)
. The capacity for
GM-CSF and IL-4 to differentiate precursors into DCs is highly
synergistic, requiring both cytokines to achieve an effect
(20)
. This type of synergism was readily apparent in our
study when the combination of GM-CSF and IL-4, but not GM-CSF alone,
dramatically increased the number of CD14+ and
CD83+ cells with APC characteristics. In terms of
tolerability, combined treatment with GM-CSF and IL-4 at doses of 4
µg/kg was similar to GM-CSF alone and was not associated with any
dose-limiting toxicity. At 6 µg/ml, patients experienced a higher
frequency and severity of constitutional symptoms, which prompted us to
shorten the treatment interval to 7 days.
Using the characteristics of in vitro monocyte-derived DCs as a guide, we carefully evaluated circulating CD14+ cells from patients for evidence of maturation along a DC pathway. Down-regulation of CD14 expression was used as a hallmark to identify monocyte-derived APCs and allowed us to identify the concurrent up-regulation of HLA-DR and CD11c on these cells. In contrast to the phenotypic changes that occurred under optimal in vitro conditions, the cells from treated patients retained modest CD14 expression and did not express high levels of either CD80 or CD86. One conclusion would be that in vivo cytokine therapy resulted in an intermediate stage of DC differentiation. This would not be surprising because cytokine-induced DC maturation occurs in a continuum that is both dose- and time-dependent. Higher dosing with IL-4 or secondary stimulation by CD40 ligand or other activating factors might be required to produce more mature DCs. Alternatively, more mature DCs that developed in response to therapy might have exited the circulation before our sampling. We obtained blood only once a week and, in each case, 24 h after the last cytokine dose. Rapid margination and trafficking of cytokine-activated cells to peripheral sites is a common phenomenon after cytokine therapy, including therapy with GM-CSF (45 , 46) . Unfortunately, biopsies from tumor sites and/or lymph nodes were not obtained in this study and could not be evaluated for DC infiltration. Finally, the failure to see full expression of CD80 and CD86 on circulating APCs could have resulted from tumor-derived immunosuppressive factors interfering with DC maturation.
To address these issues, we recovered circulating cells from treated patients and placed them into short-term culture in the absence of any exogenous cytokines. Within 24 h, we observed a dramatic up-regulation of HLA-DR, CD80, CD86, CD11c, and CD83 on the CD14dim+ population, a phenotype almost identical to that generated by culturing monocytes for 7–8 days in high concentrations of GM-CSF and IL-4. This rapid in vitro maturation is similar to that reported when naturally occurring DCs are isolated from the peripheral blood of control subjects (51 , 52) . In addition to acquiring the phenotypic characteristics of APCs, CD14+ cells from patients treated with higher doses of IL-4 also increased their capacity to take-up FITC-labeled dextran, a well-characterized marker of functionally active DCs (28) , as well as their capacity to stimulate T cells in a MLR. Collectively, these findings suggest that combined therapy with GM-CSF and higher doses of IL-4 induced myeloid precursors to differentiate into functional APCs consistent with immature DCs. Whether these circulating cells migrate into tissue and lymphoid organs and terminally differentiate, as they did when cultured for 24 h in vitro, remains to be determined.
In addition to their effects on peripheral blood monocytes, GM-CSF and IL-4 are involved in the expansion of DCs directly from bone marrow precursors. DCs generated in this manner mature either through a CD14+ intermediate stage or more directly into DCs as identified by the earlier acquisition of CD83 and CD1a in the absence of CD14 (16 , 17) . As with CD14+ cells, we observed that GM-CSF in combination with higher doses of IL-4 produced a significant increase in the number of circulating CD83+ cells. These cells lacked CD14, expressed CD1a and high levels of HLA-DR, and were active APCs in preliminary sorting experiments. Two distinct CD83+ populations emerged at higher IL-4 doses, one composed of smaller cells, and one composed of larger cells. A preliminary sorting experiment suggests that these CD83+ cells are able to stimulate T-cell proliferation, but further work is required to confirm their origins and functional role. Although increased up to 300-fold in response to GM-CSF and IL-4, the CD83+ population was still outnumbered 10-fold by CD14dim APCs.
From a clinical perspective, it appears likely that the administration of GM-CSF and IL-4 can be used to generate large numbers of APCs in situ. Although this Phase I study enrolled only four patients in each dose group, the development of objective antitumor responses in two patients is striking. The changes in PSA, lymph node involvement, and bone disease all correlated well with improvement in the responding patient’s symptoms. PSA responses have been described for patients vaccinated with GM-CSF/IL-4-generated DCs that were loaded ex vivo with prostate-specific membrane antigen (14 , 15) . It is interesting to speculate that the APCs generated in vivo in our patients were able to gain access to the patient’s own tumor antigens and initiate an antitumor response. As stated previously (42, 43, 44) , similar antitumor responses have been reported in mice when flt3 ligand was used to expand DCs in vivo. In a recent study (53) , patients with metastatic prostate cancer were treated with intermittent 2-week courses of GM-CSF alone (Leukine; 250 µg/m2/day). Ten of 22 patients treated with this regimen developed intermittent decreases in serum PSA, but their values uniformly rose to levels higher than the pretreatment levels after each treatment cycle. High levels of GM-CSF were found to directly down-regulate PSA expression when added to the LNCaP cell line in culture, offering one potential mechanism for this phenomenon. In contrast to that report with GM-CSF alone, our prostate cancer patients treated with GM-CSF and IL-4 exhibited a progressive decline in PSA and tumor burden over time. Other studies have either found minimal antitumor effects of GM-CSF therapy alone (45 , 50) or documented significantly fewer patient responses when GM-CSF was administered concurrently as an adjuvant with peptide vaccines or peptide-pulsed DCs (54 , 55) .
In conclusion, this Phase I study suggests that the combined administration of GM-CSF and IL-4 to patients with advanced cancer produces an IL-4-dependent increase in the number of circulating CD14+ and CD83+ APCs with many of the phenotypic and functional characteristics associated with DCs. Systemic cytokine administration aimed at generating APCs in situ, either alone or in combination with the administration of targeting antigens, may provide an additional approach to cancer immunotherapy.
ACKNOWLEDGMENTS
Support and recombinant human cytokines were kindly provided by
the Schering-Plough Research Institute (Kenilworth, NJ), under
the direction of Dr. Mary Ellen Rybak, past Director of Clinical
Oncology Research. We also thank Mitzi Ng for technical assistance.
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FOOTNOTES |
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1 Supported in part by the Jonsson Cancer Center
Foundation/University of California at Los Angeles (M. D. R.),
American Cancer Society Grant CRTG-97117-01-CCE (to B. J. G.), and
the Schering-Plough Research Institute. 
2 To whom requests for reprints should be
addressed, at the Division of Pulmonary and Critical Care Medicine,
Department of Medicine, University of California at Los Angeles School
of Medicine, Los Angeles, CA 90095-1690. Phone: (310) 206-7389; Fax:
(310) 267-2020; E-mail: mroth{at}mednet.ucla.edu
3 The abbreviations used are: APC,
antigen-presenting cell; GM-CSF, granulocyte macrophage
colony-stimulating factor; IL, interleukin; DC, dendritic cell; MLR,
mixed leukocyte reaction; CM, complete medium; mAb, monoclonal
antibody; FACS, fluorescence-activated cell-sorting; PSA,
prostate-specific antigen; LFI, relative mean linear fluorescence
intensity; FL, fluorescence channel; PBMC, peripheral blood mononuclear
cell; PE, phycoerythrin; PerCP, peridin chlorophyll protein.
Received 10/ 6/99. Accepted 2/ 3/00.
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, P < 0.05 compared to cohort AI or AII on the same day, unpaired
t test.
