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Leukemia-Derived Immature Dendritic Cells Differentiate into Functionally Competent Mature Dendritic Cells That Efficiently Stimulate T Cell Responses¹

Alessandro Cignetti^2,*, Antonella Vallario^*,, Ilaria Roato^*, Paola Circosta^*, Bernardino Allione, Laura Casorzo, Paolo Ghia^*, and Federico Caligaris-Cappio^3,*,

^* Laboratory of Cancer Immunology, Institute for Cancer Research and Treatment, Department of Oncological Sciences, University of Torino Medical School, and Unit of Pathology, Institute for Cancer Research and Treatment, Candiolo, Italy; and Division of Hematology, Azienda Ospedaliera Santissimi. Antonio e Biagio, Alessandria, Italy

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Primary acute myeloid leukemia cells can be induced to differentiate into dendritic cells (DC). In the presence of GM-CSF, TNF-, and/or IL-4, leukemia-derived DC are obtained that display features of immature DC (i-DC). The aim of this study was to determine whether i-DC of leukemic origin could be further differentiated into mature DC (m-DC) and to evaluate the possibility that leukemic m-DC could be effective in vivo as a tumor vaccine. Using CD40L as maturating agent, we show that leukemic i-DC can differentiate into cells that fulfill the phenotypic criteria of m-DC and, compared with normal counterparts, are functionally competent in vitro in terms of: 1) production of cytokines that support T cell activation and proliferation and drive Th1 polarization; 2) generation of autologous CD8⁺ CTLs and CD4⁺ T cells that are MHC-restricted and leukemia-specific; 3) migration from tissues to lymph nodes; 4) amplification of Ag presentation by monocyte attraction; 5) attraction of naive/resting and activated T cells. Irradiation of leukemic i-DC after CD40L stimulation did not affect their differentiating and functional capacity. Our data indicate that acute myeloid leukemia cells can fully differentiate into functionally competent m-DC and lay the ground for testing their efficacy as a tumor vaccine.

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Relapse is the main reason of treatment failure in acute myeloid leukemia (AML).⁴ Though >80% of AML patients achieve complete remission after conventional chemotherapy, >50% invariably relapse (1), underscoring the need of novel treatment strategies (2). One such strategy entails the development of immunotherapy approaches designed to obtain AML-specific CTLs potentially able to eradicate or control minimal residual disease (3).

Priming of effective CTL responses requires the presentation of relevant Ags by a professional APC (4). In this context, AML provides the unique opportunity to derive APC from leukemic cells themselves, combining the expression of all available leukemic Ags with the presence of the several accessory and costimulatory signals which are necessary to prime naive T cells (5). The simplest way to derive professional APC from myeloid malignancies is to induce the differentiation of leukemic cells into dendritic cells (DC) (6, 7), which can actually be obtained upon in vitro culture of primary AML cells in the presence of various cytokine combinations (8, 9, 10, 11). Leukemic DC have proven to be potent stimulators of allogeneic T lymphocytes in mixed leukemia-lymphocyte reactions, and a few reports also suggest that autologous CTL can be generated in vitro by stimulation with leukemic DC (9, 11, 12, 13, 14).

In chronic myeloid leukemia and in AML, leukemic DC have been initially obtained in vitro by stimulation with GM-CSF together with IL-4 and/or TNF- (8, 9, 10, 11, 15, 16, 17). With the latter cytokine combination, the leukemia-derived DC do not acquire a fully mature phenotype but rather an immature one, displaying low or intermediate levels of costimulatory molecules and low to undetectable levels of the DC maturation marker CD83 (8, 12, 13, 18, 19, 20, 21, 22, 23, 24); they also produce low to undetectable amounts of IL-12, another hallmark of DC maturation (11, 25, 26). Recent studies have suggested that normal immature DC (i-DC) or semimature DC (induced by TNF-) injected in humans might have an inhibitory effect on T cell function (27). On the contrary, normal mature DC (m-DC) are the most potent APC for efficient T cell priming in vivo (28, 29). So far, the stepwise differentiation of leukemic blasts to i-DC and then to m-DC has never been studied and characterized from a preclinical point of view (26, 30).

If DC of leukemic origin are planned to be used in vivo as a cancer vaccine, it becomes crucial to determine whether they can differentiate to m-DC not only in terms of phenotype but, most importantly, of function. To this end, we have studied 20 AML patients analyzing whether leukemic i-DC could fully differentiate to m-DC in response to CD40L and whether leukemic m-DC were functionally competent in terms of: 1) production of cytokines that support Ag-specific T cell activation, proliferation, and Th1 differentiation; 2) generation of autologous T cell effectors; 3) migration from tissues to lymph nodes; 4) amplification of Ag presentation by attraction and recruitment of monocytes and other i-DC; 5) attraction of resting and activated T cells. We here show that leukemic blasts could be induced to differentiate into cells fulfilling the phenotypic and functional criteria of m-DC in 10 of 20 cases. Irradiation of leukemic i-DC after CD40 stimulation did not affect their differentiation to m-DC in terms of phenotype, cytokine/chemokine production, and migration properties. These data lay the ground for testing the efficacy of leukemic m-DC as a vaccine aimed at eradicating minimal residual disease in a relevant number of AML patients.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
Patients

After informed consent, samples from either bone marrow (BM) or peripheral blood (PB) were obtained at diagnosis from 20 AML patients. In some cases, samples were also collected during morphological remission (<5% BM blasts) after chemotherapy. Table I shows the main clinical and diagnostic laboratory data of all cases.

BM or PBMCs were isolated after centrifugation over a density gradient (Lymphoprep; Nycomed, Oslo, Norway) and used immediately or cryopreserved. Fresh or thawed primary AML cells were seeded into 25 or 75 cm² flasks at 1 x 10⁶/ml and cultured in serum-free medium (X-Vivo15; BioWhittaker, Walkersville, MD). In four selected cases, leukemic cells were obtained by depleting PBMC with a mixture of magnetically labeled mAbs against CD3, CD19, CD16, and CD56 (Miltenyi Biotec, Auburn, CA), resulting in a purity of >98% leukemic cells (Table I). To generate leukemic i-DC, purified or unfractionated AML cells were stimulated with GM-CSF, IL-4 (both at 80 ng/ml), and TNF- (10 ng/ml) (BioSource International, Camarillo, CA). Cytokines and fresh medium were added every 4–5 days.

Normal i-DC were obtained from purified CD14⁺ cells using GM-CSF and IL-4 as described elsewhere (31). To generate leukemic and normal m-DC, cells from i-DC cultures were CD40 cross-linked using 300 ng/ml of a rCD40L-FLAG-tag fusion protein supplemented with 300 ng/ml "CD40L enhancer" (Alexis Biochemicals, San Diego, CA). After 24, 48, or 72 h, depending on the assay, cells were used for functional analysis.

Flow cytometric analysis

Unmanipulated and cultured AML samples were stained with FITC, PE, or PECy5-conjugated mouse mAbs using standard multicolor methodology, and analyzed by flow cytometry (FACS). Blasts and DC were gated based on side light scatter profile vs CD45, as previously described (31). Mouse mAbs to the following Ags were used: CD1a, CD33, CD34, CD80, CD86, CD40, HLA class I, HLA-DR (BD Pharmingen, San Diego, CA), CD13, CD14 (BD Biosciences, San Jose, CA), and CD83 (Immunotech, Westbrook, ME). Unconjugated mAb to CCR7 (IgM) was obtained from BD Pharmingen. PE-conjugated secondary goat polyclonal Ab against murine IgM was purchased from Southern Biotechnology Associates (Birmingham, AL). For other chemokine receptor mAbs, see Ref. 31 . Isotype-matched mAbs were used as negative controls. Samples were analyzed with a FACSCalibur cytometer (BD Biosciences).

Fluorescence in situ hybridization (FISH) analysis

Interphase FISH was performed on CD83⁺ purified leukemic m-DC and on naive leukemic blasts (10). Three patients were selected according to their cytogenetic abnormality and interphase nuclei were analyzed using the probes of interest. For 7q^– and 5q^– deletion (patients GE and VA), dual color FISH was performed, using probes labeled either with spectrum green or spectrum orange. A centromeric probe for chromosome 7 (7p11.1-q11.1) and a probe for 7q31 were used for the 7q^– deletion, while a probe for 5p15.2 and a probe for 5q33-34 were used for the 5q^– deletion (VYSIS, Downers Grove, IL). For monosomy 7 (patient CA), only the centromeric probe for chromosome 7 was used. Fixation, denaturation, and hybridization were conducted following manufacturers’ instruction. At least 200 nuclei were examined under fluorescence microscopy.

RNA preparation and RT-PCR

RNA was extracted from 1 to 2 x 10⁶ cells using RNAzol (Biotecx Laboratories, Houston, TX) following the manufacturer’s instructions. cDNA was prepared at 42°C using a reverse-transcription mix containing Superscript II (Invitrogen Life Technologies, Grand Island, NY). PCR amplification of cDNA samples was performed for 30 or 35 cycles. For chemokines and -actin primer sequences, see Ref. 32 . For each cytokine and chemokine, proper positive controls were used. All reactions were conducted in a PerkinElmer thermocycler (Foster City, CA).

Cytokine and chemokine production

Supernatants were obtained from cultures that, after washing, were seeded at 2 x 10⁶cells/ml and kept in serum-free medium for 24 or 72 h. Supernatants from i-DC were obtained after stimulation of i-DC cultures with GM-CSF, TNF- and IL-4; supernatants from m-DC were obtained after stimulation of i-DC cultures with CD40L. Cytokine and chemokine production was measured by ELISA using the following commercially available kits: IL-12, IL-12 HS (high sensitivity), IL-15, IL-18, macrophage- derived chemokine (MDC), thymus and activation-regulated chemokine (TARC), MCP-1, MIP-1, MIP-3, RANTES (R&D Systems, Minneapolis, MN), IFN--inducible protein 10 (IP-10) (CytImmune Sciences, College Park, MD), IL-6, IL-8, IL-10 (Bender MedSystems, Vienna, Austria), and IL-16 (Technogenetics, Milano, Italy).

Stimulatory function of the leukemic DC

Allogeneic mixed lymphocyte-tumor cultures (MLTC) were set up with PBMC from healthy donors as responders and naive leukemic cells, leukemic, and normal DC as irradiated stimulators, as previously described (10). All cultures were performed in serum-free medium (AIM-V; Invitrogen Life Technologies). Normal DC were derived from CD14⁺ purified PBMC of healthy sibling donors that were HLA-matched with the patient. Autologous MLTC were similarly performed, but responder T cells used for proliferation assay were previously cocultured with different stimulators, i.e., autologous blasts, i-DC or m-DC. Briefly, patients’ PBMC were primed with autologous blasts, i-DC, or m-DC and similarly restimulated after 7 days. Proliferation was then measured by pulsing cells with [³H]TdR 4 days after a last stimulation of all differently cultured T cells with autologous blasts.

Induction of autologous T cell effectors

AML blasts were differentiated to leukemic m-DC, irradiated at 30 Gy and used to stimulate autologous PBMC obtained after complete remission. Autologous MLTC were set up with patients’ PBMC (10⁷ nonadherent PBMC) cocultured with 2 x 10⁶ leukemic m-DC in AIM-V serum-free medium. On days 7, 14, and 21, PBMC were restimulated with autologous irradiated leukemic m-DC. rIL-2 (20 IU/ml) was added 72 h after the second stimulation and then every 3–4 days. At day 28, PBMC were separated into CD4⁺ and CD4^– cells using CD4⁺ microbeads (Miltenyi Biotec) and purified fractions were further restimulated with autologous leukemic m-DC. Assay for functional activity was performed 5–7 days after the fourth or fifth restimulation.

ELISPOT assay

The ELISPOT assay was performed using cells from MLTC as effectors (4 x 10⁴cells/well) and the following autologous targets (1 x 10⁴/well): naive leukemic blasts, CD14⁺ purified monocytes, and EBV-transformed B cells. ELISPOT assays were conducted according to manufacturer’s instruction (Mabtech, Stockholm, Sweden), with few modifications (33). Before developing the assay, T cells were routinely transferred from the ELISPOT plate to a normal 96-well plate and viability of recovered cells was verified after 2 additional days of culture (without cytokines) by checking all wells under the microscope and also by randomly counting some wells with trypan blue exclusion. This allowed us to rule out cell loss or cell death due to technical inaccuracy and also to check correspondence between cell viability and IFN- activity in individual wells. By performing this control of cell viability, in fact, we could exclude that a remarkable T cell death had occurred in those same wells where IFN- activity was detected. Blocking experiments were performed by preincubating target cells for 20' with 25 µg/ml of an anti-HLA class I (clone W6.32) or an anti-HLA class II (clone L243) and an isotype-matched irrelevant mAb (all from BD Pharmingen). Spots were counted by a computer-assisted ELISPOT reader (AID; Bioline, Torino, Italy). Indicated spot numbers per seeded PBMC represent mean values of duplicates.

Chromium release assay

Target cells (primary AML cells) were incubated with 50 µCi of Na₂⁵¹CrO₄ (Amersham Biosciences, Cologno Monzese, Italy) for 2 h at 37°C and washed. Thereafter, 5 x 10³ viable target cells were incubated with CD8⁺ effector cells from MLTC at different E:T ratios for 4 h at 37°C. After incubation, the supernatants were collected and radioactivity was measured on a gamma counter. Spontaneous release was determined by harvesting the supernatant of target cells incubated in the absence of effector cells and maximum release was determined by resuspending target cells incubated in the absence of effector cells. The percent of specific lysis was calculated as follows: (experimental release – spontaneous release/maximum release – spontaneous release) x 100%. The spontaneous release always ranged between 5 and 20% of the total label incorporated by the cells. To determine class I restriction in the recognition of target cells, blocking studies were performed by adding 25 µg/ml anti-HLA class I mAb (clone W6.32) to ⁵¹Cr-labeled targets 30 min before the assay. An Ab of the same isotype was used as negative control. The Abs were not washed out before mixing target and effector cells.

Generation of T cell subsets and migration assay

CD4⁺ and CD8⁺ T cells were purified from PBMC by depleting CD19⁺, CD56⁺, CD16⁺, CD14⁺, and CD8⁺ or CD4⁺ cells with magnetic microbeads (Miltenyi Biotec), and activated by stimulation with anti-CD3 and anti-CD28 (100 ng/ml and 1 µg/ml, respectively) (BD Pharmingen). Purity of the CD4⁺ or CD8⁺ fraction was always >95%. Activated T cells were cultured with IL-2 (2 ng/ml; Chiron, Emeryville, CA) and IL-7 (5 ng/ml; R&D Systems) in the presence of irradiated allogeneic feeder cells (PBMC) for 7 to 14 days. Naive/resting CD4⁺ and CD8⁺ T cells were prepared from nonadherent PBMC negatively depleted of CD14⁺, CD19⁺, CD56⁺, CD45RO⁺, CD25⁺, HLA-DR⁺, and CD8⁺ or CD4⁺ cells using goat anti-mouse Ig-coated magnetic microbeads (Miltenyi Biotec). Purity of the resulting CD45RA⁺/CD4⁺ or CD8⁺ cells was always >95% as controlled by FACS.

Chemotaxis assays were performed using the Transwell system (5 µm pores; Costar, Cambridge, MA), as previously described (31, 32). For chemotaxis of leukemic m-DC, escalating doses of rMIP-3 (R&D Systems) were added to X-Vivo15 and distributed on the lower compartment of the chemotaxis system; CCR7⁺ leukemic m-DC were placed into the Transwell inserts. Plates were incubated for 4 h. A control condition with no recombinant chemokine was also included. For normal monocytes and T cell subset migration, supernatants were collected from leukemic and normal DC cultures and used at 100% v/v or with serial 1/2 dilutions and distributed on the lower compartment of the chemotaxis system. Monocytes or T cells were then placed into the Transwell inserts. T cells were obtained as described above, while normal monocytes were obtained from healthy donors with the monocyte isolation kit as described for DC generation. In selected cases, monocytes or T cells were incubated before the migration assay with saturating concentrations of MCP-1 (2 µg/ml), MDC, or IP-10 (4 µg/ml) for 30 min to down-modulate CCR2, CCR4, and CXCR3 expression, respectively, as evaluated by FACS analysis. Plates were incubated for 2, 4, and 12 h (activated T cells, monocytes, and naive T cells, respectively). A negative control condition with fresh medium was always included.

In all migration assays, the liquid accumulated in the lower compartment of the chemotaxis system was carefully recovered and migrated cells were counted by flow cytometry. Results are expressed as the percentage of migrating cells, i.e., number of migrated cells/number of input cells x 100.

Statistical analysis

The statistical significance (p values) between i-DC and m-DC or between leukemic and normal i-DC and m-DC was analyzed by two-tailed Student t test for paired data.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Generation of leukemic and normal i-DC and m-DC

We analyzed 20 AML samples to determine whether AML blasts could be induced to differentiate first to i-DC and then to m-DC. When a significant proportion of cells cultured in the presence of GM-CSF, TNF-, and IL-4 started to show evidence of DC morphology, their phenotype was analyzed by flow cytometry (usually after 7–14 days) (10). As CD1a and CD83 are considered markers of i-DC and m-DC, respectively, (34, 35), AML cells were considered to have differentiated into i-DC when >50% of cells were positive for CD1a and/or CD80. Samples were then stimulated with CD40L and analyzed for CD83 expression after 24, 48, and 72 h of culture. When at least 50% of the CD80⁺ cells coexpressed CD83, cultured AML cells were considered to have become leukemic m-DC. According to these criteria, AML blasts were able to differentiate to m-DC in 10 of 20 cases (see Table I). Before differentiation, all 10 cases were CD86-positive, while only 8 of 10 cases were CD14-positive. On the contrary, cells were CD86 and CD14 positive only in 1 of 10 cases in which DC differentiation was not achieved (Table I). Fig. 1 shows how the up-modulation of CD83 and CD80 is associated with maturation of leukemic as well as of normal DC. The difference in CD83 and CD80 expression between i-DC and m-DC (both leukemic and normal) is statistically significant, when data are analyzed with a t test for paired data (p < 0.05 and p < 0.005 for CD80 and CD83, respectively). We also analyzed the expression of CD86, CD40, HLA class I, and HLA-DR in leukemic i-DC and m-DC, without finding a significant difference between the two subsets (data not shown).

View larger version (21K): [in this window] [in a new window]   FIGURE 1. Maturation of leukemic DC up-modulates CD83 and CD80 expression. Blasts, leukemic, and normal i-DC and m-DC cultures were analyzed by flow cytometry for the expression of CD83 (A) and CD80 (B). Data are expressed as percent of positive cells and each line represents data obtained with leukemic (n = 10) or normal CD14+ (n = 8) samples.

Maturation of leukemic DC induces the production of IL-12 and IL-15 and down-modulates the production of IL-10

To further evaluate the differentiation stage of leukemic DC, we measured by ELISA the production of IL-12p70 by leukemic i-DC and m-DC and compared it to CD14-derived normal counterparts. As shown in Fig. 2A, supernatants from leukemic and normal i-DC cultures showed negligible IL-12 production. On the contrary, supernatants from both normal and leukemic m-DC cultures showed significant amounts of IL-12 production. Similarly to IL-12, also levels of IL-15 production in both leukemic and normal DC subpopulations were significantly higher in m-DC than in i-DC (Fig. 2B and not shown).

View larger version (19K): [in this window] [in a new window]   FIGURE 2. Maturation of leukemic DC induces IL-12 and IL-15 production and down-modulates IL-10 production. A, Supernatants from i-DC and m-DC cultures were analyzed for IL-12p70 production by ELISA. Cells were seeded at 2 x 106 cells/ml and harvested after 24 h. Data represent the mean ± SD of 10 leukemic samples and of 4 normal samples (CD14-derived DC). *, p < 0.01; , p < 0.05. B, Supernatants from blasts, i-DC, and m-DC cultures were analyzed for IL-15 and IL-10 production by ELISA. Cells were seeded at 2 x 106 cells/ml and harvested after 72 h. Data represent the mean ± SD of 10 leukemic samples. For blasts and i-DC, IL-10 samples were divided in two subgroups, according to their different pattern of IL-10 production. *, p < 0.01; , p < 0.05.

Leukemic m-DC stimulate T cell proliferation better than i-DC in allogeneic and autologous MLTC

To prove that leukemic m-DC are capable of supporting T cell activation and proliferation better than i-DC counterparts, we investigated their ability to stimulate allogeneic T cells in primary MLTC. In three cases analyzed, we found that m-DC elicited a proliferation significantly higher than i-DC and that the stimulatory activity of leukemic i-DC and m-DC is superimposable to that of normal counterparts. CD14-derived normal i-DC and m-DC were obtained from HLA-matched sibling donors (Fig. 3A). We also compared the stimulatory activity of leukemic m-DC with that of i-DC in tertiary MLTC in three additional cases. Patients’ PBMC were repeatedly stimulated with autologous blasts, i-DC, or m-DC and their proliferation in response to naive autologous blasts was measured after 3 wk. Again, we found that m-DC elicited significantly higher proliferation of autologous T cells than i-DC counterparts (Fig. 3B).

View larger version (30K): [in this window] [in a new window]   FIGURE 3. Leukemic m-DC are better T cell stimulators than i-DC in allogeneic and autologous MLTC. A, Allogeneic MLTC was performed with 105 allogeneic PBMCs as responder cells and different numbers of thawed AML blasts or leukemic and normal i-DC and m-DC as stimulator cells at responder/stimulator ratios (R/S) ranging from 3:1 to 30:1. Figure shows data obtained at a R/S of 30:1. B, Autologous MLTC was performed similarly to allogeneic MLTC, but responder cells were patients’ T cells that had been differently stimulated at week intervals with autologous blasts, i-DC, or m-DC. Data in A and B are presented as mean ± SE of three cases in each panel for a total of six patients analyzed. *, p < 0.001 between i-DC and m-DC groups (both leukemic and normal) analyzed with a t test for paired data. All cultures were conducted in serum-free medium.

PBMC from patients were repeatedly stimulated in vitro by autologous leukemic m-DC and analyzed for their ability to recognize naive leukemic blasts. Cells were purified according to their CD4 and CD8 expression (>94% purity after selection), restimulated once more with leukemic m-DC and tested after 7 days by ELISPOT. We found that stimulation with leukemic m-DC generates CD8⁺ and CD4⁺ T cells that release IFN- (Fig. 4, A and B) in response to naive leukemia blasts in three of four and four of four cases, respectively. In one AML patient, the initial number of CD8⁺ cells was very low and the final yield of CD8⁺ T cells was not enough to perform functional assays. CD4⁺ T cells released IFN- but not IL-4 in response to naive leukemic blasts (not shown). MHC restriction of CD8⁺ and CD4⁺ effectors was proved by the inhibition of IFN- release observed with specific anti class-I or class-II mAb, respectively, but not with an irrelevant mAb of the same isotype (IgG2a). Leukemia specificity was further confirmed by the lack of recognition of autologous myeloid (PB monocytes) and lymphoid (EBV-transformed B cells) targets (Fig. 4, A and B). Finally, we also determined whether CD8⁺ T cells from MLTC could mediate effective killing of unmanipulated blasts in a traditional cytotoxicity assay. In two of two cases analyzed, CD8⁺ T cells efficiently lysed both autologous blasts and leukemic m-DC (Fig. 4C). Killing of leukemic m-DC was significantly higher than that of primary blasts. Lysis of AML blasts was blocked by a mAb to HLA-class I, but not by an irrelevant, isotype-matched, control mAb (Fig. 4C).

View larger version (11K): [in this window] [in a new window]   FIGURE 4. Leukemic m-DC generate MHC-restricted CD8+ and CD4+ T cells in autologous MLTC. CD4+ (A) and CD8+ (B and C) T cells from autologous MLTC were tested for their activity against naive autologous blasts, monocytes and EBV-transformed B cells. A and B, T cell reactivity was quantified using an ELISPOT assay for IFN-. Results shown are referred to a T cell-target ratio of 4:1, i.e., 4 x 104 CD4+ or CD8+ T cells and 1 x 104 blasts, monocytes, or EBV-transformed B cells. Figure shows mean ± SE of four experiments for CD4+ T cells and three experiments for CD8+ T cells. C, CTL activity was determined by 51Cr release assay. Figure shows mean ± SE of two independent experiments.

Leukemic m-DC but not i-DC express CCR7 and migrate in response to MIP-3

Maturing DC up-regulate CCR7 expression to acquire the ability to migrate from inflamed tissue to draining lymph nodes in response to CCR7 ligands (MIP-3, secondary lymphoid tissue chemokine). CCR7 was not expressed by leukemic i-DC while a statistically significant induction of CCR7 expression was observed in leukemic m-DC (Fig. 5A). No significant difference in CCR7 expression was observed between leukemic and normal m-DC (not shown). To prove that leukemic m-DC can effectively migrate through CCR7 engagement, we performed a chemotactic assay in vitro, where leukemic m-DC were exposed to rMIP-3. In all four samples analyzed, leukemic m-DC migrated in response to MIP-3, similarly to normal m-DC (Fig. 5B).

View larger version (24K): [in this window] [in a new window]   FIGURE 5. Mature leukemic DC express CCR7 and migrate in response to MIP-3. A, Blasts and leukemic i-DC and m-DC cultures were analyzed by flow cytometry for the expression of CCR7. A value of p of the m-DC group compared with the i-DC group is <0.01 when analyzed with a t test for paired data. B, Migration assay of CCR7+ leukemic m-DC in response to rMIP-3. Migration of four leukemic and three normal m-DC independent cases pooled together are presented.

To amplify Ag presentation, normal DC also produce chemokines to attract monocytes and other DC to inflamed tissue and to secondary lymphoid organs. We analyzed the expression of MCP-1, MIP-1, RANTES, IL-8, MIP-1, and MCP-4 by RT-PCR in both leukemic and normal DC subsets. mRNA for all these chemokines was detected in the vast majority of samples (Table II). Among these chemokines, we analyzed by ELISA the actual production of MCP-1 (Fig. 6A), MIP-1, and IL-8 (Table II) and found that they are also released at significant levels by both leukemic i-DC and m-DC. We next tested whether supernatants from DC cultures could attract in vitro normal monocytes in a chemotactic assay. Fig. 6B shows that supernatants from both leukemic i-DC and m-DC similarly induce the migration of freshly isolated PB monocytes.

View larger version (20K): [in this window] [in a new window]   FIGURE 6. Leukemic i-DC and m-DC produce inflammatory chemokines that induce the migration of PB monocytes. A, Leukemic i-DC and m-DC produce MCP-1. Supernatants from blasts, i-DC, and m-DC cultures were analyzed for MCP-1 production by ELISA. Cells were seeded at 2 x 106 cells/ml and harvested after 72 h. Data represent the mean ± SD of 10 leukemic samples and of 4 normal samples (CD14-derived DC). B, Supernatants from leukemic DC cultures induce migration of PB monocytes. Chemotaxis of PB monocyte cells in response to leukemic and normal i-DC and m-DC supernatants. Fresh PB monocytes were isolated by depleting T, B, and NK cells from PBMC. Data represent mean ± SD of four leukemic i-DC and m-DC samples and of three normal i-DC and m-DC samples (CD14-derived DC). C, Chemotaxis of PB monocytes in response to leukemic i-DC (n = 3) and m-DC-derived supernatants (n = 3) or rMCP-1, before () or after () CCR2 down-regulation by receptor desensitization with high dose rMCP-1 (2 µg/ml). The difference in migration obtained before and after desensitization is statistically significant in all three groups (p < 0.01).

Leukemic i-DC and m-DC produce chemokines that induce the migration of naive and activated T cells

Normal DC also produce chemokines that attract T cells to the sites of Ag presentation. In particular, MDC/TARC and IP-10/monokine induced by IFN- (MIG) are known to promote the interaction of DC with recently activated T cells that respectively express CCR4 and CXCR3. We found that both i-DC and m-DC of leukemic origin produce considerable amounts of MDC and TARC, similarly to CD14-derived normal DC (Fig. 7A and Table II). A trend toward a higher level of MDC and TARC production by m-DC than by i-DC was observed, which did not reach statistical significance. We also found that, in most cases, leukemic and normal DC subsets express mRNA for IP-10 and/or MIG. As documented by ELISA, IP-10 was detected at the protein level as well, without a significant difference between leukemic i-DC and m-DC.

View larger version (26K): [in this window] [in a new window]   FIGURE 7. Leukemic i-DC and m-DC produce lymphoid chemokines that induce the migration of resting and activated CD4+ and CD8+ T cells. A, Supernatants from blasts, i-DC, and m-DC cultures were analyzed for MDC production by ELISA. Cells were seeded at 2 x 106 cells/ml and harvested after 72 h. Data represent the mean ± SD of 10 leukemic samples and of 4 normal samples (CD14-derived DC). B, Chemotaxis of resting/naive and activated CD8+ T cells in response to leukemic and normal i-DC and m-DC supernatants. Purified CD8+/CD45RA+ T cells were obtained from fresh PBMC by negative selection. Activated T cells were obtained by stimulation of purified CD8+ cells with anti-CD3 and anti-CD28 (see Materials and Methods). Five independent experiments are pooled together. Data represent mean ± SD of four leukemic i-DC and m-DC samples and of three normal i-DC and m-DC samples (CD14-derived DC). The percent of cells migrating in the presence of medium alone was subtracted from each data point. C, Chemotaxis of CD8+-activated lymphocytes in response to leukemic i-DC and m-DC-derived supernatants (n = 3) or rIP-10 and rMDC, before and after CXCR3 or CCR4 down-regulation by receptor desensitization with rIP-10 (4 µg/ml) and MDC (4 µg/ml), respectively. Three independent sets of experiments are pooled together. The percent of cells migrating in the presence of medium alone was subtracted from each data point. The difference in migration obtained before and after desensitization is statistically significant in all three groups (p < 0.05).

The analysis of the chemokine secretion pattern suggests that, among the several chemokines detected at the protein level in leukemic DC supernatants, at least two chemokine/chemokine receptor pairs might be involved in the attraction of activated T cells: IP-10-MIG/CXCR3 and MDC-TARC/CCR4. To investigate this issue, we performed again desensitization experiments as described in the previous paragraph. We found that CXCR3- and CCR4-desensitized T cells migrated significantly less than normal counterpart cells in response to leukemic DC supernatants. (Fig. 7C).

Irradiation of leukemic i-DC after CD40L stimulation does not affect their differentiation to m-DC

To be used as a vaccine in vivo, leukemic DC should be irradiated to prevent proliferation of residual leukemic cells. To evaluate how irradiation affects maturation and function of leukemic DC, we stimulated i-DC cultures from three different patients with CD40L and after 4 h we exposed them to escalating doses of gamma rays. The following parameters were then assessed at different time points: viability, expression of maturation markers, cytokine/chemokine production, and chemotactic activity. As shown in Fig. 8, a progressive decrease in cell viability was observed, which was more pronounced after the third day of treatment, and complete cell death was seen at day 4. Despite cell death, remaining living cells showed induction of CD83 and CCR7 expression similar to nonirradiated cultures, which was maintained for up to 60 h. Moreover, supernatants harvested 24 h after irradiation showed no difference between treated and untreated cells in terms of chemokine and cytokine production. Fig. 8 shows ELISA data for TARC and IL-12; similar data were obtained for IL-15, IL-10, MDC, and MCP-1. Finally, irradiated CCR7⁺ m-DC were able to migrate in response MIP-3 and levels of migration were similar between irradiated and nonirradiated cells up to 48 h. Thus, irradiated leukemic DC maintain their functional properties up to at least 2 days after treatment.

View larger version (11K): [in this window] [in a new window]   FIGURE 8. Irradiation of leukemic i-DC after CD40 stimulation does not affect their differentiation to m-DC. i-DC cultures were stimulated with CD40L (day 0) and, after 4 h, were gamma-irradiated with three different doses (10, 30, and 60 Gy). Irradiated and nonirradiated cells were then analyzed at different time points as indicated. A, Viability as determined by counting cells on a Bauer chamber after staining with a 0.4% trypan blue solution to exclude dead cells; B and C, phenotype as determined by FACS analysis of residual living cells (gate on living DC was set as described in Materials and Methods); D, migration in response to rMIP-3 (200 ng/ml); E, chemokine and cytokine production as determined by testing culture supernatants harvested 24 h after irradiation. All data are referred to one representative case of three analyzed.

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

In this study, we show that leukemic i-DC can be induced to fully differentiate into m-DC using CD40L as maturating agent, and that leukemic m-DC fulfill the functional criteria listed above. For instance, leukemic m-DC (but not i-DC) up-regulate CCR7 expression and are able to migrate in vitro in response to the CCR7 ligand MIP-3. Moreover, leukemic m-DC (but not i-DC) secrete IL-12 and IL-15, two cytokines that support T cell activation, proliferation, and differentiation, and down-regulate the secretion of IL-10, which has inhibitory activity on T cells and on DC as well. This pattern of cytokine production finds functional correspondence in the superior capacity of leukemic m-DC to stimulate allogeneic as well as autologous T cell proliferation, when compared with i-DC. More importantly, the ability of m-DC to generate both CD8⁺ and CD4⁺ leukemia-specific T cells was also documented in the autologous setting. Other groups have shown that leukemia-derived DC (that were obtained otherwise than inducing maturation of i-DC with CD40L) can stimulate T cell responses in vitro (9, 11, 12, 13, 14). However, this is the first study that provides concurrent evidence that the antileukemic activity is MHC restricted, is directed against unmanipulated blasts, resides specifically in purified CD8⁺ or CD4⁺ subsets, and can be obtained using serum-free medium, which is mandatory for clinical purposes. In addition, we have shown that normal myeloid (i.e., autologous monocytes from PB) and lymphoid (EBV-transformed B cells) targets are not recognized by our T cell effectors, suggesting the existence of leukemia-associated or leukemia-specific Ags amenable to identification. In two cases, we also could provide proof of principle that our CTL were really effective in the killing of naive AML cells, suggesting that Ag recognition is not accompanied by any impairment of CTL functional properties due to tumor counterattack, such as CTL apoptosis induced by FasL-expressing AML cells.

Finally, to be used in vivo, leukemic m-DC need to be irradiated to prevent proliferation of residual undifferentiated leukemic cells. Therefore, leukemic DC should maintain their functional properties also after irradiation. Notably, leukemia-specific autologous T cells were obtained using irradiated leukemic DC as stimulators and this observation already provides evidence for their capacity to activate T cells in vitro. In addition, leukemic i-DC that were irradiated 4 h after CD40L stimulation could still differentiate into functional competent m-DC, as shown by their capacity to up-modulate maturation markers such as CD83 and CCR7, to produce cytokines/chemokines and to migrate in vitro up to 48 h after treatment. For vaccine design purposes, though, it has to be kept in mind that a decrease in cell number occurs within the same time frame. Dose, frequency, and route of DC delivery are crucial parameters for vaccine effectiveness, which have not been optimized yet in clinical trials with normal nonirradiated DC (37). Nevertheless, based on our data, it is reasonable to anticipate that a higher cell number should be used when vaccinating with leukemic m-DC as compared with normal DC, also because cells with m-DC phenotype do not represent the totality of cells in bulk cultures of leukemic DC. Tumor cell number is not a limitation in AML patients, though, considering the relative ease of tumor cell acquisition at disease presentation. The availability of large numbers of leukemic cells also overcomes the issue of the low yield of viable leukemic m-DC at the end of the differentiating culture (6, 7, 38). In our series, cells with m-DC phenotype represented 10–50% of the initial number of input cells, which would enable (considering 500 x 10⁶ as the minimum number of leukemic cells obtainable at diagnosis), the manufacturing of at least 10 vaccine preparations of 5 x 10⁶ cells/vaccine. The real limiting factor remains only the fact that differentiation into leukemic m-DC could be achieved only in half of the patients analyzed. On one hand, a possible solution would be the prediction of which patients will generate leukemic DC and which patients will not. Recently, it has been shown that the expression of CD14 (30) and of CD86 (20) on AML blasts correlates with their capacity of generating cells with DC features. In agreement with these observations, our data confirm the predictive value of both markers and particularly of CD86, which was expressed by AML blasts in all 10 cases that could differentiate to m-DC. This marker (alone or in combination with CD14) could be very helpful for determining which patients are eligible for vaccination. On the other hand, the addition of other cytokines to the GM-CSF/TNF-/IL-4 mixture (for instance, stem cell factor and Flt-3) (13, 21, 22, 23) or the use of other compounds (such as phorbol esters or bryostatin) (26, 39) might increase the number of responsive cases.

To our knowledge this is the first report that studies the differentiating potential of leukemic i-DC into m-DC and provides full functional characterization of leukemic m-DC from a preclinical point of view. In addition, we show that leukemic m-DC efficiently prime T cells from AML patients, generating MHC-restricted CD4⁺ and CD8⁺ cells that are leukemia-specific. Our data suggest that irradiated leukemic m-DC should be tested as a vaccine aimed at eradicating minimal residual disease in a significant fraction of AML patients.

	Footnotes

 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

¹ This work was supported by the "Associazione Italiana per la Ricerca sul Cancro" (AIRC), Milano, and by Ministero dell’Università e Ricerca Scientifica (MIUR), Roma, Italy.

² Address correspondence and reprint requests to Dr. Alessandro Cignetti, Laboratory of Cancer Immunology, Institute for Cancer Research and Treatment, Strada Provinciale 142, Candiolo (TO), 10060, Italy. E-mail address: alex.cignetti{at}unito.it

³ Current address: Università Vita Salute San Raffaele, 20132 Milano, Italy.

⁴ Abbreviations used in this paper: AML, acute myeloid leukemia; DC, dendritic cell; BM, bone marrow; PB, peripheral blood; i-DC, immature DC; m-DC, mature DC; FISH, fluorescence in situ hybridization; MLTC, mixed lymphocyte-tumor culture; MDC, macrophage-derived chemokine; TARC, thymus and activation-regulated chemokine; IP-10, IFN--inducible protein 10; MIG, monokine induced by IFN-; PARC, pulmonary and activation-regulated chemokine.

Received for publication December 22, 2003. Accepted for publication June 4, 2004.

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