Published online before print February 3, 2006
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* Laboratoire d’Immunologie et Histocompatibilité, INSERM U662, Institut Universitaire d’Hématologie, Paris, France;
Service des Maladies du Sang and
Laboratoire Central d’Hématologie,
¶ Unité de Thérapie Cellulaire et Clinique Transfusionnelle, AP-HP, Hôpital Saint-Louis, Paris, France; and
Service d’Onco-Hématologie, Centre Hospitalier de Versailles, Le Chesnay, France
1Correspondence: Unité de Thérapie Cellulaire et Clinique Transfusionnelle, Hôpital Saint-Louis, 1, avenue Claude Vellefaux, 75010 Paris, France. E-mail: delphine.rea{at}sls.ap-hop-paris.fr
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: CML • vaccines
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The power of cellular immunity to clear CML, demonstrated by the graft-versus-leukemia effect following allogeneic stem cell transplantation (ASCT) and by the emergence of CML-specific T cells in patients responding to interferon-
(IFN-) and ASCT, provides a rationale for developing anti-CML vaccines [5
6
7
]. Leukemic target antigens include the oncogenic p210bcr-abl fusion protein and overexpressed self-proteins such as proteinase 3, preferentially expressed antigen in melanoma, and Wilms’ tumor gene 1 [8
]. Dendritic cells (DC), the most potent antigen-presenting cells of the immune system, play a crucial role in the induction of T cell immunity against malignancies. Results from tumor-bearing animals and phase I/II clinical trials in cancer patients show that DC engineered to express tumor antigens represent highly promising cellular vaccines [9
]. In CML, the presence of the bcr-abl gene in CD34+ progenitors and monocytes [10
] provides a unique opportunity to generate DC with endogenous expression of leukemic antigens. Two pilot studies have tested bcr-abl+ DC vaccines in late CP-CML patients resistant to IFN. Ossenkoppele et al. [11] used DC obtained from total peripheral blood mononuclear cell (PBMC) cultures, and Takahashi et al. [12] performed sequential injections of blood DC and of immature and mature DC derived from adherent monocytes. Some immune responses were raised but without clinical improvement. The high leukemic burden at the time of vaccination is a potential reason, which could have precluded clinical efficacy in these settings. The addition of DC vaccines in CML patients with low leukemic burden under imatinib may yield better results by favoring further reduction in residual disease and increasing the proportion of patients with a complete molecular response. This approach requires investigations of potential immunoregulatory effects of imatinib on DC. The fact that imatinib impairs the development and maturation of healthy donors’ CD34+ progenitors and monocytes toward DC and macrophages in vitro [13
14
15
], combined with our recent finding that patients responding to imatinib show reduced blood DC in comparison with healthy individuals [16
], is indeed raising concerns about the efficacy of DC vaccines in imatinib-treated patients. Until now, the impact of imatinib on monocyte-derived DC harboring the therapeutic target of imatinib in CML, the bcr-abl oncoprotein, has never been explored.
The objectives of our study were to ask whether imatinib could affect bcr-abl+ monocyte-derived DC differentiation, maturation, and immunostimulatory functions and to compare bcr-abl+ and normal monocyte-derived DC. Toward this goal, we had to deplete granulocytic precursors abnormally present in the peripheral blood from untreated CP-CML patients prior to CD14-based monocyte purification to properly analyze bcr-abl+ monocyte-derived DC. Unlike monocytes, these granulocytic precursors indeed express low levels of CD14 but are unable to differentiate into DC. We found that imatinib had a modest inhibitory impact on bcr-abl+ and normal monocyte-derived DC differentiation and decreased interleukin (IL)-12p70 production by mature DC without altering their T cell immunostimulatory functions. However, when imatinib was present during DC-T cell interactions, the capacity of T cells to proliferate was compromised severely.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Differentiation of DC from monocytes
Monocytes were purified from PBMC by positive selection with CD14 microbeads according to the manufacturer’s instructions (Miltenyi Biotec, Auburn, CA). In CML, CD15+ granulocytic precursors circulating in the peripheral blood expressed low levels of CD14. Thus, depletion with CD15 microbeads (Miltenyi Biotec) was performed prior to monocyte isolation. Monocytes (>95% pure) were cultured for 6 days with medium change every 2 days in RPMI 1640 (Life Technologies, Gaithersburg, MD) with 2 mM glutamine, 10% fetal calf serum (FCS), 50 U/ml penicillin, 50 µg/ml streptomycin, 800 U/ml granulocyte macrophage-colony stimulating factor (GM-CSF; Leucomax, Novartis Pharma, Basel, Switzerland), and 500 U/ml IL-4 (R&D Systems, Minneapolis, MN). As previously reported [10
], 85% of CML-DC carried the bcr-abl fusion gene by fluorescence in situ hybridization (FISH) analysis (not shown). For maturation, DC were plated at a density of 106/ml and incubated with 250 ng/ml lipopolysaccharide (LPS) from Escherichia coli strain 055:B5 (Sigma Chemical Co., St. Louis, MO) for 48 h. In some experiments, 1–10 µM imatinib (kindly provided by Novartis Pharma), dissolved in culture medium from 10 mM stock solution in dimethyl sulfoxide (DMSO), was added at initiation of DC differentiation every other day during medium change and during DC maturation. Equal amounts of DMSO were added to control DC to exclude any effect of the solvent. Of note: The doses of imatinib used in this study are in the range of the concentrations pharmacologically achieved in vivo in patients treated with 400 mg daily imatinib [17
].
Quantification of bcr-abl transcripts in monocytes and DC
After purification, monocytes (2x106) were stored at –80°C in TriReagent (Molecular Research Center, Cincinnati, OH). The remaining monocytes were differentiated into immature DC in the presence of GM-CSF and IL-4 as above and stored at –80°C in TriReagent. RNA was extracted, and measurement of bcr-abl transcripts in monocytes before and after DC differentiation was performed by real-time quantitative-polymerase chain reaction (RQ-PCR), as published [18
].
Apoptosis and necrosis assay
Apoptosis of monocyte-derived DC from healthy donors and CML patients and of CD4+ T cells cultured in the presence of imatinib or control DMSO was measured by double-staining with fluorescein isothiocyanate (FITC)-conjugated annexin V-FITC and propidium iodide (PI; Bender MedSystems, Vienna, Austria), according to the instruction of the manufacturer, followed by flow cytometry analysis. Apoptotic cells were defined as annexin V-FITC+/PI–.
Analysis of DC and CD4+ T cell phenotype by flow cytometry
Cells were stained on ice for 20 min in phosphate-buffered saline (PBS)-1% FCS with FITC or phycoerythrin-(PE)-conjugated monoclonal antibodies (mAb) and analyzed by flow cytometry. The following mAb and corresponding isotype controls were used: PE-anti-CD1a (HI149), PE-anti-CD86 (FUN-1), FITC-anti-CD40 (5C3), FITC-anti-human leukocyte antigen (HLA)-A,B,C (G46-2.6), FITC-anti-HLA-DR (G46.6), FITC-anti-HLA-DQ (Tü 169), PE-anti-CD54 (HA58), FITC-anti-class II-associated invariant chain peptide (CLIP; CerCLIP), FITC-anti-CD4 (RPA-T4), and PE-anti-CD25 (M-A51; all from BD PharMingen, San Diego, CA), FITC-anti-CD14 (Miltenyi Biotec), FITC-anti-CC chemokine receptor 7 (CCR7; R&D Systems), and PE-anti-CD83 (HB15A; Immunotech, France).
Antigen-uptake experiments
Antigen-uptake experiments were performed as described by Sallusto et al. [19
]. Cells were resuspended in medium buffered with 25 mM HEPES. FITC-bovine serum albumin (BSA; Sigma Chemical Co.) was added at the final concentration of 1 mg/ml. Cells were incubated at 37°C or 4°C to determine background uptake. After 1 h, cells were washed extensively with ice-cold PBS and analyzed by flow cytometry.
Cytokine secretion by DC
To compare cytokine production during maturation in healthy donors and CML patients, DC (105), exposed or not to imatinib, were stimulated with 250 ng/ml LPS (Sigma Chemical Co.) for 6, 24, and 48 h, followed by restimulation for 24 h with 1 µg/ml CD40 ligand (CD40L; kindly provided by Amgen, Thousand Oaks, CA), as described by Langenkamp et al. [20
]. Human IL-12p70 was measured in the supernatants of DC cultures by solid-phase sandwich enzyme-linked immunosorbent assay (ELISA; Diaclone Research, Stamford, CT; sensitivity, 3 pg/ml).
DC-induced, naive CD4+ T cell polarization
Naive CD4+ T cells from healthy donors were purified by negative selection with the human CD4+/CD45RO– naive T cell subset column kit (R&D Systems) according to the manufacturer’s instructions. Immature and mature DC, exposed or not to imatinib, were plated at 104/well in 96-well U-bottom plates. Allogeneic, naive CD4+ T cells were added to the DC at a density of 5 x 104/well. After 7 days, T cells were harvested and stimulated for 6 h with 20 ng/ml phorbol 12-myristate 13-acetate (PMA) and 250 ng/ml ionomycin in the presence of 5 µg/ml brefeldin A (Sigma Chemical Co.). CD4+ T cells were fixed with 4% paraformaldehyde, permeabilized with PBS containing 0.5% BSA and 0.1% saponin, and stained for intracellular cytokines. Polarization toward T helper cell type 1 (Th1) and Th2 was measured by flow cytometry. The following mAb were used: FITC-anti-IFN-
(4SB.3), PE-anti-IL-4 (8D4-8), and PE-anti-IL-10 (JES3-19F1; BD PharMingen).
Phytohemagglutinin (PHA) and DC-driven CD4+ T cell proliferation
Immature and mature DC, exposed or not to imatinib, were plated at 104/well in 96-well U-bottom plates. Allogeneic CD4+ T cells from healthy donors were isolated after monocyte depletion of PBMC using CD14 microbeads (Miltenyi Biotec) and positive selection of CD4+ T cells with CD4 microbeads (Miltenyi Biotec), as indicated by the manufacturer. T cells were washed in serum-free medium and stained for 5 min with the PKH26 dye (Sigma Chemical Co.) at 2 x 10–6M final concentration, according to the manufacturer’s instructions. After three washes, T cells were added to the DC at 5 x 104/well. Proliferation was assessed on Day 7 by flow cytometry. The PKH26 dye is retained stably in the cell membrane of T cells and is equally divided in daughter cells upon each cell division, allowing the identification of T cells that had divided among cultured cells by flow cytometry. In some experiments, imatinib at 1 µM and 5 µM was added during DC-T cell coculture. To analyze CD4+ T cell response to PHA, PBMC from healthy donors or CML patients were stained with the PKH26 dye as above, plated at 105/well in 96-well U-bottom plates, and stimulated with 5 µg/ml PHA (Sigma Chemical Co.). Proliferation was assessed on Day 7 by flow cytometry.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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+/IL-4–) or Th2 effectors (IFN-–/IL-4+; Fig. 1D
). When cultured in GM-CSF and IL-4, purified monocytes from CML patients and healthy donors developed into homogeneous and typical immature DC as CD14– and CD1a+. The expression of CD1a did not differ statistically between monocyte-derived DC from CML patients (median MFI, 454; range, 59–2363) and from healthy donors (median MFI, 529; range, 339–2845). The median DC yield from monocytes was 33% (16.3–60) in CML patients and 34% (16–52.5) in healthy donors (not significant), which is comparable with what was reported in the literature with normal DC [21
]. In conclusion, bcr-abl+ and normal monocytes share a comparable DC differentiation potential, and the removal of CD14low granulocytic CML cells is a critical step in the generation of bcr-abl+ monocyte-derived DC.
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Figure 1. bcr-abl+ CD14high monocytes but not CD14low leukemic precursors differentiate along the DC pathway. CD14high monocytes (circle) and circulating CD14low/CD15+ granulocytic precursors (square) from CML patients were cultured for 6 days in the presence of GM-CSF and IL-4. Differentiation into DC was assessed by the measurement of CD1a and CD14 expression by flow cytometry. Specific mean fluorescence intensities (MFI) are indicated (A). The surface expression of HLA-DR, HLA-class I, CD86, and CD40 was also analyzed by flow cytometry. Specific MFI are indicated (B). Antigen capture was determined after incubation of the cells with medium containing 1 mg/ml FITC-BSA. Empty histograms show the background autofluorescence, shaded histograms show the background uptake at 0°C, and black-filled histograms show the specific uptake at 37°C. Specific MFI are indicated (C). Induction of naïve, allogeneic CD4+ T cell polarization toward Th1 (IFN-+/IL-4–) and Th2 (IFN-–/IL-4+) was determined by intracellular cytokine staining after PMA and calcium ionophore treatment of CD4+ T cells stimulated 6 days before with CD14high monocytes or CD14low/CD15+ granulocytic precursors cultured in the presence of GM-CSF and IL-4 (D). Dot-plots with double IFN--FITC and IL-4-PE staining are shown, and percentages of Th1 and Th2 CD4+ T cells are indicated. In each panel, results from one representative CML patient are shown among three independent experiments.
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Figure 2. Effect of imatinib on apoptosis and CD1a expression during normal and bcr-abl+ DC differentiation. Normal and bcr-abl+ monocytes were exposed to control DMSO and imatinib at 1 µM and 5 µM, and DC differentiation was initiated by addition of GM-CSF and IL-4. DC were analyzed after 6 days of culture. Apoptosis was assessed by annexin V-FITC/PI staining, and a representative experiment among four CML patients and four healthy donors is shown. The percentages of apoptotic cells (Annexin V-FITC+/PI–, lower right quadrants) are indicated. The 5-µM imatinib forward-scatter/side-scatter dot-plot is also shown (A). The expression of CD14 and CD1a was determined by flow cytometry, and a representative experiment among four CML patients and four healthy donors is shown. The percentage of CD1ahigh cells is indicated (B). CD1a and CD14 expression was also determined on DC differentiated for 6 days in the presence of GM-CSF and IL-4 and exposed to imatinib at 5 µM for an additional period of 48 h. A representative experiment among two CML patients and two healthy donors is shown (C). To analyze the modulation of bcr-abl during leukemic DC differentiation, bcr-abl transcripts were quantified by RQ-PCR in bcr-abl+ monocytes and monocyte-derived DC. Experiments performed in five individual CML patients are shown (D).
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Figure 3. Normal and bcr-abl+ monocyte-derived DC immunophenotype is not affected by imatinib. Immature and mature, normal and bcr-abl+ DC were compared for their cell-surface immunophenotype and for their capacity to internalize FITC-BSA by flow cytometry. Results from 20 CML patients and 10 healthy donors are shown (A). Normal and bcr-abl+ monocyte-derived DC were incubated in the presence of control DMSO or imatinib at 1 µM and 5 µM during DC development and maturation. The expression of MHC molecules, of costimulatory molecules, of the lymphoid chemokine receptor CCR7, and of the maturation marker CD83 was analyzed by flow cytometry. Results from a representative CML patient are shown among experiments performed in three CML patients and three healthy donors (B).
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+/IL-4–) and Th2 (IFN-–/IL-4+) polarization of naive CD4+ T cells (Fig. 4B)
. Upon maturation, both DC types were found to be potent inducers of Th1 polarization with Th1 cells, respectively, representing, on average, 50% (30–63) and 60% (39–83) of T cells when bcr-abl+ and normal mature DC were used as stimulators (Fig. 4B)
. It is important that bcr-abl+ and normal monocyte-derived DC were not found to prime significant numbers of IL-10-producing T cells (Fig. 4B)
. The presence of imatinib at 1 µM or 5 µM during DC development and maturation did not have a deleterious effect on the induction of Th1-type responses, whether bcr-abl+ or normal DC were used (Fig. 4C)
.
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Figure 4. Imatinib induces a dose-dependent decrease in IL-12p70 secretion by normal and bcr-abl+ monocyte-derived DC but does not impair CD4+ T cell polarization. Normal and bcr-abl+ monocyte-derived DC were incubated in the presence of control DMSO or imatinib at 1 µM and 5 µM during DC during differentiation. Maturation was induced by addition LPS for 6 h, 24 h, and 48 h, followed by restimulation for 24 h with CD40L. IL-12p70 production was measured in the culture supernatants by ELISA. Results from four CML patients and three healthy donors are shown (A). bcr-abl+ and normal monocyte-derived DC were compared for their capacity to polarize naive CD4+ T cells. Allogeneic CD4+/CD45RA+ T cells (5x104) were cultured for 7 days with immature and mature bcr-abl+ and normal monocyte-derived DC (1x104). T cells were restimulated for 6 h with calcium ionophore and PMA in the presence of brefeldin A, and expression of intracellular IFN-, IL-4, and IL-10 was determined by flow cytometry. The proportion of CD4+ T cells polarized toward Th1 (IFN-+/IL-4–), Th2 (IFN-–/IL-4+), and producing IL-10 from 11 CML patients and six healthy donors is shown. Pretreatment of immature and mature bcr-abl+ and normal monocyte-derived DC with imatinib at 1 µM and 5 µM did not affect polarization of CD4+ T cells toward Th1 (IFN-+/IL-4–). Results from three CML patients and three healthy donors are shown (C).
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Figure 5. Imatinib inhibits PHA- and DC-driven T cell proliferation. PBMC from healthy donors and from imatinib-treated CP-CML patients were stained with PKH26 and stimulated by the mitogenic lectin PHA without addition of exogenous imatinib. PKH26 is divided in daughter cells upon cell division and allows the identification of T cells, which had divided by flow cytometry. Proliferation was assessed on Day 7. Results from three healthy donors and four imatinib-treated CP-CML patients are shown. Typical PKH26 staining of T cells from a healthy donor and a CML patient are also shown, and the proportion of T cells, which have undergone proliferation, is indicated (A). Allogeneic CD4+ T cells were stained with PKH26 and cultured for 7 days with bcr-abl+ or normal, immature and mature monocyte-derived DC, which had been generated previously in the presence of control DMSO or imatinib at 1 µM and 5 µM. Proliferation of CD4+ T cells was measured at Day 7 by flow cytometry. Results performed with DC, three CML patients and two healthy donors, are shown (B, left panel). Allogeneic CD4+ T cells were stained with PKH26 and cultured for 7 days with bcr-abl+ or normal, immature and mature monocyte-derived DC in the presence of control DMSO or imatinib at 1 µM and 5 µM. Proliferation of CD4+ T cells was measured at Day 7 by flow cytometry. Results performed with DC, three CML patients and two healthy donors, are shown (B, middle panel) as well as a typical PKH26 staining of T cells after 7 days culture with bcr-abl+ immature DC in the presence of control DMSO or 5 µM imatinib (B, right panel).
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Figure 6. Imatinib does not impede DC-T cell reciprocal activation. Allogeneic CD4+ T cells were cultured with immature or mature bcr-abl+ or normal monocyte-derived DC in the presence of control DMSO or imatinib at 5 µM. Expression of the T cell activation marker CD25 was analyzed on CD4+ T cells by flow cytometry after 24 h. Results from three independent experiments are shown as well as a typical CD4/CD25 staining from one experiment (A). Expression of the costimulatory molecule CD86 was analyzed on DC by flow cytometry after 72 h. Results from three independent experiments are shown as well as a typical CD86 staining from one experiment (B).
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In this study, we demonstrate that bcr-abl+ and normal monocytes differentiating along the DC pathway share typical DC features with similar surface immunophenotype and functional properties. It is important that we show that depletion of CD15+/CD14low cells prior to isolation of leukemic monocytes is mandatory to obtain pure bcr-abl+ monocyte-derived DC for therapeutic purposes. We indeed found that circulating CD15+ leukemic granulocytic precursors expressed low levels of CD14 but failed to develop into DC when cultured with GM-CSF and IL-4, based on morphology, on the absence of CD1a and CD86 expression, on inefficient antigen internalization, and on the lack of capacity to polarize naïve T cells. In apparent contrast with our findings, Oehler et al. [29
] reported previously that granulocytic precursors from CML patients, patients with bacterial infections, and patients treated with G-CSF could differentiate into DC-like cells expressing CD1a and CD86. Our culture conditions for the generation of DC differ from those described by Oelher et al. [29] in one important aspect: We used GM-CSF and IL-4 rather than GM-CSF, IL-4, and tumor necrosis factor (TNF-), and the capacity of this latter cytokine to increase surface expression of CD1a and costimulatory molecules during DC development is well-known [30
]. Even with TNF-, only 40% of the granulocytic precursors up-regulated CD1a; thus, a large proportion of the cells remained unable to acquire critical features of DC in the Oehler et al. [29] study.
DC vaccines in CML patients may be a promising strategy after an initial phase of imatinib treatment to achieve an immune-mediated control of the residual disease. Whether normal and bcr-abl+ DC vaccines can combine effectively with imatinib is an important issue, as imatinib not only inhibits constitutively activated kinases such as bcr-abl but also affects kinases regulating key functions of immune cells such as mitogen-associated protein kinases (MAPK), Lck, and Abl/Arg. We found a limited impact of imatinib during normal and bcr-abl+ monocyte-derived DC differentiation. At most, 25% of DC did not acquire CD1a, and the expression of costimulatory and MHC molecules was preserved. It is not clear why imatinib affected bcr-abl+ and normal monocyte-derived DC development in a comparable manner, but it is interesting to speculate that the decrease in bcr-abl transcription, which we observed during DC generation, may be involved. Whether this modulation of bcr-abl transcripts lowers leukemic antigen presentation by bcr-abl+ monocyte-derived DC cannot be excluded. The differentiation of monocytes into DC under GM-CSF and IL-4 is accompanied by the activation of multiple transduction pathways including Raf/MAPK kinase/extracellular-signal regulated kinase, phosphatidylinositol-3 kinase (PI-3K)/AKT, and nuclear factor-
B, and PI-3K/AKT appears especially to play a key role [31
]. The fact that the PI-3K/AKT pathway is not a direct target of imatinib may explain the poor impact of this drug on normal and bcr-abl+ monocyte-derived DC development. Our results differ strongly from those recently obtained by Appel et al. [15
]. These authors indeed found a dramatic blockade in CD1a and costimulatory molecule up-regulation when normal, monocyte-derived DC were generated in the presence imatinib at the same concentrations as those used in our study. The reasons for these discrepancies remain unclear, although we purified monocytes through CD14-positive selection, and Appel et al. [15] obtained monocytes through plastic adherence of monocytes in serum-free medium. However, it should be noted that the level of CD1a expression in control cultures was much lower than that we obtained. We focused here on the comparison between bcr-abl+ and normal monocyte-derived DC. It is notable that we found that monocytes from imatinib-treated patients retained their capacity to differentiate into CD14–CD1ahigh DC.
Upon maturation, we showed that bcr-abl+ monocyte-derived DC, like their normal counterparts, lost their ability to capture antigens, increased costimulatory and MHC molecules at high levels, and acquired the maturation marker CD83 and the lymphoid chemokine receptor CCR7. Furthermore, the cognate maturation signal CD40L enhanced IL-12p70 production triggered by the inflammatory stimulus LPS, and the dynamic regulation of IL-12 secretion by bcr-abl+ monocyte-derived DC was comparable with that of normal DC. Although the amount of IL-12 produced by leukemic DC was reduced slightly, mature bcr-abl+ monocyte-derived DC exhibited potent immunostimulatory functions and promoted a strong polarization of naïve CD4+ T cells toward Th1. Therefore, our results are in line with those from Gabriele et al. [32 ], who found that bcr-abl+ monocyte-derived DC were fully competent to stimulate CD8+ T cell responses. When DC maturation was initiated in the presence of imatinib, immunophenotypic changes and T cell immunostimulatory capacity of bcr-abl+ and normal monocyte-derived DC were preserved, but IL-12p70 secretion was decreased in a dose-dependent manner, as also observed by Appel et al. [15] in normal monocyte-derived DC. Nevertheless, we found that the capacity of imatinib-treated DC to drive naive CD4+ T cell responses toward a Th1 profile was not affected in the experimental settings used in our study.
Imatinib was reported recently to inhibit PHA and anti-CD3-driven human T cell proliferation in vitro [25 26 27 ] by targeting TCR signal transduction through the Abl and Lck tyrosine kinases [27 , 33 ]. Our data suggest that imatinib may also be detrimental to T cell expansion in vivo, as the proliferation of CD4+ T cells from imatinib-treated CML patients upon PHA stimulation was decreased. Another striking finding in our study is that normal and bcr-abl+ monocyte-derived DC were unable to overcome the antiproliferative effect of imatinib in vitro. Indeed, exposure to imatinib of CD4+ T cells cocultured with DC dramatically hampered T cell proliferation induced by normal and bcr-abl+ monocyte-derived DC. It is interesting that imatinib did not inhibit DC-mediated T cell activation, as CD4+ T cells from imatinib-treated and untreated cultures demonstrated a similar increase in the T cell activation marker CD25. In turn, up-regulation of the DC costimulatory molecule CD86 upon interaction with responding CD4+ T cells was not suppressed by imatinib. Altogether, our results show that imatinib does not prevent cognate interactions between DC and T cells and suggest that T cells and not monocyte-derived DC are the main targets of imatinib-suppressive action. However, we cannot exclude imatinib-mediated modifications of the immunological DC-T cell synapse. It is indeed important to keep in mind that imatinib was shown recently to alter the cross-talk between DC and other immune effectors such as natural killer cells [34 ]. Moreover, the tyrosine kinase Abl, a target of imatinib, is critical for optimal neuronal and neuromuscular synapse formation [35 ], and neurological and immunological synapses share many similarities [35 36 37 ].
Further research is needed to understand how imatinib may affect immunity more generally and to determine how immunological control of CML in imatinib-treated patients can be achieved and how imatinib should be integrated in the treatment of residual disease in the post-transplantation setting. Our findings readily demonstrate progress toward the clinical applicability of DC vaccines in imatinib-treated patients. As a result of imatinib-induced suppression of T cell proliferation, imatinib-free therapeutic windows during monocyte-derived DC vaccination may be required to enable expansion of leukemia-specific T cell responses.
Received July 28, 2005; revised November 16, 2005; accepted December 8, 2005.
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plus cytarabine in newly diagnosed chronic myeloid leukemia N. Engl. J. Med. 349,1423-1432, but not the ABL-kinase inhibitor imatinib (STI571), induces expression of myeloblastin and a specific T-cell response in chronic myeloid leukemia Blood 101,259-264 induces dendritic cell differentiation of CML mononuclear cells in vitro and in vivo Leukemia 16,1484-1489[CrossRef][Medline]B and Akt signaling pathways Clin. Cancer Res. 11,1928-1940 J. Exp. Med. 179,1109-1118 promotes the rapid differentiation of monocytes from patients with chronic myeloid leukaemia into activated dendritic cells tuned to undergo maturation after LPS treatment Blood 103,980-987This article has been cited by other articles:
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