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Angiotensin-converting enzyme (CD143) is abundantly expressed by dendritic cells and discriminates human monocyte-derived dendritic cells from acute myeloid leukemia-derived dendritic cells
Experimental Hematology 31 (2003) 1301–1309
Angiotensin-converting enzyme (CD143) is abundantly expressed by dendritic cells and discriminates human monocyte-derived dendritic cells from acute myeloid leukemia-derived dendritic cells Sergei M. Danilova, Elena Sadovnikovab, Nicole Scharenborgc, Irina V. Balyasnikovaa, Daria A. Svinarevab, Elena L. Semikinad, Elena N. Parovichnikovab, Valery G. Savchenkob, and Gosse J. Ademac a Department of Anesthesiology, University of Illinois at Chicago, Chicago, Ill., USA; National Center for Hematology, Moscow, Russia; cLaboratory of Tumor Immunology, University Medical Center, Nijmegen, The Netherlands; dChild Health Center, Moscow, Russia b
(Received 21 January 2003; revised 30 July 2003; accepted 14 August 2003)
Objectives. The pattern of angiotensin-converting enzyme (ACE) expression in dendritic cells (DC) and macrophages derived from normal monocytes vs that in DC derived from acute myeloid leukemia blasts was investigated. Materials and Methods. ACE expression was quantified by flow cytometry using a set of monoclonal antibodies (mAbs) directed against five different epitopes on the ACE molecule and by enzyme activity measurement. Results. The binding pattern of a set of anti-ACE mAbs to the surface of blood cells and their progeny, as revealed by FACS, showed lineage and epitope specificity. Differentiation of monocytes to macrophages and DC was accompanied by a dramatic increase in ACE expression. ACE activity was 50-fold higher in macrophages and 150-fold higher in DC than in monocytes. ACE level normalized per cell revealed that DC expressed 1300-fold more ACE than did monocytes. In contrast, DC derived from acute myeloid leukemia blasts did not show an elevated level of ACE, although they acquired DC markers CD80, CD40, and CD86 upon cytokine or calcium ionophore treatment. Conclusions. ACE expression becomes the first marker to functionally distinguish DC generated from monocytes and leukemic blast cells. Given that ACE plays an important role in the hydrolysis of many peptides, as well as in the presentation of some antigens to immune cells, these data suggest that elevated ACE expression on the surface of DC is not just a reflection of the general activation of monocytes during differentiation; rather, it may be physiologically important for the functioning of these cells. 쑖 2003 International Society for Experimental Hematology. Published by Elsevier Inc.
Dendritic cells (DC) play a key role in the induction of the adaptive immune response because they are equipped for efficient antigen presentation and costimulation of naive lymphocytes. It has been demonstrated that cytokines or modulators of calcium metabolism (calcium ionophore) induce a cascade of maturation events in DC precursors leading to the expression of a diverse spectrum of differentiation markers characteristic for DC and required for their immune stimulatory function, including major histocompatibility
Offprint requests to: Sergei M. Danilov, M.D., Ph.D., Anesthesiology Research Center, University of Illinois at Chicago, 1819 W. Polk Street (M/ C 519), Chicago, IL 60612; E-mail: [email protected]
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complex (MHC) up-regulation, expression of costimulatory molecules, adhesion receptors, chemokines, chemokine receptors, and cytokines (for review, see [1,2]). We have demonstrated a high level of angiotensin-converting enzyme (ACE) expression in blood-derived DC [3]. ACE (kininase II), a zinc-metalloprotease with a broad substrate specificity, is generally known as an important regulator of blood pressure, fluid, and electrolyte homeostasis. High levels of ACE expression were demonstrated in endothelial cells, on the epithelium of brush borders of intestine, and the proximal tubules of the kidney [4–6]. Recently, ACE was assigned as new CD marker: CD143 [7]. ACE also has been found in tissue macrophages, especially those in human atherosclerotic plaques [8–10]. Differentiation of monocytes
쑖 2003 International Society for Experimental Hematology. Published by Elsevier Inc.
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into macrophages in culture is accompanied by an increase in ACE expression [11–14]. ACE also is expressed in T lymphocytes, albeit at much lower level [15]. Due to its carboxydipeptidase activity, ACE enhances the presentation of certain endogenously synthesized peptides to MHC class I-restricted cytotoxic T lymphocytes, by generation of optimally sized class I-binding peptides from larger protein fragments [16–20]. Because ACE may play a role in antigen presentation, its expression in DC is of particular interest for exploiting in vitro generated DC as natural adjuvant for anti-cancer vaccines [21]. In this respect, adequate antigen processing and presentation of exogenous antigens used for DC sensibilization by transfected DC or DC derived from leukemic blasts may depend on ACE expression by these cells. Leukemic DC are characterized by high levels of costimulatory molecule expression, cytokine production, and potent antigen-presenting capacity [22,23]. To date, little difference has been shown in the pattern of DC marker expression between monocyte-derived DC and DC derived from leukemic blasts. In the present study, a systematic quantitative analysis of ACE expression during monocytes/macrophage/DC differentiation was performed by flow cytometry using a representative panel of anti-ACE monoclonal antibodies (mAbs) [24], as well as by enzymatic analysis. Differentiation of monocytes into DC was demonstrated to be accompanied by a dramatic increase in ACE expression: more than 150fold (normalized to cell protein) or more than 1300-fold (normalized to cell number). The unusually high level of ACE expression in DC may indicate that this enzyme plays an important physiologic role in antigen-presenting cells. In contrast to their normal counterparts, DC derived from leukemic blasts consistently fail to up-regulate ACE in response to cytokines or the calcium ionophore A23187.
Materials and methods Blood cell isolation and cultivation Peripheral blood mononuclear cells (PBMC) were isolated from heparinized peripheral blood of healthy volunteer donors by Percoll gradient centrifugation. PBMC were allowed to adhere to culture flasks. To obtain immature DC, the adherent fraction (⬎95% of the cells are CD14⫹) was cultured for 3 to 7 days in RPMI-1640 medium with 5% heat-inactivated fetal calf serum (FCS), 800 U/mL granulocyte-macrophage colony-stimulating factor (GM-CSF), and 500 U/mL interleukin-4 (IL-4) [25]. Alternatively, adherent cells were cultured in medium with 5% FCS in the presence of 180 ng/ mL calcium ionophore A23187 (Sigma, St. Louis, MO, USA) for 4 days [26,27]. Mature DC were obtained by the activation of immature DC with lipopolysaccharide (LPS; 2 µg/mL) or tumor necrosis factor-α (TNF-α; 15 ng/mL) (both from R&D Systems, Minneapolis, MN, USA). For leukemia DC generation, peripheral blood cells obtained from leukemic patients were cultured in the presence of GM-CSF, IL-4, and TNF-α for 8 to 10 days or in the presence of calcium ionophore for 5 days. As a positive control for that experiment, we used DC generated from monocytes (⬎95%
CD14⫹) of healthy donors, which were cultivated under identical conditions. Macrophages were obtained from healthy donor PBMC by incubation of the adherent cell fraction with macrophage colonystimulating factor (M-CSF; 50 U/mL; R & D Systems) according to [28]. The nonadherent fraction of PBMC was activated for 3 days with phytohemagglutinin (PHA; 1 µg/mL) and IL-2 (20 U/ mL) or with GM-CSF/IL-4 as described earlier. Immunocytometry Phenotypic analysis, i.e., surface expression of the antigen markers, was performed by flow cytometry. Immunolabeling of cells tested in this study was performed using saturating concentrations of the primary mouse mAbs against different surface markers: anti–HLAA, B; DR; DQ; anti-CD3, 14, 40, 80, 83, 86, 115 (all Becton Dickinson, Mountain View, CA, USA), anti–ACE-CD143 (Chemicon International, Temecula, CA, USA). Fluorescein isothiocyanate-conjugated goat anti-mouse IgG F(ab′)2 (Zymed, San Francisco, CA, USA) was used as secondary antibody. The following fluorochrome-conjugated antibodies were used: anti-CD14 and CD80 (Becton Dickinson), anti-CD83 (Coulter), anti–ACE-CD143 (Serotec, Oxford, UK). Flow cytometric analysis was performed on a FACScan flow cytometer (Becton Dickinson). Propidium iodide was used to set life gates. ACE activity measurements Prior to harvesting, the cells were thoroughly washed to remove fetal bovine serum, which, even after heat inactivation, contained trace bovine ACE activity. Fluorometric assay of ACE activity in lysates of cultured cells was performed by measuring the release of His-Leu from the substrates Hip-His-Leu and Z-Phe-His-Leu [29]. ACE level was normalized to the number of cells and amount of total protein in cell lysates treated with detergent CHAPS (Sigma). Patient samples After obtaining informed consent, blood samples were collected from patients diagnosed with acute myeloid leukemia (AML), before the induction of chemotherapy. The investigation was approved by the Institutional Human Research Committee. Patients with a high blast cell count in peripheral blood were selected for this study. Patient’s characteristics are listed in Table 1. An additional blood sample was obtained from patient R during hematologic remission when his blood count became normal 4 weeks after completing consolidation chemotherapy. Table 1. Patient characteristics Patient code L R K I Ag A D Dn O Kr
Sex/age (y) F/27 M/45 F/62 F/42 M/37 F/37 M/63 F/45 F/43 M/31
Disease/ FAB type AML AML AML AML AML AML AML AML AML AML
M0 M5 M4 M2 M4 M4 M4 M4 M4 M4
WBC count (×103 µL)
Blasts in PB (%)
260 129 25 48 83 120 184 2 120 30
60 91 66 90 ⬎50 90 68 51 65 49
M0 through M5 indicate FAB (French-American-British) type. AML ⫽ acute myeloid leukemia; PB ⫽ peripheral blood; WBC ⫽ white blood cell.
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Results Optimal conditions for measuring ACE expression on DC by FACS analysis In our previous study, flow cytometric analysis of ACE expression on immature DC was performed using antiACE mAb 9B9 [3]. In order to perform a systematic quantitative study of ACE expression on different cells and to find the most sensitive mAb for ACE detection, in preliminary experiments we compared the saturation curves of the surface binding of five different anti-ACE mAbs [24] to the surface of immature DC (not shown). Based on these data, all of the following experiments were performed with an mAb concentration of 10 µg/mL. Of the antibodies tested, the combination of mAbs 9B9 and i2H5, which are directed against nonoverlapping ACE epitopes [24], was the most sensitive method of ACE detection (see Figs. 1–3).
Figure 2. Surface phenotype of the macrophages during differentiation. (A) FACS analysis of surface antigens. For ACE expression quantification, the mixture of mAb 9B9 and i2H5 was used (see also panel B). (B) Binding of anti-ACE mAbs to ACE expressed in macrophages. Macrophages, which were obtained by treatment with M-CSF alone from monocytes or with subsequent treatment with LPS, were stained using a panel of mAbs against ACE Other details as in Figure 1.
Figure 1. Surface phenotype of the DC during differentiation. (A) Flow cytometry of surface expression of DC-specific markers. For ACE expression quantification, the mixture of mAb 9B9 and i2H5 was used (see also panel B). (B) Binding of anti-ACE mAbs to ACE expressed in maturing DC. DC at several stages of differentiation (see Material and methods section) were stained with a panel of mouse mAbs. ACE detection on the cell surface is indicated as a mean of fluorescence (⫾SD). The value of fluorescence intensity of cells stained with isotype control antibodies was subtracted from the value of fluorescence intensity of cells in experimental samples.
Kinetics of ACE expression during DC maturation (FACS analysis) Monocytes from healthy donors stimulated for 3 days with GM-SF and IL-4 became loosely adherent, developed protrusions characteristic of DC morphology, and tended to make clusters. The cells were characterized by high levels of MHC class I, MHC class II, and CD86 expression, and were negative for CD3 and CD14 (Fig. 1A). The expression of DC specific antigens was up-regulated upon further culture (up to 6 days) and was further increased by treatment with LPS or TNF-α. ACE expression revealed by the combination of mAb 9B9 and i2H5 also was gradually increased during the process of DC differentiation. The general pattern of DC staining with mAbs recognizing different ACE epitopes was similar for all five antibodies used (Fig. 1B). The level of ACE expression in DC obtained by cytokine treatment (GM-CSF/IL-4. TNF-α) was comparable to that detected in A-23187 stimulated monocytes (data not shown).
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of mAb i1A8 with the surface of monocytes and macrophages was 4- to 5-fold higher than the binding of any other mAbs ( p ⬍ 0.05). The monocytic cell lines THP-1 and U937 demonstrated the same pattern of binding (data not shown). The ratio of mAbs i1A8/9B9 binding was in the range from 5 to 7 for monocytes, macrophages, THP-1, and U937, but was in the range from 0.6 to 0.9 for other tested cells. Therefore, we conclude the best mAb for ACE detection on cells of monocyte/macrophage lineage is i1A8. In lymphocytes, binding with mAb i2H5 was greater than with the other tested mAbs ( p ⬍ 0.05; Fig. 3). The same epitope-dependent pattern of binding was characteristic for DC (Figs. 1 and 3). Effect of ACE inhibition on the human DC phenotype To study whether ACE enzymatic activity affects differentiation of DC, we incubated monocytes during their differentiation to DC (in the presence of GM-CSF/IL-4) with the ACE inhibitor lisinopril (100 µM). At this concentration, ACE activity in monocytes as well as in DC was inhibited by more than 95%. Lisinopril was added at 0, 3, or 6 days of cultivation. Fig. 4 shows a representative histogram of the effect of lisinopril on the distribution of surface marker in immature DC. In two independent donors, the differentiation of monocytes to immature DC was not affected by lisinopril. However, in 2 of 3 experiments with cells from donor B and 3 of 3 from donor C, we observed an increase in CD83 expression (Fig. 4). In 3 of 6 experiments, the expression of CD80 slightly increased.
Figure 3. Cell-surface ACE expression in different types of hematopoietic cells. Immunofluorescence analysis of cultured PBMC (gated for monocytes and lymphocytes separately), PBL, and immature DC was performed on the cells isolated from seven donors. Analysis of anti-ACE binding with macrophages was performed on the cells isolated from two donors. Cellsurface ACE expression is indicated as mean fluorescence intensity (⫾ SD). (Mean fluorescence intensity of cells stained with isotype control antibodies was subtracted from each value).
Macrophages generated by culturing monocytes from peripheral blood of healthy donors in the presence of M-CSF were characterized by significant levels of CD14 and CD115 expression (Fig. 2A). Analysis of ACE expression on monocytes and lymphocytes was performed by gating on relative populations in freshly isolated peripheral blood cells from healthy donors. The level of ACE expression on monocytes as revealed by the mixture of mAbs 9B9 and i2H5 was significantly lower than in cultured macrophages or DC, but was higher compared to ACE expression on lymphocytes ( p ⬍ 0.05 in both cases; Fig. 3). Epitope specificity of ACE detection in different cell types Fig. 3 shows the binding pattern of the tested set of anti-ACE mAbs with different cells used in this study. The binding
ACE expression in leukemic DC It has been demonstrated previously that AML blasts acquire certain characteristics of DC upon stimulation with cytokines and modulators of calcium metabolism. Significant up-regulation of CD40, CD80, and CD86 expression is observed in DC derived from AML blasts [22,23,27]. In order to investigate whether the coordinated up-regulation of ACE expression is observed in leukemic cells during their DCtype differentiation, we performed both FACS analysis and ACE activity tests on cultured AML blasts. Leukemic DC derived from 10 AML patients displayed the phenotypic characteristics of DC in these cells (data not shown). As detected by staining with a mixture of 9B9 and i2H5, antiACE mAb cells from eight patients were ACE negative (data not shown). Only a low level of ACE expression was observed in leukemic DC derived from patients L and R (Fig. 5A and D). Because a remission sample was available from patient R, we compared ACE expression in DC derived from AML blasts and monocytes from the same patient. Leukemic DC derived from blast cells of patient R (Fig. 5B and E) had lower levels of ACE that observed in untreated peripheral blood monocytes from healthy donors. In contrast, DC cells derived from peripheral blood monocytes from the same patient harvested during remission produced high levels of ACE (Fig. 5C and F), which were comparable
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observed in cells cultured in the presence of cytokines (GMCSF, IL-4, TNF-α) and calcium ionophore A23187. Double staining of leukemic DC from patients R and L revealed ACE expression only in a small proportion of CD80⫹ cells (Fig. 5D and E). When DC were obtained from patient R monocytes harvested during remission, the CD80 fraction of these cells stained brightly with anti-ACE antibodies (Fig. 5F).
Figure 4. Effect of the ACE inhibitor lisinopril on DC-type monocyte differentiation. Immature DC cells were obtained from monocytes by culturing with cytokines for 6 days (see Material and methods section) in the absence (open histogram) or presence (filled histogram) of lisinopril 100 µM and stained with specific mAbs. Results of a representative experiment are shown.
with the ACE levels observed in DC obtained from monocytes of healthy donors (Fig. 1). In quantitative terms, the median fluorescence intensity of DC from blasts with anti-ACE mAbs was 3.6 ⫾ 5.3 (n ⫽ 10 patients), whereas that value for DC from monocytes was 192 ⫾ 92 (n ⫽ 5). The difference (54-fold) was statistically significant ( p ⫽ 0.010). The level of ACE expression in leukemic DC (as in DC from monocytes) was not dependent on the culture conditions used for DC generation: a low level of ACE was
ACE activity of monocytes, macrophages, and DC derived from healthy donor monocytes and DC from AML blasts To confirm the data obtained by FACS analysis, we measured the level of total ACE activity (intracellular plus cell surface ACE) in the lysates from these cells (Table 2). Specificity of ACE activity assay was demonstrated by the inhibition of measurable ACE activity using either ACE inhibitors or a set of blocking mAbs to ACE (data not shown). Generally, the results of ACE activity measurements confirmed that of flow cytometric analysis. The lowest (but still measurable) level of ACE activity was demonstrated for nonstimulated PBMC containing a mixture of resting monocytes and lymphocytes), and in an isolated population of lymphocytes stimulated with a IL-4/GM-CSF or IL-2/PHA. M-CSF–induced macrophage-type differentiation of monocytes was accompanied by a significant (55-fold) increase in ACE activity. Stimulation of monocytes with IL-4/GM-CSF, which leads to DC-type differentiation, induced an even greater level of ACE activity (150-fold). The effect of GMCSF and IL-4 on induction of ACE expression appears specific for monocytes because the combination of IL-4/ GM-CSF only slightly increases ACE activity in purified lymphocytes. DC derived from AML blasts showed negligible ACE activity. In addition, activation of lymphocytes with IL-2 and PHA did not significantly increase ACE activity. ACE activity in mature DC stimulated with LPS or TNF-α slightly decreased in comparison with immature DC. It is important to note that the level of ACE expression in DC (expressed as enzymatic activity per milligram of protein in cell lysate) is comparable with the highest level of somatic ACE expression in cultured human cells (human endothelial cells from umbilical vein): 5 to 15 mU/mg [30]. The highest ACE activity recorded in a human cell under the natural ACE promoter was found in spermatozoa: 650 mU/mg (Balyasnikova and Danilov, manuscript in preparation). However, these cells express the testicular form of ACE that are driven by “testicular” ACE promoter localized in twelfth intron of the ACE gene [31,32]. ACE activity in CHO cells transfected with the ACE cDNA driven by the CMV promoter ranged (depending on particular clone tested) from 200 to 1500 mU/mg [33]. However, to normalize ACE level in different cells per cell number, the relative value of ACE expression in DC will be changed dramatically. We found that the yield of protein in supernatant from lysate of DC was 270 µg/106 cells, CHO-ACE 125 µg/106 cells, PBMC/peripheral blood lymphocyte (PBL) 22 µg/106 cells,
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Figure 5. ACE and CD80 expression in dendritic cells derived from blasts of AML patients. Cells from AML M2 patient L were obtained at diagnosis (A, D). Cells from AML M5 patient R were harvested at diagnosis (B, E) and in remission (C, F). (A–C) Cells were stained with anti-ACE antibodies (mixture 9B9⫹i2H5; filled histograms) or with control anti-mouse IgG (clear histograms) Figures correspond to values of median fluorescence intensity. (D–F) Double staining with anti-ACE antibodies: mixture 9B9⫹i2H5 and anti-CD80 antibodies.
and spermatozoa 2.1 µg/106 cells. Therefore, ACE activity in DC expressed per 106 cells (2.5 mU/106 cells) is 1300 times higher than in PBMC or lymphocytes and twice as high as the expression of testicular form of ACE in spermatozoa (1.4 mU/106 cells). Thus, we can consider that DC expressed the highest level of ACE among human cells tested.
Discussion We previously demonstrated that a high level of ACE expression could be induced in immature DC [3]. Here we present a systematic quantitative study of ACE expression during the different stages of DC differentiation and maturation. We also show that AML blasts fail to up-regulate ACE during their DC-type differentiation induced by cytokine or calcium ionophore treatment. Somatic ACE has two highly homologous domains (Nand C-domains), each bearing a functional catalytic site [34]. ACE exists in both membrane bound and soluble forms. It is constitutively expressed on the surface of endothelial cells, on the epithelium of intestinal brush borders, on the epithelial cells of the proximal tubule of the kidney, on the tubular epithelial cells of the epididymis, and on alveolar macrophages. The interest of hematologists in ACE has dramatically increased with the discovery that N-acetylseryl-aspartyl-lysyl-proline (Ac-SDKP), a regulatory factor in hematopoiesis, is a natural substrate for the N-terminal active center of ACE [35,36]. Ac-SDKP reversibly prevents the recruitment of pluripotent hematopoietic stem cells and
normal early progenitors into the S-phase of the cell cycle by maintaining them in the G0 phase [37,38]. The differentiation of monocytes into macrophages was accompanied by a dramatic (60-fold) increase in ACE expression [11]. ACE expression also can be increased upon induction with T lymphocytes [13,39], with dexamethasone [11], or with GM-CSF [14]. ACE overexpression also was demonstrated in giant and epithelioid cells in granulomas during sarcoidosis, Gaucher disease, and leprosy resulting in increased ACE blood level in these patients [40–42]. ACE participates in antigen presentation (of certain peptides, at least) by altering antigen processing. The efficient presentation of 147–158/R-peptide of influenza virus nucleoprotein results from removal of the COOH-terminal dipeptide by ACE [16]. T-cell stimulation by the HIV-1 gp160-derived peptide p18 presented by H-2D class I MHC molecules also was found to require proteolytic cleavage of the peptide by ACE [17]. ACE acts to enhance antigen presentation exclusively in an exocytic intracellular compartment, and this pathway is TAP (transporter associated with antigen presentation) dependent [18,19]. The present study demonstrated that DC-type differentiation of monocytes is accompanied by a dramatic 150-fold increase in ACE activity normalized to cellular protein, which is equivalent to a 1300-fold increase in ACE activity per cell. This fact was confirmed by two independent assays: flow cytometric analysis with a set of mAbs to ACE (Figs. 1 and 3) and measurement of enzymatic activity (Table 2). The absolute values of ACE activity in human monocytes
S.M. Danilov et al. / Experimental Hematology 31 (2003) 1301–1309 Table 2. Angiotensin-converting enzyme activity in different cell types derived from human blood Cells PBMC Immature DC 3 days in culture Immature DC 6 days in culture Mature DC (LPS treatment) 8 days in culture Mature DC (TNF-α treatment) 8 days in culture Leukemic DC (CI or TNF-α treatment) 5 or 8 days in culture Macrophages 6 days in culture Macrophages (⫹LPS) 6 days in culture PBL (⫹IL-2/PHA) 3 days PBL (⫹IL-4/GM-CSF) 3 days
ACE activity (mU/mg) 0.061 ⫾ 0.02 5.3 ⫾ 2.3 9.34 ⫾ 2.0 9.07 ⫾ 2.5 9.2 ⫾ 0.6 0.090 ⫾ 0.040 3.4 ⫾ 0.8 2.4 ⫾ 1.1 0.086 ⫾ 0.036 0.09 ⫾ 0.05
Angiotensin-converting enzyme (ACE) activity was measured in the detergent lysates of the indicted cells fluorimetrically using Hip-His-Leu as substrate. ACE activity was normalized per milligram of protein in cell lysate and expressed as mean ⫾ SD (n ⫽ 4). CI ⫽ calcium ionofore A23187; DC ⫽ dendritic cells; GM-CSF ⫽ granulocyte-macrophage colony-stimulating factor; IL-2 ⫽ interleukin-2; IL4 ⫽ interleukin-4; LPS ⫽ lipopolysacharide; PBL = peripheral blood lymphocytes; PBMC = peripheral blood mononuclear cells; PHA ⫽ phytohemagglutinin; TNF-α ⫽ tumor necrosis factor-α.
and macrophages presented in this study (Table 2) were practically identical to the data published elsewhere [11,15], thus validating our measurements and new data on the absolute value of ACE activity in DC. Generally, the ACE activity increase during differentiation of monocytes into macrophages and DC corresponds to level of ACE expression revealed by flow cytometry with anti-ACE antibodies (see Table 2: ACE activity; Figs. 1–3: FACS scan). However, we found that the binding pattern of a set of mAbs to ACE differs in different cells. These results (Fig. 3) clearly show the epitope specificity of ACE revealed in different cell types. Binding of mAb i1A8 to the surface of monocytes, macrophages, and some other related cells (THP-1 and U937) was 4- to 5-fold higher than the binding of any other mAb and thus might be considered to be more specific for monocyte/macrophage ACE. However, for quantification of ACE expression on the cell surface by FACS scan, perhaps more correct would be use of the mixture of two mAbs: 9B9 and i2H5, or i1A8 and i2H5. These pairs of mAbs bind to different nonoverlapping ACE epitopes [23], thus allowing more complete ACE detection. Moreover, these combinations—at least 9B9 and i2H5 (Figs. 1– 3)—are less influenced by culture conditions. A high level of ACE expression in DC may have practical significance for gene therapy and vaccination via DC. Retargeting of an adenoviral vector to a marker expressed on DC (CD40) dramatically enhanced gene transfer to monocytederived DC [43]. Retargeting of an adenoviral vector to
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ACE-expressing cells and efficient gene transfer was demonstrated in vitro and in vivo using anti-ACE antibody 9B9 [44]. Therefore, based on the surface density of ACE in DC and on the effectiveness of gene transfer with anti-ACE mAb, we expect that gene transfer to DC via ACE would significantly improve the efficacy of DC-based vaccination. Results of this work, as well as previous studies cited earlier, demonstrate the low level of ACE activity in mononuclear blood cells. The ACE activity in THP-1 (human monocytic leukemia cell line) and the macrophage-like diffuse histiocytic lymphoma U937 also were very low (albeit measurable) and comparable to the ACE activity in monocytes, PBMC, and PBL (Table 2). These cell lines are used widely as a model system for the study of monocyte/macrophage differentiation. The addition of a stimulus such as phorbol myristate acetate (PMA) results in phenotypic changes characteristic for macrophages. However, the amplitude of PMA-induced ACE induction in these cells (3-fold for THP-1 [45]) is incomparable with the ACE 55-fold induction during monocyte maturation (Table 2). This suggests that leukemic cell lines display abnormal regulation of ACE expression during their macrophage-type differentiation similar to that of AML blasts during their DC-type differentiation. We showed that DC expressed the highest level of ACE per cell among the human cells tested. The biologic function of ACE in DC, as well as in macrophages, remains largely unknown. ACE inhibition in maturing DC caused a significant increase in CD83 expression and slight increase in CD80 expression by DC (Fig. 4). CD83 is a marker for DC maturation [46]. Up-regulation of CD83 expression upon ACE inhibition can be explained either by the fact that ACE is involved in maintaining the immature DC phenotype and blocking ACE activity promotes DC maturation, or by a direct action of lisinopril on CD83 production. Recently, the effect of ACE inhibitor (captopril) on the phenotype of DC was studied [47]. The authors did not find any changes of commonly used DC markers. They incubated mature DC with captopril for 24 hours, in contrast to the 5 days in our experiments. The authors demonstrated, however, that their short treatment of DC with captopril significantly decreased the levels of secreted TNF-α, IL-1α, IL-10, IL-12, and IL-18. The choice of captopril as the ACE inhibitor for their study may have led to misleading results, because captopril has an SH-group, which in addition to providing ACE inhibition also demonstrates significant anti-oxidant effects. Therefore, part of the effects of captopril on cytokine secretion by DC might be ACE independent. It also was shown in this study that preincubation of DC with angiotensin II increased production of TNF-α and IL-6, but not other cytokines. Therefore, we consider the secretion of at least TNF-α and IL-6 by DC to be ACE dependent. ACE in mononuclear cells may hydrolyze regulatory peptides at the site of inflammation, e.g., ACE hydrolyzes bradykinin, substance P, and other tachykinins that are implicated
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in inflammatory and immunologic responses [48]. Participation of ACE in antigen presentation and the fact that the negative regulator of stem cell proliferation (Ac-SDKP) is a natural substrate for ACE lead us to speculate that the high ACE expression on the surface of DC is not just a reflection of the general activation of monocytes during differentiation, but might have physiologic significance for effective functioning of these cells. The conditions required for DC generation in vitro were shown to be sufficient for the induction of DC differentiation markers in AML blast cells arrested at the early stages of hematopoiesis [22,26]. Leukemic DC express immune costimulatory molecules, IL-12, and are fully competent as antigen-presenting cells in allogeneic and autologous antigen-specific lymphocyte stimulation [49,50]. The results obtained indicate that the transformation process affects the regulation of ACE gene expression in AML blasts. Interestingly, a lack of ACE up-regulation was noted for cells derived from patients with various AML subtypes, characterized by different genetic abnormalities. Surprisingly, none of the publications describing profiling changes in gene expression during differentiation of monocyte-derived DC mentioned a statistically significant (more than 2-fold) increase in ACE mRNA expression, albeit the probes for ACE mRNA were present on microchips used for these studies [51–55]. Moreover, in a separate study, where the changes in ACE mRNA expression during DC differentiation was specifically studied using RT-PCR analysis, the authors also did not find significant up-regulation of ACE mRNA [56]. These results allow us to suggest that ACE up-regulation during differentiation from monocytes to DC (as well as failure of up-regulation in the case of leukemiaDC) might be due to posttranscriptional processing, like redistribution of inactive intracellular ACE to the surface where ACE become catalytically active (and becomes accessible for mAbs). The presence of an intracellular pool of immature inactive ACE was demonstrated previously [30]. The exact mechanism of the regulation of ACE expression in DC during differentiation, as well as the link between the regulation of ACE expression and leukemogenesis, remains to be investigated. It has been proposed that leukemic DC may prove useful for anti-cancer vaccine design because these cells express tumor-specific antigens endogenously, being progeny of a leukemic clone. The observed deficiency of leukemic DC to up-regulate ACE may interfere with the ability of leukemic DC to provide adequate processing and presentation of tumor antigens and should be considered during the development of strategies for anti-cancer therapy.
Acknowledgments The authors thank Professor Ronald F. Albrecht, Head of the Department of Anesthesiology, University of Illinois at Chicago, for his continuous encouragement and support of this project. The
authors also thank Dr. Richard D. Minshall, University of Illinois at Chicago, for critical reading of this manuscript.
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Angiotensin-converting enzyme (CD143) is abundantly expressed by dendritic cells and discriminates human monocyte-derived dendritic cells from acute myeloid leukemia-derived dendritic cells Sergei M. Danilova, Elena Sadovnikovab, Nicole Scharenborgc, Irina V. Balyasnikovaa, Daria A. Svinarevab, Elena L. Semikinad, Elena N. Parovichnikovab, Valery G. Savchenkob, and Gosse J. Ademac a Department of Anesthesiology, University of Illinois at Chicago, Chicago, Ill., USA; National Center for Hematology, Moscow, Russia; cLaboratory of Tumor Immunology, University Medical Center, Nijmegen, The Netherlands; dChild Health Center, Moscow, Russia b
(Received 21 January 2003; revised 30 July 2003; accepted 14 August 2003)
Objectives. The pattern of angiotensin-converting enzyme (ACE) expression in dendritic cells (DC) and macrophages derived from normal monocytes vs that in DC derived from acute myeloid leukemia blasts was investigated. Materials and Methods. ACE expression was quantified by flow cytometry using a set of monoclonal antibodies (mAbs) directed against five different epitopes on the ACE molecule and by enzyme activity measurement. Results. The binding pattern of a set of anti-ACE mAbs to the surface of blood cells and their progeny, as revealed by FACS, showed lineage and epitope specificity. Differentiation of monocytes to macrophages and DC was accompanied by a dramatic increase in ACE expression. ACE activity was 50-fold higher in macrophages and 150-fold higher in DC than in monocytes. ACE level normalized per cell revealed that DC expressed 1300-fold more ACE than did monocytes. In contrast, DC derived from acute myeloid leukemia blasts did not show an elevated level of ACE, although they acquired DC markers CD80, CD40, and CD86 upon cytokine or calcium ionophore treatment. Conclusions. ACE expression becomes the first marker to functionally distinguish DC generated from monocytes and leukemic blast cells. Given that ACE plays an important role in the hydrolysis of many peptides, as well as in the presentation of some antigens to immune cells, these data suggest that elevated ACE expression on the surface of DC is not just a reflection of the general activation of monocytes during differentiation; rather, it may be physiologically important for the functioning of these cells. 쑖 2003 International Society for Experimental Hematology. Published by Elsevier Inc.
Dendritic cells (DC) play a key role in the induction of the adaptive immune response because they are equipped for efficient antigen presentation and costimulation of naive lymphocytes. It has been demonstrated that cytokines or modulators of calcium metabolism (calcium ionophore) induce a cascade of maturation events in DC precursors leading to the expression of a diverse spectrum of differentiation markers characteristic for DC and required for their immune stimulatory function, including major histocompatibility
Offprint requests to: Sergei M. Danilov, M.D., Ph.D., Anesthesiology Research Center, University of Illinois at Chicago, 1819 W. Polk Street (M/ C 519), Chicago, IL 60612; E-mail: [email protected]
0301-472X/03 $–see front matter. Copyright doi: 1 0. 10 1 6 / j .e x p he m.2 0 03 .0 8 .0 1 8
complex (MHC) up-regulation, expression of costimulatory molecules, adhesion receptors, chemokines, chemokine receptors, and cytokines (for review, see [1,2]). We have demonstrated a high level of angiotensin-converting enzyme (ACE) expression in blood-derived DC [3]. ACE (kininase II), a zinc-metalloprotease with a broad substrate specificity, is generally known as an important regulator of blood pressure, fluid, and electrolyte homeostasis. High levels of ACE expression were demonstrated in endothelial cells, on the epithelium of brush borders of intestine, and the proximal tubules of the kidney [4–6]. Recently, ACE was assigned as new CD marker: CD143 [7]. ACE also has been found in tissue macrophages, especially those in human atherosclerotic plaques [8–10]. Differentiation of monocytes
쑖 2003 International Society for Experimental Hematology. Published by Elsevier Inc.
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into macrophages in culture is accompanied by an increase in ACE expression [11–14]. ACE also is expressed in T lymphocytes, albeit at much lower level [15]. Due to its carboxydipeptidase activity, ACE enhances the presentation of certain endogenously synthesized peptides to MHC class I-restricted cytotoxic T lymphocytes, by generation of optimally sized class I-binding peptides from larger protein fragments [16–20]. Because ACE may play a role in antigen presentation, its expression in DC is of particular interest for exploiting in vitro generated DC as natural adjuvant for anti-cancer vaccines [21]. In this respect, adequate antigen processing and presentation of exogenous antigens used for DC sensibilization by transfected DC or DC derived from leukemic blasts may depend on ACE expression by these cells. Leukemic DC are characterized by high levels of costimulatory molecule expression, cytokine production, and potent antigen-presenting capacity [22,23]. To date, little difference has been shown in the pattern of DC marker expression between monocyte-derived DC and DC derived from leukemic blasts. In the present study, a systematic quantitative analysis of ACE expression during monocytes/macrophage/DC differentiation was performed by flow cytometry using a representative panel of anti-ACE monoclonal antibodies (mAbs) [24], as well as by enzymatic analysis. Differentiation of monocytes into DC was demonstrated to be accompanied by a dramatic increase in ACE expression: more than 150fold (normalized to cell protein) or more than 1300-fold (normalized to cell number). The unusually high level of ACE expression in DC may indicate that this enzyme plays an important physiologic role in antigen-presenting cells. In contrast to their normal counterparts, DC derived from leukemic blasts consistently fail to up-regulate ACE in response to cytokines or the calcium ionophore A23187.
Materials and methods Blood cell isolation and cultivation Peripheral blood mononuclear cells (PBMC) were isolated from heparinized peripheral blood of healthy volunteer donors by Percoll gradient centrifugation. PBMC were allowed to adhere to culture flasks. To obtain immature DC, the adherent fraction (⬎95% of the cells are CD14⫹) was cultured for 3 to 7 days in RPMI-1640 medium with 5% heat-inactivated fetal calf serum (FCS), 800 U/mL granulocyte-macrophage colony-stimulating factor (GM-CSF), and 500 U/mL interleukin-4 (IL-4) [25]. Alternatively, adherent cells were cultured in medium with 5% FCS in the presence of 180 ng/ mL calcium ionophore A23187 (Sigma, St. Louis, MO, USA) for 4 days [26,27]. Mature DC were obtained by the activation of immature DC with lipopolysaccharide (LPS; 2 µg/mL) or tumor necrosis factor-α (TNF-α; 15 ng/mL) (both from R&D Systems, Minneapolis, MN, USA). For leukemia DC generation, peripheral blood cells obtained from leukemic patients were cultured in the presence of GM-CSF, IL-4, and TNF-α for 8 to 10 days or in the presence of calcium ionophore for 5 days. As a positive control for that experiment, we used DC generated from monocytes (⬎95%
CD14⫹) of healthy donors, which were cultivated under identical conditions. Macrophages were obtained from healthy donor PBMC by incubation of the adherent cell fraction with macrophage colonystimulating factor (M-CSF; 50 U/mL; R & D Systems) according to [28]. The nonadherent fraction of PBMC was activated for 3 days with phytohemagglutinin (PHA; 1 µg/mL) and IL-2 (20 U/ mL) or with GM-CSF/IL-4 as described earlier. Immunocytometry Phenotypic analysis, i.e., surface expression of the antigen markers, was performed by flow cytometry. Immunolabeling of cells tested in this study was performed using saturating concentrations of the primary mouse mAbs against different surface markers: anti–HLAA, B; DR; DQ; anti-CD3, 14, 40, 80, 83, 86, 115 (all Becton Dickinson, Mountain View, CA, USA), anti–ACE-CD143 (Chemicon International, Temecula, CA, USA). Fluorescein isothiocyanate-conjugated goat anti-mouse IgG F(ab′)2 (Zymed, San Francisco, CA, USA) was used as secondary antibody. The following fluorochrome-conjugated antibodies were used: anti-CD14 and CD80 (Becton Dickinson), anti-CD83 (Coulter), anti–ACE-CD143 (Serotec, Oxford, UK). Flow cytometric analysis was performed on a FACScan flow cytometer (Becton Dickinson). Propidium iodide was used to set life gates. ACE activity measurements Prior to harvesting, the cells were thoroughly washed to remove fetal bovine serum, which, even after heat inactivation, contained trace bovine ACE activity. Fluorometric assay of ACE activity in lysates of cultured cells was performed by measuring the release of His-Leu from the substrates Hip-His-Leu and Z-Phe-His-Leu [29]. ACE level was normalized to the number of cells and amount of total protein in cell lysates treated with detergent CHAPS (Sigma). Patient samples After obtaining informed consent, blood samples were collected from patients diagnosed with acute myeloid leukemia (AML), before the induction of chemotherapy. The investigation was approved by the Institutional Human Research Committee. Patients with a high blast cell count in peripheral blood were selected for this study. Patient’s characteristics are listed in Table 1. An additional blood sample was obtained from patient R during hematologic remission when his blood count became normal 4 weeks after completing consolidation chemotherapy. Table 1. Patient characteristics Patient code L R K I Ag A D Dn O Kr
Sex/age (y) F/27 M/45 F/62 F/42 M/37 F/37 M/63 F/45 F/43 M/31
Disease/ FAB type AML AML AML AML AML AML AML AML AML AML
M0 M5 M4 M2 M4 M4 M4 M4 M4 M4
WBC count (×103 µL)
Blasts in PB (%)
260 129 25 48 83 120 184 2 120 30
60 91 66 90 ⬎50 90 68 51 65 49
M0 through M5 indicate FAB (French-American-British) type. AML ⫽ acute myeloid leukemia; PB ⫽ peripheral blood; WBC ⫽ white blood cell.
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Results Optimal conditions for measuring ACE expression on DC by FACS analysis In our previous study, flow cytometric analysis of ACE expression on immature DC was performed using antiACE mAb 9B9 [3]. In order to perform a systematic quantitative study of ACE expression on different cells and to find the most sensitive mAb for ACE detection, in preliminary experiments we compared the saturation curves of the surface binding of five different anti-ACE mAbs [24] to the surface of immature DC (not shown). Based on these data, all of the following experiments were performed with an mAb concentration of 10 µg/mL. Of the antibodies tested, the combination of mAbs 9B9 and i2H5, which are directed against nonoverlapping ACE epitopes [24], was the most sensitive method of ACE detection (see Figs. 1–3).
Figure 2. Surface phenotype of the macrophages during differentiation. (A) FACS analysis of surface antigens. For ACE expression quantification, the mixture of mAb 9B9 and i2H5 was used (see also panel B). (B) Binding of anti-ACE mAbs to ACE expressed in macrophages. Macrophages, which were obtained by treatment with M-CSF alone from monocytes or with subsequent treatment with LPS, were stained using a panel of mAbs against ACE Other details as in Figure 1.
Figure 1. Surface phenotype of the DC during differentiation. (A) Flow cytometry of surface expression of DC-specific markers. For ACE expression quantification, the mixture of mAb 9B9 and i2H5 was used (see also panel B). (B) Binding of anti-ACE mAbs to ACE expressed in maturing DC. DC at several stages of differentiation (see Material and methods section) were stained with a panel of mouse mAbs. ACE detection on the cell surface is indicated as a mean of fluorescence (⫾SD). The value of fluorescence intensity of cells stained with isotype control antibodies was subtracted from the value of fluorescence intensity of cells in experimental samples.
Kinetics of ACE expression during DC maturation (FACS analysis) Monocytes from healthy donors stimulated for 3 days with GM-SF and IL-4 became loosely adherent, developed protrusions characteristic of DC morphology, and tended to make clusters. The cells were characterized by high levels of MHC class I, MHC class II, and CD86 expression, and were negative for CD3 and CD14 (Fig. 1A). The expression of DC specific antigens was up-regulated upon further culture (up to 6 days) and was further increased by treatment with LPS or TNF-α. ACE expression revealed by the combination of mAb 9B9 and i2H5 also was gradually increased during the process of DC differentiation. The general pattern of DC staining with mAbs recognizing different ACE epitopes was similar for all five antibodies used (Fig. 1B). The level of ACE expression in DC obtained by cytokine treatment (GM-CSF/IL-4. TNF-α) was comparable to that detected in A-23187 stimulated monocytes (data not shown).
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of mAb i1A8 with the surface of monocytes and macrophages was 4- to 5-fold higher than the binding of any other mAbs ( p ⬍ 0.05). The monocytic cell lines THP-1 and U937 demonstrated the same pattern of binding (data not shown). The ratio of mAbs i1A8/9B9 binding was in the range from 5 to 7 for monocytes, macrophages, THP-1, and U937, but was in the range from 0.6 to 0.9 for other tested cells. Therefore, we conclude the best mAb for ACE detection on cells of monocyte/macrophage lineage is i1A8. In lymphocytes, binding with mAb i2H5 was greater than with the other tested mAbs ( p ⬍ 0.05; Fig. 3). The same epitope-dependent pattern of binding was characteristic for DC (Figs. 1 and 3). Effect of ACE inhibition on the human DC phenotype To study whether ACE enzymatic activity affects differentiation of DC, we incubated monocytes during their differentiation to DC (in the presence of GM-CSF/IL-4) with the ACE inhibitor lisinopril (100 µM). At this concentration, ACE activity in monocytes as well as in DC was inhibited by more than 95%. Lisinopril was added at 0, 3, or 6 days of cultivation. Fig. 4 shows a representative histogram of the effect of lisinopril on the distribution of surface marker in immature DC. In two independent donors, the differentiation of monocytes to immature DC was not affected by lisinopril. However, in 2 of 3 experiments with cells from donor B and 3 of 3 from donor C, we observed an increase in CD83 expression (Fig. 4). In 3 of 6 experiments, the expression of CD80 slightly increased.
Figure 3. Cell-surface ACE expression in different types of hematopoietic cells. Immunofluorescence analysis of cultured PBMC (gated for monocytes and lymphocytes separately), PBL, and immature DC was performed on the cells isolated from seven donors. Analysis of anti-ACE binding with macrophages was performed on the cells isolated from two donors. Cellsurface ACE expression is indicated as mean fluorescence intensity (⫾ SD). (Mean fluorescence intensity of cells stained with isotype control antibodies was subtracted from each value).
Macrophages generated by culturing monocytes from peripheral blood of healthy donors in the presence of M-CSF were characterized by significant levels of CD14 and CD115 expression (Fig. 2A). Analysis of ACE expression on monocytes and lymphocytes was performed by gating on relative populations in freshly isolated peripheral blood cells from healthy donors. The level of ACE expression on monocytes as revealed by the mixture of mAbs 9B9 and i2H5 was significantly lower than in cultured macrophages or DC, but was higher compared to ACE expression on lymphocytes ( p ⬍ 0.05 in both cases; Fig. 3). Epitope specificity of ACE detection in different cell types Fig. 3 shows the binding pattern of the tested set of anti-ACE mAbs with different cells used in this study. The binding
ACE expression in leukemic DC It has been demonstrated previously that AML blasts acquire certain characteristics of DC upon stimulation with cytokines and modulators of calcium metabolism. Significant up-regulation of CD40, CD80, and CD86 expression is observed in DC derived from AML blasts [22,23,27]. In order to investigate whether the coordinated up-regulation of ACE expression is observed in leukemic cells during their DCtype differentiation, we performed both FACS analysis and ACE activity tests on cultured AML blasts. Leukemic DC derived from 10 AML patients displayed the phenotypic characteristics of DC in these cells (data not shown). As detected by staining with a mixture of 9B9 and i2H5, antiACE mAb cells from eight patients were ACE negative (data not shown). Only a low level of ACE expression was observed in leukemic DC derived from patients L and R (Fig. 5A and D). Because a remission sample was available from patient R, we compared ACE expression in DC derived from AML blasts and monocytes from the same patient. Leukemic DC derived from blast cells of patient R (Fig. 5B and E) had lower levels of ACE that observed in untreated peripheral blood monocytes from healthy donors. In contrast, DC cells derived from peripheral blood monocytes from the same patient harvested during remission produced high levels of ACE (Fig. 5C and F), which were comparable
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observed in cells cultured in the presence of cytokines (GMCSF, IL-4, TNF-α) and calcium ionophore A23187. Double staining of leukemic DC from patients R and L revealed ACE expression only in a small proportion of CD80⫹ cells (Fig. 5D and E). When DC were obtained from patient R monocytes harvested during remission, the CD80 fraction of these cells stained brightly with anti-ACE antibodies (Fig. 5F).
Figure 4. Effect of the ACE inhibitor lisinopril on DC-type monocyte differentiation. Immature DC cells were obtained from monocytes by culturing with cytokines for 6 days (see Material and methods section) in the absence (open histogram) or presence (filled histogram) of lisinopril 100 µM and stained with specific mAbs. Results of a representative experiment are shown.
with the ACE levels observed in DC obtained from monocytes of healthy donors (Fig. 1). In quantitative terms, the median fluorescence intensity of DC from blasts with anti-ACE mAbs was 3.6 ⫾ 5.3 (n ⫽ 10 patients), whereas that value for DC from monocytes was 192 ⫾ 92 (n ⫽ 5). The difference (54-fold) was statistically significant ( p ⫽ 0.010). The level of ACE expression in leukemic DC (as in DC from monocytes) was not dependent on the culture conditions used for DC generation: a low level of ACE was
ACE activity of monocytes, macrophages, and DC derived from healthy donor monocytes and DC from AML blasts To confirm the data obtained by FACS analysis, we measured the level of total ACE activity (intracellular plus cell surface ACE) in the lysates from these cells (Table 2). Specificity of ACE activity assay was demonstrated by the inhibition of measurable ACE activity using either ACE inhibitors or a set of blocking mAbs to ACE (data not shown). Generally, the results of ACE activity measurements confirmed that of flow cytometric analysis. The lowest (but still measurable) level of ACE activity was demonstrated for nonstimulated PBMC containing a mixture of resting monocytes and lymphocytes), and in an isolated population of lymphocytes stimulated with a IL-4/GM-CSF or IL-2/PHA. M-CSF–induced macrophage-type differentiation of monocytes was accompanied by a significant (55-fold) increase in ACE activity. Stimulation of monocytes with IL-4/GM-CSF, which leads to DC-type differentiation, induced an even greater level of ACE activity (150-fold). The effect of GMCSF and IL-4 on induction of ACE expression appears specific for monocytes because the combination of IL-4/ GM-CSF only slightly increases ACE activity in purified lymphocytes. DC derived from AML blasts showed negligible ACE activity. In addition, activation of lymphocytes with IL-2 and PHA did not significantly increase ACE activity. ACE activity in mature DC stimulated with LPS or TNF-α slightly decreased in comparison with immature DC. It is important to note that the level of ACE expression in DC (expressed as enzymatic activity per milligram of protein in cell lysate) is comparable with the highest level of somatic ACE expression in cultured human cells (human endothelial cells from umbilical vein): 5 to 15 mU/mg [30]. The highest ACE activity recorded in a human cell under the natural ACE promoter was found in spermatozoa: 650 mU/mg (Balyasnikova and Danilov, manuscript in preparation). However, these cells express the testicular form of ACE that are driven by “testicular” ACE promoter localized in twelfth intron of the ACE gene [31,32]. ACE activity in CHO cells transfected with the ACE cDNA driven by the CMV promoter ranged (depending on particular clone tested) from 200 to 1500 mU/mg [33]. However, to normalize ACE level in different cells per cell number, the relative value of ACE expression in DC will be changed dramatically. We found that the yield of protein in supernatant from lysate of DC was 270 µg/106 cells, CHO-ACE 125 µg/106 cells, PBMC/peripheral blood lymphocyte (PBL) 22 µg/106 cells,
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Figure 5. ACE and CD80 expression in dendritic cells derived from blasts of AML patients. Cells from AML M2 patient L were obtained at diagnosis (A, D). Cells from AML M5 patient R were harvested at diagnosis (B, E) and in remission (C, F). (A–C) Cells were stained with anti-ACE antibodies (mixture 9B9⫹i2H5; filled histograms) or with control anti-mouse IgG (clear histograms) Figures correspond to values of median fluorescence intensity. (D–F) Double staining with anti-ACE antibodies: mixture 9B9⫹i2H5 and anti-CD80 antibodies.
and spermatozoa 2.1 µg/106 cells. Therefore, ACE activity in DC expressed per 106 cells (2.5 mU/106 cells) is 1300 times higher than in PBMC or lymphocytes and twice as high as the expression of testicular form of ACE in spermatozoa (1.4 mU/106 cells). Thus, we can consider that DC expressed the highest level of ACE among human cells tested.
Discussion We previously demonstrated that a high level of ACE expression could be induced in immature DC [3]. Here we present a systematic quantitative study of ACE expression during the different stages of DC differentiation and maturation. We also show that AML blasts fail to up-regulate ACE during their DC-type differentiation induced by cytokine or calcium ionophore treatment. Somatic ACE has two highly homologous domains (Nand C-domains), each bearing a functional catalytic site [34]. ACE exists in both membrane bound and soluble forms. It is constitutively expressed on the surface of endothelial cells, on the epithelium of intestinal brush borders, on the epithelial cells of the proximal tubule of the kidney, on the tubular epithelial cells of the epididymis, and on alveolar macrophages. The interest of hematologists in ACE has dramatically increased with the discovery that N-acetylseryl-aspartyl-lysyl-proline (Ac-SDKP), a regulatory factor in hematopoiesis, is a natural substrate for the N-terminal active center of ACE [35,36]. Ac-SDKP reversibly prevents the recruitment of pluripotent hematopoietic stem cells and
normal early progenitors into the S-phase of the cell cycle by maintaining them in the G0 phase [37,38]. The differentiation of monocytes into macrophages was accompanied by a dramatic (60-fold) increase in ACE expression [11]. ACE expression also can be increased upon induction with T lymphocytes [13,39], with dexamethasone [11], or with GM-CSF [14]. ACE overexpression also was demonstrated in giant and epithelioid cells in granulomas during sarcoidosis, Gaucher disease, and leprosy resulting in increased ACE blood level in these patients [40–42]. ACE participates in antigen presentation (of certain peptides, at least) by altering antigen processing. The efficient presentation of 147–158/R-peptide of influenza virus nucleoprotein results from removal of the COOH-terminal dipeptide by ACE [16]. T-cell stimulation by the HIV-1 gp160-derived peptide p18 presented by H-2D class I MHC molecules also was found to require proteolytic cleavage of the peptide by ACE [17]. ACE acts to enhance antigen presentation exclusively in an exocytic intracellular compartment, and this pathway is TAP (transporter associated with antigen presentation) dependent [18,19]. The present study demonstrated that DC-type differentiation of monocytes is accompanied by a dramatic 150-fold increase in ACE activity normalized to cellular protein, which is equivalent to a 1300-fold increase in ACE activity per cell. This fact was confirmed by two independent assays: flow cytometric analysis with a set of mAbs to ACE (Figs. 1 and 3) and measurement of enzymatic activity (Table 2). The absolute values of ACE activity in human monocytes
S.M. Danilov et al. / Experimental Hematology 31 (2003) 1301–1309 Table 2. Angiotensin-converting enzyme activity in different cell types derived from human blood Cells PBMC Immature DC 3 days in culture Immature DC 6 days in culture Mature DC (LPS treatment) 8 days in culture Mature DC (TNF-α treatment) 8 days in culture Leukemic DC (CI or TNF-α treatment) 5 or 8 days in culture Macrophages 6 days in culture Macrophages (⫹LPS) 6 days in culture PBL (⫹IL-2/PHA) 3 days PBL (⫹IL-4/GM-CSF) 3 days
ACE activity (mU/mg) 0.061 ⫾ 0.02 5.3 ⫾ 2.3 9.34 ⫾ 2.0 9.07 ⫾ 2.5 9.2 ⫾ 0.6 0.090 ⫾ 0.040 3.4 ⫾ 0.8 2.4 ⫾ 1.1 0.086 ⫾ 0.036 0.09 ⫾ 0.05
Angiotensin-converting enzyme (ACE) activity was measured in the detergent lysates of the indicted cells fluorimetrically using Hip-His-Leu as substrate. ACE activity was normalized per milligram of protein in cell lysate and expressed as mean ⫾ SD (n ⫽ 4). CI ⫽ calcium ionofore A23187; DC ⫽ dendritic cells; GM-CSF ⫽ granulocyte-macrophage colony-stimulating factor; IL-2 ⫽ interleukin-2; IL4 ⫽ interleukin-4; LPS ⫽ lipopolysacharide; PBL = peripheral blood lymphocytes; PBMC = peripheral blood mononuclear cells; PHA ⫽ phytohemagglutinin; TNF-α ⫽ tumor necrosis factor-α.
and macrophages presented in this study (Table 2) were practically identical to the data published elsewhere [11,15], thus validating our measurements and new data on the absolute value of ACE activity in DC. Generally, the ACE activity increase during differentiation of monocytes into macrophages and DC corresponds to level of ACE expression revealed by flow cytometry with anti-ACE antibodies (see Table 2: ACE activity; Figs. 1–3: FACS scan). However, we found that the binding pattern of a set of mAbs to ACE differs in different cells. These results (Fig. 3) clearly show the epitope specificity of ACE revealed in different cell types. Binding of mAb i1A8 to the surface of monocytes, macrophages, and some other related cells (THP-1 and U937) was 4- to 5-fold higher than the binding of any other mAb and thus might be considered to be more specific for monocyte/macrophage ACE. However, for quantification of ACE expression on the cell surface by FACS scan, perhaps more correct would be use of the mixture of two mAbs: 9B9 and i2H5, or i1A8 and i2H5. These pairs of mAbs bind to different nonoverlapping ACE epitopes [23], thus allowing more complete ACE detection. Moreover, these combinations—at least 9B9 and i2H5 (Figs. 1– 3)—are less influenced by culture conditions. A high level of ACE expression in DC may have practical significance for gene therapy and vaccination via DC. Retargeting of an adenoviral vector to a marker expressed on DC (CD40) dramatically enhanced gene transfer to monocytederived DC [43]. Retargeting of an adenoviral vector to
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ACE-expressing cells and efficient gene transfer was demonstrated in vitro and in vivo using anti-ACE antibody 9B9 [44]. Therefore, based on the surface density of ACE in DC and on the effectiveness of gene transfer with anti-ACE mAb, we expect that gene transfer to DC via ACE would significantly improve the efficacy of DC-based vaccination. Results of this work, as well as previous studies cited earlier, demonstrate the low level of ACE activity in mononuclear blood cells. The ACE activity in THP-1 (human monocytic leukemia cell line) and the macrophage-like diffuse histiocytic lymphoma U937 also were very low (albeit measurable) and comparable to the ACE activity in monocytes, PBMC, and PBL (Table 2). These cell lines are used widely as a model system for the study of monocyte/macrophage differentiation. The addition of a stimulus such as phorbol myristate acetate (PMA) results in phenotypic changes characteristic for macrophages. However, the amplitude of PMA-induced ACE induction in these cells (3-fold for THP-1 [45]) is incomparable with the ACE 55-fold induction during monocyte maturation (Table 2). This suggests that leukemic cell lines display abnormal regulation of ACE expression during their macrophage-type differentiation similar to that of AML blasts during their DC-type differentiation. We showed that DC expressed the highest level of ACE per cell among the human cells tested. The biologic function of ACE in DC, as well as in macrophages, remains largely unknown. ACE inhibition in maturing DC caused a significant increase in CD83 expression and slight increase in CD80 expression by DC (Fig. 4). CD83 is a marker for DC maturation [46]. Up-regulation of CD83 expression upon ACE inhibition can be explained either by the fact that ACE is involved in maintaining the immature DC phenotype and blocking ACE activity promotes DC maturation, or by a direct action of lisinopril on CD83 production. Recently, the effect of ACE inhibitor (captopril) on the phenotype of DC was studied [47]. The authors did not find any changes of commonly used DC markers. They incubated mature DC with captopril for 24 hours, in contrast to the 5 days in our experiments. The authors demonstrated, however, that their short treatment of DC with captopril significantly decreased the levels of secreted TNF-α, IL-1α, IL-10, IL-12, and IL-18. The choice of captopril as the ACE inhibitor for their study may have led to misleading results, because captopril has an SH-group, which in addition to providing ACE inhibition also demonstrates significant anti-oxidant effects. Therefore, part of the effects of captopril on cytokine secretion by DC might be ACE independent. It also was shown in this study that preincubation of DC with angiotensin II increased production of TNF-α and IL-6, but not other cytokines. Therefore, we consider the secretion of at least TNF-α and IL-6 by DC to be ACE dependent. ACE in mononuclear cells may hydrolyze regulatory peptides at the site of inflammation, e.g., ACE hydrolyzes bradykinin, substance P, and other tachykinins that are implicated
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in inflammatory and immunologic responses [48]. Participation of ACE in antigen presentation and the fact that the negative regulator of stem cell proliferation (Ac-SDKP) is a natural substrate for ACE lead us to speculate that the high ACE expression on the surface of DC is not just a reflection of the general activation of monocytes during differentiation, but might have physiologic significance for effective functioning of these cells. The conditions required for DC generation in vitro were shown to be sufficient for the induction of DC differentiation markers in AML blast cells arrested at the early stages of hematopoiesis [22,26]. Leukemic DC express immune costimulatory molecules, IL-12, and are fully competent as antigen-presenting cells in allogeneic and autologous antigen-specific lymphocyte stimulation [49,50]. The results obtained indicate that the transformation process affects the regulation of ACE gene expression in AML blasts. Interestingly, a lack of ACE up-regulation was noted for cells derived from patients with various AML subtypes, characterized by different genetic abnormalities. Surprisingly, none of the publications describing profiling changes in gene expression during differentiation of monocyte-derived DC mentioned a statistically significant (more than 2-fold) increase in ACE mRNA expression, albeit the probes for ACE mRNA were present on microchips used for these studies [51–55]. Moreover, in a separate study, where the changes in ACE mRNA expression during DC differentiation was specifically studied using RT-PCR analysis, the authors also did not find significant up-regulation of ACE mRNA [56]. These results allow us to suggest that ACE up-regulation during differentiation from monocytes to DC (as well as failure of up-regulation in the case of leukemiaDC) might be due to posttranscriptional processing, like redistribution of inactive intracellular ACE to the surface where ACE become catalytically active (and becomes accessible for mAbs). The presence of an intracellular pool of immature inactive ACE was demonstrated previously [30]. The exact mechanism of the regulation of ACE expression in DC during differentiation, as well as the link between the regulation of ACE expression and leukemogenesis, remains to be investigated. It has been proposed that leukemic DC may prove useful for anti-cancer vaccine design because these cells express tumor-specific antigens endogenously, being progeny of a leukemic clone. The observed deficiency of leukemic DC to up-regulate ACE may interfere with the ability of leukemic DC to provide adequate processing and presentation of tumor antigens and should be considered during the development of strategies for anti-cancer therapy.
Acknowledgments The authors thank Professor Ronald F. Albrecht, Head of the Department of Anesthesiology, University of Illinois at Chicago, for his continuous encouragement and support of this project. The
authors also thank Dr. Richard D. Minshall, University of Illinois at Chicago, for critical reading of this manuscript.
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