Published online before print June 26, 2007
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* Department of Clinical Immunology and Transplantation, Polish-American Institute of Pediatrics, Jagiellonian University Medical College; and
Department of Immunology, Faculty of Biotechnology, Jagiellonian University, Cracow, Poland
1 Correspondence: Department of Clinical Immunology and Transplantation, Polish-American Institute of Pediatrics, Jagiellonian University Medical College, Wielicka str. 265, 30-663 Cracow, Poland. E-mail: mizembal{at}cyf-kr.edu.pl
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: CD34+ cells • monocyte subpopulations • macrophages
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Several protocols for ex vivo expansion and generation of DC from CD34+ progenitors of hematopoietic cells were described [4 , 5 ]. However, little is known about in vitro expansion and differentiation of CD34+ cells into monocytes/macrophages. CD34+ cells differentiate without expansion to functional mature monocytic cells in the presence of M-CSF or combination of M-CSF, IL-6, and mast cell growth factor (MSF), affecting CD34+ cells at precursor level without distinct effect on the more mature stages [6 ]. On the other hand, G-CSF-mobilized CD34+ peripheral blood cells cultured in the presence of SCF and IL-2 allows generation of phenotypically novel (CD56+CD33+) minor subset of monocytes [7 ].
In the present study, a different approach for obtaining monocytes was designed. CD34+ cord blood cells were first expanded for 3–10 days to increase the cell yield, and then the cells were transferred to the medium with various differentiating factors in order to obtain mature monocytes.
Human blood monocytes are highly heterogeneous population of cells. There are two main subpopulations of blood monocytes: the major one consists of CD14++ "classical" monocytes and a minor one (5–10% of total monocytes) formed by CD14+CD16+ cells [8 ], which have some similarity to previously described CD64+ and CD64– monocytes [9 , 10 ]. CD14+CD16+ cells are regarded as more mature monocytes with enhanced proinflammatory and antitumor activity [8 , 11 ]. However, other rare (2.0–3.7%) subpopulations of monocytes: CD14++CD16+, CD14+CD16–CD33++, and CD14++CD56+ were also described, although little is known about their function, except that CD14++CD16+ preferentially binds enzymatically degraded low-density lipoproteins [12 13 14 ]. Finally, there is also a rare CD14+CD16++ subpopulation of monocytes that undergoes expansion in various diseases, e.g., HIV infection, or in vitro in the presence of GM-CSF, IL-4, IL-10, that exhibits phenotypic and functional DC-like characteristics [15 , 16 ].
The present study shows that using two-step approach, i.e., expansion of CD34+ hematopoietic progenitors isolated from cord blood, and then differentiating them in the presence of M-CSF, SCF, IL-3, and Flt-3L, it is possible to obtain two novel subpopulations of monocytes (CD14+CD16– and CD14++CD16+), which occur at the ratio of approximately 2:1, show several differences in the immunophenotype and biological activity and are distinct from known subpopulations of blood monocytes.
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Immunophenotyping
The following pairs of mAbs for characterization of cell phenotype were used: anti-CD14 (clone M5.E2) allophycocyanin (APC)-conjugated and anti-CD16 (clone Leu 11c) PE-conjugated or anti-CD14 FITC and anti-CD16 phycoerythrin-Cyanine 5 (PE-Cy5)-conjugated (all from BD Biosciences, San Diego, CA, USA). Single staining of cultured cells with the following mAbs was used: PE-labeled anti-CD15, CD19, CD29, CD41, CD62p, and FITC-labeled anti-CD1a, CD83, CD38, CD3, and CD40 (all from BD Biosciences). To further characterize monocyte subpopulations in some experiments, three color flow cytometry analysis was performed after staining with the following mAbs against: HLA-DR, CD11a, CD40, CD64, all FITC-labeled (BD Biosciences) and CD11b, CD13, CD33, CD80, CD86, CD163, CCR2, CCR5, CXCR4 PE-conjugated (BD Biosciences), and CCR2-PE (R & D). Unlabeled purified anti-CD56 mAb, following staining with FITC-conjugated rabbit anti-mouse immunoglobulins (Dako A/S, Glostrup, Denmark) was also used. In parallel, staining with appropriate isotype matched mouse IgG were used as negative controls. Monocytes after incubation for 30 min at 4°C with mAb or isotype controls were washed, resuspended in 0.3 ml of PBS containing 0.1% sodium azide, and analyzed by flow cytometry (FACS Canto; BD Biosciences Immunocytometry Systems, San Jose, CA, USA) using FACS DiVa software. List mode data for 10,000 events were acquired, and statistical analysis was performed according to green, orange, or red fluorescence of cells stained with isotype controls.
Isolation of monocyte subpopulations
Cells cultured in the differentiation medium were harvested, washed, and suspended at the concentration of 10x106/ml. After staining with anti-CD14 APC and anti-CD16 PE cells were sorted into CD14+ (total population of monocytes) and CD14+CD16– and CD14++CD16+ subpopulations with the use of FACS Vantage SE (BD Biosciences IS) flow cytometer, equipped with TurboSort option (BD Biosciences IS) and Aerosol Protection System (Flexoduct International ApS, Greve, Denmark). The ion laser Innova Enterprise II (Coherent, Santa Clara, CA, USA) operating at 488 nm was used as a light source. Sorting was performed using a 70-µm nozzle tip with a drop drive frequency of 65 kHz, 1.5 drop envelopes and "normal-R" sort mode. Sorted cells were collected into water-cooled (constant temperature circulator; Neslab Instruments, Portsmouth, NH, USA) polystyrene Falcon 2057 tubes (BD Biosciences) precoated with FCS to avoid plastic charging and cell attachment to the wall. The purity of sorted cells was checked by flow cytometry and exceeded 95%.
Analysis of cell morphology
FACS-sorted monocyte subpopulations (CD14+CD16– and CD14++CD16+) were centrifuged on microscope slides in a Cytospin 2 (Shannon, Cheshire, UK), then fixed in 96% ethanol for 30 min and stained with HE. The images at 1000x magnification were taken using light microscope BX51 (Olympus, Tokyo, Japan), a Camedia 30 camera and DP software (Olympus).
Production and measurement of cytokines
Sorted total CD14+ cells and CD14+CD16– and CD14++CD16+ subpopulations were cultured in flat-bottom 96-microtiter plates (Nunc, Roskilde, Denmark) at 1x105/100 µl/well in RPMI 1640 medium (PAA Laboratories GmbH, Pasching, Austria) with 5% FCS. Cells were cultured alone or were stimulated with 400 U/ml of human recombinant IFN
(Sigma) and 100 ng/ml of LPS from Salmonella minessota (Sigma), or human pancreatic carcinoma cells (HPC-4 line) [17
] at the ratio 1:0.3 for 18 h at 37°C in a humidified 5% CO2 atmosphere. The levels of TNF, IL-10, and IL-12p40 in the culture supernatants were determined by ELISA. The following matched mAbs pairs (BD Biosciences) were used: for TNF
Mab1 (capture) and Mab11 (detection); for IL-10: JES3-9d7 (capture) and JES-12G8 (detection); for IL-12p40: C8.3 (capture) and C8.6 (detection). Recombinant human cytokines (all from BD Biosciences) were used as the standards. Tests were performed, according to the manufacturer's protocol (sensitivity for TNF and IL-12p40: 20 pg/ml; for IL-10: 10 pg/ml), and results were determined with ELISA reader (universal microplate reader, Bio-Tek Instruments, Vinooski, VT, USA) at 492 vs. 630 nm wavelength. All samples and standards were run in duplicate. For determination of intracellular IL-12 production, expanded and differentiated cells were stimulated with IFN and LPS for 18 h in the presence of monensin (Golgi Stop, 2 µm; BD), harvested and stained with anti-CD14-APC and anti-CD16-FITC mAbs, as described above. Cells were washed with ice cold PBS, fixed and permeabilized with Cytofix/Cytoperm (BD) reagent (20 min at 4°C). Then cells were washed twice with Perm/Wash solution (BD) and pelleted cells were stained (30 min, 4°C) with anti-IL-12p40/70 (clone C11.5, BD) PE-conjugated or appropriate isotype control. After three washes, cells were suspended in PBS with 0.1% sodium azide and analyzed by flow cytometry.
MLR
PBMC isolated from EDTA-blood of healthy donors by standard Ficoll/Isopaque density gradient centrifugation were used as responding cells. 1 x 105/well of allogeneic PBMC were added to irradiated (2500 cGy) monocytes or their subpopulations (1x104 to 1x105/well) and cultured in triplicate in RPMI 1640 medium with 10% of pooled human AB serum for 6 days. 1 µCi/well of [3H]-thymidine (3H-TdR, Amersham, Aylesbury, UK) was added for the final 18 h of culture, then cells were harvested on fiberglass filter and uptake of isotope was determined in ß-scintillation counter (Beckman, Palo Alto, CA, USA). The results were expressed as cpm of incorporated 3H-TdR.
Phagocytosis
Phagocytosis was measured by flow cytometry (FACS Canto) using fluorescent bacteria, as described previously [18
]. Briefly, after staining with anti-CD14 APC and anti-CD16 PE-Cy5, the cells (1x106/ml) were incubated in Falcon 2054 tubes (BD Biosciences) at 37°C for 30 min in RPMI 1640 medium, with FITC-labeled Staphylococcus aureus ATCC 25923 opsonized (30 min, 37°C) in the presence of 10% pooled fresh human serum (cells-to-bacteria ratio 1:20), and cells were incubated for additional 15 min at 37°C. Afterward, cells were analyzed by flow cytometry using FACS Canto and DiVa software, taking into account cells emitting green–FL1 (phagocytosis of bacteria) fluorescence among far red–FL4 positive only (CD14+CD16–), and far red/red–FL4/FL3 positive cells (CD14++CD16+).
Statistics
The differences were analyzed by Student’s t test with the use of Microcal Origin version 5.0 software (Northampton, MA, USA). The differences were considered significant at P < 0.05.
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Figure 1. Expansion of CD34+ cord blood hematopoietic progenitor cells. CD34+ cells isolated by immunomagnetic sorting from cord blood mononuclear cells were cultured for 3–14 days in X-VIVO 10 medium+FCS with SCF, TPO, IL-3, and Flt-3L at 1x105/ml/well. Every 3 or 4 days, cells were harvested and transferred at the same concentration to new wells with fresh medium. (A) The mean fold increase (±SD) over cell number plated on day 0. Results from four independent experiments are shown at each time point. (B) The percentage of cells expressing indicated determinants at different time points ± SD is shown.
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300-fold by day 14. Flow cytometry analysis of the whole cultures showed a lack of cells expressing the following determinants: CD1a, CD3, CD15, CD19, CD41, CD56, CD62p, CD83, almost all cells (92–98%) expressed CD11a, CD29, and CD33, approximately 64% were CD38+, and 72% HLA-DR+ (Table 1
). Around 40% of cells expressed CD14 and CD64 and 20% were CD16+. The proportion of CD34+ cells was usually below 5.0%. Determination of CD14 and CD16 coexpression indicated that among CD14+ cells, two subpopulations of CD14+CD16– and CD14++CD16+ cells were observed (Fig. 2B)
. By days 7–10 on average, up to 60% (range 20–70%) were CD14+, and these two subpopulations were present at approximately 2:1 ratio (Fig. 2C)
. Over the next days, the decrease of other subpopulations, in particular CD14++CD16+, was observed with simultaneous increase of CD14–CD16+ cells. Isolated CD14+ monocytes cultured for 7–10 days in the differentiation medium did not proliferate as judged by a lack of increase of the cell yield (data not shown).
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Figure 2. Characterization of cells cultured in the differentiation medium. CD34+ cells expanded for 7 days in X-VIVO 10 medium (see Fig. 1
) were harvested and cultured at 1x105/ml/well in the differentiation medium: IMDM supplemented with 20% FBS, SCF, M-CSF, and Flt-3L for 3–14 days. (A) The mean fold increase (± SD) over cell number plated on day 0, (B) FACS analysis of CD14 and CD16 expression on cells harvested on day 10 shows the percentage of CD14+CD16– and CD14++CD16+ cells and MFI of CD14 (right panel), isotype controls (left and middle panels). (C) Kinetic analysis of CD14 and CD16 expression (percentage of positively stained cells) over 14 days of culture is shown. Data are based on six independent experiments.
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Table 1. Immunophenotypic of CD34+ Cells Expanded for 3 Days and Differentiated for 7 Daysa
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Figure 3. The kinetics of CD14+ cell yield in the differentiation cultures. The cells from expansion cultures were harvested on days 3, 7, and 10 and then cultured in the differentiation medium for 3–14 days. The mean fold increase of CD14+ cells over CD34+ cell number plated on day 0 of expansion culture is shown at each time point of differentiation culture. Data are based on four experiments performed.
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Table 2. Immunophenotypic Characteristics of CD14+CD16– and CD14++CD16+ Monocyte Subsetsa
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Figure 4. Morphological features of CD34+ cell-derived monocyte subsets. CD34+ cells were expanded for 7 days and then cultured in the differentiation medium for 10 days. Cells were stained with anti-CD14 and anti-CD16 mAbs and FACS sorted into CD14+CD16– (A) and CD14++CD16+ (B) monocyte subsets. Cytospin slides were made and cells were stained with HE and examined under light microscope (x1000). Photos were made with Camedia 30 camera (Olympus). Data acquisition was made using DP software (Olympus).
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Figure 5. Effect of HVD addition to the differentiation culture on the occurrence of monocyte subpopulations. Following 7 days expansion, the cells were transferred to the differentiation medium without (A) or with 10–8 M of 1.25(OH)2 Vitamin D3 (B) and cultured for 3–14 days. FACS analysis of cells harvested at day 7 stained with anti-CD14 and anti-CD16 mAbs is shown. The FSC/SSC parameters (left) and the percentage of CD14+CD16– and CD14++CD16+ cells and MFI of CD14+ cells (right) are shown. One representative experiment out of four performed is presented.
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/LPS or cancer cells. Both subpopulations secreted TNF and IL-12 but little or no IL-10. CD14++CD16+ cells released higher quantity of TNF and IL-12p40, but only the latter was significantly different (Table 3
). However, the release of cytokines by cells obtained in separate experiments was highly variable. To exclude possible activation of sorted cells, triple staining of all cells in the culture for CD14, CD16 and intracellular cytokines was performed. Fig. 6
shows the data for IL-12p40/70. It confirmed that CD14++CD16+ subset produced more of this cytokine. |
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Table 3. Cytokine Secretion by Initial Population of Monocytes and Their Subsetsa
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Figure 6. Intracellular IL-12 expression by CD14+CD16– and CD14++CD16+ monocyte subsets. Expanded cells cultured in differentiation medium for 7 days were harvested and stained with anti-CD14 APC and anti-CD16 FITC mAbs, washed, fixed, permeabilized, and stained with anti-IL-12 APC mAb. After washing, cells were analyzed by flow cytometry.
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Figure 7. Stimulation of allogeneic MLR by monocyte subsets. Subpopulations of monocytes were isolated as described in Fig. 4
, irradiated and added in different numbers (indicated at the bottom) to allogeneic PBMC. Cells were cultured for 6 days with terminal pulse of 3H-TdR for 18 h. Data are obtained in four separate experiments are expressed as cpm ± SD.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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In the first step, four different media were used for CD34+ cell expansion: X-VIVO 10 ± FCS, SFEM and STEMLINE II. Within 14 days of culture, the highest increase (40–80 fold) of the cell number was achieved in cultures with X-VIVO 10+FCS medium. Therefore, this medium was used for initial expansion and subsequent differentiation to monocytes. However, we noticed a rapid drop of CD34+ cells cultured in this medium from day 3 onward, although in other media, CD34+ cells were present for longer periods of time. Almost all of the cells cultured in X-VIVO 10+FCS were CD33+ CD38+, while CD14+ and CD15+ were not found. This is in keeping with other observations showing a rapid loss of CD34+ cells at day 7 cultured in Iscove’s medium+FCS with slightly different cytokine combination [21
].
In the second step, the cells from expansion cultures, containing myeloid progenitors (CD13+, CD33+, HLA-DR+) were transferred to differentiation medium and cultured for 3–14 days. The further increase in cell numbers (up to 300-fold at day 14) was observed. In these cultures, no lymphocytic or granulocytic precursors were observed. On average, on day 7, 45 ±18% of cells were CD14+ and 22±10% CD16+. Majority of cells (>90%) were CD11a+, CD29+ CD13+, CD33+, while CD38+ and HLA-DR were in range 64–82%. Few (below 5%) CD34+ cells were also detected. Double staining with anti-CD14 and anti-CD16 mAbs revealed that two subpopulations of monocytes were observed: CD14+ (MFI 964) CD16– and CD14++ (MFI 2756) CD16+. At day 14, the proportion of CD14++CD16+ decreased while CD14+CD16– was similar as at day 7. Absolute fold increase, i.e., increase during expansion for 10 days and differentiation for 14 days, of CD14+ cells was from 160-fold at day 7 to over 600-fold at day 14.
To our knowledge, such subpopulations of monocytes were not generated previously from cord blood CD34+ hematopoietic progenitors in vitro, although CD14++CD16+ cells were obtained in vitro by culturing blood monocytes in the presence of IL-10 [16 ]. Differentiation, but not expansion, of functional mature monocytic cells from CD34+ progenitor cells in the presence of M-CSF or its combination with MGF and IL-6 was described, but occurrence of subpopulations was not observed [6 ]. Expansion of myeloid progenitors was achieved by culturing CD34+ cell from cord blood in the presence of steel factor, IL-6, GM-CSF, and G-CSF [21 ]. A phenotypically novel, small population (2.5%) of monocytes (CD56dimCD33+) cells with macrophage morphology was generated from G-CSF mobilized CD34+ cells cultured in the presence of IL-2 and SCF [7 ]. Also, with the use of 1.25(OH)2 Vitamin D3 differentiation, but not expansion, of CD34+ progenitors to CD14+CD11b++ cells was observed [19 ]. Hence, none of these protocols is similar to our system that allows significant increase of the number of CD14+ cells and differentiation into two subpopulations (CD14+CD16– and CD14++CD16+) of monocytes from CD34+ progenitors.
Blood monocytes are heterogeneous populations of cells and their two main subpopulations were described: CD64+, CD64– cells [9 , 10 ], and CD14++ ("classical"), CD14+CD16+ monocytes [22 ]. However, subpopulations of monocytes generated in the present system have little immunophenotypic similarities to the CD14+CD16+ and CD14++ cells present in the blood, not only by different CD14+, but also other determinants expression. Thus, while blood CD14+CD16+ monocytes are characterized by low expression of CD33 and CD64 [8 , 12 , 14 , 23 ], it is not the case for CD14++CD16+. On the other hand, both blood CD14+CD16+ and CD34+-derived CD14++CD16+ (in comparison to CD14+CD16–) show an enhanced expression of CD86, HLA-DR [23 ]. The CD14+CD16– cells also differ from "classical" CD14++CD16– blood monocytes not only by a low CD14 but also low CD11b and CD33 expression and the presence of CCR5 [8 , 12 ]. No DC markers (CD1a, CD83) were detected on both cell subsets, which make them distinct from blood CD14+CD16++ cells [15 ]. Hence, monocyte subpopulations described here are phenotypically different from two major blood monocyte subsets: CD14++ and CD14+CD16+ [22 ]. Since we have also not observed expression of CD56 on both monocyte subpopulations (not shown), they seem to be distinct from a small subset of CD56low, CD33+, CD14+, HLA-DR+, and CD11bhigh monocytes present in the peripheral blood [24 ], and distinct from CSF-induced proliferating subpopulation of monocytes, which do not express CD86 [25 ]. The small monocyte-derived subpopulation of CD14++CD16+ blood monocytes (3.6±1.5% of total) and similar subset derived from monocytes by culturing in vitro in the presence of IL-10 have been described that is characterized by decreased CCR2 and CD64 and increased CCR5 and CX3CR1 expression (by comparison to CD14+CD16– monocytes) that preferentially reacts with fractalkine [14 , 15 ]. CD14++CD16+ subpopulation, as defined by Rothe et al [12 ], also preferentially binds enzymatically degraded LDL, show an increased expression of CD11a and CD11b, and exhibit similar phagocytosis of E. coli and MPO expression as CD14++ cells [12 ]. However, CD14++CD16+ monocytes described by us differ from this subset present in the blood as although CCR5 was increased, CCR2, CD11a expression (by comparison to CD14+CD16– cells) was comparable, while CD40 and CD64 was increased, although an enhanced expression of CD11b makes this subset similar to blood CD14++CD16+ cells [12 ]. We have also observed that CD14++CD16+ subset, like CD14+CD16+ monocytes is characterized by enhanced expression of CXCR4 [21 ], which may imply its higher migratory activity to SDF and significantly increased CD163 (hemoglobin scavenger receptor). The latter suggests that these cells are more mature macrophage-like cells [26 ], which is also compatible with their macrophage-like morphology. CD34+ cell-derived subsets also differed in their morphology. CD14++CD16+ cells were large with macrophage-like appearance, while CD14+CD16– were monocyte-like cells. Finally, CD40 was also preferentially expressed on CD14++CD16+ cells, which may implicate their higher potential in interactions with T cells and B cells. Therefore, although CD34+ cell-derived monocyte subpopulations share the expression of some determinants with blood monocytes subsets, they are clearly distinct from them in other markers expression.
Since 1.25(OH)2 Vitamin D3 induces the monocytic commitment of CD34+ progenitor cells [19 ], their effect on generation of monocyte subsets was studied. 1,25(OH)2 Vitamin D3 did not enhance proliferation of cells but up-regulated HLA-DR and CD11b expression on both subpopulations, which is in keeping with previous observations [17 ]. The novel finding was that 1.25(OH)2 Vitamin D3 down-regulated CD16 expression, resulting in an increased proportion of CD14+CD16– cells. Hence, 1.25(OH)2 Vitamin D3 seems to regulate reciprocally proportions of these monocyte subpopulations or their maturation.
Subpopulations of monocytes generated in the present system showed also functional differences. Both subpopulations stimulated either with IFN/LPS or tumor cells did not release significant amount of IL-10, which is in contrast to the classical blood monocytes [11
, 20
] but produced TNF and IL-12. The TNF and IL-12p40 release by CD14++CD16+ cells was higher (IL-12 significantly) than by the other subset. In this regard, the CD14++CD16+ subset is similar to the behavior of CD14+CD16+ cells from the blood [11
, 20
] and may be regarded as proinflammatory. The CD14++CD16+ cells were more potent stimulators of allo-MLR, which make them functionally similar to CD14+CD16+ blood monocytes [10
]. The increased stimulatory potential of CD14++CD16+ cells may be associated with an enhanced expression of HLA-DR and costimulatory CD86 molecule. In contrast, phagocytic potential of CD14+CD16– subset was substantially higher than CD14++CD16+, which makes the former more similar to "classical" CD14++CD16– blood monocytes [12
].
In conclusion, the present paper shows that CD34+ cord blood hematopoietic progenitors expansion in vitro followed by differentiation to monocytes leads to generation of high numbers of CD14+ cells, which consists of two novel subpopulations of monocytes: CD14++CD16+ and CD14+CD16– that differ in immunophenotype and biological functions from monocyte subpopulations generated so far ex vivo from CD34+ cells, or present in the peripheral blood. It seems likely that the different monocytes subsets described in the present and other studies reflect developmental stages with distinct physiological roles, allowing dissection of functional relevance. This may have implications for the development of therapeutic strategies targeted to modify particular subpopulations of monocytes [23 ]. Alternatively, as present protocol allows the generation of proliferating monocytes/macrophages, these cells can potentially be used in gene therapy as cellular delivery vehicles [3 ].
Received February 19, 2007; revised May 24, 2007; accepted May 24, 2007.
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J. Exp. Med. 184,695-706
glarczyk, K., Ruggiero, I., Hajto, B., Zembala, M. (2004) Antitumour response of CD14+/CD16+ monocyte subpopulation Exp. Hematol. 32,748-755[CrossRef][Medline]
oszyn, M., Szatanek, R., Baj-Krzyworzeka, M., Siedlar, M., Ruggiero, I., Wickiewicz, J., Zembala, M. (2003) Tumour cell-induced deactivation of human monocytes J. Leukoc. Biol. 74,1094-1101glarczyk, K., Mysiak, M., Guzik, K., Ernst, M., Flad, H.-D., Pryjma, J. (2001) Fas (CD95)-Fas ligand interactions are responsible for monocyte apoptosis occurring as a result of phagocytosis and killing of Staphylococcus aureus Infect. Immun. 69,1287-1297,25-dihydroxyvitamin D3 induce the monocytic commitment of CD34+ hematopoietic progenitors J. Leukoc. Biol. 71,641-651This article has been cited by other articles:
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