Published online before print August 3, 2004
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* Department of Medicine I and
Institute of Pathology, University Hospital Carl Gustav Carus at the University of Dresden Technical Center, Germany
1 Correspondence: Medizinische Klinik und Poliklinik I an der Technischen Universität Dresden, Fetscherstr. 48, 01307 Dresden, Germany. E-mail: alexander.kiani{at}uniklinikum-dresden.de
 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Key Words: cyclosporin A • bone marrow • neutrophil granulocytes • megakaryocytes • proliferation
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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To date, four calcium-regulated NFAT proteins with partly distinct but overlapping tissue distribution have been identified: NFATc2 (also termed NFAT1), NFATc1 (NFAT2), NFATc4 (NFAT3), and NFATc3 (NFAT4); a fifth family member, NFAT5, is regulated by osmotic stress. NFAT family members share many target genes; however, knockout mice and other models have also revealed unique and unexpected functions of individual proteins [9 10 11 ]. Notably, expression of NFAT proteins is not restricted to T cells but is detectable in a number of other immune as well as nonimmune cells. Whereas in lymphocytes and other cell types, NFAT proteins predominantly regulate the rapid activation of inducible genes of fully differentiated cells, in some tissues, they exert a second biological function, namely the control of tissue differentiation and adaptation [4 12 13 ]. Specifically, NFAT family members are involved in the regulation of chondrocyte [14 ], myocyte [15 ], adipocyte [16 ], keratinocyte [17 ], and endothelial cell [18 ] differentiation, as well as in the development of cardiac valves [19 ] and vessels [20 ] and the hypertrophy reaction of skeletal [21 ] and heart muscle cells [22 ].
CD34+ cells of bone marrow, cord blood (CB), and granulocyte-colony stimulating factor (G-CSF)-mobilized peripheral blood (PBL) are known to be enriched for hematopoietic stem cells (HSCs) and provide long-term repopulating activity when used as a source for allogeneic stem cell transplantation [23
24
25
]. A network of extrinsic and intrinsic factors including cytokines, cytokine receptors, and transcription factors [26
27
28
29
] controls two fundamental properties of HSCs, self-renewal and differentiation. For example, differentiation of HSCs into the diverse myeloid lineages requires the presence of specific transcription factors such as CCAAT/enhancer-binding protein 
(C/EBP; for the differentiation of neutrophils), PU-1 (for monocytes/macrophages), and GATA-1 (for erythrocytes, eosinophils, and megakaryocytes). Most of these transcription factors are coexpressed at low levels in early CD34+ CD38– progenitor cells [30
]; once differentiation of a given myeloid lineage is initiated, the expression of lineage-specific factors is up-regulated, and the expression of others is suppressed. By experimental manipulation of these mechanisms, the route or efficacy of differentiation can be affected [28
29
31
].
Although regulation and function of NFAT proteins in lymphocytes have been studied extensively, little is known about their expression and potential biological relevance in myeloid cells. Furthermore, no reports exist about whether hematopoietic stem and progenitor cells express NFAT genes or proteins or if they are specifically regulated during myeloid differentiation. In the current study, we analyzed the expression and regulation of NFAT family members in CD34+ cells, as well as in myeloid cells of various lineages and at different stages of maturation. We found that NFATc2 mRNA and protein are expressed at surprisingly high levels in CD34+ cells and are rapidly down-regulated during granulocytic differentiation. Furthermore, the NFAT inhibitor CsA profoundly inhibits proliferation of CD34+ cells but not their differentiation into neutrophil granulocytes.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Cell isolation
CD34+ cells and monocytes were purified from the blood of G-CSF-mobilized, healthy stem cell donors; in selected experiments, CD34+ cells were isolated from umbilical CB scheduled for discard. Total bone marrow was obtained by iliac crest aspiration from healthy stem cell donors. Neutrophil granulocytes and mononuclear cells were isolated from the PBL of healthy volunteers. The study was approved by the institutional review board of the medical faculty of the Technical University Dresden (Germany), and informed consent was obtained from the donors.
For CD34+ cell isolation, a sample of the leukapheresis product was centrifuged and resuspended in phosphate-buffered saline (PBS; PAA Laboratories, Pasching, Austria) containing 0.4% acid citrate dextrose (Baxter, München-Unterschleissheim, Germany) and 0.5% human serum albumin (Baxter). CD34+ cells and/or CD14+ monocytes were positively isolated by immunomagnetic selection using the CD34 progenitor cell isolation kit and/or CD14 MicroBeads (Miltenyi Biotec GmbH, Bergisch-Gladbach, Germany), following the instructions of the manufacturer. The purities of the resulting CD34+ and CD14+ populations, respectively, were assessed by fluorescence-activated cell sorting (FACS) and routineously exceeded 95%. Neutrophil granulocytes were isolated by a first-step sedimentation of whole blood on 1.5% dextran T500 (Carl Roth GmbH, Karlsruhe, Germany) followed by centrifugation of the granulocyte-rich supernatant on Ficoll-Hypaque (density 1.077 g/ml; Biochrom AG, Berlin, Germany). The resulting CD15+ population of neutrophil granulocytes was subjected to hypotonic lysis, washed, and resuspended in PBS containing 1% fetal calf serum (FCS; Invitrogen GmbH, Karlsruhe, Germany). Purity was analyzed by FACS using mAb to CD15, CD14, CD3, and CD19 and routineously exceeded 95%. Peripheral blood mononuclear cells (PBMC) and total bone marrow cells were isolated by density gradient centrifugation using Ficoll-Hypaque (1.077 g/ml).
Culture and ex vivo stimulation of CD34+ cells
For short-term ex vivo stimulation, CD34+ cells and PBMC were seeded at 1 x 106/ml in RPMI 1640 (Invitrogen GmbH), supplemented with 2 mM CaCl2 (Sigma, Deisenhofen, Germany), 10% FCS, 1% penicillin/streptomycin, 10 mM Hepes (PAA Laboratories), and 50 µM 2-mercaptoethanol (Merck, Darmstadt, Germany), and stimulated for 30 min at 37°C with the calcium ionophore ionomycin (1 µM, Calbiochem, Merck Biosciences, Bad Soden, Germany). Where indicated, stimulation was done in the presence of CsA (1 µM, Sigma), which was added 30 min prior to ionomycin. At the end of the stimulation, the cells were harvested and directly used for cytospin preparations or protein isolation (see below).
For differentiation experiments, freshly isolated CD34+ cells were seeded at 1 x 105/ml in Cellgro medium (CellGenix, Freiburg, Germany) and cultured for 14 days in the presence of a cytokine cocktail containing rhSCF (50 ng/ml), FLT3L (100 ng/ml), IL-3 (5 ng/ml), GM-CSF (5 ng/ml), and G-CSF (30 ng/ml; days 1–5). On days 5, 8, 11, and 14, cells were harvested, washed, counted, and expanded with fresh media supplemented with cytokines (day 5: IL-3 and G-CSF; day 8 and 11: G-CSF) or used directly for further analysis (see below). Where indicated, CsA was added to the cultures.
Morphologic analysis and flow cytometry
The morphology of the cells was analyzed by staining cytospin preparations with May-Grünwald-Giemsa. The phenotype and/or differentiation efficacy was analyzed by FACS.
Immunofluorescence of cytospins
After ex vivo stimulation, cytospin preparations were fixed in 3% neutrally buffered paraformaldehyde, washed, blocked with PBS/10% FCS, and permeabilized with 0.5% Nonidet P-40 (NP-40; Boehringer Mannheim, Mannheim, Germany). The cells were then incubated for 30 min with the mAb against NFATc2, diluted 1:1000 in blocking buffer (PBS, 10% FCS, 0.5% NP-40), washed, and incubated for 30 min with the secondary antibody (Cy3-labeled sheep anti-mouse immunoglobulin G, Sigma), diluted 1:1000 in blocking buffer. Slides were mounted using the SlowFade antifade kit (Molecular Probes, Eugene, OR) containing 4',6-diamidine-2'-phenylindole dihydrochloride, analyzed with a Nikon Labophot-2 fluorescence microscope and processed by ISISF software (MetaSystems GmbH, Altlussheim, Germany).
Immunohistochemistry and immunofluorescence of bone marrow biopsies
Bone marrow biopsies from routine pathology cases were fixed in 4% neutrally buffered formaldehyde for 24 h, decalcified with 270 mM EDTA (pH 7.4) for 12 h, and embedded in paraffin. For all immunolabelings, antigen recovery was performed by microwave pretreatment. For immunohistochemistry, the mAb to NFATc2 was used at a dilution of 1:100. Peroxidase-based detection of the antibody was performed with the LSAB2 horseradish peroxidase (HRP) kit (Dako, Hamburg, Germany), according to the manufacturer’s instructions. For immunofluorescence, the mAb to NFATc2 was used at a dilution of 1:50; the polyclonal antiserum to NFATc2 was used at a dilution of 1:5. The mAb to CD34 (Dako) was used at a dilution of 1:50, and the mAb to glycophorin A (Dako) was used at a dilution of 1:500. The secondary antibodies (dichlorotriazinylaminofluorescein-labeled anti-mouse and anti-rabbit; Texas red-labeled anti-mouse and anti-rabbit) were used at a dilution of 1:80. Immunofluorescence pictures were taken at a Zeiss fluorescence microscope equipped with FluoroArc fluorescence control and Optimas 6.5 software.
Western blot analysis
Cells (5–10x106) were washed with ice-cold PBS, resuspended with Totex buffer (100 µl/ 5x106 cells) containing 20 mM Hepes buffer (PAA Laboratories), 1% NP-40, 0.42 M NaCl, 20% glycerol, 1 mM MgCl2, 10 mM EDTA, 10 µg/ml leupeptin, and 0.5 M dithiothreitol (DTT; Sigma), and freshly supplemented with 1 mM phenylmethylsulfonyl fluoride and 20 mM sodium pyrophosphate (Sigma) and vortexed for 20 min at 4°C. Protein was quantified in duplicate with the bicinchoninic acid kit (Pierce, Rockford, IL) using bovine serum albumin as standard.
From each sample, 30 µg total protein was separated by 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE; BioRad, Hercules, CA) and transferred to a nitrocellulose membrane [0.2 µm Hybond enhanced chemiluminescence (ECL), Amersham Biosciences, Freiburg, Germany]. Membranes were incubated with the monoclonal anti-NFATc2 antibody (1:2500), the monoclonal anti-NFATc1 antibody (1:2000), the polyclonal anti-NFATc3 antibody (1:200), or the monoclonal anti-actin antibody (1:2000–1:3500, clone AC-40, Sigma) followed by HRP-conjugated secondary antibody, goat anti-mouse or goat anti-rabbit (1:2000, Dako), as appropriate. Immunoreactive bands were visualized by using the ECL detection kit (Amersham Pharmacia Biotech, Little Chalfont, UK).
Reverse transcriptase-polymerase chain reaction (RT-PCR) analysis
Total RNA was prepared using Trizol reagent (Invitrogen Corp., Carlsbad, CA), according to the recommendations of the manufacturer. Target RNA (1 µg) was reverse-transcribed using 200 IU Superscript II RT (Invitrogen Corp.) at 42°C for 40 min in the presence of 10 mmol/L Tris-HCl (pH 8.3), 50 mmol/L KCl, 1.5 mmol/L MgCl2, 10 mM DTT, 0.625 mmol deoxyribonucleoside triphosphate (dNTP), 40 IU RNasin (Promega, Madison, Wisconsin), and 5 µM random hexamer.
For nonquantitative analysis of NFATc1-4, PCR was performed in a 50-µl vol containing 1 µl cDNA, 2.5 IU AmpliTaq Gold polymerase (Applied Biosystems, Foster City, CA), 200 µM dNTPs (Invitrogen), GeneAmp 10x PCR buffer (Applied Biosystems), 1.5 mM MgCl2, and 300 nM primers. The following primers were used: NFATc2 forward, 5'-AGAAACTCGGCTCCAGAATCC-3'; NFATc2 reverse, 5'-TGGTTGCCCTCATGTTGTTTTT-3'; NFATc1 forward, 5'-GCCGCAGCACCCCTACCAGT-3'; NFATc1 reverse, 5'-TTCTTCCTCCCGATGTCCGTCTCT-3'; NFATc4 forward, 5'-TCAGAAGACACGGCGGACTTCC-3'; NFATc4 reverse, 5'-TGAACATCTGTAGGGTCAGTGG-3'; NFATc3 forward, 5'-ACCAGCC-CGGGAGACTTCAATAGA-3'; NFATc3 reverse, 5'-AAATACCTGCACAATCAATACTGG-3'. The PCR conditions were as follows: one cycle at 94°C for 10 min, followed by 30 cycles at 94°C for 30 s, 60°C (NFATc2, NFATc3, and NFATc4) or 64°C (NFATc1) for 30 s, 72°C for 30 s, and a final extension cycle at 72°C for 10 min. For the control of RNA and cDNA preparations, we used RT-PCR for ß-actin. Amplified products were analyzed electrophoretically on a 2% agarose gel and were visualized under ultraviolet rays after ethidium bromide staining.
Real-time RT-PCR was performed on an ABI-Prism 7700 PCR SDS (Applied Biosystems). Validated 6-FAM-labeled PCR primers and Taqman probes for NFATc2 (assay ID: Hs00234855_m1) were used, and VIC-labeled glyceraldehyde 3-phosphate dehydrogenase (GAPDH; ID: 4310884E) served as an endogenous control. PCR mix was prepared according to the manufacturer’s instructions (assay-on-demand, Applied Biosystems). PCR was always performed in duplicates. Thermal cycler conditions were as follows: 2 min 50°C, 10 min 95°C, 40 cycles denaturation (15 s, 95°C), and combined annealing/extension (1 min, 60°C). For the quantification of NFATc2 mRNA expression in differentiating CD34+ cells, mRNA/cDNA levels of NFATc2, after normalization against the housekeeping gene GAPDH, were expressed relative to the value obtained at day 0, calculated by the formula 1/2x–y (x: threshold cycle [CT] of each sample; y: Ct at day 0). For the quantification of NFATc2 mRNA expression in different cell types, PCR fragments of NFATc2 (amplified with the above TaqMan primers) were cloned into the pCR 2.1-TOPO vector (Invitrogen) and quantified, and copy numbers were calculated based on the molecular weight. NFATc2 mRNA copy numbers of each sample were then determined by plotting the Ct of each well against a reference curve prepared by amplifying serial dilutions of the corresponding plasmid. As populations of CD34+ cells, neutrophil granulocytes, monocytes, and Jurkat cells considerably differ in RNA content and GAPDH expression, the respective copy numbers were expressed per µg of total cellular RNA.
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Figure 1. Expression of NFAT family members in CD34+ cells, isolated to a purity of at least 95% by immunomagnetic separation from the PBL of G-CSF-mobilized, healthy donors (A–D) and CB (D). Total cellular RNA and protein were extracted and analyzed for the expression of NFAT family members by RT-PCR (A) and Western blot (B–D). Jurkat T cells were used as a positive control. Note that differences in the migration of the NFATc2 band (B–D) may represent differences in the phosphorylation status of the protein (*, completely dephosphorylated form) or indicate the presence of different isoforms of NFATc2 in CD34+ and T cells [32
]. (B) The three bands (middle panel) correspond to the three known isoforms of NFATc1 [33
].
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60%) consists of immature myeloid precursor cells at various stages of differentiation (Fig. 2 A
). This suggested that expression of NFATc2 in CD34+ progenitor cells is lost in differentiating and mature myeloid cells of the neutrophilic/monocytic lineages.
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Figure 2. Expression of NFATc2 in myeloid cells at different stages of maturation. (A) PBL neutrophil granulocytes (Neutr) and monocytes (Mono; purity >95%), total bone marrow, and G-CSF-mobilized PBL CD34+ cells were isolated from healthy donors as described in Materials and Methods and analyzed for the expression of NFATc2 by Western blot. Jurkat T cells were used as a positive control. Note that the migration of the NFATc2 band corresponds to the phosphorylation status of the protein (lower band, dephosphorylated form). (B) Total cellular RNA was extracted from CD34+ cells (n=7), neutrophil granulocytes (n=4), monocytes (n=4), and Jurkat T cells (n=2) and analyzed for the expression of transcript levels of NFATc2 by quantitative Taqman RT-PCR, as described in Materials and Methods. Shown are means ± SEM.
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Figure 3. NFATc2 expression in normal human bone marrow. Paraffin-embedded normal human bone marrow biopsies were analyzed by immunofluorescence (A, C) and immunohistochemistry (B) for expression of NFATc2. (A, C) Immunofluorescence double-labeling with polyclonal NFATc2 antiserum (red) and mAb to (A) CD34 and (C) glycophorin A (green). Note that (A) the NFATc2 fluorescence (red) colocalizes with the stem cell marker CD34 (green) but (C) not with the erythroid marker glycophorin A (green). (B) Peroxidase immunohistochemistry, using a mAb to NFATc2, demonstrates positive staining of bone marrow megakaryocytes (arrowhead), whereas granulocytes identified by their typical nuclear morphology (arrows) are negative.
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2 weeks; furthermore, it was accompanied by considerable proliferation of the cells (Fig. 4B)
. Strikingly, as early as 5 days after the start of differentiation, the expression of NFATc2 protein (Fig. 4C)
and mRNA (Fig. 4D)
in the differentiating cells was almost completely suppressed. The time-point of down-regulation of NFATc2 correlated with the development of a promyelocytic phenotype (data not shown) and the loss of CD34 expression (Fig. 4A) . Thus, expression of NFATc2 in CD34+ cells is rapidly suppressed upon progression of the cells into the promyelocytic stage, consistent with the observed expression profile of NFATc2 in immature and mature myeloid cells isolated from bone marrow and PBL.
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Figure 4. Rapid suppression of NFATc2 expression in CD34+ cells differentiating along the neutrophilic lineage. CD34+ cells were isolated from the PBL of G-CSF-mobilized, healthy donors and cultured in the presence of a sequentially added cytokine cocktail inducing neutrophilic differentiation of the cells (see Materials and Methods for details). Aliquots of the cells were harvested before and at different time-points after the start of the culture. (A) Efficacy of neutrophilic differentiation of the cells, indicated by changes in the percentage of cells positive for surface expression of the stem cell marker CD34 and the neutrophil marker CD15. (B) Proliferation of the cells, indicated by the increase in cell number, relative to the start of the culture (day 0). (C) NFATc2 protein expression, analyzed by Western blot. (D) NFATc2 mRNA expression, analyzed by quantitative Taqman RT-PCR. NFATc2 transcript levels were normalized for expression of the reference gene GAPDH and expressed relative to the value obtained at the start of the culture (day 0; for details, see Materials and Methods). Shown are means ± SEM of three experiments.
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[40
], and PU-1 [41
], but the function of NFATc2 in CD34+ cells is unknown. In T lymphocytes and most other cell types tested, NFATc2 is target for the calcineurin inhibitor CsA, which completely abrogates NFAT-mediated gene expression and therefore has been useful as a tool to evaluate NFAT functions in these cells [2
6
]. Before we could similarly test the effect of CsA on CD34+ cells, we had to establish whether NFATc2 regulation in these cells follows the same calcium-dependent/CsA-sensitive pattern as in T cells. As shown in Figure 5 A
, a 30-min treatment of CD34+ cells with the calcium ionophore ionomycin resulted in increased migration of the NFATc2 band in SDS-PAGE/Western blot, which in earlier studies using T cells had been demonstrated to correspond to dephosphorylation of the protein [42
]. Similarly, ionomycin treatment induced complete translocation of NFATc2 from the cytoplasm into the nucleus of the cells (Fig. 5B)
. Ionomycin-induced dephosphorylation and nuclear translocation of NFATc2 were completely blocked by the addition of CsA (Fig. 5A
and 5B)
. Therefore, NFATc2 regulation in CD34+ cells is dependent on calcium/calcineurin activation and sensitive to inhibition by CsA.
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Figure 5. Regulation of NFATc2 in CD34+ cells, isolated from the PBL of G-CSF-mobilized donors and left unstimulated or treated for 30 min with ionomycin (Iono), CsA, or both. (A) Calcineurin-dependent dephosphorylation of NFATc2. Total cellular protein was extracted, separated by SDS-PAGE, blotted, and stained for NFATc2. Note that the migration of the NFATc2 band in the gel under these conditions corresponds to the phosphorylation status of the protein (lower band, dephosphorylated form) [42
]. (B) Calcineurin-dependent nuclear translocation of NFATc2. Cytospin preparations of CD34+ cells were prepared, and the subcellular localization of NFATc2 was visualized by immunofluorescence using a mAb to NFATc2. PBMC were used as a control.
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Figure 6. CsA inhibits the proliferation of CD34+ cells but not their differentiation into neutrophil granulocytes. CD34+ cells were isolated from the PBL of G-CSF-mobilized, healthy donors and cultured in the presence or absence of the indicated cytokine or combination of cytokines. CsA or solvent was added to the cultures in the indicated concentrations (in µM). Aliquots of the cells were harvested before and at different time-points after the start of the culture. Shown are means ± SEM of three experiments, respectively. (A) Proliferation of the cells in response to a sequentially added cytokine cocktail inducing neutrophilic differentiation of the cells (see Materials and Methods for details). Indicated is the increase in cell number relative to the start of the culture. The inset represents a magnified section (days 0–5) of the graph. (B) Proliferation of the cells in response to individual cytokines or combinations of cytokines. Indicated is the increase in cell number after a culture of 5 days relative to the start of the culture. (C) Efficacy of neutrophilic differentiation, indicated by changes in the surface expression of the neutrophil marker CD15 [mean fluorescence intensity (MFI), gated on live cells] over time.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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CD34+ cells are known to coexpress several transcription factors at low levels, and specific combinations of transcription factors determine the survival, proliferation, and differentiation responses of CD34+ cells to extrinsic or intrinsic signals [29 30 47 ]. The observation that NFATc2 is expressed in CD34+ cells is novel and may indicate unexpected functions for this transcription factor family in the hematopoietic system. Expression levels for NFATc2 mRNA and protein in CD34+ cells were surprisingly high and comparable with those found in Jurkat cells, a T cell line that contains considerable amounts of NFATc2 [34 ]. As we used CD34+ cells of G-CSF-mobilized PBL in most of our experiments, we considered it important to confirm NFATc2 expression in CD34+ cells of bone marrow (Fig. 3A) and CB (Fig. 1D) . CD34+ cells isolated from these sources differ in their frequency of primitive precursor cells [48 ], proliferative potential [49 ], and cell-cycle status [35 ]. The results show that CD34+ cell expression of NFATc2 is independent of the source and furthermore in PBL not notably influenced by potentially confounding effects of the mobilizing cytokine G-CSF. An interesting, yet open, point is the question of whether NFATc2 is differentially expressed in subgroups of the CD34+ population, especially if it can be detected in primitive hematopoietic precursor cells (e.g., CD34+CD38–).
Another important aspect was to work out the mechanisms by which NFATc2 is regulated in CD34+ cells. In most cell types, NFAT activation occurs via the "classical" Ca2+/calcineurin-dependent, CsA-sensitive pathway of dephosphorylation and nuclear translocation; however, calcineurin-/CsA-independent pathways also exist [50 51 52 ]. Furthermore, to our knowledge, the presence of calcineurin (phosphatase 2B) in CD34+ cells has not yet been reported. Our results indicate that as in T lymphocytes, NFATc2 in CD34+ cells is constitutively cytoplasmic and responds alike to pharmacological stimulation and inhibition of calcineurin. The physiological stimulus for NFATc2 activation in CD34+ cells remains presently unclear; the most likely candidates are cytokines and growth factors, which in other cell types are known to be capable of increasing intracellular calcium levels sufficient for calcineurin activation [53 54 ]. Exemplary for the hematopoietic system are eosinophils, in which stimulation with IL-4 and IL-5 has recently been demonstrated to result in NFAT activation [43 ].
The predominant task of future studies will be to define the biological role of NFATc2 in CD34+ cells. As a first step, we approached this question indirectly by assessing the effect of CsA on CD34+ cells in vitro, taking advantage of our observation that the activation of NFATc2 in CD34+ cells was sensitive to this drug. We therefore postulated that any effect of NFATc2 on CD34+ cell self-renewal/proliferation, apoptosis/death, or differentiation/commitment would be neutralized by the presence of CsA; conversely, CD34+ cell functions found to be resistant to CsA would unlikely be mediated by NFATc2. CsA profoundly suppressed the proliferative response of CD34+ cells to a cocktail of hematopoietic growth factors in liquid culture in a dose-dependent manner (Fig. 6A)
. Inhibition of CD34+ cell proliferation was evident at even low and clinically relevant levels of CsA. Few reports exist in the literature addressing the effect of CsA on the proliferation and/or differentiation of hematopoietic precursor cells, most of which are hampered by the problem that as a result of the presence of lymphocytes in the cellular culture, direct and indirect effects of CsA on CD34+ cells could not be adequately separated [55
56
]. In one study by Perry et al. [57], an antiproliferative effect of CsA was reported on the colony growth of highly enriched, murine hematopoietic precursor cells (Thy-1.1lowSca1+Lin–) stimulated in methylcellulose cultures with SCF, IL-3, IL-6, G-CSF, and erythropoietin. At first sight, the antiproliferative effect of CsA on CD34+ cells in vitro may be surprising, as CsA is commonly used in patients after allogeneic stem cell transplantation, and in this situation does not appear to affect the normal hematopoietic recovery of the patients after the conditioning therapy. However, we also show that CsA does not suppress (and even may facilitate) the differentiation of myeloid precursor cells; furthermore, the antiproliferative effect of CsA at concentrations achieved in vivo (200 ng/ml) is relatively weak (Fig. 6A)
and probably outweighed by suppression of T cell-derived interferon-
, which is known to inhibit the colony formation of hematopoietic stem cells [58
59
]. Nevertheless, the negative effect of CsA on the proliferation of CD34+ cells characterizes this drug as less than optimal for the immunosuppressive treatment of patients after allogeneic stem cell transplantation, and the search for agents acting more specifically on lymphocytes appears to be desirable for this patient group.
Received April 26, 2004; revised June 25, 2004; accepted July 1, 2004.
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-deficient mice Proc. Natl. Acad. Sci. USA 94,569-574-(immune) interferon antiserum in vitro Exp. Hematol. 14,621-625[Medline] Transplantation 51,1276-1282[Medline]This article has been cited by other articles:
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