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Aggressive characteristics of myeloblasts expressing CD7 in myelodysplastic syndromes
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Leukemia Research 33 (2009) 326–331
Aggressive characteristics of myeloblasts expressing CD7 in myelodysplastic syndromes Chikako Satoh a,b , Hideto Tamura a , Taishi Yamashita a,c , Takashi Tsuji c , Kazuo Dan a , Kiyoyuki Ogata a,∗ a
Division of Hematology, Department of Medicine, Nippon Medical School, 1-1-5 Sendagi, Bunkyo-ku, Tokyo 113-8603, Japan b Department of Bioregulation, Institute of Gerontology, Nippon Medical School, Kanagawa, Japan c Department of Industrial Science and Technology, Tokyo University of Science, Chiba, Japan Received 24 May 2008; received in revised form 26 June 2008; accepted 8 July 2008 Available online 20 August 2008
Abstract Clinical data suggest that CD7+ myeloblasts are linked with poor prognosis in myeloid malignancies including myelodysplastic syndromes (MDS). To explore the biology behind this, we compared cell characteristics between CD34+CD7+ and CD34+CD7− myeloblasts from an MDS cell line and fresh samples from MDS patients. Compared with CD34+CD7− myeloblasts, CD34+CD7+ myeloblasts showed greater proliferative capacity, more active cell cycling, and less apoptosis. In analyses of a cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of interleukin-8 and chemokine (C–C motif) ligand 2 genes, suggesting immaturity of these cells. These findings might underlie the clinical aggressiveness in CD7+ MDS. © 2008 Elsevier Ltd. All rights reserved. Keywords: Myelodysplastic syndromes; CD7; CD34; Myeloblasts; Apoptosis; IL-8; CCL2
1. Introduction Human CD7 molecules are expressed on thymocytes, T and natural killer cells, and progenitors of lymphoid and myeloid cells in healthy individuals [1]. The physiological ligand and function of CD7 have not been completely established [1]. In T cell proliferation, CD7-mediated signals may be costimulatory or inhibitory depending on the experimental conditions [2,3]. In one study, CD7mediated signals induced granulocyte-macrophage colonystimulating factor (GM-CSF) production in two myeloid cell lines [4]. Myeloblasts expressing CD7 may have a pathophysiological role in myeloid malignancies. In acute myeloid leukemia (AML), we and others showed that CD7+ cases (defined as patients in whom more than 20% of myeloblasts express CD7 in this paper) were very common in patients with unfavorable cytogenetics but not in others [5,6]. In chronic ∗
Corresponding author. Tel.: +81 3 3822 2131; fax: +81 3 5685 1793. E-mail address: [email protected] (K. Ogata).
0145-2126/$ – see front matter © 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.leukres.2008.07.006
myeloid leukemia, patients whose myeloblasts express CD7 in flow cytometry (FCM) or gene expression analyses have poor prognoses [7,8]. In myelodysplastic syndromes (MDS), we reported that CD7+ cases (actually CD34+CD7+ cases because most MDS myeloblasts express CD34) were virtually confined to high-grade MDS and showed an especially poor prognosis [9]. Furthermore, myeloblasts gain CD7 expression with disease progression in some MDS patients [9,10]. Finally, another group has recently reported that lowgrade MDS patients, in whom even a small fraction of myeloblasts expressed CD7, were transfusion dependent or showed disease progression in the follow-up period [11]. However, the mechanism explaining these clinical observations remains unknown. By examining an MDS cell line and fresh samples from patients with MDS or AML transformed from MDS, this study showed that CD34+CD7+ myeloblasts were more aggressive, in terms of proliferation and avoiding apoptosis, compared with CD34+CD7− myeloblasts. In analyses of the cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of interleukin-8
C. Satoh et al. / Leukemia Research 33 (2009) 326–331
(IL-8) and chemokine (C–C motif) ligand 2 (CCL2) genes, suggesting immaturity of these cells.
2. Materials and methods 2.1. Cells A CD34+ myeloblastic MDS cell line (F-36P), which was established from a patient with refractory anemia with excess blasts (RAEB) in transformation [12] and a small fraction of which expressed CD7, was obtained from Riken Cell Bank (Ibaraki, Japan). Blood or marrow samples from 7 patients, which contained both CD34+CD7+ and CD34+CD7− myeloblasts and were collected after obtaining informed consent, were also used. The diagnoses of the patients (French–American–British classification [13]) were 3 RAEB, 2 RAEB in transformation, and 2 AML transformed from MDS. Cytogenetic analysis was successful for all patients, and abnormal karyotypes were detected in 5 (2 RAEB, 2 RAEB in transformation, and 1 AML transformed from MDS). These karyotypes were classified as intermediateand poor-risk groups in 2 and 3 of the 5 patients, respectively, according to the previous categorization [14]. CD34+CD7+ and CD34+CD7− myeloblasts were purified using fluorescenceactivated cell sorting (FACS, Fig. 1A) as described previously [15]. In some experiments, CD34+ cells were purified from the patient samples using magnetic cell sorting [15]. Isolated cells with more than 97% purity were used. This study was approved by the Institutional Review Board of Nippon Medical School. 2.2. Liquid cultures for examining cell proliferation The purified CD34+CD7+ and CD34+CD7− F-36P cells (2 × 104 cells/mL) were cultured in RMPI 1640 medium containing fetal bovine serum (FCS, 5%) and human IL-3 (5 ng/mL, Immuno-Biological Laboratories [IBL], Gunma, Japan). The viable cell numbers were determined using the trypan blue dye-exclusion test every 3–7 days to maintain cell concentrations appropriate for survival (2 × 104 to 1 × 105 mL−1 ). In single-cell cultures, an irradiated HESS-5 stromal cell layer, needed for single F-36P cells to proliferate, was prepared in 96-well microtiter plates as described previously [15,16]. Purification and automated deposition of a single CD34+CD7+ or CD34+CD7− F-36P cell into each well was performed with FACS. Patient cells (5 × 104 cells/mL), suspended in ␣-MEM containing 12.5% FCS, 12.5% horse serum, and human cytokines (IL-3 [10 ng/mL], thrombopoietin [Kirin Brewery, Takasaki, Japan, 50 ng/mL], stem cell factor [IBL, 50 ng/mL], and Flk2 ligand [IBL, 50 ng/mL]), were cultured on irradiated HESS-5 stromal cell layers. 2.3. Colony-forming assay MethoCult H4230 and GF H4434 methylcellulose media (StemCell Technologies, Vancouver, BC, Canada) were used for F-36P and patient cells, respectively [15]. IL-3 (10 ng/mL) was added to the former. Colonies (aggregates of 20 or more cells) were scored on day 14 of culture.
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2.4. Analyses of immunophenotypes, cell cycle, and apoptosis Multicolor FCM was used [9,17]. The antibodies included anti-CD45-peridin chlorophyll, anti-CD34-fluorescein isothiocyanate (FITC), anti-CD13-phycoerythrin (PE), anti-CD33-PE (BD PharMingen, San Diego, CA, USA), and anti-CD7-PE (BeckmanCoulter, Marseille, France). For cell cycle analysis, cells were stained with anti-CD7-PE, fixed in 70% cold ethanol, and stained with PI [18]. In experiments analyzing apoptosis, F-36P cells were or were not exposed to cytarabine (Ara-C, 0.13 M) for 6 h, washed, and then cultured for 3 days in RMPI 1640 medium containing 5% FCS and human IL-3 (5 ng/mL). Meanwhile, patient cells were cultured for 12 h in ␣-MEM containing 12.5% FCS, 12.5% horse serum, and various human cytokines (10 ng/mL of IL-3 and 50 ng/mL of thrombopoietin, stem cell factor and Flk2 ligand) with or without Ara-C (0.13 M). This shorter culture period was used due to the fragility of fresh MDS cells. These cells were stained with antiCD7-PE, annexin V-FITC, and propidium iodide (PI) (Trevigen, Gaithersburg, MD, USA) for detecting apoptosis [19]. 2.5. Gene microarray RNA from purified CD34+CD7+ and CD34+CD7− F-36P cells was analyzed with microarray hybridization [20]. Total RNA (5 g) was reverse-transcribed using an oligo-dT primer containing a T7 RNA polymerase promoter sequence to synthesize the first-strand cDNA. The double-stranded cDNA that contained the T7 RNA polymerase promoter at the 3 end was transcribed into cRNA by T7 RNA polymerase with allylamine-derivative nucleotides. cRNA was then reacted with N-hydroxy succinimide esters of Cy3 or Cy5 (PerkinElmer, Foster City, CA, USA). Dye molecules were separated from labeled products using the RNeasy mini kit (Qiagen, Valencia, CA, USA). The mixture was then applied to the microarray (Hitachi Human Oligo DNAchip, Hitachi Life Science, Saitama, Japan) and hybridization was carried out for 17 h at 45 ◦ C. After hybridization, slides were washed and scanned using a confocal laser scanner (PerkinElmer). Fluorescence intensities on scanned images were quantified, corrected for background, and normalized using global normalization methods based on the assumption that the median value of fluorescence intensities of both samples should be the same. The DNAchip used covers 1606 genes of various functional classes, including cytokines/growth factors and their receptors, apoptosis regulators, oncogenes, transcription factors, signal transducers, transporters, and cell cycle regulators. 2.6. Real-time quantitative PCR Total RNA from purified CD34+CD7+ and CD34+CD7− F-36P cells was transcribed into cDNA using Super Script FirstStrand Synthesis System for real-time PCR (Invitrogen, Carlsbad, CA, USA). Real-time PCR reactions were performed using the Gene Amp 5700 Sequence Detection System (Applied Biosystems [ABI], Foster City, CA, USA) as described previously [21]. The following Assays on-Demand probes (ABI) were used: actin, Hs 99999903 m1; IL-8, Hs 00174103 m1; and CCL2, Hs 00234140 m1. The relative standard curve experiments were used to measure the expression level of the 2 target genes relative to the -actin gene expression level.
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2.7. Statistics The two-sided Wilcoxon signed-rank test was used.
3. Results First, purified CD34+CD7+ and CD34+CD7− F-36P cells were compared for their growth potential. Compared with CD34+CD7− F-36P cells, CD34+CD7+ F-36P cells proliferated more rapidly in liquid cultures (Fig. 1B) and formed more colonies in semisolid cultures (Fig. 1C). Consistent with these findings, CD34+CD7+ F-36P cells had
fewer G0/G1-phase and more G2/M-phase cells compared with CD34+CD7− F-36P cells (Fig. 1D). Furthermore, CD34+CD7+ F-36P cells had fewer apoptotic cells compared with CD34+CD7− F-36P cells (Fig. 1E, without Ara-C). Finally, the addition of cytarabine to the cultures increased apoptosis in CD34+CD7− F-36P cells but not in CD34+CD7+ F-36P cells (Fig. 1E). Although CD34+CD7+ F-36P cells showed a proliferation advantage, they never dominate in cultures of unseparated F36P cells. Therefore, CD34+CD7+ F-36P cells may generate CD34+CD7− F-36P cells in culture. To examine this possibility, CD34+CD7+ and CD34+CD7− F-36P cells were subjected to single-cell culture. Considerable cell prolifera-
Fig. 1. A distinct difference was seen in cell characteristics of CD34+CD7+ and CD34+CD7− F-36P cells. (A) Purification of CD34+CD7+ and CD34+CD7− F-36P cells using FACS. (B) CD34+CD7+ F-36P cells (black circles) proliferated more rapidly than CD34+CD7− F-36P cells (white circles) in liquid cultures. Data are means of duplicate cultures of a representative experiment (S.D. values were too small to be shown here). Eight separate experiments showed essentially the same results. (C) CD34+CD7+ F-36P cells (black bar) formed more colonies than CD34+CD7− F-36P cells (white bar) in semisolid cultures (P < 0.05). Data are mean ± S.D. of four separate duplicate cultures. (D) Cell cycle status of CD34+CD7+ and CD34+CD7− F-36P cells. The upper panels show representative analyses of the cell cycle and the lower panels show a comparison of cell cycle between CD34+CD7+ (black bars) and CD34+CD7− (white bars) F-36P cells. Data are mean ± S.D. of six separate experiments in which CD7+ and CD7− cells were gated in FCM of unseparated F-36P cells in exponential cell growth. Asterisks indicate significant differences between black and white bars (P < 0.05). Experiments using the purified CD34+CD7+ and CD34+CD7− F-36P cells showed essentially the same results. (E) Apoptosis of CD34+CD7+ and CD34+CD7− F-36P cells. The upper left panel shows representative analyses of apoptosis and the lower left panel shows that caspase-3 activation was observed in most annexin V+ cells. The middle and the right panels are comparisons of annexin V+ and PI+ cells between CD34+CD7+ (black bars) and CD34+CD7− (white bars) F-36P cells. Data are mean ± S.D. from eight separate experiments in which CD7+ and CD7− cells were gated in FCM of unseparated F-36P cells. * P < 0.01, ** P < 0.05. Experiments using the purified CD34+CD7+ and CD34+CD7− F-36P cells showed essentially the same results. (F) Single CD34+CD7+ (four bars on the left) and CD34+CD7− (four bars on the right) F-36P cell cultures in microtiter wells. The numbers of wells, in which cells were immunophenotyped on days 12, 21, and 31 of culture, are shown under the bars. The wells were categorized according to the results of immunophenotyping, i.e., wells in which more than 95% of cells were CD7+ (black), wells in which more than 95% of cells were CD7− (white), and wells in which CD7+ and CD7− cells comprised more than 5%, respectively (spotted). The Y-axis shows the proportion of wells meeting these categories, e.g., the second bar from the left shows that two of 14 wells examined were in the black category and other 12 wells were in the spotted category.
C. Satoh et al. / Leukemia Research 33 (2009) 326–331
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Table 1 Differences in gene expression profile between CD7-positive and CD7negative F-36P cells determined in microarray analysis Accession no.
Gene name
Ratioa
M17017.1 S71513.1 NM 000129.2 NM 030812.1
Interleukin-8 Chemokine (C–C motif) ligand 2 Coagulation factor XIII, A1 polypeptide Actin-like 8
0.3 0.39 0.45 2.02
a
Gene expression ratios between CD7-positive and CD7-negative F-36P cells (the former data/the latter data).
tion (generating more than 100 cells from a single cell) was observed in approximately one-third of wells plated with single cells. Immunophenotyping of the proliferated cells in each well showed clearly that CD34+CD7− cells were generated in most wells plated with single CD34+CD7+ cells (Fig. 1F, left). Meanwhile, few CD34+CD7+ cells were generated in wells plated with single CD34+CD7− cells (Fig. 1F, right). In microarray analysis, four genes (for actin-like 8, IL8, CCL2 and coagulation factor XIII, A1 polypeptide) showed a greater than 2-fold difference in expression between CD34+CD7+ and CD34+CD7− F-36P cells (Table 1) (data below the threshold for detection were excluded). The expression of the actin-like 8 gene was increased and that of the other three was decreased in CD34+CD7+ F-36P cells compared
Fig. 2. A difference was seen in IL-8 and CCL2 gene expressions between CD34+CD7+ and CD34+CD7− F-36P cells determined using real-time PCR. Gene expression levels of IL-8 and CCL2 relative to that of -actin are shown (data are mean ± S.D. of three separate experiments, with each assay performed in triplicate). * P < 0.01.
with CD34+CD7− F-36P cells. We confirmed the lower IL8 and CCL2 gene expressions in CD34+CD7+ F-36P cells using real-time PCR (Fig. 2). Although CD7-mediated signals induced GM-CSF production in one report [4], there was no difference in GM-CSF gene expression and GM-CSF content in the culture supernatant between CD34+CD7+ and CD34+CD7− F-36P cells (data not shown).
Fig. 3. CD34+CD7+ myeloblasts have advantages in growth and avoiding apoptosis over CD34+CD7− myeloblasts in patients. (A) Purified CD34+CD7+ myeloblasts (black circles) proliferated more rapidly than purified CD34+CD7− myeloblasts (white circles) in liquid cultures. Data are mean ± S.D. of duplicate cultures. (B) Purified CD34+CD7+ myeloblasts (black bars) formed more colonies than purified CD34+CD7− myeloblasts (white bars) in semisolid cultures. Data are mean ± S.D. of duplicate cultures. (C) Apoptosis of CD34+CD7+ (black bars) and CD34+CD7− (white bars) myeloblasts, which were or were not exposed to Ara-C. The two cell populations were gated in FCM of the purified CD34+ cells. Data for the annexin V+ and PI+ cells are shown.
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Finally, we examined CD34+CD7+ and CD34+CD7− myeloblasts from patients. CD34+CD7+ myeloblasts again showed a growth advantage over CD34+CD7− myeloblasts in liquid cultures and colony-forming assays (Fig. 3A and B). Furthermore, CD34+CD7+ myeloblasts had fewer apoptotic cells compared with CD34+CD7− myeloblasts in liquid cultures in either the presence or the absence of Ara-C (Fig. 3C).
4. Discussion Although several clinical observations indicate that CD7+ myeloblasts are associated with poor prognosis in myeloid malignancies, the biological background explaining this notion was not known. Using the F-36P MDS cell line and fresh samples from patients, this study for the first time showed that CD34+CD7+ myeloblasts were more aggressive in terms of proliferation and avoiding apoptosis compared with CD34+CD7− myeloblasts in MDS. Based on these data, we speculate that the aggressive clinical behavior in CD7 + MDS patients is due to the cell characteristics of CD7+ myeloblasts rather than an epiphenomenon. In analyses of the F-36P cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of IL-8 and CCL2 genes compared with CD34+CD7− myeloblasts. Interestingly, it was reported that IL-8 and CCL2 production increased with cell maturation in AML blasts and that CCL2 was involved in apoptosis induction [22–24]. It was hypothesized that CD7+ myeloblasts are more immature than CD7− myeloblasts, because CD7+ cases are more common in immature leukemia such as AML-M0 and stem cell leukemia [25,26]. Taken together, all of the above findings suggest that CD34+CD7+ myeloblasts are more immature than CD34+CD7− myeloblasts. Do CD34 + CD7 + MDS myeloblasts have stem cell potential? In our previous study, CD34+ cells including CD34+CD7+ cells from MDS patients lacked long-term culture-initiating cell activity and did not reconstitute hematopoiesis in NOD/Shi-scid/scid (NOD/SCID) mice [15]. However, because MDS stem cells are difficult to detect, probably due to the instability of MDS cells [27], the definitive answer to this question remains unknown. This topic is especially interesting in view of the fact that agents targeting leukemic stem and progenitor cells have been reported recently [28,29]. It is unknown whether currently available therapies, such as demethylating agents and stem cell transplantation for eligible patients, improve the prognosis of CD7 + MDS patients. We recommend that the status of CD7 expression on myeloblasts be examined in clinical trials of such therapies. Furthermore, development of a therapy specific for patients with CD34+CD7+ myeloblasts might be beneficial. In summary, the data obtained here can explain the aggressive clinical behavior of CD7 + MDS cases. Monitoring and targeting of CD34+CD7+ myeloblasts are worth considering in MDS.
Acknowledgments This work was supported in part by a Grant-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (No. 17591015). Contributions: C.S. performed experiments, analyzed data and wrote the manuscript. H.T. designed the research. T.Y. and T.T supported technical aspect. K.D. provided administrative and clinical support. K.O. designed the research, analyzed data, and wrote the manuscript.
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Leukemia Research 33 (2009) 326–331
Aggressive characteristics of myeloblasts expressing CD7 in myelodysplastic syndromes Chikako Satoh a,b , Hideto Tamura a , Taishi Yamashita a,c , Takashi Tsuji c , Kazuo Dan a , Kiyoyuki Ogata a,∗ a
Division of Hematology, Department of Medicine, Nippon Medical School, 1-1-5 Sendagi, Bunkyo-ku, Tokyo 113-8603, Japan b Department of Bioregulation, Institute of Gerontology, Nippon Medical School, Kanagawa, Japan c Department of Industrial Science and Technology, Tokyo University of Science, Chiba, Japan Received 24 May 2008; received in revised form 26 June 2008; accepted 8 July 2008 Available online 20 August 2008
Abstract Clinical data suggest that CD7+ myeloblasts are linked with poor prognosis in myeloid malignancies including myelodysplastic syndromes (MDS). To explore the biology behind this, we compared cell characteristics between CD34+CD7+ and CD34+CD7− myeloblasts from an MDS cell line and fresh samples from MDS patients. Compared with CD34+CD7− myeloblasts, CD34+CD7+ myeloblasts showed greater proliferative capacity, more active cell cycling, and less apoptosis. In analyses of a cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of interleukin-8 and chemokine (C–C motif) ligand 2 genes, suggesting immaturity of these cells. These findings might underlie the clinical aggressiveness in CD7+ MDS. © 2008 Elsevier Ltd. All rights reserved. Keywords: Myelodysplastic syndromes; CD7; CD34; Myeloblasts; Apoptosis; IL-8; CCL2
1. Introduction Human CD7 molecules are expressed on thymocytes, T and natural killer cells, and progenitors of lymphoid and myeloid cells in healthy individuals [1]. The physiological ligand and function of CD7 have not been completely established [1]. In T cell proliferation, CD7-mediated signals may be costimulatory or inhibitory depending on the experimental conditions [2,3]. In one study, CD7mediated signals induced granulocyte-macrophage colonystimulating factor (GM-CSF) production in two myeloid cell lines [4]. Myeloblasts expressing CD7 may have a pathophysiological role in myeloid malignancies. In acute myeloid leukemia (AML), we and others showed that CD7+ cases (defined as patients in whom more than 20% of myeloblasts express CD7 in this paper) were very common in patients with unfavorable cytogenetics but not in others [5,6]. In chronic ∗
Corresponding author. Tel.: +81 3 3822 2131; fax: +81 3 5685 1793. E-mail address: [email protected] (K. Ogata).
0145-2126/$ – see front matter © 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.leukres.2008.07.006
myeloid leukemia, patients whose myeloblasts express CD7 in flow cytometry (FCM) or gene expression analyses have poor prognoses [7,8]. In myelodysplastic syndromes (MDS), we reported that CD7+ cases (actually CD34+CD7+ cases because most MDS myeloblasts express CD34) were virtually confined to high-grade MDS and showed an especially poor prognosis [9]. Furthermore, myeloblasts gain CD7 expression with disease progression in some MDS patients [9,10]. Finally, another group has recently reported that lowgrade MDS patients, in whom even a small fraction of myeloblasts expressed CD7, were transfusion dependent or showed disease progression in the follow-up period [11]. However, the mechanism explaining these clinical observations remains unknown. By examining an MDS cell line and fresh samples from patients with MDS or AML transformed from MDS, this study showed that CD34+CD7+ myeloblasts were more aggressive, in terms of proliferation and avoiding apoptosis, compared with CD34+CD7− myeloblasts. In analyses of the cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of interleukin-8
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(IL-8) and chemokine (C–C motif) ligand 2 (CCL2) genes, suggesting immaturity of these cells.
2. Materials and methods 2.1. Cells A CD34+ myeloblastic MDS cell line (F-36P), which was established from a patient with refractory anemia with excess blasts (RAEB) in transformation [12] and a small fraction of which expressed CD7, was obtained from Riken Cell Bank (Ibaraki, Japan). Blood or marrow samples from 7 patients, which contained both CD34+CD7+ and CD34+CD7− myeloblasts and were collected after obtaining informed consent, were also used. The diagnoses of the patients (French–American–British classification [13]) were 3 RAEB, 2 RAEB in transformation, and 2 AML transformed from MDS. Cytogenetic analysis was successful for all patients, and abnormal karyotypes were detected in 5 (2 RAEB, 2 RAEB in transformation, and 1 AML transformed from MDS). These karyotypes were classified as intermediateand poor-risk groups in 2 and 3 of the 5 patients, respectively, according to the previous categorization [14]. CD34+CD7+ and CD34+CD7− myeloblasts were purified using fluorescenceactivated cell sorting (FACS, Fig. 1A) as described previously [15]. In some experiments, CD34+ cells were purified from the patient samples using magnetic cell sorting [15]. Isolated cells with more than 97% purity were used. This study was approved by the Institutional Review Board of Nippon Medical School. 2.2. Liquid cultures for examining cell proliferation The purified CD34+CD7+ and CD34+CD7− F-36P cells (2 × 104 cells/mL) were cultured in RMPI 1640 medium containing fetal bovine serum (FCS, 5%) and human IL-3 (5 ng/mL, Immuno-Biological Laboratories [IBL], Gunma, Japan). The viable cell numbers were determined using the trypan blue dye-exclusion test every 3–7 days to maintain cell concentrations appropriate for survival (2 × 104 to 1 × 105 mL−1 ). In single-cell cultures, an irradiated HESS-5 stromal cell layer, needed for single F-36P cells to proliferate, was prepared in 96-well microtiter plates as described previously [15,16]. Purification and automated deposition of a single CD34+CD7+ or CD34+CD7− F-36P cell into each well was performed with FACS. Patient cells (5 × 104 cells/mL), suspended in ␣-MEM containing 12.5% FCS, 12.5% horse serum, and human cytokines (IL-3 [10 ng/mL], thrombopoietin [Kirin Brewery, Takasaki, Japan, 50 ng/mL], stem cell factor [IBL, 50 ng/mL], and Flk2 ligand [IBL, 50 ng/mL]), were cultured on irradiated HESS-5 stromal cell layers. 2.3. Colony-forming assay MethoCult H4230 and GF H4434 methylcellulose media (StemCell Technologies, Vancouver, BC, Canada) were used for F-36P and patient cells, respectively [15]. IL-3 (10 ng/mL) was added to the former. Colonies (aggregates of 20 or more cells) were scored on day 14 of culture.
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2.4. Analyses of immunophenotypes, cell cycle, and apoptosis Multicolor FCM was used [9,17]. The antibodies included anti-CD45-peridin chlorophyll, anti-CD34-fluorescein isothiocyanate (FITC), anti-CD13-phycoerythrin (PE), anti-CD33-PE (BD PharMingen, San Diego, CA, USA), and anti-CD7-PE (BeckmanCoulter, Marseille, France). For cell cycle analysis, cells were stained with anti-CD7-PE, fixed in 70% cold ethanol, and stained with PI [18]. In experiments analyzing apoptosis, F-36P cells were or were not exposed to cytarabine (Ara-C, 0.13 M) for 6 h, washed, and then cultured for 3 days in RMPI 1640 medium containing 5% FCS and human IL-3 (5 ng/mL). Meanwhile, patient cells were cultured for 12 h in ␣-MEM containing 12.5% FCS, 12.5% horse serum, and various human cytokines (10 ng/mL of IL-3 and 50 ng/mL of thrombopoietin, stem cell factor and Flk2 ligand) with or without Ara-C (0.13 M). This shorter culture period was used due to the fragility of fresh MDS cells. These cells were stained with antiCD7-PE, annexin V-FITC, and propidium iodide (PI) (Trevigen, Gaithersburg, MD, USA) for detecting apoptosis [19]. 2.5. Gene microarray RNA from purified CD34+CD7+ and CD34+CD7− F-36P cells was analyzed with microarray hybridization [20]. Total RNA (5 g) was reverse-transcribed using an oligo-dT primer containing a T7 RNA polymerase promoter sequence to synthesize the first-strand cDNA. The double-stranded cDNA that contained the T7 RNA polymerase promoter at the 3 end was transcribed into cRNA by T7 RNA polymerase with allylamine-derivative nucleotides. cRNA was then reacted with N-hydroxy succinimide esters of Cy3 or Cy5 (PerkinElmer, Foster City, CA, USA). Dye molecules were separated from labeled products using the RNeasy mini kit (Qiagen, Valencia, CA, USA). The mixture was then applied to the microarray (Hitachi Human Oligo DNAchip, Hitachi Life Science, Saitama, Japan) and hybridization was carried out for 17 h at 45 ◦ C. After hybridization, slides were washed and scanned using a confocal laser scanner (PerkinElmer). Fluorescence intensities on scanned images were quantified, corrected for background, and normalized using global normalization methods based on the assumption that the median value of fluorescence intensities of both samples should be the same. The DNAchip used covers 1606 genes of various functional classes, including cytokines/growth factors and their receptors, apoptosis regulators, oncogenes, transcription factors, signal transducers, transporters, and cell cycle regulators. 2.6. Real-time quantitative PCR Total RNA from purified CD34+CD7+ and CD34+CD7− F-36P cells was transcribed into cDNA using Super Script FirstStrand Synthesis System for real-time PCR (Invitrogen, Carlsbad, CA, USA). Real-time PCR reactions were performed using the Gene Amp 5700 Sequence Detection System (Applied Biosystems [ABI], Foster City, CA, USA) as described previously [21]. The following Assays on-Demand probes (ABI) were used: actin, Hs 99999903 m1; IL-8, Hs 00174103 m1; and CCL2, Hs 00234140 m1. The relative standard curve experiments were used to measure the expression level of the 2 target genes relative to the -actin gene expression level.
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2.7. Statistics The two-sided Wilcoxon signed-rank test was used.
3. Results First, purified CD34+CD7+ and CD34+CD7− F-36P cells were compared for their growth potential. Compared with CD34+CD7− F-36P cells, CD34+CD7+ F-36P cells proliferated more rapidly in liquid cultures (Fig. 1B) and formed more colonies in semisolid cultures (Fig. 1C). Consistent with these findings, CD34+CD7+ F-36P cells had
fewer G0/G1-phase and more G2/M-phase cells compared with CD34+CD7− F-36P cells (Fig. 1D). Furthermore, CD34+CD7+ F-36P cells had fewer apoptotic cells compared with CD34+CD7− F-36P cells (Fig. 1E, without Ara-C). Finally, the addition of cytarabine to the cultures increased apoptosis in CD34+CD7− F-36P cells but not in CD34+CD7+ F-36P cells (Fig. 1E). Although CD34+CD7+ F-36P cells showed a proliferation advantage, they never dominate in cultures of unseparated F36P cells. Therefore, CD34+CD7+ F-36P cells may generate CD34+CD7− F-36P cells in culture. To examine this possibility, CD34+CD7+ and CD34+CD7− F-36P cells were subjected to single-cell culture. Considerable cell prolifera-
Fig. 1. A distinct difference was seen in cell characteristics of CD34+CD7+ and CD34+CD7− F-36P cells. (A) Purification of CD34+CD7+ and CD34+CD7− F-36P cells using FACS. (B) CD34+CD7+ F-36P cells (black circles) proliferated more rapidly than CD34+CD7− F-36P cells (white circles) in liquid cultures. Data are means of duplicate cultures of a representative experiment (S.D. values were too small to be shown here). Eight separate experiments showed essentially the same results. (C) CD34+CD7+ F-36P cells (black bar) formed more colonies than CD34+CD7− F-36P cells (white bar) in semisolid cultures (P < 0.05). Data are mean ± S.D. of four separate duplicate cultures. (D) Cell cycle status of CD34+CD7+ and CD34+CD7− F-36P cells. The upper panels show representative analyses of the cell cycle and the lower panels show a comparison of cell cycle between CD34+CD7+ (black bars) and CD34+CD7− (white bars) F-36P cells. Data are mean ± S.D. of six separate experiments in which CD7+ and CD7− cells were gated in FCM of unseparated F-36P cells in exponential cell growth. Asterisks indicate significant differences between black and white bars (P < 0.05). Experiments using the purified CD34+CD7+ and CD34+CD7− F-36P cells showed essentially the same results. (E) Apoptosis of CD34+CD7+ and CD34+CD7− F-36P cells. The upper left panel shows representative analyses of apoptosis and the lower left panel shows that caspase-3 activation was observed in most annexin V+ cells. The middle and the right panels are comparisons of annexin V+ and PI+ cells between CD34+CD7+ (black bars) and CD34+CD7− (white bars) F-36P cells. Data are mean ± S.D. from eight separate experiments in which CD7+ and CD7− cells were gated in FCM of unseparated F-36P cells. * P < 0.01, ** P < 0.05. Experiments using the purified CD34+CD7+ and CD34+CD7− F-36P cells showed essentially the same results. (F) Single CD34+CD7+ (four bars on the left) and CD34+CD7− (four bars on the right) F-36P cell cultures in microtiter wells. The numbers of wells, in which cells were immunophenotyped on days 12, 21, and 31 of culture, are shown under the bars. The wells were categorized according to the results of immunophenotyping, i.e., wells in which more than 95% of cells were CD7+ (black), wells in which more than 95% of cells were CD7− (white), and wells in which CD7+ and CD7− cells comprised more than 5%, respectively (spotted). The Y-axis shows the proportion of wells meeting these categories, e.g., the second bar from the left shows that two of 14 wells examined were in the black category and other 12 wells were in the spotted category.
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Table 1 Differences in gene expression profile between CD7-positive and CD7negative F-36P cells determined in microarray analysis Accession no.
Gene name
Ratioa
M17017.1 S71513.1 NM 000129.2 NM 030812.1
Interleukin-8 Chemokine (C–C motif) ligand 2 Coagulation factor XIII, A1 polypeptide Actin-like 8
0.3 0.39 0.45 2.02
a
Gene expression ratios between CD7-positive and CD7-negative F-36P cells (the former data/the latter data).
tion (generating more than 100 cells from a single cell) was observed in approximately one-third of wells plated with single cells. Immunophenotyping of the proliferated cells in each well showed clearly that CD34+CD7− cells were generated in most wells plated with single CD34+CD7+ cells (Fig. 1F, left). Meanwhile, few CD34+CD7+ cells were generated in wells plated with single CD34+CD7− cells (Fig. 1F, right). In microarray analysis, four genes (for actin-like 8, IL8, CCL2 and coagulation factor XIII, A1 polypeptide) showed a greater than 2-fold difference in expression between CD34+CD7+ and CD34+CD7− F-36P cells (Table 1) (data below the threshold for detection were excluded). The expression of the actin-like 8 gene was increased and that of the other three was decreased in CD34+CD7+ F-36P cells compared
Fig. 2. A difference was seen in IL-8 and CCL2 gene expressions between CD34+CD7+ and CD34+CD7− F-36P cells determined using real-time PCR. Gene expression levels of IL-8 and CCL2 relative to that of -actin are shown (data are mean ± S.D. of three separate experiments, with each assay performed in triplicate). * P < 0.01.
with CD34+CD7− F-36P cells. We confirmed the lower IL8 and CCL2 gene expressions in CD34+CD7+ F-36P cells using real-time PCR (Fig. 2). Although CD7-mediated signals induced GM-CSF production in one report [4], there was no difference in GM-CSF gene expression and GM-CSF content in the culture supernatant between CD34+CD7+ and CD34+CD7− F-36P cells (data not shown).
Fig. 3. CD34+CD7+ myeloblasts have advantages in growth and avoiding apoptosis over CD34+CD7− myeloblasts in patients. (A) Purified CD34+CD7+ myeloblasts (black circles) proliferated more rapidly than purified CD34+CD7− myeloblasts (white circles) in liquid cultures. Data are mean ± S.D. of duplicate cultures. (B) Purified CD34+CD7+ myeloblasts (black bars) formed more colonies than purified CD34+CD7− myeloblasts (white bars) in semisolid cultures. Data are mean ± S.D. of duplicate cultures. (C) Apoptosis of CD34+CD7+ (black bars) and CD34+CD7− (white bars) myeloblasts, which were or were not exposed to Ara-C. The two cell populations were gated in FCM of the purified CD34+ cells. Data for the annexin V+ and PI+ cells are shown.
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Finally, we examined CD34+CD7+ and CD34+CD7− myeloblasts from patients. CD34+CD7+ myeloblasts again showed a growth advantage over CD34+CD7− myeloblasts in liquid cultures and colony-forming assays (Fig. 3A and B). Furthermore, CD34+CD7+ myeloblasts had fewer apoptotic cells compared with CD34+CD7− myeloblasts in liquid cultures in either the presence or the absence of Ara-C (Fig. 3C).
4. Discussion Although several clinical observations indicate that CD7+ myeloblasts are associated with poor prognosis in myeloid malignancies, the biological background explaining this notion was not known. Using the F-36P MDS cell line and fresh samples from patients, this study for the first time showed that CD34+CD7+ myeloblasts were more aggressive in terms of proliferation and avoiding apoptosis compared with CD34+CD7− myeloblasts in MDS. Based on these data, we speculate that the aggressive clinical behavior in CD7 + MDS patients is due to the cell characteristics of CD7+ myeloblasts rather than an epiphenomenon. In analyses of the F-36P cell line, CD34+CD7+ myeloblasts produced CD34+CD7− myeloblasts and showed lower expressions of IL-8 and CCL2 genes compared with CD34+CD7− myeloblasts. Interestingly, it was reported that IL-8 and CCL2 production increased with cell maturation in AML blasts and that CCL2 was involved in apoptosis induction [22–24]. It was hypothesized that CD7+ myeloblasts are more immature than CD7− myeloblasts, because CD7+ cases are more common in immature leukemia such as AML-M0 and stem cell leukemia [25,26]. Taken together, all of the above findings suggest that CD34+CD7+ myeloblasts are more immature than CD34+CD7− myeloblasts. Do CD34 + CD7 + MDS myeloblasts have stem cell potential? In our previous study, CD34+ cells including CD34+CD7+ cells from MDS patients lacked long-term culture-initiating cell activity and did not reconstitute hematopoiesis in NOD/Shi-scid/scid (NOD/SCID) mice [15]. However, because MDS stem cells are difficult to detect, probably due to the instability of MDS cells [27], the definitive answer to this question remains unknown. This topic is especially interesting in view of the fact that agents targeting leukemic stem and progenitor cells have been reported recently [28,29]. It is unknown whether currently available therapies, such as demethylating agents and stem cell transplantation for eligible patients, improve the prognosis of CD7 + MDS patients. We recommend that the status of CD7 expression on myeloblasts be examined in clinical trials of such therapies. Furthermore, development of a therapy specific for patients with CD34+CD7+ myeloblasts might be beneficial. In summary, the data obtained here can explain the aggressive clinical behavior of CD7 + MDS cases. Monitoring and targeting of CD34+CD7+ myeloblasts are worth considering in MDS.
Acknowledgments This work was supported in part by a Grant-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (No. 17591015). Contributions: C.S. performed experiments, analyzed data and wrote the manuscript. H.T. designed the research. T.Y. and T.T supported technical aspect. K.D. provided administrative and clinical support. K.O. designed the research, analyzed data, and wrote the manuscript.
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