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Stem Cells 2005;23:619-630 www.StemCells.com
© 2005 AlphaMed Press

Identification and Hematopoietic Potential of CD45– Clonal Cells with Very Immature Phenotype (CD45–CD34–CD38–Lin–) in Patients with Myelodysplastic Syndromes

Kiyoyuki Ogataa, Chikako Satoha,b, Mikiko Tachibanaa, Hideya Hyodoa, Hideto Tamuraa, Kazuo Dana, Takafumi Kimurac, Yoshiaki Sonodad, Takashi Tsujie

a Division of Hematology, Third Department of Internal Medicine, and
b Department of Bioregulation, Nippon Medical School, Tokyo, Japan;
c Department of Molecular-Targeting Cancer Prevention, Graduate School of Medical Science, Kyoto Prefectural University of Medicine, Kyoto, Japan;
d Department of Hygiene, Kansai Medical University, Osaka, Japan;
e Department of Industrial Science and Technology, Tokyo University of Science, Chiba, Japan

Key Words. Myelodysplastic syndromes • CD45 • Hematopoietic stem cells

Correspondence: Kiyoyuki Ogata, M.D., Division of Hematology, Third Department of Internal Medicine, Nippon Medical School, 1-1-5 Sendagi, Bunkyo-ku, Tokyo 113-8603, Japan. Telephone: 81-3-3822-2131; Fax: 81-3-5685-1793; e-mail: ogata{at}nms.ac.jp


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
CD45 is a hematopoietic lineage-restricted antigen that is expressed on all hematopoietic cells except for some mature cell types. Cells expressing CD45 and CD34 but lacking CD38 and lineage antigens (CD45+CD34+CD38–Lin– cells) are well-documented hematopoietic stem cells (HSCs), and CD45+CD34–CD38–Lin– cells are probably less mature HSCs. In myelodysplastic syndromes (MDS), the malignant transformation site is a matter of debate, and CD45+CD34+CD38–Lin– HSCs were recently reported to be clonal. In the study reported here, we detected CD45–CD34–CD38–Lin– cells in the peripheral blood and bone marrow of patients with MDS and isolated them by successive application of density centrifugation, magnetic cell sorting, and fluorescence-activated cell sorting. Fluorescence in situ hybridization showed that CD45–CD34–CD38–Lin– cells had the same chromosomal aberration as the myeloblasts. In addition to CD45– and CD34–, they lacked CD117 and CD133 expression. Generally, MDS cells have extremely reduced hematopoietic potential compared with normal hematopoietic cells, but we documented the following in some patients. Freshly isolated CD45–CD34–CD38–Lin– cells did not form any hematopoietic colonies but had long-term culture-initiating cell activity. When cocultured with stroma cells, CD45–CD34–CD38–Lin– cells showed only weak potential for proliferation and differentiation, yet they differentiated into CD34+ cells and then mature myeloid cells. This newly identified cell population represents the most immature immunophenotype so far identified in the hematopoietic lineage and is involved in the malignant clone in MDS.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Hematopoietic stem cells (HSCs) are rare cell populations that are capable of self-renewal and blood cell production [1]. Therefore, they maintain hematopoiesis throughout life. Further, HSCs might be able to differentiate into cells of other tissues such as muscle cells [2]. Cells that express CD45 and CD34 but lack CD38 and lineage antigens (CD45+CD34+CD38–Lin–) are a well-documented HSC population [3, 4]. Recent data from multiple groups have indicated that cells which express CD45 but lack expression of CD34, CD38, and lineage antigens (CD45+CD34–CD38–Lin–) are probably less mature HSCs than are CD45+CD34+CD38–Lin– HSCs [5, 6]. CD45 is a hematopoietic lineage-restricted cell-surface marker that is expressed on all hematopoietic cells, from HSCs to mature blood cells, except for erythroid cells, platelets, and plasma cells, which lose this antigen during maturation [7].

Myelodysplastic syndromes (MDS) are hematological neoplasms in which neoplastic myeloid cells (i.e., neutrophilic, monocytic, megakaryocytic, and erythroid cells) proliferate in the bone marrow (BM). There is a hypothesis that the MDS malignant transformation occurs at the committed myeloid progenitor cell level [8], but recent data suggest that the transformation occurs in CD45+CD34+CD38– HSCs in MDS [9]. The neoplastic hematopoietic cells in MDS have various degrees of defective differentiation capability in each patient, and thus the percentage of immature blast cells in the BM differs among patients. MDS is classified into several subgroups, based mainly on the blast percentages in the BM and peripheral blood (PB) [10]. During the clinical course, patients with MDS often show a transition from the original MDS subtype to another subtype with a higher blast percentage and, finally, to secondary acute myeloid leukemia (AML) [11]. Recently, based on the findings that immature cells are lighter than mature cells [12], a new density-centrifugation method for enriching blastoid immature cells from PB and BM samples was developed [13, 14]. In a prior study, we used this method to prepare blast-rich MDS specimens for immunophenotyping and found that MDS blasts have the immunophenotype of committed myeloid precursors (CD45+CD34+CD38+CD13+CD33+) [15]. Further, we showed that, in accordance with disease progression (increase in blast percentage), the phenotype of MDS blasts became more immature (e.g., gain of CD7 and c-kit expression), at least in some patients [15, 16].

Here we report detection of CD45–CD34–CD38–Lin– blastoid cells having a chromosomal aberration in the PB and BM samples of MDS. These cells were detected in the advanced disease stages of MDS. The freshly isolated CD45–CD34–CD38–Lin– cells did not form any hematopoietic colonies but differentiated into hematopoietic colony-forming cells and fully mature myeloid cells when cultured together with murine stroma cells. This newly identified cell population has the most immature immunophenotype so far identified in the hematopoietic lineage and is involved in the MDS clone.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Subjects
Patients with MDS (either of two subtypes: refractory anemia with excess blasts [RAEB] or RAEB in transformation [RAEB-t]) or acute leukemia transformed from MDS (AL-MDS), diagnosed according to the French-American-British criteria [10, 17], were the subjects of this study. Patients who had previously undergone cytotoxic chemotherapy and those with a secondary MDS were excluded. Cytogenetic analyses were performed using standard G-banding with trypsin-Giemsa staining. Karyotypes were interpreted using the International System for Cytogenetic Nomenclature criteria [18].

Blastretriever (BR) Density Centrifugation
Heparinized BM cells, which were aspirated for clinical diagnosis, and heparinized PB were obtained from the patients. The study was approved by the Institutional Review Board of Nippon Medical School, and informed consent was obtained from all subjects. Samples were enriched for immature blastoid cells by density-gradient centrifugation using BR (Japan Immunoresearch Laboratories Co., Takasaki, Japan, http://www.jimro.com) according to the manufacturer’s instructions, as reported previously [13–16]. The blastoid cell–enriched samples were subjected to flow cytometry (FCM) to detect CD45–CD34–CD38–Lin– cells. The cell differential of these samples was determined for cytospin preparations (after Wright-Giemsa staining, 100 nucleated cells were examined for each cytospin).

FCM
Immunophenotyping was performed by three-color FCM, in which the optimal quantity of antibodies to be used was determined in preliminary experiments and each cell population was gated by a CD45 gating method, as described previously [15, 19, 20]. In brief, the cells were stained with anti-CD45 antibody labeled with peridin chlorophyll (PerCP) (clone 2D1; Becton, Dickinson, San Jose, CA, http://www.bd.com), and pairs of antibodies were conjugated with either fluorescein isothiocyanate (FITC) or phycoerythrin (PE). These antibodies were directed to CD3, CD11b, CD13, CD15, CD16, CD19, CD34, CD41a, CD44, HLA-class I, HLA-DR (FITC- or PE-conjugated; Becton, Dickinson); CD2, CD7, CD33, CD38, CD56 (FITC- or PE-conjugated; Pharmingen, San Diego, http://www.bdbiosciences.com/pharmingen); glycophorin A (GPA), CD10, CD117 (FITC- or PE-conjugated; Beckman Coulter, Fullerton, CA, http://www.beckman.com); and CD133 (PE-conjugated; Miltenyi Biotec, Bergisch Gladbach, Germany, http://www.miltenyibiotec.com). An antibody against vascular endothelial growth factor receptor 2 (KDR) (Immuno-Biological Laboratories Co., Takasaki, Gunma, Japan, http://www.ibl-japan.co.jp) was also used, and in this case an FITC-conjugated second antibody was used. Anti-CD45 antibodies conjugated with FITC or PE-Cy7 (clone HI30; Pharmingen), which recognize a different epitope from the above-mentioned anti-CD45 antibody, were also used to confirm the CD45– of cells and to sort cells. Single-labeled cells were used to compensate for fluorescence emission overlap of each fluorochrome into inappropriate channels. Isotype-matched negative controls were used in all assays. At least 30,000 events were acquired for most samples after the BR density centrifugation. Analysis was performed on a FACScan and a FACSVantage (Becton, Dickinson).

Isolation of CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– Blastoid Cells
We applied magnetic cell sorting (MACS) and fluorescence-activated cell sorting (FACS) to purify cells, as reported previously [6, 21, 22]. The cell samples, which had been subjected to BR density centrifugation and immunophenotyped using their aliquots, were subjected to MACS. The antibody-bound magnetic colloids used included CD3, CD15, CD33, CD34, CD56, CD61, and GPA (Miltenyi Biotec). The kind of magnetic colloid used for each case was determined on the basis of the immunophenotype of the CD45– cells. The magnetic colloid–bound cells (positive fraction) were separated from the unbound cells (negative fraction) on a MACS column (Miltenyi Biotec). Aliquots of both fractions were again subjected to immunophenotyping by FCM before FACS. When the antibody bound to the magnetic colloid used for MACS and the antibody used for FCM and FACS were directed against the same molecule, we confirmed beforehand that the antibodies did not interfere with each other under the experimental conditions (e.g., anti-CD34 antibodies for MACS and FCM [and FACS] recognized class II and III antigens of CD34, respectively, and no interference was observed in preliminary experiments).

Based on the immunophenotype data, cells in the negative fraction were stained with a combination of PE-conjugated antibodies (i.e., CD34, CD38, and lineage antibodies) and FITC-conjugated CD45 (Becton, Dickinson), and then CD45–CD34–CD38–Lin– cells were sorted using an EPICS ALTRA (Beckman Coulter). In some cases, cells in the negative fraction were stained with PE-conjugated antibodies (i.e., CD34, CD38, and lineage antibodies), FITC-conjugated other lineage antibodies, and PE-Cy7-conjugated CD45, and then CD45–CD34–CD38–Lin– cells were sorted using a FACSVantage (Becton, Dickinson). CD34+ myeloblasts were sorted from the positive fraction after staining with PE-conjugated CD34 and FITC-conjugated CD45 (or PE-Cy7-conjugated CD45). Similarly, CD45–CD34+CD38–Lin– cells were obtained from the positive fraction using appropriate antibody combinations.

Cell viability, determined by trypan blue dye exclusion, was at least 95% in all purified cell fractions.

Fluorescence In Situ Hybridization (FISH)
Isolated cells were subjected to simultaneous morphological and FISH analyses as described previously [14, 23]. In brief, microscopic images of Giemsa-stained cells on cytospin glass slides were saved in a computer, and the location of the cells was recorded. Then the slides were treated with 70% ethanol for a few seconds, 75 mmol/L KCl for 10 minutes, and Carnoy’s fixation solution for 5 minutes. Next, the slide was treated with trypsin digestion (0.005% trypsin in phosphate-buffered saline [PBS], pH 7.5) for 10 minutes at 20°C, washed with PBS, dehydrated with 70%, 85%, and 100% ethanol, immersed in 2x standard saline citrate solution containing 0.1% Nonidet P-40, and dehydrated through 70%, 85%, and 100% ethanol. The probe was designed to hybridize the centromere region of chromosomes X, Y, and 7, respectively (CEP X, CEP Y, and CEP 7) (Vysis, Downers Grove, IL, http://www.vysis.com). Denaturation, hybridization, and posthybridization wash were performed according to the manufacturer’s instructions. DAPI (4,6 diamidino-2-phenylindole) was used as a counterstain.

Cell Coculture with HESS-5 Murine Stroma Cells
The hematopoietic-supportive stromal cell line HESS-5 was previously established from murine BM [24], and its application to hematopoietic cell cultures was reported [21, 22]. HESS-5 cells were maintained in minimum essential medium (MEM) supplemented with 10% horse serum at 37°C under 5% CO2 in humidified air. We prepared irradiated HESS-5 cell layers in microtiter 96-well plates, 24-well plates, and 35-mm2 plates and then cultured the isolated CD45–CD34–CD38–Lin– cells, CD45–CD34+CD38–Lin– cells, or CD34+ myeloblasts on the layers at 37°C under 5% CO2 in humidified air. The kind of plate used was decided based on the available cell number (e.g., 1 x 103 cells per 96-well plate and 3 x 104 cells per 35-mm2 plates). The culture medium was MEM supplemented with 12.5% horse serum, 12.5% fetal calf serum (FCS), and various combinations of human cytokines. The final concentrations of cytokines were as follows: interleukin-3 (IL-3), 10 ng/mL; thrombopoietin (TPO), 50 ng/mL; stem cell factor (SCF), 50 ng/mL; Flk2 ligand, 50 ng/mL; vascular endothelial growth factor (VEGF), 10 ng/mL. IL-3, SCF, and TPO were provided by the Kirin Brewery Co. (Tokyo, http://www.kirin.co.jp/english). Flk2 ligand and VEGF were purchased from Immuno-Biological Laboratories. After an appropriate interval (3–7 days, depending on cell growth), half of the volume of the culture medium was harvested and replaced with fresh medium as follows. The cells in the harvested medium were collected by centrifugation, suspended in fresh medium, and returned to the culture. When cell growth had become extensive, the cell culture was scaled up. When the cultured cells were analyzed, all cells were harvested by vigorous pipetting and washed in PBS before use.

In Vitro Colony-Forming Assay and Long-Term Culture-Initiating Cell (LTC-IC) Assay
The colony-forming potential of the freshly isolated CD45–CD34–CD38–Lin– cells, CD45–CD34+CD38–Lin– cells, and CD34+ myeloblasts was examined in a methylcellulose medium (MethoCult GF H4434; Stem Cell Technologies, Vancouver, British Columbia, Canada, http://www.stemcell.com) as described previously [21, 22]. The medium was supplemented with optimal concentrations of human recombinant cytokines such as IL-3 (10 ng/mL), SCF (50 ng/mL), granulocyte-macrophage colony-stimulating factor (10 ng/mL), and erythropoietin (3 U/mL). MethoCult cultures were incubated at 37°C in a humidified atmosphere of 5% CO2, and colonies were scored after 14 days of culture.

To assess the LTC-IC activity, the CD45–CD34–CD38–Lin– cells and CD34+ myeloblasts that had been cultured with HESS-5 cells for 5 weeks were similarly examined for their colony-forming potential.

SCID-Repopulating Cell Assay by the Intra-BM Injection (IBMI) Method
The animal experiments were approved by the Animal Care Committee of Kyoto Prefectural University of Medicine and carried out at that university. Severe combined immunodeficient (SCID)–repopulating cell assay by the IBMI method was performed as reported previously [6, 25]. Five-week-old nonobese diabetic/severe combined immunodeficient (NOD/SCID) mice (Central Institute for Experimental Animals, Kawasaki, Japan) were handled under sterile conditions and maintained in germ-free isolators located in the Central Laboratory Animal Facility. Purified cells were transplanted by IBMI into sublethally irradiated (250 cGy) 8- to 12-week-old mice. Briefly, after sterilization of the skin around the left knee joint, the knee was flexed to 90 degrees, and the proximal side of the tibia was drawn to the anterior. A 27-gauge needle was inserted into the joint surface of the tibia through the patellar tendon and then inserted into the BM cavity. Using a Hamilton microsyringe, cells suspended in 10 µL of {alpha}-medium were carefully injected via the bone hole into the BM cavity. The mice were killed 10–14 weeks after transplantation, and the BMs from the bilateral femurs, tibiae, and humeri were flushed out using {alpha}-medium containing 10% FCS. The presence of human cells was analyzed by FCM. Mice were scored as positive if more than 0.1% of the total murine BM cells were positive for human CD45. In some experiments, the mice received an intraperitoneal injection of 20 µL of anti-asialo GM1 antiserum (Wako, Osaka, Japan, http://www.wako-chem.co.jp) in 400 µL of PBS to suppress natural killer cell activity. CD45+CD34–Lin– cells, which were obtained from cord blood as described previously [6], were used as a positive control.

Statistical Analyses
Differences between three or more groups of data of continuous variables were analyzed by one-way analysis of variance. Differences in categorical variables were evaluated using the chi-square test. A p value of less than .05 was considered to be statistically significant.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Detection of CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– Clonal Cells in MDS
Figure 1Go shows a representative example of flow cytometric analysis of a sample after BR density centrifugation. In the CD45 versus side scatter (SSC) display, we detected a cell cluster that lacked CD45 expression and had low SSC (R3 in Fig. 1BGo). The CD45– was confirmed by using another antibody that recognizes a different epitope of the CD45 molecule. The forward scatter (FSC) showed that the size of cells in R3 ranged from lymphocyte size to myeloblast size, but cells smaller than myeloblasts were predominant (Fig. 1DGo). Immunophenotyping of the cells in R3 revealed that, in addition to the CD45–, the majority of cells were negative for hematopoietic lineage antigens (CD2, CD3, CD10, CD11b, CD13, CD15, CD16, CD19, CD20, CD33, CD41a, CD56, and GPA) and stem cell–related antigens (CD34, CD38, CD117, and CD133). They were negative for HLA-DR and KDR but positive for HLA-class I antigen and CD44. Only minor subpopulations of the cells in R3 were weakly positive for CD7, CD34, CD38, and CD133. Therefore, in this case the dominant cells in R3 were CD45–CD34–CD38–Lin–. Part of the flow cytometric immunophenotyping for CD45– cells as well as myeloblasts is shown in Figure 1EGo (CD45– cells, blue dots; myeloblasts [cells in R2], gray dots). The cardinal data of the antigen profiles for these two cell populations are presented in Table 1Go (patient 1) and clearly differ considerably between CD45– cells and myeloblasts. Figure 2Go shows the plots for another patient, and it is seen that one third of the CD45– cells (cells in R3, Fig. 2BGo) expressed CD34 and a few expressed CD38 and myeloid antigens (blue dots in Fig. 2CGo). Expression of other hematopoietic lineage antigens was sparse. Therefore, for this patient the dominant cells in R3 were CD45–CD34+CD38–Lin–. Again, the antigen profile differs considerably between the CD45– cells and myeloblasts (gray dots in Fig. 2CGo) in this patient (Table 1Go, patient 8).



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  		Figure 1. Representative example of CD45–CD34–CD38–Lin– cell detection in a sample of myelodysplastic syndromes. (A): Forward scatter (FSC) versus side scatter (SSC) display of bone marrow cells after blastretriever density centrifugation (patient 1 of Table 1Go). (B): CD45 versus SSC display of the cells gated by R1 in panel A. The bold vertical line, the left side of which shows CD45–, was obtained from panel C. R2, R3, and R4 indicate myeloblasts, CD45– cells, and lymphocytes, respectively (the immunophenotype data for myeloblasts and CD45– cells are shown in Table 1Go). (C): The cells were stained with isotype-matched control immunoglobulin G (IgG) conjugated with peridin chlorophyll (PerCP). (D): Cell size of the cells gated by R3 in panel B (FSC versus SSC display). Similar results were obtained when CD7+ or CD38+ cells were excluded from the analysis. (E): Part of antigen-expression analysis of myeloblasts (gray dots) and CD45– cells (blue dots). Abbreviations: FITC, fluorescein isothiocyanate; GPA, glycophorin A; PE, phycoerythrin.

 

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  		Table 1. Phenotypic characteristics of myeloblasts and CD45– cells
 


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  		Figure 2. Representative example of CD45–CD34+CD38–Lin– cell detection in a sample of myelodysplastic syndromes. (A): Forward scatter (FSC) versus side scatter (SSC) display of bone marrow cells after blastretriever density centrifugation (patient 8 in Table 1Go). (B): CD45 versus SSC display of the cells gated by R1 in panel A. The left side of the bold vertical line shows CD45–. R2 and R3 indicate myeloblasts and CD45– cells, respectively. (C): Part of antigen-expression analysis of myeloblasts (gray dots) and CD45– cells (blue dots). Abbreviations: FITC, fluorescein isothiocyanate; PE, phycoerythrin; PerCP, peridin chlorophyll.

 
In total, we examined BR-treated samples from 60 patients by FCM. The BR treatment enriches immature blastoid cells and depletes other cells. However, when original samples contain large numbers of erythroblasts (Ebls) that are CD45–, the BR treatment often cannot deplete Ebls to a level that does not interfere with CD45–CD38–Lin– cell detection. For this reason, we omitted nine of the 60 patients from the analyses. In eight of the remaining 51 patients, we detected CD45–CD38–Lin– cells in the CD45– cell clusters, as exemplified in Figures 1Go and 2Go. Tables 1Go and 2Go show the characteristics of these eight patients and their phenotypic data for CD45– cells and myeloblasts. In all patients, CD45– cells almost completely lacked expression of myeloid antigens (CD11b, CD13, CD15, and CD33) and lymphoid-specific antigens (CD2, CD3, CD10, and CD19). Compared with the myeloblasts in each patient, CD45– cells had much lower expression of stem cell–related antigens (CD34, CD38, CD117, and CD133), but CD34 expression on CD45– cells varied greatly among the patients. MDS myeloblasts often express CD7 and/or CD56 aberrantly. The expression of these antigens was reduced on CD45– cells in all patients whose myeloblasts expressed them, but in three patients (patients 1, 4, and 7) more than 10% of CD45– cells expressed CD7 weakly. CD45–CD38–Lin– cells were generally smaller than myeloblasts in all patients. The ratio of the cell number between CD45– cells and myeloblasts, which was determined by FCM ([cells in R3]/[cells in R2] x 100), ranged from 3.1%–26.1% in these eight patients (median 6.5%). Interestingly, CD45–CD38–Lin– cells were detected not only in BM samples but also in PB samples (Table 1Go). For patients 1 and 6, we obtained both BM and PB samples and detected CD45–CD38–Lin– cells in both samples. As shown in Table 3Go, a CD45– cell cluster that contained CD45–CD38–Lin– cells was detected only in RAEB-t and AL-MDS patients, and not in any of the RAEB patients, even though the percentages of enriched blastoid cells and contaminating Ebls in the analyzed samples were similar in all three disease groups.


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  		Table 2. Characteristics of patients in whose samples CD45–CD38–Lin– cells were detected
 

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  		Table 3. Relationships between disease subtypes and CD45–CD38–Lin– cell detection
 
Next, we isolated the CD45–CD34–CD38–Lin– cells from six of the eight patients and CD45–CD34+CD38–Lin– cells from two of the eight patients by successive application of BR density centrifugation, MACS, and FACS. CD45+CD34+ cells expressing myeloid antigens (CD34+ myeloblasts) were similarly isolated from the same patients as control cells. The purity of the isolated cells was at least 98% when assessed by FCM (Fig. 3AGo). The isolated CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– cells were blastoid cells that had scanty cytoplasm with no granules (Fig. 3BGo). Based on the findings that CD45–CD38–Lin– blastoid cells appeared after the disease stage of MDS had progressed (RAEB-t and AL-MDS) and were present not only in BM but also in PB, it is highly probable that these cells were clonal in origin. We performed FISH analysis of purified CD45–CD38–Lin– cells and CD34+ myeloblasts in four of the eight patients and confirmed that CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– cells had the same chromosomal aberration as the myeloblasts in each patient (Fig. 3CGo and Table 4Go).



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  		Figure 3. Isolation of CD45–CD34–CD38–Lin– cells and their morphological and cytogenetic analyses. (A): The top panel shows the CD34 versus CD45 display of myelodysplastic syndromes (MDS) cells after blastretriever (BR) density centrifugation. The middle two panels show the cells after magnetic cell sorting (MACS) treatment. The rectangles are gates for fluorescence-activated cell sorting (FACS). The bottom two panels show the isolated CD34+ myeloblasts (right) and CD45–CD34–CD38–Lin– cells (left) after FACS. In each MDS patient, the antibody-coated colloids used for MACS and the antibodies used for FACS were selected based on the immunophenotypes of CD45– cells. (B): Isolated CD45–CD34–CD38–Lin– cells (Wright-Giemsa stain). (C): Isolated CD45–CD34–CD38–Lin– cells from patient 4 were subjected to Giemsa staining (left) and then to fluorescence in situ hybridization (right). The red spot and green spots show X- and Y-chromosome signals. Abbreviations: FITC, fluorescein isothiocyanate; PE, phycoerythrin; PerCP, peridin chlorophyll.

 

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  		Table 4. Results of fluorescence in situ hybridization analysis
 
Proliferation and Differentiation of CD45–CD34–CD38–Lin– Cells Cultured with HESS-5 Stroma Cells
We previously established a murine BM stromal cell line, HESS-5 [24], and showed that in combination with cytokines it dramatically supported the proliferation and differentiation of normal human HSCs, including CD45+CD34+38–Lin– cells and more immature CD45+CD34–CD38–Lin– cells [21, 22, 26]. Thus, we examined whether HESS-5 and cytokines could expand and/or differentiate the newly identified CD45–CD34–CD38–Lin– cells of MDS. First, we cultured CD45–CD34–CD38–Lin– cells and CD34+ myeloblasts from one patient (patient 4) under various conditions. We confirmed that cytokines alone were much inferior to HESS-5 plus cytokines in expanding and/or differentiating these cells (data not shown). Among the examined cytokine combinations, a combination of IL-3, TPO, SCF, and Flk2 ligand was the most suitable for CD45–CD34–CD38–Lin– cell culture (Fig. 4AGo). Therefore, using this cytokine combination and HESS-5 cells, we cultured CD45–CD34–CD38–Lin– cells and CD34+ myeloblasts, which had been isolated from six patients, for up to 5 weeks and examined the time-course changes in the number, morphology, and immunophenotypes of the cells in the cultures. When normal CD34+ myeloblasts obtained from granulocyte colony-stimulating factor (G-CSF)–mobilized PB (normal control) were cultured under the present culture conditions, the cell number had increased 370-fold (mean of four experiments) on day 7 of the culture. In contrast, when CD34+ myeloblasts from six MDS patients were cultured, the increase in cell number was much less. In two patients (patients 2 and 3), almost all the cultured cells died during the first 7–10 days of culture. The data for the other four MDS patients are shown in Figure 4BGo. Among these four patients, the highest increase in cell number on day 7 was 50-fold (patient 4; the solid circles in Fig. 4BGo). These inferior results in MDS were expected because we and others had documented that the expansion and differentiation abilities of HSCs and hematopoietic progenitors were often markedly reduced in MDS patients compared with normal subjects [9, 27–30]. Moreover, the fold increase in the cell number of cultured CD45–CD34–CD38–Lin– MDS cells was much less compared with the cultured CD34+ myeloblasts in MDS. In addition to patients 2 and 3, in one other case (patient 5, the open star in Fig. 4BGo) almost all the cells in CD45–CD34–CD38–Lin– cell culture died during the first 10 days of culture. For the remaining three patients, the cell number decreased during the first 7–10 days, while small clusters of cells (aggregates of two to four cells) began to appear on days 7–10 of culture. These cell clusters later produced more cells. This is in contrast to the CD34+ myeloblast culture, in which cell clusters began to appear on day 3 of culture. The fold increase in cell number was always less in the CD45–CD34–CD38–Lin– cell cultures compared with CD34+ myeloblast cultures for each patient (Fig. 4BGo). Time-course photographs of the cultured cells from a representative patient are shown in Figure 4CGo.



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  		Figure 4. Proliferation and differentiation of CD45–CD34–CD38–Lin– cells cocultured with HESS-5 stroma cells. (A): CD34+ myeloblasts (solid symbols) and CD45–CD34–CD38–Lin– cells (open symbols) obtained from patient 4 were cocultured with HESS-5 cells in the presence of various combinations of cytokines (circles: IL-3, TPO, SCF, and Flk2 ligand; rectangles: IL-3, TPO, SCF, Flk2 ligand, and vascular endothelial growth factor; triangles: TPO, SCF, and Flk2 ligand). (B): CD34+ myeloblasts (solid symbols) and CD45–CD34–CD38–Lin– cells (open symbols) obtained from four patients (shown as circles, rectangles, triangles, and stars) were cocultured with HESS-5 cells in the presence of IL-3, TPO, SCF, and Flk2 ligand. (C): Time-course photographs of the cultured cells from patient 4. On day 7 of culture, a marked increase in cells was observed in the myeloblast culture but not in the CD45–CD34–CD38–Lin– cell culture. Abbreviations: IL-3, interleukin-3; SCF, stem cell factor; TPO, thrombopoietin.

 
In two patients from whom relatively large numbers of cells were obtained for culture, we examined the immunophenotypes of the cultured cells at various time points. Representative results are shown in Figure 5Go. Regarding CD34+ myeloblast culture, the percentage of CD34+ cells in the culture decreased with time and became 0% on day 35 of culture (Fig. 5AGo, lower panel). Throughout the culture period, almost all cells expressed myeloid antigens (data not shown). On day 7 of CD45–CD34–CD38–Lin– cell culture, the CD45–CD34–CD38–Lin– cell percentage had decreased markedly, and CD45+CD34+ cells and CD45+CD34– cells expressing myeloid antigens had appeared (Fig. 5AGo, upper panel). These data, together with the finding that the cell number in CD45–CD34–CD38–Lin– cell cultures decreased during the first 7–10 days while small clusters of cells began to appear on days 7–10, indicate that most CD45–CD34–CD38–Lin– MDS cells could not survive in our culture system, and a minor subpopulation of CD45–CD34–CD38–Lin– cells contributed to the production of CD45+CD34+ cells and CD45+CD34– cells expressing myeloid antigens. On days 15 and 35 of the CD45–CD34–CD38–Lin– cell culture, the percentage of CD45+CD34+ cells had decreased, but a small percentage of CD45+CD34+ cells still existed on day 35. Throughout the culture period, almost all CD45+ cells expressed myeloid antigens (Fig. 5BGo). The findings for Wright-Giemsa–stained cytospin preparations at various time points were consistent with the FCM data and confirmed that both cultures produced myeloid cells in various stages of maturation (Fig. 5CGo). FISH analyses also confirmed that these cells were clonal in origin (data not shown).



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  		Figure 5. Time courses of immunological and morphological characteristics of cultured CD45–CD34–CD38–Lin– cells. CD45–CD34–CD38–Lin– cells and CD34+ myeloblasts were cocultured with HESS-5 cells in the presence of interleukin-3, thrombopoietin, stem cell factor, and Flk2 ligand. (A): Expression of CD45 and CD34 on the cultured cells at various time points. The number in the upper right corner of each dot plot indicates the percentage of CD45+CD34+ cells. (B): Staining with lineage-specific antibodies on day 15 of CD45–CD34–CD38–Lin– cell culture showed most cells expressed myeloid antigens. (C): A Wright Giemsa–stained cytospin on day 15 of CD45–CD34–CD38–Lin– cell culture showed myeloid cells in various stages of maturation. The arrowhead and arrow indicate a neutrophil and a blast, respectively. Abbreviations: FITC, fluorescein isothiocyanate; GPA, glycophorin A; PE, phycoerythrin; PerCP, peridin chlorophyll.

 
Colony-Forming Activity and LTC-IC Activity of CD45–CD34–CD38–Lin– Cells
The results are shown in Figure 6Go. It has been reported that colony-forming activity is often defective in MDS [27, 28]. In the experiments reported here, we observed clear colony formation from freshly isolated CD34+ myeloblasts in only two of the five MDS patients examined. In contrast, freshly isolated CD45–CD34–CD38–Lin– cells did not form any colonies in any of the five MDS patients. Conversely, when these two cell populations from the five MDS patients were cultured with HESS-5 cells for 5 weeks and then examined, LTC-IC activity was detected in the cultured CD45–CD34–CD38–Lin– cells (clear activity in two patients) but not in the cultured CD34+ myeloblasts.



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  		Figure 6. Colony-forming activity and long-term culture-initiating cell (LTC-IC) activity of CD45–CD34–CD38–Lin– cells. (A): Freshly isolated CD45–CD34–CD38–Lin– cells (light columns) and CD34+ myeloblasts (dark columns) were analyzed for colony-forming activity. (B): CD45–CD34–CD38–Lin– cells (light columns) and CD34+ myeloblasts (dark columns) were cocultured with HESS-5 stroma cells for 5 weeks and then analyzed for LTC-IC activity. Data in panels A and B are the mean ± SD of triplicate cultures. Forty colonies from the freshly isolated CD34+ myeloblasts of patient 4 (panel A) were from BFU-E. Other colonies in panels A and B were from granulocyte-macrophage colony-forming units.

 
In Vitro Hematopoietic Potential of CD45–CD34+CD38–Lin– Cells
For patient 4, we compared the in vitro hematopoietic potential among CD45–CD34–CD38–Lin– cells, CD45–CD34+CD38–Lin– cells, and CD34+ myeloblasts. When freshly isolated populations of these cells were cocultured with HESS-5 under the above-mentioned conditions, the proliferation and differentiation kinetics of CD45–CD34+CD38–Lin– cells were similar to those of CD34+ myeloblasts. That is, cell clusters began to appear on day 3 of culture, and the cell number had increased significantly by day 7 of culture (Fig. 7AGo). Further, freshly isolated CD45–CD34+CD38–Lin– cells had clear colony-forming potential (Fig. 7BGo).



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  		Figure 7. In vitro hematopoietic potential of CD45–CD34+CD38–Lin– cells. (A): CD45–CD34+CD38–Lin– cells (gray circles), CD34+ myeloblasts (black circles), and CD45–CD34–CD38–Lin– cells (white circles) obtained from patient 4 were cocultured with HESS-5 cells in the presence of interleukin-3, thrombopoietin, stem cell factor, and Flk2 ligand. (B): Freshly isolated CD45–CD34+CD38–Lin– cells (middle column labeled CD45–CD34+), CD34+ myeloblasts (left column labeled Mbl), and CD45–CD34–CD38–Lin– cells (right column labeled CD45–CD34–) from patient 4 were analyzed for colony-forming activity. Data are the mean ± SD of duplicate cultures. Fifty-three and 19 colonies from CD34+ myeloblasts and CD45–CD34+CD38–Lin– cells, respectively, were from BFU-E. Other colonies were from granulocyte-macrophage colony-forming units.

 
NOD/SCID Repopulating Activity of CD45–CD34–CD38–Lin– Cells
Demonstration of human hematopoiesis in NOD/SCID mice has been used to confirm HSC activity of human cell samples [31]. However, the published data indicate that MDS stem cells seldom or never establish hematopoiesis in NOD/SCID mice [9, 32, 33]. Therefore, in this study we used IBMI of MDS cells to NOD/SCID mice. The IBMI technique detects HSC activity more sensitively compared with the conventional method—intravenous injection of HSCs to NOD/SCID. That is, IBMI, but not the conventional method, can detect HSC activity in CD34– HSCs, and it needs fewer HSCs to reconstitute human hematopoiesis than does the conventional method [6]. Moreover, we treated some NOD/SCID mice with anti-asialo GM1 antiserum to facilitate HSC engraftment. Despite these approaches, we have so far been unable to detect any NOD/SCID repopulation activity in CD45–CD34–CD38–Lin– MDS cells (Table 5Go).


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  		Table 5. Intrabone marrow injection of CD45–CD34–CD38–Lin– cells to nonobese diabetic/severe combined immunodeficient mice
 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Whether HSCs are actually involved in MDS remains controversial [34]. There is a hypothesis that MDS malignant transformation occurs at the committed myeloid progenitor cell level [8]. However, it was recently reported that CD45+CD34+CD38– HSCs are involved in a malignant clone of MDS [9]. In this study, we detected cells with unreported immunophenotypes (CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– cells) and found that these cells were clonal in origin. The freshly isolated CD45–CD34–CD38–Lin– cells did not form any hematopoietic colonies but differentiated to CD34+ cells and fully matured myeloid cells when cultured with HESS-5 stroma cells. We observed that when the CD45–CD34–CD38–Lin– cells were cocultured with HESS-5 cells, hematopoietic colony-forming activity appeared on day 7 of culture (data not shown), which was consistent with the FCM data; that is, we detected CD34+ cells in the same culture on day 7 (Fig. 5AGo). Further, the CD45–CD34–CD38–Lin– cells had LTC-IC activity, while CD34+ myeloblasts did not. The cell proliferation kinetics of CD45–CD34–CD38–Lin– cells differed from those of CD34+ myeloblasts and CD45–CD34+CD38–Lin– cells. The number of cells in CD45–CD34–CD38–Lin– cell cultures decreased during the first 7–10 days, while clusters consisting of two to four cells began to appear on days 7–10. In contrast, in the CD34+ myeloblast culture and CD45–CD34+CD38–Lin– cell culture, cell clusters began to appear on day 3 and a marked increase in cells was observed on day 7. These data indicate that CD45–CD34–CD38–Lin– cells have hematopoietic potential and are at least one of the precursors of clonal CD34+ myeloblasts in MDS. Therefore, our data support the idea that at least in some MDS patients the transformation site is considerably upstream from the committed myeloid precursors having an immunophenotype of CD45+CD34+CD38+CD13+CD33+ [35, 36].

Using exactly the same approach, we examined normal BM cells, PB of lymphoma patients receiving G-CSF, cord blood, and samples (PB and BM cells) from de novo AML patients for the presence of CD45–CD34–CD38–Lin– cells, as detected in MDS samples. However, we have not yet found such cells in those sources (data not shown). One possible explanation for this failure is that the frequency of CD45–CD34–CD38–Lin– cells in normal and de novo AML samples is too low to be detected. De novo AML has quite different biological characteristics from those of MDS and AL-MDS [37, 38], and thus this failure in de novo AML is not surprising. A second possibility is that derangement of the cell-surface antigens of malignant cells in MDS samples produces CD45–CD34–CD38–Lin– cells, which do not exist in normal or de novo AML samples. If this is correct, although CD45–CD34–CD38–Lin– MDS cells had the potential to produce CD45+CD34+ cells, their relation to other immature cells, such as CD45+CD34–CD38–Lin– cells and CD45+CD34+CD38–Lin– cells, needs to be carefully investigated. It might be speculated that the CD45–CD34–CD38–Lin– cells detected in this study arise due to loss of antigen expression during ex vivo cell processing. In particular, it is known that CD34 expression can change according to the cell activation status [39]. In this study, the treatment with BR density centrifugation alone, which is a 10-minute centrifugation method that can speedily enrich immature cells [13–16], was sufficient to detect CD45–CD34–CD38–Lin– cells (Figs. 1Go and 2Go; Table 1Go). We showed previously that the immunophenotype of cells, including CD34 and CD45 expressions, did not change after BR density centrifugation [15]. Further, as described above, we could not detect CD45–CD34–CD38–Lin– cells in samples obtained from subjects other than MDS and treated by BR density centrifugation. These findings weigh against the speculation that ex vivo cell processing created the CD45–CD34–CD38–Lin– cells. However, more detailed characterization of the CD45–CD34–CD38–Lin– MDS cells and detection and characterization of these cells in other samples are needed to answer those various issues.

BR density centrifugation was developed based on the finding that immature blastoid cells are lighter than mature cells [12]. It is also known that hematopoietic progenitors and HSCs are lighter than most other BM cells [40], which is probably the reason we were able to detect CD45–CD34–CD38–Lin– cells after BR density centrifugation. Why do clonal CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– cells increase to detectable levels in RAEB-t and AL-MDS? We speculate that in the process of disease progression of MDS, a loss of differentiation capacity in transformed MDS HSCs (or switching from the initial MDS clone to a subclone having much less differentiation capacity) might lead to an increase in the CD45–CD34–CD38–Lin– cells, CD45–CD34+CD38–Lin– cells, and CD34+ myeloblasts in RAEB-t and AL-MDS. Documentation of the site of transformation of hematopoietic cells in MDS is important for understanding the pathophysiology of MDS and designing effective therapies. Based on our present findings, we conclude that transplantation of autologous CD34– HSCs can never guarantee engraftment of normal clones in MDS.

From the technical point of view, the most problematic point in conducting the study reported here was that the proliferating and differentiating capacities of MDS cells are much less than those of normal cells. Therefore, we used HESS-5 cells and the IBMI technique, which are powerful tools for examining in vitro and in vivo hematopoietic potentials, respectively. Nevertheless, CD45–CD34–CD38–Lin– cells from only a fraction of our patients proliferated even weakly in vitro, and none showed repopulation of hematopoiesis in NOD/SCID mice. Our in vitro data indicated that most CD45–CD34–CD38–Lin– cells could not survive in the present culture system and that a minor subpopulation of CD45–CD34–CD38–Lin– cells contributed to the production of CD45+CD34+ cells and more mature myeloid cells. Because most CD45–CD34–CD38–Lin– cells died early in the present cultures, we could not obtain enough cells to study the early phase of cell differentiation from CD45–CD34–CD38–Lin– cells such as whether these cells convert to CD45–CD34+CD38–Lin– cells (or CD45+CD34–CD38–Lin– cells) before generating CD45+CD34+ cells. It remains unclear whether the in vitro and in vivo data presented here reflect the true characteristics of CD45–CD34–CD38–Lin– cells or the extremely reduced hematopoietic potential of MDS cells when compared with normal cells. Further, our experimental conditions might be inadequate for CD45–CD34–CD38–Lin– cells.

In conclusion, CD45–CD34–CD38–Lin– cells, which we newly identified here, are phenotypically the most immature among hematopoietic cells so far reported and involved in the malignant clone in MDS. Although we could not show NOD/SCID repopulating activity in these cells, our in vitro data suggest that these cells have a role in the clonal hematopoiesis in MDS. Verification of the presence of these cells in normal condition and elucidation of their precise role(s) in normal and MDS hematopoiesis await further studies.


   ACKNOWLEDGMENTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
This work was supported in part by a Grant-in-Aid for Scientific Research from the Japan Society for the Promotion of Science (No. 14571002) and a research fund from Kirin Brewery Co., Ltd. to K.O.


   REFERENCES
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Received October 14, 2004; accepted for publication January 7, 2005.



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