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DOI: 10.1634/stemcells.2004-0280 2005;23;619-630 Stem Cells

Dan, Takafumi Kimura, Yoshiaki Sonoda and Takashi Tsuji Kiyoyuki Ogata, Chikako Satoh, Mikiko Tachibana, Hideya Hyodo, Hideto Tamura, Kazuo

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

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Original Article

Identification and Hematopoietic Potential of CD45– Clonal Cells with

Very Immature Phenotype (CD45–CD34–CD38–Lin–) in Patients with

Myelodysplastic Syndromes

Kiyoyuki Ogata,a Chikako Satoh,a,b Mikiko Tachibana,a Hideya Hyodo,a Hideto Tamura,a Kazuo Dan,a Takafumi Kimura,c Yoshiaki Sonoda,d Takashi Tsujie

aDivision of Hematology, Third Department of Internal Medicine, and bDepartment of Bioregulation, Nippon Medical

School, Tokyo, Japan; cDepartment of Molecular-Targeting Cancer Prevention, Graduate School of Medical Science, Kyoto

Prefectural University of Medicine, Kyoto, Japan; dDepartment of Hygiene, Kansai Medical University, Osaka, Japan; eDepartment 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: [email protected] Received October 14, 2004; accepted for publication January 7, 2005. ©AlphaMed Press 1066-5099/2005/$12.00/0 doi: 10.1634/stemcells.2004-0280

AbstractCD45 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 trans-formation 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 cen-trifugation, 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. Gen-erally, 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 hematopoi-etic colonies but had long-term culture-initiating cell activ-ity. 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 popula-tion represents the most immature immunophenotype so far identified in the hematopoietic lineage and is involved in the malignant clone in MDS. Stem Cells 2005;23:619–630

IntroductionHematopoietic stem cells (HSCs) are rare cell populations that

are capable of self-renewal and blood cell production [1]. There-

fore, 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-docu-

mented 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-sur-

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620 CD45– Clonal Cells in MDS

face 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 neo-

plasms 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 pro-

genitor cell level [8], but recent data suggest that the transfor-

mation 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 imma-

ture 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 imma-

ture (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 immunopheno-

type so far identified in the hematopoietic lineage and is involved

in the MDS clone.

Materials and Methods

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), diag-

nosed 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 CentrifugationHeparinized BM cells, which were aspirated for clinical diag-

nosis, and heparinized PB were obtained from the patients. The

study was approved by the Institutional Review Board of Nip-

pon Medical School, and informed consent was obtained from

Table 1. Phenotypic characteristics of myeloblasts and CD45– cells

Cell-surface antigens (% positive)Patient HLA- HLA-no. Sample Cells CD7 CD11b CD13 CD15 CD33 CD34 CD38 CD41a CD44 CD56 CD117 CD133 GPA class I DR KDR1 BM Myeloblasts 85.9 5.5 58.5 26.1 <1 67.1 99.5 1.5 89.6 <1 1.7 99.0 <1 99.9 96.6 7.6

CD45– cells 18.4 <1 <1 <1 <1 1.5 15.7 9.1 94.2 <1 <1 7.1 2.8 94.6 3.2 <1

2 BM Myeloblasts 5.3 13.7 33.1 26.9 53.4 55.4 87.7 4.8 81.5 28.7 15.8 27.2 1.5 99.8 ND 5.1

CD45– cells <1 <1 4.1a 6.1a 6.2a 9.3 9.3a 9.8 83.3 7.9a <1 <1 3.2 83.8 ND 3.2

3 PB Myeloblasts 30.6 13.3 99.5 71.3 98.7 99.1 51.3 5.4 98.3 3.3 64.7 10.3 <1 100 99.4 <1

CD45– cells <1 <1 <1 1.4a <1 10.5 7.4a 6.2 85.2 <1 <1 2.1 9.5 94.5 2.6a <1

4 PB Myeloblasts 18.6 12.4 77.1 29.5 11.6 89.0 90.0 8.8 88.5 6.7 63.3 50.8 6.4 99.1 ND 5.8

CD45– cells 11.5a <1 1.2a <1 <1 10.5 14.2a 9.9 88.2 <1 <1 <1 9.5 100 ND <1

5 PB Myeloblasts 4.5 41.6 95.3 42.8 89.0 90.1 85.2 22.6 ND 28.1 57.1 ND 8.6 99.2 83.4 ND

CD45– cells 1.2a 3.5a 3.3a 5.2a 1.7a 25.7 10.7a 7.8 ND 8.0a <1 ND 5.6 94.7 2.7a ND

6 PB Myeloblasts 25.8 10.1 94.8 27.6 89.7 97.6 78.7 14.5 85.8 3.9 78.7 1.6 2.2 99.8 99.1 <1

CD45– cells <1 2.1 <1 <1 <1 27.6 7.1 5.6 60.0 2.0 2.4 <1 8.3 75.5 5.2a <1

7 BM Myeloblasts 36.3 <1 97.8 ND 99.2 98.1 25.2 <1 ND <1 ND 92.7 1.8 ND ND ND

CD45– cells 19.5a <1 1.2a ND 5.9a 46.3 1.7a <1 ND <1 ND <1 8.2 ND ND ND

8 BM Myeloblasts <1 4.2 99.3 55.9 49.2 98.9 14.0 <1 ND <1 72.4 89.5 <1 ND 53.9 ND

CD45– cells <1 <1 14.8a <1 1.5a 65.5 8.1a <1 ND <1 4.8a 31.3a 5.1 ND 3.3a ND

CD45– cells positive for CD2, CD3, CD10, CD16, and CD19 were <1% in all patients.aIn the combined staining with CD34, we confirmed that CD34+ cells coexpressed these antigens. In patient 6, only HLA-DR was costained with CD34.Abbreviations: BM, bone marrow; GPA, glycophorin A; ND, not determined; PB, peripheral blood.



Ogata, Satoh, Tachibana et al. 621

all subjects. Samples were enriched for immature blastoid cells

by density-gradient centrifugation using BR (Japan Immunore-

search 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 sub-

jected 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).

FCMImmunophenotyping was performed by three-color FCM, in

which the optimal quantity of antibodies to be used was deter-

mined 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 f luorescein

isothiocyanate (FITC) or phycoerythrin (PE). These antibod-

ies 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); glycopho-

rin A (GPA), CD10, CD117 (FITC- or PE-conjugated; Beck-

man 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 over-

lap 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 CellsWe applied magnetic cell sorting (MACS) and fluorescence-acti-

vated cell sorting (FACS) to purify cells, as reported previously

[6, 21, 22]. The cell samples, which had been subjected to BR den-

sity 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 condi-

tions (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 anti-

bodies (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 stain-

ing 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 morphologi-

cal 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 2× 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 manu-




facturer’s instructions. DAPI (4,6 diamidino-2-phenylindole) was

used as a counterstain.

Cell Coculture with HESS-5 Murine Stroma CellsThe hematopoietic-supportive stromal cell line HESS-5 was

previously established from murine BM [24], and its applica-

tion 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 lay-

ers 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 ×

103 cells per 96-well plate and 3 × 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 combina-

tions 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 ana-

lyzed, 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) AssayThe 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, Vancou-

ver, 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-macro-

phage colony-stimulating factor (10 ng/mL), and erythropoi-

etin (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 Injec-tion (IBMI) MethodThe animal experiments were approved by the Animal Care

Committee of Kyoto Prefectural University of Medicine and car-

ried out at that university. Severe combined immunodeficient

(SCID)–repopulating cell assay by the IBMI method was per-

formed 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 irradi-

ated (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 ante-

rior. 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 α-medium were carefully injected via the bone hole into the

BM cavity. The mice were killed 10–14 weeks after transplanta-

tion, and the BMs from the bilateral femurs, tibiae, and humeri

were flushed out using α-medium containing 10% FCS. The pres-

ence of human cells was analyzed by FCM. Mice were scored as

positive if more than 0.1% of the total murine BM cells were posi-

tive 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 previ-

ously [6], were used as a positive control.

Statistical AnalysesDifferences between three or more groups of data of continuous

variables were analyzed by one-way analysis of variance. Dif-

ferences in categorical variables were evaluated using the chi-

square test. A p value of less than .05 was considered to be statis-

tically significant.

Results

Detection of CD45–CD34–CD38–Lin– and CD45–

CD34+CD38–Lin– Clonal Cells in MDSFigure 1 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. 1B). 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. 1D). 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 posi-

tive for HLA-class I antigen and CD44. Only minor subpopula-

tions 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 1E (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 1 (patient 1) and

clearly differ considerably between CD45– cells and myeloblasts.

Figure 2 shows the plots for another patient, and it is seen that one

third of the CD45– cells (cells in R3, Fig. 2B) expressed CD34 and

a few expressed CD38 and myeloid antigens (blue dots in Fig. 2C).

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 consider-

ably between the CD45– cells and myeloblasts (gray dots in Fig.

2C) in this patient (Table 1, patient 8).

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

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 1). (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 1). (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, fluores-

cein isothiocyanate; GPA, glycophorin A; PE, phycoerythrin.




clusters, as exemplified in Figures 1 and 2. Tables 1 and 2 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 anti-

gens (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] × 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

1). 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 3, 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.

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 express-

ing 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. 3A). The iso-

lated CD45–CD34–CD38–Lin– and CD45–CD34+CD38–Lin– cells

Figure 2 . Representat ive example of CD45–

CD34+CD38–Lin– cell detection in a sample of myelo-

dysplastic syndromes. (A): Forward scatter (FSC)

versus side scatter (SSC) display of bone marrow cells

after blastretriever density centrifugation (patient 8 in

Table 1). (B): CD45 versus SSC display of the cells

gated by R1 in panel A. The left side of the bold verti-

cal 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, fluo-

rescein isothiocyanate; PE, phycoerythrin; PerCP,

peridin chlorophyll.

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

Patient no. Age/Sex Diagnosis Karyotype

1 62/M AL-MDS 46,XY [20]

2 72/M RAEB-t 46,XY [20]

3 37/F AL-MDS 45,XX,add(1)(q32),add(2)(q11), –5, –7,+8,add(11)(p15), –12, –15,dic(18;21)(p1?;p1?), –19,+der(?)t(?;12)(?;q1?),+mar1,+mar2,+mar3 [4] /46,idem, –mar3,+mar4,+mar5 [5] / 45,idem.add(X)(p11),–mar3,+mar [2]

4 68/M RAEB-t 47,XY,+Y [20]

5 82/F RAEB-t 46,XX,inv(3)(p25q21), –5,del(6)(q13q23), –7, –9,+3mar,inc [14]

6 73/M RAEB-t 50,XY,-5,del(5)(q13q33),+6,der(7;15)(p10;q10),+8,+8,+11,+13,–14,add(17)(p11), +2mar [11] /51,idem,+3mar[8]/46,XY [1]

7 66/M AL-MDS 45,XY,del(5)(q13q33),-7,add(19)(p13) [20]

8 72/F AL-MDS 43,XX,del(5)(q23q32),-7,add(10)(p11),-13,del(16)(q13),–18 [20]

Number of cells showing each karyotype is in square brackets.Abbreviations: AL-MDS, acute leukemia transformed from myelodysplastic syndromes; RAEB-t, refractory anemia with excess blasts in transformation.




were blastoid cells that had scanty cytoplasm with no granules

(Fig. 3B). 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. 3C and Table 4).

Proliferation and Differentiation of CD45–CD34–

CD38–Lin– Cells Cultured with HESS-5 Stroma CellsWe previously established a murine BM stromal cell line, HESS-

5 [24], and showed that in combination with cytokines it dramati-

cally 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 infe-

rior to HESS-5 plus cytokines in expanding and/or differentiat-

ing 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. 4A). 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 col-

ony-stimulating factor (G-CSF)–mobilized PB (normal control)

Figure 3. Isolation of CD45–CD34–CD38–Lin– cells and their mor-

phological 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) treat-

ment. 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 immunophe-

notypes 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 flu-

orescence in situ hybridization (right). The red spot and green spots

show X- and Y-chromosome signals. Abbreviations: FITC, fluores-

cein isothiocyanate; PE, phycoerythrin; PerCP, peridin chlorophyll.

Table 3. Relationships between disease subtypes and CD45–CD38–

Lin– cell detection

Disease

RAEB RAEB-t AL-MDS p value

CD45–CD38–Lin– cellsa 0/21 4/17 4/13 .03

Blastoid cellsb 70.9 ± 20.8 76.6 ±21.9 83.2 ± 18.0 .24

Erythroblastsb 1.3 ± 3.6 2.1 ± 5.5 0 ± 0 .40

aPositive number of patients per analyzed number of patients.bMean ± SD of the percentages of blastoid cells and erythroblasts in samples used for flow cytometry; determined in Wright Giemsa–stained cytospins.Abbreviations: AL-MDS, acute leukemia transformed from myelo-dysplastic syndromes; RAEB, refractory anemia with excess blasts; RAEB-t, RAEB in transformation.

Table 4. Results of fluorescence in situ hybridization analysis Detection of chromosomal aberrationa

Patient no. Lymphocytes Myeloblasts CD45–CD38–Lin– cellsb

4 0/35 17/31 16/25

5 1/20 18/20 7/10

7 ND 19/20 6/6

8 0/20 15/19 5/9

aNumber of cells having chromosomal aberration per number of cells examined. Analyzed chromosomal aberrations were +Y in patients 4 and –7 in the other three patients.bAnalyzed cells were CD45–CD34–CD38–Lin– cells for patients 4, 5, and 7 and CD45–CD34+CD38–Lin– cells for patient 8.Abbreviation: ND, not determined.




were cultured under the present culture conditions, the cell num-

ber 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 4B. Among these four patients,

the highest increase in cell number on day 7 was 50-fold (patient

4; the solid circles in Fig. 4B). These inferior results in MDS were

expected because we and others had documented that the expan-

sion and differentiation abilities of HSCs and hematopoietic pro-

genitors 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. 4B) 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. 4B). Time-course pho-

tographs of the cultured cells from a representative patient are

shown in Figure 4C.

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 5. Regarding CD34+ myeloblast culture, the

percentage of CD34+ cells in the culture decreased with time and

became 0% on day 35 of culture (Fig. 5A, lower panel). Through-

out the culture period, almost all cells expressed myeloid antigens

(data not shown). On day 7 of CD45–CD34–CD38–Lin– cell cul-

ture, the CD45–CD34–CD38–Lin– cell percentage had decreased

markedly, and CD45+CD34+ cells and CD45+CD34– cells

Figure 4. Proliferation and differentiation of CD45–CD34–CD38–

Lin– cells cocultured with HESS-5 stroma cells. (A): CD34+ myelo-

blasts (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.

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, thrombopoi-

etin, 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 antibod-

ies 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, phyco-

erythrin; PerCP, peridin chlorophyll.




expressing myeloid antigens had appeared (Fig. 5A, 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. 5B). The findings for Wright-

Giemsa–stained cytospin preparations at various time points

were consistent with the FCM data and confirmed that both cul-

tures produced myeloid cells in various stages of maturation (Fig.

5C). FISH analyses also confirmed that these cells were clonal in

origin (data not shown).

Colony-Forming Activity and LTC-IC Activity of CD45–CD34–CD38–Lin– CellsThe results are shown in Figure 6. It has been reported that colony-

forming activity is often defective in MDS [27, 28]. In the experi-

ments 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.

In Vitro Hematopoietic Potential of CD45–

CD34+CD38–Lin– CellsFor 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-men-

tioned 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 cul-

ture, and the cell number had increased significantly by day 7 of

culture (Fig. 7A). Further, freshly isolated CD45–CD34+CD38–

Lin– cells had clear colony-forming potential (Fig. 7B).

NOD/SCID Repopulating Activity of CD45–CD34–

CD38–Lin– CellsDemonstration 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 sen-

sitively compared with the conventional method—intravenous

injection of HSCs to NOD/SCID. That is, IBMI, but not the con-

ventional 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 5).

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 activ-

ity. (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 pan-

els 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 gran-

ulocyte-macrophage colony-forming units.

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 col-

umn labeled Mbl), and CD45–CD34–CD38–Lin– cells (right column

labeled CD45–CD34–) from patient 4 were analyzed for colony-form-

ing 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.




DiscussionWhether HSCs are actually involved in MDS remains controver-

sial [34]. There is a hypothesis that MDS malignant transforma-

tion 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. 5A). Further, the CD45–CD34–CD38–Lin– cells had LTC-

IC activity, while CD34+ myeloblasts did not. The cell prolif-

eration 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 cul-

ture, 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 dif-

ferent 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-min-

ute centrifugation method that can speedily enrich immature cells

[13–16], was sufficient to detect CD45–CD34–CD38–Lin– cells

(Figs. 1 and 2; Table 1). We showed previously that the immuno-

phenotype 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 sam-

ples obtained from subjects other than MDS and treated by BR den-

sity 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 den-

sity 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 dis-

ease 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 hema-

topoietic cells in MDS is important for understanding the patho-

physiology of MDS and designing effective therapies. Based on

our present findings, we conclude that transplantation of autolo-

gous 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

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

Patient no. Injected cell number (× 104) Engraftment

1 4.3 0/12 1.4, 3.5a 0/24 1.8, 3.0, 3.0, 5.3, 6.0, 6.8a 0/65 2.0 0/16 0.3 0/1Normal cellsb 5.0 7/7

aMice were treated with anti-asialo GM1 antiserum. bCD45+CD34–Lin– cells from cord blood were used as a positive control. In this case, the engraftment was documented in all mice (10.0%–52.6% [median, 19.3%] of human cells in the bone marrow of recipient mice).




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 pro-

liferated 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 pres-

ent 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 differentia-

tion 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 hemato-

poiesis await further studies.

AcknowledgmentsThis 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.

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DOI: 10.1634/stemcells.2004-0280 2005;23;619-630 Stem Cells

Dan, Takafumi Kimura, Yoshiaki Sonoda and Takashi Tsuji Kiyoyuki Ogata, Chikako Satoh, Mikiko Tachibana, Hideya Hyodo, Hideto Tamura, Kazuo

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

This information is current as of February 12, 2007

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