1 Department of Pediatrics, Graduate School of Medicine, Kyoto University, 54
Kawahara-cho, Shogoin, Sakyo-ku, Kyoto 606-8507, Japan
2 Department of Development and Differentiation, Institute for Frontier Medical
Science, Kyoto University, 53 Kawahara-cho, Shogoin, Sakyo-ku, Kyoto 606-8507,
Japan
3 Department of Medicine, Boston University School of Medicine, 88 East Newton
Street, Boston, MA 02118, USA
4 Research Center for Animal Life Science, Shiga University of Medical Science,
Tsukinowa-cho, Seta, Ohtsu, Shiga 520-2192, Japan
5 Division of Genetics, Institute of Medical Science, University of Tokyo, 4-6-1
Shirokane-dai, Minato-ku, Tokyo 108-8639, Japan
* Author for correspondence (e-mail: tnakaha{at}kuhp.kyoto-u.ac.jp)
Accepted 8 January 2004
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SUMMARY |
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Key words: ES cells, Primate, Primitive hematopoiesis, Definitive hematopoiesis
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Introduction |
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Recently, it has been reported that primate hematopoiesis occurs in a
manner similar to that of mice, based on immunohistochemical studies of human
embryos (Tavian et al., 1996;
Tavian et al., 1999
). In the
erythrocytes of primates, the embryonic (
and
), fetal (
)
and adult (ß and
) globin genes are expressed sequentially during
development, although small amounts of fetal and adult globin chains are
detected even during primitive hematopoiesis
(Johnson et al., 2000
;
Stamatoyannopoulos et al., 2001). Concomitant switches in the
cluster
(replacement of the
globin gene by the
) and ß cluster
(replacement of the
globin gene by the
, and replacement of the
globin gene by the ß) occur during development, and coincide with
the transition from yolk sac to fetal liver, and finally to bone marrow
hematopoiesis.
The hematopoietic development of primates remains to be elucidated, in part
because of the ethical restrictions on experiments using their embryos. Old
World monkeys, such as the cynomolgus monkey (Macaca fascicularis),
are widely used for medical research
(Hanazono et al., 2000) and
have globin gene expression that is similar to that of humans
(Johnson et al., 2000
).
Therefore, their ES cells might be used as a model for elucidating primate
hematopoietic development. Recently primate ES cell lines were established
(Thomson et al., 1995
;
Thomson et al., 1996
;
Thomson et al., 1998
;
Suemori et al., 2001
), and
hematopoietic differentiation from primate ES cells was also induced
successfully in vitro (Kaufman et al.,
2001
; Li et al.,
2001
; Lu et al.,
2002
; Chadwick et al.,
2003
). However, compared with the murine system, little work has
been done to precisely analyze primitive and definitive hematopoietic
development.
To address this problem, we induced hematopoietic differentiation in cynomolgus monkey ES cells by co-culture with OP9 stromal cells. This is the first report to demonstrate that primitive hematopoiesis and its transition to definitive hematopoiesis can be induced from primate ES cells in vitro.
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Materials and methods |
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Cytokines and growth factors
Recombinant human granulocyte colony-stimulating factor (G-CSF; CSF2
Human Gene Nomenclature Database), EPO, interleukin 3 (IL3), stem cell
factor (SCF) and thrombopoietin (THPO) were kindly provided by Kirin Brewery
(Tokyo, Japan). Recombinant human VEGF, basic fibroblast growth factor (bFGF),
and bone morphogenetic protein (BMP) 4 were all purchased from R&D Systems
(Minneapolis, Minnesota).
Antibodies
The primary antibodies used in this study were: mouse anti-human CD41
(clone 5B12) and CD45 (clone T16) antibodies (Dako, Kyoto, Japan); mouse
anti-human CD34 (clone 563) antibody (Becton-Dickinson, San Jose, CA); KIT
antibody (Nichirei, Tokyo, Japan); rabbit anti-human hemoglobin (Hb)
polyclonal antibody (Cappel, Aurora, Ohio); and sheep anti-human fetal
hemoglobin (HbF) polyclonal antibody (Bethyl, Montgomery, TX). The mouse
anti-human embryonic -globin (HbEmb) monoclonal antibody, and anti-human
fetal liver kinase (FLK) 1 monoclonal antibody were used as described
previously (Luo et al., 1999
;
Sawano et al., 2001
).
Cy3-conjugated donkey anti-mouse IgG, Cy3-conjugated donkey anti-sheep IgG,
fluorescein isothiocyanate (FITC)-conjugated donkey anti-rabbit IgG and
alkaline phosphatase (ALP)-conjugated donkey anti-mouse IgG (all purchased
from Jackson ImmunoResearch Laboratories, West Grove, PA), and allophycocyanin
(APC)-conjugated goat anti-mouse IgG (Becton-Dickinson), were used as
secondary antibodies.
In vitro hematopoietic differentiation from ES cells
OP9 stromal cells were kindly provided by Dr Hiroaki Kodama, and were
maintained in MEM (Gibco BRL) supplemented with 20% fetal calf serum
(FCS) (EQUITECH-BIO, Kerrville, TX). Trypsintreated ES cells
(4x103 cells/well) were transferred onto confluent OP9
stromal cells in
MEM supplemented with 10% FCS and 50 µM 2ME, in the
presence or absence of VEGF, bFGF or BMP4. On day 6 of differentiation, the
induced cells were harvested with cell dissociation buffer (Invitrogen,
Carlsbad, CA). Then the cells were filtered through a 70 µm nylon cell
strainer (Falcon, Lincoln Park, NJ), and 1x105 cells/well
were transferred onto fresh confluent OP9 cells in 6-well plates and cultured
in
MEM supplemented with 10% FCS and 50 µM 2ME, in the presence or
absence of EPO (10 U/ml). The medium was changed every 2 or 3 days during the
induction of differentiation. Adherent hematopoietic cell clusters, which
consisted of more than 20 round blast-like cells, were counted using an
inverted microscope. The same series of experiments was performed at least
three times.
Staining
For cytochemical staining, the floating cells were centrifuged onto glass
slides and analyzed by microscopy after May-Giemsa or myeloperoxidase
staining.
For immunostaining, floating cells spun onto glass slides were fixed in 4%
paraformaldehyde and permeabilized with phosphate buffered saline (PBS)
containing 5% skim milk (Becton-Dickinson) and 0.1% Triton X-100 for 30
minutes. The cells were then incubated with primary antibodies overnight,
washed three times with PBS containing 5% skim milk, and then incubated with
FITC- or Cy3-conjugated secondary antibodies for 30 minutes. Nuclei were
labeled with Hoechst 33342 (Molecular Probes, Eugene, Oregon). The cells were
then washed three times with PBS and observed by fluorescence microscopy
(Olympus, Tokyo, Japan). In the human erythroblastic cell line K562, which is
known to express ,
,
and
globins
(Rutherford et al., 1981
), all
erythroid cells were positive for Hb, HbF and HbEmb. In adult cynomolgus bone
marrow, all erythrocytes were positive for Hb and a few were positive for HbF,
whereas Hb Emb-positive erythrocytes were rarely detected (data not shown).
The adherent cells were fixed and incubated with primary and ALP-conjugated
secondary antibodies, as described above, and positive cells detected using a
Vector Blue Alkaline Phosphatase Substrate Kit III (Vector Laboratories,
Burlingame, CA). Endogenous ALP activity was blocked by 2 mM levamisole (Wako,
Osaka, Japan).
Methylcellulose colony forming assays
The medium was replaced with a fresh semisolid medium consisting of
MEM, 0.9% methylcellulose, 30% FCS, 10% bovine serum albumin and 50
µM 2ME, and a mixture of human G-CSF 10 ng/ml, EPO 2 U/ml, IL3 20 ng/ml,
SCF 100 ng/ml and THPO 10 ng/ml, as previously reported
(Sui et al., 1995
). All
cultures were incubated at 37°C in a humidified atmosphere flushed with 5%
CO2 in air. Seven days later, individual colonies were lifted with
an Eppendorf micropipette under direct microscopic visualization, washed twice
with PBS, and processed for May-Giemsa staining, immunostaining and RT-PCR
analysis. Colonies (
50 cells) were counted using an inverted microscope
according to the criteria previously reported
(Nakahata and Ogawa, 1982
;
Tajima et al., 1996
).
Reverse transcription-polymerase chain reaction (RT-PCR)
Total RNA was prepared by using TRIzol (Invitrogen) according to the
manufacturer's protocol. Each total RNA sample was then reverse-transcribed
using a SuperScript first-strand synthesis system for RT-PCR (Invitrogen)
according to the manufacturer's instructions. The cDNA was amplified in a
final volume of 20 µl PCR buffer containing 2.5 mM MgCl2 and 250
µM dNTP, using Taq DNA polymerase (Takara Shuzo, Kyoto, Japan). As
cynomolgus monkey-specific sequences were unavailable, we employed those of
corresponding human or other Old World monkey genes to design our PCR primers,
based on the generally close homology between human and cynomolgus monkey gene
sequences. The EPO receptor (EPOR) and -fetoprotein (AFP)-specific
primers have been described previously (Schuldiner et al., 2000;
Yokomizo et al., 2002
).
Samples were initially denatured at 94°C for 5 minutes, followed by
amplification rounds consisting of 94°C for 1 minute (denaturing),
57-66°C for 1 minute (annealing) and 72°C for 1 minute (extension),
and then a final extension at 94°C for 7 minutes. The oligonucleotide
primers were:
The PCR reactions were carried out as follows: -globin, 35 cycles;
-globin, 35 cycles; ß-globin, 35 cycles;
-globin, 35 cycles;
-globin, 35 cycles; EPOR, 40 cycles; Brachyury, 45 cycles; FLK1, 40
cycles; SCL, 45 cycles; LMO2, 40 cycles; MYB, 40 cycles; GATA2, 35 cycles;
Nestin, 35 cycles; AFP, 35 cycles; REX1; 35 cycles; ß-actin, 35 cycles.
The analysis of globin gene expression in individual erythroid colonies was
performed for 40 cycles. PCR products were visualized by 1.5% agarose gel
electrophoresis using ethidium bromide staining. cDNA from cynomolgus monkey
bone marrow or K562 cells was used as a positive control. For
semi-quantitative comparisons, samples were normalized by dilution to give
equivalent signals for ß-actin. DNA sequencing was performed for genes
from which we were unable to obtain adequate PCR products from the positive
control.
Flow cytometric analysis and cell sorting
The cells induced from CMK6 were harvested sequentially with cell
dissociation buffer (Invitrogen), filtered through a 70 µm nylon cell
strainer (Falcon), and incubated in human IgG for 30 minutes to block the
non-specific binding of FC-receptors. The cells were then incubated
with PE-conjugated CD34, or unconjugated FLK1 and KIT antibodies for 30
minutes. Samples staining with unconjugated antibodies were then incubated
with APC-conjugated goat anti-mouse antibody (BD PharMingen) for 30 minutes.
The cells were then washed and analyzed using a FACScaliber with the CellQuest
program (Becton-Dickinson). Forward- and side-scatter plots were used to
exclude MEF or OP9 stromal cells, and propidium iodide costaining was used to
exclude non-viable cells. Mouse IgG1 (Dako) was used as an isotype control. On
day 10, the cultured cells were harvested and labeled with CD34 antibody as
described above. CD34-positive cells were collected using a FACSVantage flow
cytometer (Becton-Dickinson) and re-plated at a concentration of
2x104 cells per well onto fresh OP9 stromal cells in
MEM supplemented with 10% FCS and 50 µM 2ME, in the presence or
absence of EPO (10 U/ml). Floating cells were processed for May-Giemsa
staining, immunostaining and RT-PCR analysis.
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Results |
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CD41-positive megakaryocytes appeared on day 8 (Fig. 2A,B), and MPO-positive myeloid lineage cells appeared on day 12 (Fig. 2C,D). Time-course analysis demonstrated that the number of megakaryocytes peaked on day 10 but decreased thereafter, whereas myeloid lineage cells gradually increased until they comprised more than half of the total number of hematopoietic cells on day 14 and thereafter (Fig. 2E).
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Exogenous VEGF increased the number of erythrocytes in a dose-dependent manner until day 14, in the presence or absence of EPO (Fig. 8A). These erythrocytes consisted exclusively of EryP. EryD were rarely observed in the presence of VEGF alone (data not shown), whereas exogenous VEGF plus EPO enhanced EryD production more prominently with time (Fig. 8B).
|
More efficient primitive and definitive hematopoiesis is induced by re-plating sorted CD34-positive cells
As previously shown, other lineages developed concomitantly in our culture
system. Consequently, we purified the CD34-positive cells in the cultures and
seeded them onto fresh OP9 stromal cells on day 10
(Fig. 9A-C). In the presence of
EPO, approximately on day 25 (i.e. 15 days after the cell sorting), adherent
hematopoietic cell clusters grew larger and had a cobblestone appearance
(Fig. 9D). The number of
floating cells, most of which were erythrocytes, increased with a peak on day
28 (Fig. 9E,H,I). Nearly 40% of
these erythrocytes were definitive, and many were enucleated
(Fig. 9E-G,J,K). In the absence
of EPO, hematopoietic development after cell sorting was barely observed.
These results indicate that CD34-positive cells in cultures contained
progenitors of both primitive and definitive hematopoiesis.
|
![]() |
Discussion |
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The hematopoietic development of primates, however, is different from that
of mice. For example, expression of the gene during the fetal period
is an event that occurs only among primates
(TomHon et al., 1997
).
Therefore, in vitro and in vivo studies of primate hematopoietic development
should be performed using primate-derived materials. In recent studies in
primate ES cells, Kaufman et al. and Li et al. demonstrated that the
definitive hematopoiesis, but not the primitive hematopoiesis, of in vivo
differentiation is recapitulated (Kaufman
et al., 2001
; Li et al.,
2001
). Lu et al. revealed that the coexistence of primitive and
definitive hematopoiesis is recapitulated at the mRNA level
(Lu et al., 2002
). In this
study, we have demonstrated for the first time the transition from primitive
to definitive hematopoietic development in primate ES cells at both the
transcriptional and the translational level in vitro. Immunostaining using
human hemoglobin antibodies demonstrates that embryonic and definitive (fetal)
erythrocytes appear on day 8 and day 16, respectively. Sequential RT-PCR
analysis of globin genes demonstrates upregulation of primitive (
and
) globin gene expression on day 8 and of definitive (
and
) globin genes on day 12, which indicates that the erythropoietic
transition can be recapitulated in ES cells at the mRNA level. Therefore, our
in vitro system is superior in precisely reflecting the ontogeny of
hematopoietic cells in vivo, and should be a useful tool to define the
mechanisms of primate hematopoiesis. The generation of adherent hematopoietic
cell clusters containing CD34-positive cells onto the OP9 cell layer indicates
that this induction system also recapitulates hematopoietic development at the
progenitor level from primate ES cells, as has been observed in murine ES
cells (Nakano et al., 1996
;
Suwabe et al., 1998
;
Era et al., 2000
;
Kitajima et al., 2002
).
Our results from the colony assays also demonstrate for the first time that
primitive erythroid colonies are generated with the aid of stromal cells, but
that definitive colonies do not emerge in the presence or absence of stromal
cells. By contrast, a recent study has shown that definitive erythroid
colonies are generated from primate ES cells under stromal-free conditions,
but that primitive colonies are not
(Kaufman et al., 2001). These
differences may be partially due to differences in the culture conditions, the
colony assays, and/or the ES cells and stromal cells that were used for the
induction of differentiation. Notably, our individual primitive erythroid
colonies express not only embryonic but also fetal and adult globin genes,
which is consistent with the results obtained by plating human embryonic or
fetal cells (Peschle et al., 1984;
Stamatoyannopoulos et al.,
1987
). Fetal and adult hemoglobin synthesis, and factors
regulating their synthesis, have been intensively analyzed in human cord
blood, and in neonatal and adult erythroid colonies (Stamatoyannopoulos et
al., 2001). However, precise analysis of hemoglobin synthesis in primitive
erythroid colonies has not been performed. Thus, our culture system will also
serve as a powerful tool for elucidating the regulatory mechanisms of
primitive hematopoiesis.
Co-culture on OP9 stromal cells alone induces hematopoietic development
less efficiently in primate ES cells than in murine ES cells. VEGF, bFGF and
BMP4 have been reported to promote primitive or definitive hematopoietic
development in previous studies in murine ES cells
(Johansson and Wiles, 1995;
Faloon et al., 2000
;
Nakayama et al., 2000
).
Therefore, we quantified the stimulatory effects of these growth factors on
both types of hematopoiesis. Unexpectedly, exogenous BMP4 fails to induce
hematopoietic differentiation in our culture system. There are two possible
explanations for this discrepancy. One is that human BMP4 does not work on the
cynomolgus ES cell line we used. However, considering that human BMP4
functions in both murine and primate ES cells
(Johansson and Wiles, 1995
;
Nakayama et al., 2000
;
Li et al., 2001
;
Chadwick et al., 2003
) the
possibility seems unlikely. Another possibility is that BMP4 causes the OP9
stromal cells to differentiate and thereby impairs their interaction with ES
cells. Supporting this notion is the fact that we observed that exogenous BMP4
resulted in an increase of adipocytes on the OP9 cell layer (data not shown).
This observation is also consistent with a previous report that showed that
BMP4 induces the differentiation of mesenchymal progenitors into distinct
various mesenchymal cell lineages including adipocytes
(Ahrens et al., 1993
).
Of course, there is a common requirement for cytokines or growth factors
during hematopoietic differentiation from both primate and murine ES cells. We
demonstrated that in primate erythropoiesis, exogenous EPO is required for
EryD development, whereas EryP develop independently of EPO, despite
substantial expression of EPO receptor. This result is consistent with reports
showing that murine primitive and definitive erythrocytes have different
requirements for EPO (Wu et al.,
1995; Lin et al.,
1996
). As previously reported in murine ES cells
(Heberlein et al., 1992
;
Keller et al., 1993
), the EPOR
is expressed in undifferentiated primate ES cells. However, it is unlikely
that erythrocytes are contained in undifferentiated ES cells, because no
globin gene expression is detected before the induction of differentiation.
Further studies will be required to analyze the function of EPOR expressed in
undifferentiated ES cells.
Among growth factors examined in this study, VEGF, a ligand for FLK1, was
the only one to stimulate both primitive and definitive hematopoiesis. FLK1 is
required for the development of primitive and definitive hematopoietic cells,
as well as endothelial cells, in the murine embryo
(Shalaby et al., 1995;
Shalaby et al., 1997
). Recent
studies on the differentiation of murine ES cells in vitro also indicate that
primitive and definitive hematopoietic and endothelial cell lineages can be
generated from FLK1-positive cells (Choi et
al., 1998
; Nishikawa et al.,
1998
; Faloon et al.,
2000
). In our study, the expression of FLK1 was upregulated on day
6, before hematopoietic development. This result is consistent with the recent
report on vascular progenitor cell differentiation from cynomolgus monkey ES
cells onto OP9 stromal cells (Sone et al.,
2003
). Furthermore, we observed that exogenous VEGF also enhances
the development of vascular endothelial cadherin-positive endothelial colonies
under the same culture conditions (K.U., T.H. and T.N., unpublished). Taken
together, these results strongly suggest that primitive and definitive
hematopoietic, as well as endothelial, lineage progenitors are derived from
FLK1-positive cells in culture. Further studies, by single cell culture of
FLK1-positive cells to differentiate into both lineage cells, will be needed
to confirm this possibility.
We also examined indispensable genes associated with hematopoietic
development. GATA2 has been reported to be necessary for the proliferation and
survival of both primitive and definitive hematopoietic progenitors
(Tsai et al., 1994). Its
expression in our system supports the proposed role it plays in the generation
of hematopoietic progenitors. The expression of Brachyury, an early mesodermal
marker (Herrmann et al., 1994), was upregulated on day 4, and was followed by
the upregulation of SCL, MYB and LMO2 expression on day 6, before
hematopoietic development. SCL (Robb et
al., 1996
; Porcher et al.,
1996
) and LMO2 (Warren et al.,
1994
; Yamada et al.,
1998
) are required for both primitive and definitive hematopoietic
development, whereas MYB is essential for the development of definitive
hematopoiesis only (Mucenski et al.,
1991
). SCL is also crucial for the development of hemangioblasts
(Faloon et al., 2000
;
Chung et al., 2002
). These
results suggest that a similar profile of genes is involved in hematopoiesis
in culture as is involved in early hematopoiesis in vivo. These observations
will also facilitate the genetic manipulations of ES cells that may shed light
on the unresolved molecular mechanisms behind hematopoietic development.
As sequential RT-PCR analysis of Nestin, AFP and REX1 indicated that other lineage cells and undifferentiated ES cells also grow during the differentiation induction process, we purified the CD34-positive cells in the cultures and seeded them onto fresh OP9 stromal cells. Analyses after cell sorting indicated that enhanced definitive hematopoiesis was generated on day 25 and thereafter, although primitive hematopoiesis was still produced. These results indicate that both hematopoietic processes originate from the sorted CD34-positive population. Further experiments to quantitatively analyze definitive hematopoiesis will be performed using this improved assay.
In conclusion, the sequential development of primitive and definitive hematopoiesis can be induced from primate ES cells by co-culture with OP9 stromal cells. This induction system will provide new approaches for elucidating the mechanisms regulating primate hematopoietic development and differentiation during embryogenesis.
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ACKNOWLEDGMENTS |
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