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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 94 No. 8 (October 15), 1999:
pp. 2686-2695
Unique Differentiation Programs of Human Fetal Liver Stem Cells
Shown Both In Vitro and In Vivo in NOD/SCID Mice
By
Franck E. Nicolini,
Tessa L. Holyoake,
Johanne D. Cashman,
Pat
P.Y. Chu,
Karen Lambie, and
Connie J. Eaves
From the Terry Fox Laboratory, British Columbia Cancer Agency, and
the Department of Medical Genetics, University of British Columbia,
Vancouver, British Columbia, Canada.
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ABSTRACT |
Comparative measurements of different types of hematopoietic
progenitors present in human fetal liver, cord blood, and adult marrow
showed a large (up to 250-fold), stage-specific, but
lineage-unrestricted, amplification of the colony-forming cell (CFC)
compartment in the fetal liver, with a higher ratio of all types of CFC
to long-term culture-initiating cells (LTC-IC) and a lower ratio of
total (mature) cells to CFC. Human fetal liver LTC-IC were also found
to produce more CFC in LTC than cord blood or adult marrow LTC-IC, and
more of the fetal liver LTC-IC-derived CFC were erythroid. Human fetal liver cells regenerated human multilineage hematopoiesis in NOD/SCID mice with the same kinetics as human cord blood and adult marrow cells,
but sustained a high level of terminal erythropoiesis not seen in adult
marrow-engrafted mice unless exogenous human erythropoietin (Epo) was
injected. This may be due to a demonstrated 10-fold lower activity of
murine versus human Epo on human cells, sufficient to distinguish
between a differential Epo sensitivity of fetal and adult erythroid
precursors. Examination of human LTC-IC, CFC, and erythroblasts
generated either in NOD/SCID mice and/or in LTC showed the types of
cells and hemoglobins produced also to reflect their ontological
origin, regardless of the environment in which the erythroid precursors
were generated. We suggest that ontogeny may affect the behavior of
cells at many stages of hematopoietic cell differentiation through key
changes in shared signaling pathways.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
HEMATOPOIESIS IS FIRST detectable in the
extravascular regions of the developing human fetal liver during week 6 of gestation, which then remains the major site of hematopoiesis until
birth.1 In addition to morphologically recognizable
hematopoietic cells, the human fetal liver has been shown to contain
the full hierarchy of more primitive progenitors detectable by
functional endpoints. These include erythroid, megakaryopoietic,
granulopoietic, and multilineage colony-forming cells (CFC) detectable
in semisolid culture assays,2-7 their more primitive
precursors, referred to as long-term culture-initiating cells (LTC-IC)
because they are able to proliferate and differentiate in vitro for
more than 5 weeks on stromal feeder layers,7 and
transplantable stem cells able to engraft xenogeneic8-11 as
well as allogeneic recipients.12 Comparisons of analogous
cell types in murine fetal tissues and adult marrow have shown
differences in their rate of turnover,13 surface
phenotype,14,15 rate of regeneration
posttransplant,16-21 responsiveness to specific growth
factors,22-27 and differentiation programs executed by
their lineage-restricted progeny.28-30 Previous studies
have provided some indication that similar differences between fetal
and adult hematopoietic cells of human origin also exist.31-37
In vivo, the fetal liver is characterized by a predominance of
terminally differentiating erythroid cells. This could reflect an
intrinsically increased probability of pluripotent fetal liver stem
cells to commit to this pathway or the acquisition by their erythroid-committed progenitors of features that favor expansion of
their differentiating progeny in vivo. To investigate these possibilities, we examined the erythropoietic potential and types of
hemoglobin produced by the erythroid progeny of very primitive (uncommitted) human fetal liver cells stimulated to differentiate under
a variety of conditions either in vivo (in NOD/SCID mice) and/or in
vitro (in CFC and LTC-IC assays). We then compared the results with
parallel measurements for cells derived from newborn and adult sources
of human hematopoietic cells (ie, cord blood and adult marrow).
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MATERIALS AND METHODS |
Human cells.
Human bone marrow cells were either aspirate samples obtained from
normal individuals donating marrow for allogeneic marrow transplantation or were from cyropreserved cadaveric marrow samples obtained from the Northwest Tissue Center (Seattle, WA). Cord blood
samples were obtained from mothers undergoing cesarean delivery of
normal, full-term infants and livers were from 14- to 21-week-old aborted fetuses. The age of the embryo was determined by a foot length
measurement. (There was no evidence of a consistent change in any
parameter assessed in cells obtained from fetal livers over this range
in gestational age and, accordingly, data from different fetal livers
have been pooled.) For all human samples, approved institutional
procedures for obtaining informed consent were followed. Single-cell
suspensions were obtained from the human fetal livers by pushing the
sample through a coarse sieve. The low-density (< 1.077 g/mL) cells
were then isolated by density centrifugation on Ficoll-Hypaque
(Pharmacia Biotech, Uppsala, Sweden). To obtain a population that was
further enriched in CD34+ cells, cells bearing lineage
markers expressed by different types of mature cells (lin+
cells) were removed using a StemSep column according to the suppliers' directions (StemCell Technologies, Vancouver, British Columbia, Canada).
Animals.
NOD/LtSz-scid/scid (NOD/SCID) mice38 were bred and
maintained in microisolators under defined sterile conditions in the animal facility of the British Columbia Cancer Research Centre (Vancouver, British Columbia, Canada). Six- to 8-week-old mice were
sublethally irradiated with 350 cGy from a 137Cs source the
day before being intravenously injected with human cells. In some
experiments, mice were injected intraperitoneally 3 times a week for 2 weeks from day 15 to day 30 posttransplant with the following
combination of human growth factors: 10 µg per mouse of Steel factor
(SF; Amgen, Thousand Oaks, CA), 6 µg per mouse of interleukin-3
(IL-3; Novartis, Basel, Switzerland), 6 µg per mouse of
granulocyte-macrophage colony-stimulating factor (GM-CSF; Novartis),
and 10 U per mouse of erythropoietin (Epo; StemCell) per injection.
These mice were killed 2 hours after the last injection of growth
factors. Cells were flushed from the shafts of the 4 hind leg long
bones of all mice using a syringe and 21-g needle prefilled with cold
Hanks' balanced salt solution supplemented with 5% fetal calf serum
(HF; StemCell) plus 5% pooled normal human serum (HF/5% HS), and a
single-cell suspension was obtained by gentle aspiration.
Flow cytometry.
For immunophenotyping of human cells, suspensions were first incubated
for 10 minutes at 4°C with HF/5% HS supplemented with 3 mg/mL of
an antimouse Fc receptor antibody (2.4 G2) to block Fc receptors and
prevent nonspecific antibody binding39 and were then
labeled for 30 minutes at 4°C with antihuman CD34-fluorescein isothiocyanate (FITC; 8G12)40 and CD19-phycoerythrin (PE)
and CD20-PE (Becton Dickinson [BD], San Jose, CA), or antihuman
CD71-PE (OKT-9), CD45-PE, CD15-FITC, and CD66b-FITC (Pharmingen,
Mississauga, Ontario, Canada), or glycophorin-A-FITC (10F7) and
antimouse Ter-119-PE (Pharmingen), or CD41 (3H2) only, or CD34-FITC
(8G12) and CD38-PE (BD), as described.41,42 Various
phenotypes within the viable (propidium iodide-negative
[PI ], Sigma Chemical Co, St Louis, MO) fraction
were determined using a FACSort (BD) and LYSIS II software (BD).
Assessment of CD34+ cells was further restricted to cells
with medium-to-high forward light scattering (FSC) and low side light
scattering (SSC) properties. Positivity in all cases was defined as
fluorescence that exceeded 99.98% (ie, >5 such events in 20,000 analyzed) of that obtained with irrelevant isotype-matched control
antibodies labeled with the corresponding fluorochromes. For in vitro
assays of human progenitor activity, cells labeled with both antihuman
CD45-PE and CD34-FITC were isolated by fluorescence-activated cell
sorting (FACS) as described.41,42
For intracellular staining and analysis of human cells producing
different hemoglobins (Hb), similar appearing, well-hemoglobinized, erythroblast-containing colonies generated in direct methylcellulose assays or in methylcellulose assays of 6-week LTC were harvested after
12 to 14 days of growth, pooled, and suspended in phosphate-buffered saline (PBS; StemCell). Because erythroid colonies from fetal liver and
cord blood progenitors develop slightly faster, these were generally
harvested on day 12, whereas those derived from adult marrow were
usually harvested 2 days later. Cells obtained from engrafted NOD/SCID
mice were washed 1 to 3 times in PBS and then incubated for 1 hour at
room temperature in 1 mL of 4% formaldehyde in PBS (BDH Inc, Toronto,
Ontario, Canada). Glutaraldehyde (EM grade) in PBS (Polyscience Inc,
Warrington, PA) was then added (final concentration, 0.01%), and the
cells were vortexed vigorously before centrifugation and resuspension
in 250 µL of 5% nonfat dry milk in PBS for 10 minutes at room
temperature. Cells were finally resuspended in 500 µL of 0.01%
Triton X-100 (Bio-Rad, Richmond, CA) in PBS/0.1% bovine serum albumin
(BSA; StemCell) and stained with FITC- and PE-labeled isotype control
antibodies (or streptavidin-PE [SAv-PE; Pharmingen] alone in some
cases) or antihuman Hb F-FITC (or HbF-biotin + SAv-PE in some cases; Isolab Inc, Akron, OH) plus anti-glycophorin A-PE (30 minutes at room
temperature). The cells were then washed once, resuspended in PBS, and
analyzed by FACS. To maximize specificity, gates were set first to
exclude cell debris (very low FSC events) and then to select positive
cells within the glycophorin A+ population.
In vitro progenitor assays.
Assays for colony-forming unit-erythroid (CFU-E), burst-forming
unit-erythroid (BFU-E), colony-forming unit-granulocyte-macrophage (CFU-GM), and colony-forming unit granulocyte, erythroid, monocyte, megakaryocyte (CFU-GEMM) were performed in methylcellulose
medium (H4330; StemCell) supplemented with 50 ng/mL of human SF and 20 ng/mL each of IL-3 and GM-CSF (Novartis), IL-6 (Cangene, Mississauga, Ontario, Canada), granulocyte colony-stimulating factor (G-CSF; StemCell), and 3 U/mL Epo as described.43
Colony-forming units-megakaryocyte (CFU-Mk) were assayed
separately in a serum-free agarose culture medium supplemented with 50 ng/mL of human thrombopoietin (TPO; Genentech, San Francisco, CA) and
10 ng/mL each of IL-3 and IL-6 to allow specific identification by
APAAP staining of pure Mk ( 3 CD41+ cells/colony) as well
as mixed Mk colonies (containing other lineages as well as
CD41+ Mk), as previously described.44
Cells were assayed for LTC-IC in 6-week cocultures that contained mixed
feeders of M2-10B4 and Sl/Sl fibroblasts genetically engineered to
produce human IL-3 (10 ng/mL), human G-CSF (130 ng/mL), and human SF
(10 ng/mL), as previously described.43 LTC-IC frequencies
were determined by limiting dilution analysis of the proportions of
negative cultures (<1 CFC produced per culture) in groups of cultures
seeded with different numbers of test cells (3 to 4 dilutions spanning
a 30-fold range, 13 to 24 cultures per dilution). Otherwise, LTC-IC
values were determined by dividing the total CFC output obtained from
bulk culture assays by the CFC per LTC-IC value determined for the
respective type of LTC-IC (see Table 3).
Proliferation bioassays of murine and human Epo.
Purified recombinant murine and human Epo were gifts from Dr E. Goldwasser (University of Chicago, Chicago, IL) and Stem Cell, respectively. Murine BAF/3 cells expressing the murine Epo
receptor45 were obtained from Dr G. Krystal (Terry Fox
Laboratory, Vancouver, British Columbia, Canada) and maintained in RPMI
with 10% fetal calf serum (FCS; Stem Cell) supplemented with 0.5 ng/mL
murine IL-3 (Terry Fox Laboratory). Human TF-1 cells46 were
obtained from Dr R. Kay (Terry Fox Laboratory) and maintained in RPMI
with 10% FCS and 2.5 ng/mL human GM-CSF. To assay the response of
BAF/3 cells to murine and human Epo, 104 log phase cells
were incubated for 24 hours at 37°C in 100 mL of RPMI with 5% FCS
containing various concentrations of Epo. To assay the response of TF-1
cells, log phase cells were first incubated for 24 hours at 37°C in
RPMI with 10% FCS and were then transferred at 104 per 100 mL RPMI plus 5% FCS plus Epo for another 48 hours. In both cases,
3H-thymidine (8 µCi/mL, 2.0 Ci/mmol; NEN Life Science
Product, Inc, Boston, MA) was added 4 hours before harvesting the cells for measurement of their radioactivity.
Statistical analysis.
All values shown are the mean ± SEM unless otherwise indicated.
Significant differences (P < .05) were established using the Student's t-test.
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RESULTS |
Frequencies of different types of CFC in human fetal liver by
comparison with cord blood and adult marrow.
Table 1 shows a comparison of the
frequencies of different types of CFC in the light density fraction of
9 independently assessed human fetal liver samples. For comparison, the
same reagents and procedures were used to analyze the light density
cells from 12 human cord bloods and 7 normal adult human marrow
samples. The frequency of every progenitor type measured (relative to
the total nucleated cell population) was found to be higher in fetal liver than in either cord blood or adult marrow, although none of the
differences was statistically significant (P > .05) due to
the extensive variability in CFC frequencies in individual samples. The increased frequency of fetal liver CFC was most
pronounced for CFU-GEMM (60- and 250-fold higher, respectively, in
fetal liver), intermediate for BFU-E/CFU-E and CFU-GM (~50-fold and 100-fold higher, respectively, in fetal liver), and least for CFU Mk
(~3-fold higher in fetal liver).
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Table 1.
Comparison of the Frequencies of Different Types of CFC
Per 105 Light-Density Human Fetal Liver, Cord Blood, and
Adult Marrow Cells and Their Representation in the Total CFC
Compartment
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Assessment of the frequency and CFC output of human fetal liver
LTC-IC.
Preliminary experiments demonstrated that LTC-IC assays could not be
performed directly on low-density human fetal liver cells, because this
resulted in the rapid production of large numbers of macrophages, which
was usually followed within the first 2 weeks by a complete destruction
of the feeder layer (data not shown). However, initial removal of
lin+ cells from the input fetal liver population allowed
the presence of LTC-IC to be shown. Limiting dilution assays were used
to determine the frequency of LTC-IC in the lin
cells (62% ± 8% CD34+ cells) obtained from 4 different fetal livers and in the
CD34+CD38 cells isolated by FACS from 2 of these (same samples as used to obtain CFC frequencies in Table 1).
This showed that 1.0% ± 0.2% of the lin fetal
liver cells were LTC-IC. Calculation of the LTC-IC content of the
original low-density fetal liver cell population (assuming all LTC-IC
were recovered in the lin fraction) gives a
frequency (7.6 ± 1.7 LTC-IC per 104 light-density fetal
liver cells) which is approximately twice that previously
determined for low-density adult marrow cells.43 In the 2 experiments in which a direct comparison was made of the frequency of
LTC-IC in the lin (70% CD34+) and
CD34+CD38 cells obtained from the same
fetal liver samples, isolation of the
CD34+CD38 cells gave a further
approximately 4-fold enrichment of the LTC-IC, at a final purity of
4%, and showed that approximately 79% of the LTC-IC were recovered in
the CD34+CD38 cell fraction.
Assessment in 1 of these experiments of the frequency of fetal liver
LTC-IC using parental M2-10B4 cells (not producing human growth
factors) as feeders showed that the number of LTC-IC detected was
2.5-fold lower than the value obtained with the human
growth-factor-producing feeders. This difference is similar to what we
have previously found for adult marrow and cord blood
LTC-IC.43
The LTC-IC frequency data obtained from the limiting dilution analyses
were also used to calculate the average 6-week output per LTC-IC of
each type of CFC assessed. Table 2 shows
the results obtained together with those we have previously reported
for adult marrow and cord blood LTC-IC assayed using the same reagents
and procedures.43 The results for lin
CD34+ and CD34+CD38 fetal
liver LTC-IC were not different (data not shown) and have been
combined. By comparison with cord blood and adult marrow, fetal liver
LTC-IC were found to produce, on average, significantly more (P < .05) CFC of all types, as well as proportionately more CFU-E and
BFU-E (P = .06 for fetal liver v cord blood LTC-IC and P = .05 for fetal liver v adult marrow LTC-IC).
Examination of the CFC produced by single LTC-IC (cultures initiated
with a cell number containing on average  1 LTC-IC) showed a wide
range of values (from 0 to 464 CFC per well). A broad range of CFC
outputs was previously also described for individual adult marrow
LTC-IC detected using human marrow feeders.47 On average,
20% of the wells plated with a single fetal liver LTC-IC in the
present experiments contained both CFU-GM and BFU-E (and/or CFU-GEMM)
as compared with 11% and less than 4% for single cord blood and adult
marrow LTC-IC, respectively, assayed under the same conditions.
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Table 2.
Output of Different Types of CFC in LTC From LTC-IC
Changes According to the Ontological Origin of the LTC-IC
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Analysis of the hematopoietic cell types regenerated in sublethally
irradiated NOD/SCID mice transplanted with human fetal liver cells.
Figure 1 and Table 3 show
the time course of changes in the numbers of various types of human
hematopoietic cells detectable in the marrow of sublethally irradiated
NOD/SCID mice assessed up to 16 weeks after the intravenous injection
of 107 low-density human fetal liver cells. For comparison,
comparable data (where available) from previously published time course
studies of the patterns of engraftment seen with transplants of
107 low-density human cord blood42 or 2 × 107 low-density normal adult human marrow
cells48 are shown. As found for other sources of human
hematopoietic stem cells, less than 0.1% of any of the types of cells
in human fetal liver that were injected could be found in the marrow of
the mice 2 to 3 days later. After this, there was a rapid (within 2 to
4 weeks) increase in the marrow of the mice of many primitive and
mature human hematopoietic cell types. All of these reached peak levels approximately 6 to 8 weeks posttransplant and were then sustained for
the duration of the experiment, ie, up to 16 weeks
posttransplant. Human populations detected included
CD45/71+ cells, CD34+ cells, CD41+
(Mk-lineage) cells, BFU-E, CFU-E, CFU-GM, and LTC-IC.
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| Fig 1.
Kinetics of appearance of different types of human
hematopoietic cells in NOD/SCID mice transplanted with 107
low-density human fetal liver cells. Values shown represent the numbers
calculated to be present in each entire mouse, assuming 2 femurs
and 2 tibias comprise 25% of this value.60 Results from 2 independent experiments have been pooled (2 to 6 mice per time point).
Previously published data for grafts of 107 light-density
human cord blood cells (dashed line)42 or 2 × 107 light-density human adult marrow cells (dotted
line)48 are shown for comparison.
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Table 3.
Kinetics of LTC-IC Production in NOD/SCID Mice
Transplanted With 107 Light-Density Human Fetal Liver or
Cord Blood or 2 × 107 Light-Density Adult Marrow Cells
Are Similar
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Analysis of the distribution of human CFC subtypes among those present
at different times posttransplant (Table 4)
indicates that there was a slight shift between week 2 and week 6 in
favor of granulopoietic CFC, as seen previously in NOD/SCID mice
engrafted with human cord blood and adult marrow cells. Also of note is the observation that fetal liver grafts regenerated relatively more
progeny LTC-IC (8-fold over input) than was achieved previously with
transplants of cord blood (5-fold over input) or adult marrow (decreased 2-fold below input). Assessment of the number of glycophorin A+ cells (erythroblasts) showed large numbers of these to
be present in the fetal liver and cord blood-engrafted mice, but none
was detected in the marrow-engrafted mice
(Fig 2). A 2-week course of 6 injections of
human Epo, SF, IL-3, and GM-CSF restored the ability of adult human
erythroid progenitors to mature into erythroblasts in vivo (as
previously reported for similar transplants in SCID mice49), but had no effect (P > .05) on terminal
human erythropoiesis in the mice engrafted with fetal liver or cord
blood cells (Fig 2; P = .4 and P = .2, respectively) or
on any other type of human progenitor in any of the groups (data not
shown).
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Table 4.
Relative Frequencies of Different Types of CFC in Human
Fetal Liver-Engrafted NOD/SCID Mice Assessed at Different Time Points
Posttransplant
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| Fig 2.
Total human (CD45/71+) and mature erythroid
(glycophorin A+) cells in NOD/SCID mice transplanted with
equivalent grafts of human fetal liver, cord blood, or adult marrow
cells and then injected with recombinant human growth factors ( ) or
not ( ) as described in the text. The difference plus and minus
growth factors for all pairs was not significant (P > .05)
except for glycophorin A+ cells in the mice transplanted
with adult marrow (P << .001).
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Because we had found that freshly isolated human fetal liver LTC-IC
display an enhanced erythropoietic activity in vitro by comparison with
those present in cord blood or adult marrow, it was of interest to also
examine the types of CFC produced by the human LTC-IC regenerated in
fetal liver-engrafted mice. As shown in
Table 5, these were characterized by the
same increased output of CFU-E and BFU-E as the LTC-IC present in
freshly isolated fetal liver suspensions.
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Table 5.
Comparison of Different Types of CFC Produced by the
LTC-IC Regenerated in Human Fetal Liver-Engrafted NOD/SCID Mice at
Different Time Points Posttransplant
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Murine Epo has a reduced activity on human cells.
A reduced ability of adult versus fetal and neonatal human erythroid
cells to undergo terminal maturation in NOD/SCID mice would be
anticipated from a slight differential (decreased) responsiveness of
adult cells to Epo if the level of murine Epo in the mice were limiting. We therefore compared the ability of murine and human Epo to
stimulate murine and human Epo receptor-expressing cells. When murine
targets were used (murine Epo receptor-expressing BAF/3
cells),45 both preparations gave superimposable
dose-response curves (data not shown). The unitage of the human Epo
preparation could thus be used to assign a unitage to the murine Epo
preparation. The 2 preparations were then tested for their
abilities to stimulate the proliferation of human TF-1
cells.46 As shown in Fig 3, concentrations of murine Epo that had an equivalent ability to human
Epo to stimulate murine cells were approximately 10-fold less active on
human cells. Thus, levels of murine Epo that are sufficient to sustain
murine erythropoiesis might also be adequate to stimulate human
erythropoietic cells of fetal (and neonatal) but not adult origin,
which could thus explain the unique requirement of adult human
erythroid precursors for additional (exogenously administered) human
Epo to terminally differentiate in an in vivo murine environment.
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| Fig 3.
Dose-response curves showing the differential mitogenic
effects of murine and human Epo on human TF-1 cells. The specific
activities of both preparations were standardized using murine
Epo-expressing BAF/3 cells.
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Fidelity of globin gene expression in the human erythroid progeny of
cells produced under different conditions from ontologically distinct
sources.
Terminally differentiating human erythroblasts generated in vitro from
progenitors produced in LTC or in NOD/SCID mice or in LTC initiated
with cells from NOD/SCID mice were assessed for the type of Hb they
contained by FACS analysis of intracytoplasmically stained fixed cell
preparations (as described in Materials and Methods). For each sample,
the preparation was counter-stained with antihuman glycophorin A
antibody labeled with a different fluorochrome to enhance the
specificity of the procedure. Representative profiles are shown in
Fig 4. The data for all sources of
erythroblasts are summarized in Table 6.
Most of the erythroblasts within, or derived in vitro from progenitors
from, suspensions of fetal liver or cord blood expressed high levels of
HbA and HbF. As expected, most of the erythroblasts present in freshly
obtained adult marrow contained HbA and very few contained HbF,
although increased production of HbF+ cells was seen when
marrow progenitors were stimulated to generate erythroblasts in vitro.
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| Fig 4.
FACS profiles of glycophorin A+ cells
obtained directly from fetal liver, cord blood, and adult marrow or
from directly plated 12- to 14-day-old erythroid colonies generated in
methylcellulose assays of human fetal liver, cord blood, or adult
marrow cells or from human CFU-E/BFU-E produced in NOD/SCID mice
engrafted for 6 weeks with human fetal liver, cord blood, or adult
marrow cells. Harvested cells were stained with anti-glycophorin A and
anti-Hb antibodies as described in the Materials and Methods. Note that
the intensity of staining with the 2 different anti-Hb antibodies
provide only a relative measure of the amount of HbA and HbF in each
cell and therefore cannot be used to compare absolute HbA and HbF
expression levels.
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Table 6.
Proportion of Erythroid Cells Produced From Different
Sources of Progenitors Containing Different Types of Hb
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Analysis of the erythroblast progeny of progenitors generated from
human fetal liver cells within the microenvironment of the adult
NOD/SCID mouse bone marrow, or in vitro from LTC-IC generated in human
fetal liver-engrafted mice, showed most of these to continue to be
HbF+. Similarly, the relatively low proportion of
HbF+ erythroblasts produced in vitro from progenitors
generated in adult human marrow-engrafted mice was the same as for
those generated from freshly isolated adult human marrow progenitors.
Interestingly, in the human cord blood-engrafted mice, the proportion
of HbF+ erythroblasts obtained from progenitors produced 6 weeks posttransplant had declined to half the level seen in
erythroblasts obtained from freshly isolated progenitors (P < .04).
To examine the dual expression of HbA and HbF in the same cells (which
precluded the use of glycophorin A-staining), cells were harvested from
maturing erythroid colonies generated in vitro from progenitors of
different sources. These provide an enriched, although not pure,
population of erythroblasts. As can be seen in
Fig 5, most of the Hb+ cells
obtained from colonies of fresh fetal liver origin contained both HbA
and HbF. Similar results were obtained for colonies generated from
human BFU-E isolated from fetal liver-engrafted mice. In contrast, many
more of the Hb+ cells in colonies produced by adult marrow
contained exclusively HbA.
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| Fig 5.
Demonstration of cells containing both HbF and HbA in 12- to 14-day-old erythroid colonies generated in methylcellulose assays of
human fetal liver or adult marrow CFC and CFC obtained from NOD/SCID
mice transplanted 6 weeks previously with human fetal liver cells.
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DISCUSSION |
The present study shows a number of important features of human fetal
liver hematopoiesis that identify differences in how this process is
regulated in the fetus and in the adult. At both stages of ontogeny,
terminally differentiating erythroid and myeloid cells make up the
majority of the light-density population in the hematopoietic tissues
in which they are produced. However, erythroid cells predominate in the
fetal liver, whereas myeloid cells predominate in the adult. In
contrast, more primitive cells, detected in vitro as erythroid and
granulopoietic CFC, although present at a higher overall frequency
(relative to the total number of cells) in fetal liver, were also found
to be present in both fetal liver and adult marrow at similar
frequencies to each other (Table 1).5,33 The lack of a
selective increase in the number of erythroid progenitors accumulated
in vivo does not argue in favor of a model of increased commitment of
fetal hematopoietic stem cells to the erythroid pathway to explain the
predominance of erythroid cells seen in the terminally differentiating
fetal liver populations. Nevertheless, a shift in that direction was suggested here by the increased output of erythroid progenitors from
individual fetal liver (as compared with adult marrow) LTC-IC under LTC
conditions. These conditions are nonpermissive for erythropoiesis and
hence allow detection of the first progenitors to commit to that pathway.
We also found that the frequency of megakaryocyte progenitors was
increased, although to a much smaller degree, whereas the frequency of
multipotent CFC was increased to a greater extent than the
granulopoietic and erythroid progenitors. As noted by others,7,50 we observed the maximum size of colonies
produced in vitro by all types of CFC to be larger than those produced by their adult marrow counterparts. However, this increased CFC proliferative potential of fetal liver CFC is not realized in vivo,
because the ratio of CFC to mature cells is decreased. This could occur
either because the microenvironment of the fetal liver promotes the
more rapid differentiation of fetal progenitors and/or because a higher
rate of apoptosis occurs among their progeny. Because the composition
of the mature cell compartment in adult marrow and fetal liver also
differ, the mechanisms responsible for affecting these changes in cell
output must be distinct for each lineage.
Given the large increases seen in the frequency of CFC in the human
fetal liver (overall, ~70-fold relative to adult marrow), the
discovery that the frequency of LTC-IC is only 2-fold higher was
unanticipated. This means that the ratio of CFC to LTC-IC in the fetal
liver is much higher than in adult marrow. Interestingly, this feature
was reproduced in the LTC system, in which we also noted an increased
(4-fold higher) 6-week output of CFC by comparison with adult marrow
LTC-IC (72 v 18 CFC per LTC-IC). A similarly enhanced output of
CFC by fetal liver LTC-IC was noted by Roy et al7 using a
shorter (5-week) assay on parental M2-10B4 cell feeders that may detect
a less primitive LTC-IC subset.43 Similarly, a greater
output of progeny from fetal versus adult sources of CD34+
CD45RACD7131 or
CD34+CD38 cells51 in
stroma-free cytokine-supplemented cultures has been reported. Such a
result could be explained by a decreased self-renewal probability of
primitive fetal cells resulting in an expanded yield of CFC or a
reduced cell cycle time resulting in an accumulation of more CFC
through more cell generations. It is also possible that the progeny of
fetal LTC-IC may acquire an ability to form colonies in semisolid media
without such a precipitous loss of proliferative potential, or they may
be less susceptible to conditions that cause apoptosis of their adult
counterparts. Current evidence argues against the first possibility,
because fetal stem cells of both human52 and
murine17,19 origin have been found to display increased
rather than decreased self-renewal properties. In addition, evidence of
a shorter cell cycle time and faster differentiation of fetal
hematopoietic cells has been reported.3,16,53
The present studies also provide evidence of the intrinsic
determination of a number of other properties of fetal human
hematopoietic cells that are linked to their embryonic status. These
include an increased output of erythroid CFC by fetal LTC-IC, an
increased sensitivity of fetal CFU-E to factors that support their
terminal differentiation, and an increased production of HbF by their
erythroblast progeny. All of these characteristics were shown to be
stably transmitted to the progeny of fetal stem cells stimulated to
proliferate and differentiate in the marrow of NOD/SCID mice; in some
cases, even those generated after transfer of in vivo-derived LTC-IC to
cultures containing murine stromal feeder layers engineered to produce
human G-CSF, IL-3, and SF. Less stability of HbF production was seen
previously when fetal sheep cells were transplanted into adult
sheep54 or when human fetal cells were cultured for
prolonged periods in the absence of stroma.55 From such
results it has been suggested that the type of hemoglobin produced can
be influenced by the environment in which the cells are generated. On
the other hand, a fetal human erythroid program was seen in the
erythroid cells produced in both patients56,57 and
sheep58 transplanted with fetal cells, consistent with the
results reported here. The continued production in the presence of
murine stroma of human progenitors that remain committed to a high
level of HbF and low HbA synthesis appears to represent only one of
several manifestations of a preserved fetal hematopoietic program that
affects multiple stages of differentiation. It seems unlikely that the
different behaviors of corresponding stages of differentiation of fetal and adult cells will reflect a single common underlying intrinsic molecular alteration. However, it is interesting to note that, even in
adult erythroid precursors, HbF expression is enhanced by strong growth
factor stimulation34,59 and that a feature of fetal
erythroid precursors appears to be their enhanced sensitivity to growth
factor stimulation. Such findings thus invite speculation as to whether
some basic differences in the intracellular signaling machinery of
fetal hematopoietic cells may account for their distinct biology.
Investigations of this possibility are now underway.
| |
ACKNOWLEDGMENT |
The authors thank the staff of the Stem Cell Assay Service for their
assistance in the initial preparation of primary human cell samples,
Jessyca Maltman and Maya Sinclaire for their assistance in the animal
studies, Gayle Thornbury and Giovanna Cameron for operating the FACS,
and Tara Palmater for secretarial assistance. We are also grateful to
E. Goldwasser (University of Chicago), P. Lansdorp, G. Krystal, and R. Kay (Terry Fox Laboratory), and Amgen, Cangene, Isolab, Novartis, and
Stem Cell for valuable gifts of cells or reagents.
| |
FOOTNOTES |
Submitted February 11, 1999; accepted June 14, 1999.
Supported by grants from the National Institutes of Health (PO1
HL55435), Novartis (Basel, Switzerland), and the National Cancer
Institute of Canada (NCIC) with funds from the Terry Fox Run. T.L.H.
holds a United Kingdom Leukemia Research Fund Senior Lectureship and
P.P.Y.C. holds a Roman Babicki Graduate Studentship from the University
of British Columbia. C.J.E. is a Terry Fox Cancer Research Scientist of
the NCIC.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Connie J. Eaves, PhD, Terry Fox Laboratory,
601 W 10th Ave, Vancouver, BC, V5Z 1L3 Canada; e-mail:
connie{at}terryfox.ubc.ca.
| |
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ABSTRACT
INTRODUCTION