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Next Article 
Blood, Vol. 94 No. 7 (October 1), 1999:
pp. 2161-2168
Direct Evidence for Multiple Self-Renewal Divisions of Human In Vivo
Repopulating Hematopoietic Cells in Short-Term Culture
By
H. Glimm and
C.J. Eaves
From the Terry Fox Laboratory, British Columbia Cancer Agency; and
the Department of Medical Genetics, University of British Columbia,
Vancouver, BC, Canada.
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ABSTRACT |
Recently, culture conditions that stimulate the proliferation of
primitive hematopoietic cells defined by various phenotypic and
functional endpoints in vitro have been identified. However, evidence
that they support a high probability of self-renewal leading to a large
net expansion in vitro of transplantable cells with lympho-myeloid
repopulating ability has been more difficult to obtain. The present
study was designed to investigate whether the low overall expansion of
human repopulating hematopoietic cells seen in vitro reflects a
selective unresponsiveness of these rare cells to the growth factors
currently used to stimulate them or, alternatively, whether they do
proliferate in vitro but lose engrafting potential. For this, we used a
high-resolution procedure for tracking and reisolating cells as a
function of their proliferation history based on the loss of
cellular fluorescence after staining with (5- and 6-)
carboxyfluorescein diacetate succinimidyl ester. The results show that
the vast majority of long-term culture-initiating cells and in vivo
lympho-myeloid competitive repopulating units present in 5-day
suspension cultures initiated with CD34+ human cord blood
and fetal liver cells are the progeny of cells that have divided at
least once in response to stimulation by interleukin-3, interleukin-6,
granulocyte colony-stimulating factor, Steel factor, and Flt3-ligand.
Thus, most human repopulating cells from these two sources are
stimulated to undergo multiple divisions under currently used
short-term suspension culture conditions and a proportion of these
retain engraftment potential.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
HEMATOPOIESIS originates in a small
number of hematopoietic stem cells. These cells are defined by their
ability to differentiate into all of the blood cell lineages as well as
to generate progeny with the same unrestricted hematopoietic potential. Because specific markers of these unique functional properties of stem
cells have not yet been identified, their detection and enumeration
require the use of retrospective assays. These involve demonstrating an
ability to repopulate all compartments of the hematopoietic system
after intravenous injection of the cells under study into suitable
myeloablated hosts.1 In the murine system, the use of
histocompatible but genetically distinguishable donor and host
combinations has been combined with limiting dilution analysis to allow
many properties of stem cells to be elucidated.2-5 A
similar approach to the quantitation of transplantable human stem cells
has been possible using sublethally irradiated immunodeficient nonobese
diabetic-scid/scid (NOD/SCID) mice as
recipients.6,7
Self-renewal divisions are believed to be responsible for the expansion
of the hematopoietic stem cell compartment that occurs both during
fetal life and posttransplant, as well as for the stable maintenance of
this population throughout normal adult life.8,9 However,
the precise mechanisms that regulate the outcome of hematopoietic stem
cell divisions are largely unknown. Recent progress in the development
of methods for obtaining highly enriched populations of stem cells and
the availability of an increasing number of recombinant growth factors
to which they can respond has stimulated a plethora of studies to
identify conditions that will support a net expansion of stem cells in
vitro. These have shown that factors like Flt3-ligand (FL), Steel
factor (SF), interleukin-6 (IL-6), IL-11, and thrombopoietin (TPO)
are important synergizing growth factors active on these
cells.7,10-12 However, to date, large (>10-fold) and
continuing net expansions of cells with retention of stem cell activity
have not been shown. One possible explanation for this may lie in the
reversible loss of engraftment activity that might be related to the
transit of stem cells through specific phases of the cell
cycle.13 Thus, the activation of human stem cells from
G0 into G1 might be expected to cause a similar
rapid loss of their transplantability, as recently observed.14 If this reflects a transient change in the
homing properties of stem cells,15 rather than an intrinsic
alteration in their growth and differentiation potential, it should be
possible to demonstrate the passage of some stem cells through multiple self-renewal divisions in vitro. To investigate this possibility, we
have used a high-resolution procedure for tracking successive generations of hematopoietic cells in asynchronously dividing populations.16 In this procedure, cells are labeled at
the beginning of the experiment with (5- and 6-)
carboxyfluorescein diacetate succinimidyl ester (CFSE). The precise
halving of fluorescence at each successive mitosis then allows multiple
daughter generations to be reproducibly resolved. The application of
this procedure to human CD34+ cell populations
isolated from both cord blood and fetal liver and then
cultured for 5 days in serum-free medium containing FL, SF, IL-3,
IL-6, and granulocyte colony-stimulating activity (G-CSF) has
shown that the majority of stem cells detectable after culture have
already undergone multiple self-renewal divisions within the 5 days in vitro.
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MATERIALS AND METHODS |
Cells.
Samples of human fetal liver were obtained from elective abortuses of
10 to 16 weeks of gestation and dispersed either by pressing minced
tissue fragments through a sieve or using dispase as
described.17 Samples of heparinized cord blood were
obtained from normal, full-term infants delivered by cesarean section. In both cases, institutional guidelines were adhered to for access and
use of human material. Low-density cells (<1.077 g/mL) were isolated
after centrifugation of the cells on Ficoll-hypaque (Pharmacia, Piscataway, NJ) and cryopreserved in 10% dimethyl sulfoxide plus 90%
fetal calf serum (FCS; StemCell Technologies, Vancouver, BC, Canada)
until required. Cells were then thawed, pooled ( 5 cord blood samples
and 10 fetal liver samples per experiment), and cells expressing
surface antigens characteristic of more mature hematopoietic cells
(lin+ cells) were removed using StemSep columns
(StemCell Technologies) according to the manufacturer's instructions.
CFSE labeling and isolation of labeled cells.
Cells were washed, resuspended at 5 × 106 cells/mL in
phosphate-buffered saline (PBS), and CFSE (Molecular Probes, Eugene, OR) was added at 20 µmol/L. After 10 minutes at 37°C, further uptake of the dye was stopped by addition of ice-cold Hanks' balanced salt solution (HBSS) supplemented with 20% FCS. The cells were then
washed twice, the second time in HBSS without FCS, and finally resuspended at 5 to 10 × 105 cells/mL in Iscove's
medium supplemented with BIT (BIT9500; StemCell Technologies),
10 4 mol/L 2-mercaptoethanol (Sigma Chemicals, St
Louis, MO), 40 µg/mL low-density lipoproteins (Sigma), 50 ng/mL human
TPO (Genentech, Palo Alto, CA), and 0.1 µg/mL colcemid (GIBCO-BRL,
Burlington, Canada). Cells were then cultured overnight at 37°C to
allow the efflux of all CFSE not stably bound to intracellular
protein.16
After washing in HBSS with 2% FCS and 5% human serum (kindly provided
by D. Hogge, Terry Fox Laboratory), CFSE-labeled cells were stained
with Cy5-conjugated 8G12 (anti-CD34) antibody (kindly provided by P.M.
Lansdorp, Terry Fox Laboratory) for 30 minutes at 4°C. The cells
were then washed 2 times with HBSS containing 2% FCS with 1 µg/mL
propidium iodide (PI; Sigma Chemicals) being added to the second wash.
A narrow gate (32 channels wide using a 1,024-channel log amplifier)
was then used to define a sort gate for isolating a subset of
homogeneously CFSE+ PI CD34+
cells with low side-scattering characteristics, as
described.16 Using a FACStar Plus cell sorter equipped with
a 5-W argon laser and a 30-mW helium neon laser (Becton Dickinson, San
Jose, CA), 2 adjacent populations of CFSE-stained CD34+
cells were isolated and subsequently manipulated identically, but
in parallel, to allow the majority of CD34+ cells to be used.
Short-term suspension cultures.
CFSE-stained CD34+ cells were cultured for 5 days at 5 to
10 × 104 cells/mL in 35-mm petri dishes (StemCell
Technologies) in serum-free Iscove's medium containing the same
supplements described above, but with replacement of the TPO by 20 ng/mL IL-3 (Novartis, Basel, Switzerland), 20 ng/mL IL-6 (Cangene,
Mississauga, Ontario, Canada), 20 ng/mL G-CSF (StemCell Technologies),
100 ng/mL SF (prepared and purified from cDNA transfected COS cells in
our laboratory), and 100 ng/mL FL (Immunex Corp, Seattle, WA). After 72 hours an equal volume of fresh medium and growth factors was added to
each culture. Another 2 days later, the cells were harvested, washed in
HBSS with 2% FCS, restained with Cy5-conjugated 8G12 (anti-CD34) antibody, and the cells then resorted according to their CFSE fluorescence relative to a parallel aliquot of cells that had been
cultured under the same conditions and for the same time but in the
presence of 0.1 µg/mL colcemid (to inhibit cell division). This made
it possible to fix the positions of the fluorescence peaks
corresponding to the undivided cells and first-, second-, and
third-generation cells present after 5 days in culture. Comparison of
the colony-forming cell (CFC) content (see below) of the cultured cells
before and after the second sort showed no significant selective progenitor loss or enrichment as a result of this procedure (recovery of cord blood CFC = 99% ± 9%, n = 4; recovery of fetal liver CFC = 86% ± 24%, n = 3).
In vitro progenitor assays.
CFC and 6-week long-term culture-initiating cell (LTC-IC) assays (on
murine fibroblast feeders engineered to produce human SF, IL-3, and
G-CSF) were performed as previously described.18
Competitive repopulation unit (CRU) assay.
Nonobese diabetic/severe combined immunodeficient (NOD/SCID) mice were
bred and maintained in the animal facility of the British Columbia
Cancer Research Centre (Vancouver, BC, Canada) under sterile conditions
in microisolator cages and were provided exclusively with autoclaved
food and water. To assay human CRU, test cells plus 106
irradiated (15 Gy) normal human bone marrow (carrier) cells
were injected intravenously into 6- to 12-week-old mice that had just been given 350 cGy total body 137Cs irradiation. Mice were
killed 6 to 8 weeks later and the contents of both tibiae and femurs
suspended in HBSS plus 2% FCS. To minimize nonspecific binding of
antibodies, human and murine Fc receptors were blocked first by
incubating the cells in 5% human serum and 2.4G2 (an anti-mouse Fc
receptor antibody19). Separate aliquots of cells were then
stained for 30 minutes at 4°C with fluorescein isothiocyanate
(FITC)-conjugated human anti-CD34 (8G12) and phycoerythrin (PE)-conjugated human anti-CD19 and anti-CD20 antibodies (Becton Dickinson) for the detection of human (CD34
CD19/20+) B-lineage cells, or with FITC-conjugated human
anti-CD15 (Becton Dickinson), anti-CD66b (Pharmacia), and PE-conjugated
human anti-CD45 (Becton Dickinson) and anti-CD71 (OKT9) for the
detection of human (CD45/71+ CD15/66b+) myeloid
cells. Mice with detectable human lymphoid and myeloid engraftment (5 positive events each per 20,000 assessed) were counted as positive. All
other mice were considered negative. Levels of specific staining were
set based on parallel analyses of additional aliquots of the same test
cells similarly incubated with irrelevant isotype-matched control
antibodies labeled with the corresponding fluorochromes. Gates were
then set to exclude levels of fluorescence which included greater than
99.99% of these negative control samples.
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RESULTS |
Expansion of committed and primitive human hematopoietic progenitors in
short-term cultures of CFSE-stained CD34+ cord blood and
fetal liver cells.
Table 1 shows the changes in total cell,
CFC, and LTC-IC numbers seen when cord blood or fetal liver cells were
first incubated overnight in the presence of TPO and colcemid, a
CFSE+ CD34+ subpopulation then isolated, and
the cells cultured for an additional 5 days in the presence of FL, SF,
IL-3, IL-6, and G-CSF. During the 5 days of culture, the total
cell expansion and the increase in CFC numbers in both
types of culture were similar ( 35-fold). Recoveries of LTC-IC
numbers were more variable but they also generally increased several
fold. These results are similar to previously published results for
cells cultured without prior CFSE staining.7,20
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Table 1.
Changes in CFC and LTC-IC Numbers After Culturing
CD34+ Fetal Liver and Cord Blood Cells for Five
Days in the Presence of IL-3, IL-6, SCF, G-SCF, and FL
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CD34+ cord blood and fetal liver cells proliferate
rapidly but asynchronously in response to stimulation by FL, SF, IL-3,
IL-6, and G-CSF.
Fluorescence-activated cell sorting (FACS) analysis of cells harvested
from 5-day cultures of CD34+ cord blood or fetal liver
cells reproducibly resolved 6 generations of progeny, in addition to a
small peak of residual undivided (very bright) cells (0.2% ± 0.1%, n = 6, and 0.3% ± 0.5%, n = 5, respectively, of the final
cells in the cord blood and fetal liver cell cultures). These undivided
cells represented, in both cases, 4% of the initial
CD34+ cells originally seeded into the cultures. The
results of a representative experiment are illustrated in
Fig 1. The average distribution of cells
among successive generations of progeny for all experiments (5 with
fetal liver, 6 with cord blood) is shown in
Fig 2A. Companion data for cells still
expressing CD34 are shown in Fig 2B. As can be seen, the results for
the cord blood and fetal liver cultures were similar, the majority of
the cells in both types of culture being derived from cells that had
divided at least 5 times, although a higher proportion of fetal liver
cells divided more than 6 times. These data were used to calculate the
proportion of the original cells that produced each of the different
sized clones seen, assuming that all cell divisions were symmetric
(with respect to subsequent proliferative activity) and that there was
also no cell loss (Fig 3). The
percentage of starting cells contributing to clones of different sizes
was calculated by the equation ([a/2x]/b) where a
is the number of cells derived from x divisions in vitro,
x is the number of divisions, and b is the total number of initial cells. According to this calculation (and the underlying assumptions), the proportion of initial CD34+ cord blood or
fetal liver cells that undergo only 1 to 3 divisions under the culture
conditions used here is small and of similar magnitude for both tissues
(3.5% to 4.0%). However, the proportion of initial CD34+
cells able to complete >6 divisions was significantly higher for
fetal liver cells than for cord blood cells (P < .05).
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| Fig 1.
Representative FACS profiles of CFSE-stained cord blood
cells before (A) and after (B and C) culture for 5 days in the presence
of IL-3, IL-6, SCF, G-CSF, and FL. The initial sort gate used to
isolate the cells placed in culture is indicated in (A). The arrows in
(B) and (C) indicate the persisting undivided cells identified by
comparison to a control aliquot of the cells cultured under the same
conditions but in the presence of colcemid to suppress cell division.
The profile shown in (C) was derived by gating on the highly
fluorescent (CFSE+) cells to allow a better
discrimination of the undivided population.
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| Fig 2.
Distribution of total cells produced (A) and total
CD34+ cells (B) produced by fetal liver (FL,  ) and
cord blood cells (CB,  ) according to their proliferation history (0 to >6 divisions) during 5 days of suspension culture in the presence
of IL-3, IL-6, SF, G-CSF, and FL. Values shown are the mean ± SEM
from 5 (FL) and 6 (CB) experiments. Note change of scale on the
ordinate axis: the lefthand side indicates the scale for cells that
underwent  3 divisions, the righthand side indicates the scale for
cells that underwent 4 divisions.
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| Fig 3.
Minimal percentages of initial CD34+ fetal
liver (FL, ) and cord blood cells (CB, ) with the ability to
divide from 0 to 6 times in short-term cultures containing 5 growth
factors (same experiments as shown in Fig 2). A similar calculation
could not be made for cells that underwent more than 6 divisions;
however, by subtraction this can be estimated as 78% ± 3% for fetal
liver cells and 45% ± 5% for cord blood cells. Values shown are the
mean ± SEM from 5 (FL) and 6 (CB) experiments.
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The rate of loss of CD34 expression as function of the number of
divisions completed within 5 days is shown in
Fig 4. Here it can be seen that greater
than 15% of all cells in the clones (and single undivided cells)
present at the end of the 5-day culture were still CD34+.
This proportion was, as expected, lowest in the largest clones (>64
cells) and highest (50%) in the smaller clones (2 to 8 cells/clone), again assuming that asymmetric divisions or cell loss are
not significant during the period studied.
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| Fig 4.
Proportion of CD34+ cells in clones of
different sizes derived from CD34+ fetal liver (FL, )
and cord blood cells (CB, ) after 5 days in culture with 5 growth
factors (same experiments as shown in Figs 2 and 3). Values shown are
the mean ± SEM.
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Most LTC-IC in short-term cultures of CD34+ cord blood
and fetal liver cells are confined to the smallest clones.
Because of previous reports that primitive human hematopoietic
progenitors detected as LTC-IC are enriched in the
PKH2bright (undivided) cell fraction of CD34+
cells that have been stimulated for 7 days with IL-3, IL-6, and SF,21,22 we sorted the cultured cells in the above
experiments according to the number of cell cycles they had completed
(on the basis of their CFSE fluorescence) and then assayed each
fraction for its LTC-IC (and CRU, see below) content. In a first set of experiments, the 2 fractions isolated consisted of 1 fraction of
undivided cells and a second fraction containing all of the rest (with
a gap 25 channels wide to minimize cross-contamination of the 2 fractions). As shown in Table 2, all or
most (>88%) of the LTC-IC were consistently found in the fraction
that had undergone at least 1 division in cultures initiated with
either CD34+ cord blood or fetal liver cells. In the second
series of experiments, the cells harvested from the 5-day cultures were
separated into those that had undergone 2 divisions, and those that
had undergone 3 divisions, again with a gap of 25 channels to
minimize cross-contamination. In these latter experiments, most of the
LTC-IC in the cord blood cultures were in the populations that had
undergone 2 cell divisions, but a somewhat greater proportion of the
fetal-liver-derived LTC-IC were found among later generations of
progeny that had divided more rapidly
(Table 3).
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Table 2.
Distribution of LTC-IC Between the Undivided and
Postmitotic Fractions of Cells Obtained From Five-Day Cultures of
CD34+ Fetal Liver and Cord Blood Cells That Had Not
Divided During This Period (Undivided Cells) and That Had
(Postmitotic)
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Table 3.
Distribution of LTC-IC Between Populations Generated in
Five-Day Cultures of Fetal Liver and Cord Blood Cells Isolated
According to Their Proliferation History
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Distribution of human CRU among different generations of
growth-factor-stimulated CD34+ cord blood and fetal liver
cells.
In the above 2 series of experiments, only a small proportion (<20%)
of all the cultured cells were used to perform cell counts, phenotyping
studies, and CFC and LTC-IC assays. The remaining 80% to 93% were
used to examine the CRU content by injecting the cells from each of the
2 fractions collected into sublethally irradiated NOD/SCID mice (the
approximate proportion of the culture injected per mouse for each group
can be derived directly from the data shown in Fig 3). As
shown in Table 4, in 4 of 4 experiments the
majority of human cells with any kind of NOD/SCID repopulating activity
had completed at least 1 cell division within 5 days of culture. This
included all of those with lympho-myeloid potential. Moreover, in 5 of
5 experiments a greater proportion of the CRU was found in the
populations that had undergone at least 3 cell cycles
(Table 5). Limiting dilution analysis of
the data pooled from all like experiments was used to determine the
frequencies of CRU in the subpopulations of cord blood cells isolated
according to their pretransplant cycling history in culture. The
average frequency of CRU in the population of cells that had divided
less than 3 times was 1 per 14,000 cells (range defined by ± SEM, 1 per 9,600 cells to 1 per 20,700 cells). The corresponding value for
cells that had divided 3 or more times was 1 CRU per 1,300,000 cells
(range, 1 per 900,000 to 1 per 1,800,000 cells), with 63% the CRU
detected in the cultured cells being found in this latter group. The
calculated change in total CRU numbers in these experiments was an
2.4 fold increase. This compares favorably to our previous results
with cord blood cells using similar culture conditions7 and, again, argues against any likelihood of a toxic effect of the CFSE
staining and sorting procedure.
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Table 4.
Distribution of CRU Between the Undivided and
Postmitotic Fractions of Cells Obtained From Five-Day Cultures of
CD34+ Fetal Liver and Cord Blood Cells That Had Not
Divided During This Period (Undivided Cells) and That Had
(Postmitotic)
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Table 5.
Distribution of CRU Between Populations Generated in
Five-Day Cultures of Fetal Liver and Cord Blood Cells Isolated
According to Their Proliferation History
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DISCUSSION |
In this study we show that the majority of transplantable hematopoietic
stem cells present in 5-day cultures of human fetal liver or cord blood
CD34+ cells stimulated with IL-3, IL-6, G-CSF, SF, and FL
have executed at least 3 full cell cycles. These results demonstrate
unequivocally that human hematopoietic stem cell activity assessed by
their ability to regenerate lymphoid and myeloid progeny after 6 to 8 weeks in irradiated NOD/SCID mice7,23 is not lost
inevitably or irreversibly when cells are activated mitogenically with
soluble growth factors. In addition, our findings show that most, if
not all, of the stem cells present in CD34+ populations of
human cord blood and fetal liver can be stimulated to rapidly
proliferate in vitro by this 5-growth factor combination.
At first glance, these findings appear to contradict those of others
who have reported rapid losses of human14,22 and
murine24 stem cell activity in the first hours after
exposure of such cells to a similar combination of growth factors, and
who have detected retained stem cell activity among those cells thought
to have remained quiescent in 7-day cultures (because they did not show a major loss of PKH2 fluorescence). From such observations, it was
concluded that retention of functions essential to the integrity of
stem cell activity were lost rapidly upon cell activation and could be
preserved by cells that were able to resist activation over prolonged
periods of time in vitro. A similar model has recently been proposed
based on measurements of CD34+CD38 cell
outputs by sequentially replated single fetal liver
cells.25 On the other hand, we previously showed that the
magnitude of LTC-IC amplification within individual clones generated
over a 10-day period in culture by adult human marrow
CD34+CD38 cells is clone
size-independent.26 Also, time-course studies of
single-cell cultures of CD34+CD38 adult
human marrow and cord blood cells have shown that greater than 95% of
those able to respond to the combination of growth factors studied here
will complete a first division within 5 to 8 days,
respectively.27 Similarly, we and others have shown that
these conditions can also support the retroviral infection of up to
30% of the human fetal liver or cord blood stem cells (CRU) present at
the end of a 5- to 6-day infection protocol.28-30 Taken
together, these findings argue strongly against the concept of a
significant persisting quiescent population of transplantable stem
cells in cultures of human cord blood and fetal liver cells stimulated
by FL plus SF and IL-3, IL-6, and G-CSF. However, they do not rule out
the possibility that such cells may exist at low frequency. One
possible explanation for the apparent discrepancy between this
conclusion and the results reported by Traycoff et al22 may
lie in the different methodologies used to separate divided and
undivided cells. It is possible that the lower resolution of PKH2
staining to discriminate cell division history allows some cells that
divide up to 4 times to be misclassified as undivided, which could be
sufficient to reconcile the present and previously published findings.
In addition, recent evidence of cell-cycle-associated variations in
engraftment potential has been presented.13 Such variations, if confirmed, would imply that the homing and
differentiation/self-renewal properties of stem cells can be
independently regulated and, hence, may be separately manipulated.
Additional support for this hypothesis is provided by evidence of an
involvement of  4 1 integrins31-33 and the
chemokine receptor CXCR415 in stem cell homing, but not in
the intrinsic ability of stem cells to proliferate and differentiate
per se.33,34 The implication of such a mechanism for the
findings reported in this study is that the CRU activity measured would
have underestimated the true stem cell content of the various
populations assessed because only those in G0 or G1 may have been detectable.
The present findings raise a number of other issues of interest to
understanding the control of human stem cell self-renewal. Recently,
much attention has been raised concerning the long-term sustainability
of hematopoiesis by CD34+ cells.35-38 In the
present studies, all cultures were initiated with highly purified
CD34+ FACS-sorted cells. Hence the likelihood that the
results obtained could be attributed to contaminating
CD34 cells seems low. Moreover, the culture
conditions used in this study were similar to conditions that have not
been found to support the generation or maintenance of human
hematopoietic repopulating activity by activated
CD34 cells.38 Thus, it can be concluded
that few, if any, of the stem cell progeny we have found to be the
result of divisions in vitro originated from contaminating
CD34 (lin) cells. However,
further studies will be required to investigate whether the in vivo
self-regenerative activity39 of human stem cells amplified
in vitro has also been retained, as has been shown in the murine
system.11 It is also important to note that the yield of
LTC-IC (and CRU) is lower than that predicted by the number of
divisions they have undergone. Part of this decreased yield is likely
due to loss of LTC-IC (and/or CRU) activity during asymmetric cell
divisions in which one of the progeny differentiates. In addition, as
noted above, CRU enumeration may be subject to cell-cycle changes
leading to underestimates of stem cell numbers in proliferating populations.
Another interesting point is the discrepant rate of loss we observed in
LTC-IC and in vivo lympho-myeloid repopulating activities. Given the
evidence that some cells detectable as LTC-IC may represent "later" cell types than those identified as CRU,40
the finding that repopulating activity persisted through more cell
generations than LTC-IC activity (or was selectively associated with
more rapidly cycling cells) was unexpected. However, other examples of
a dissociation in LTC-IC and CRU function in murine cells have been
reported,41,42 indicative of differences in the molecular mechanisms required for primitive cells to be detected in these 2 assays. It is also possible that the efficiencies of LTC-IC and CRU
detection may be differentially affected by the culture conditions to
which they were exposed here (eg, by changes in their cycling status,
as discussed above). If cells with these differences were also to be
differently distributed between progeny that had undergone different
numbers of cell divisions, this would impact on the assumed numerical
differences in CRU and LTC-IC ascribed to their cell-cycle history.
Therefore, a next step will be to examine these issues by other
experimental strategies.43
For clinical gene-therapy applications using retroviral vectors, a
culture system that supports stem cell self-renewal divisions at
practically useful frequencies is essential. At the same time, completion of a single division should be sufficient if access of the
virus into the cell is not limiting. At least for human cord blood
targets, our findings now show that prolonging the overall exposure of
these cells to FL, SF, IL-3, IL-6, and G-CSF for more than 5 days is
not advantageous. Improved yields of retrovirally transduced
hematopoietic stem cells are, therefore, likely to benefit more from
the pursuit of strategies for manipulating homing activity
postinfection. Studies to investigate such possibilities are now underway.
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ACKNOWLEDGMENT |
The authors thank Maya Sinclaire and Jessica Maltman for assistance
with the animal work, Gayle Thornbury, Giovanna Cameron, and Rick Zapf
for assistance in cell sorting, the staff of Stem Cell Assay Service
for initial hematopoietic cell processing, and Caroline Lonsdale and
Tara Palmater for manuscript preparation. The authors also thank
Cangene, Immunex, Novartis, StemCell, and Dr Peter
Lansdorp (Terry Fox Laboratory) for generous gifts of reagents.
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FOOTNOTES |
Submitted March 11, 1999; accepted May 25, 1999.
Supported by the National Cancer Institute of Canada (NCIC) with funds
from the Terry Fox Run, the National Institutes of Health
(POI-HL55435), and Novartis (Canada), and a grant from the Dr. Mildred
Scheel Stiftung für Krebsforschung, Bonn, Germany, to H.G. 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 C.J. Eaves, PhD, Terry Fox Laboratory, 601 W 10th Ave, Vancouver, British Columbia, Canada V5Z 1L3; e-mail:
connie{at}terryfox.ubc.ca.
| |
REFERENCES |
1.
Eaves CJ, Eaves AC:
Anatomy and physiology of hematopoiesis, in
Pui C-H
(ed):
Childhood Leukemias. New York, NY, Cambridge University Press, 1999, p 53.
2.
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ABSTRACT
INTRODUCTION