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Blood, Vol. 93 No. 3 (February 1), 1999:
pp. 838-848
Notch1-Induced Delay of Human Hematopoietic Progenitor Cell
Differentiation Is Associated With Altered Cell Cycle Kinetics
By
Nadia Carlesso,
Jon C. Aster,
Jeffrey Sklar, and
David T. Scadden
From the Department of Experimental Hematology, Partners AIDS
Research Center and MGH Cancer Center, Massachusetts General Hospital,
Harvard Medical School; and the Department of Pathology, Brigham and
Women's Hospital, Harvard Medical School, Boston, MA.
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ABSTRACT |
Hematopoiesis is a balance between proliferation and differentiation
that may be modulated by environmental signals. Notch receptors and
their ligands are highly conserved during evolution and have been shown
to regulate cell fate decisions in multiple developmental systems. To
assess whether Notch1 signaling may regulate human hematopoiesis to
maintain cells in an immature state, we transduced a vesicular
stomatitis virus G-protein (VSV-G) pseudo-typed bicistronic murine stem
cell virus (MSCV)-based retroviral vector expressing a
constitutively active form of Notch1 (ICN) and green fluorescence
protein into the differentiation competent HL-60 cell line and primary
cord blood-derived CD34+ cells. In addition, we observed
endogenous Notch1 expression on the surface of both HL-60 cells and
primary CD34+ cells, and therefore exposed cells to Notch
ligand Jagged2, expressed on NIH3T3 cells. Both ligand-independent and
ligand-dependent activation of Notch resulted in delayed acquisition of
differentiation markers by HL-60 cells and cord blood
CD34+ cells. In addition, primary CD34+
cells retained their ability to form immature colonies, colony-forming unit-mix (CFU-mix), whereas control cells lost this
capacity. Activation of Notch1 correlated with a decrease in the
fraction of HL-60 cells that were in G0/G1
phase before acquisition of a mature cell phenotype. This enhanced
progression through G1 was noted despite preservation of
the proliferative rate of the cells and the overall length of the cell
cycle. These findings show that Notch1 activation delays human
hematopoietic differentiation and suggest a link of Notch
differentiation effects with altered cell cycle kinetics.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
BLOOD CELL DEVELOPMENT is a process of
highly regulated proliferation and differentiation during which a small
number of multipotent cells are sequentially recruited to
generate tens of billions of mature circulating cells daily.
Organization of the bone marrow includes stem cells that are capable of
self renewal with some daughter cells differentiating into highly
proliferative precursor cells ultimately yielding the cellular
components of blood.1,2 Acquisition of lineage-specific
differentiation features is associated with an irreversible loss of
multipotentcy and declining proliferative potential. A common model of
hematopoietic regulation poses differentiation as inversely related to
proliferative capacity as cells pass through the intermediate stages of
blood cell maturation. The pool of highly proliferative precursor cells responds to physiological stress with consequent expansion in the
numbers of differentiated mature blood elements. The proliferation of
this precursor pool has been shown to be induced by cytokines, many of
which also have prodifferentiative effects. However, the spectrum of
action of cytokines is broad, with cytokines such as kit ligand and
flt-3 ligand enhancing survival of primitive cells as a function
distinct from that of augmenting proliferation.3,4 Because
the maintenance of a primitive precursor cell population is critical
for amplification of proliferation, it may be hypothesized that there
are also regulated environmental cues within hematopoietic tissue that
impede differentiation. These signals may be envisioned to delay
acquisition of a terminally differentiated phenotype, thereby
permitting ongoing proliferation.
One signaling pathway that seems likely to affect the balance between
proliferation and differentiation involves Notch, which has been shown
to influence the cell fate of diverse types of progenitor cells in a
wide range of multicellular animals. Notch genes encode large, highly
conserved type 1 transmembrane glycoprotein receptors composed of a
series of iterated structural motifs, including epidermal growth factor
(EGF)-like repeats that have been implicated in ligand
binding and an intracellular ankyrin-like repeat region that is
critical for downstream signaling events.5 Physiological
activation of Notch signaling occurs through binding of one of a series
of ligands, such as Jagged1, Jagged2, and Delta, that are also
transmembrane proteins.6-11 Of the four Notch genes described thus far in vertebrates,12-15 most studies have
been conducted with Notch1, which was originally identified as a gene that is rearranged by a recurrent (7;9) chromosomal translocation associated with a subset of human T lymphoblastic
leukemias.16 Subsequent studies have shown that
constitutive activation of Notch1 results in blocked differentiation
and expansion of chick retinal precursor cells7,17 and in
the prevention of murine myoblast8 and neuroblast
differentiation.18 Recent studies also suggest a
physiological role for Notch1 in the regulation of hematopoiesis.
Notch1 is expressed by CD34+ bone marrow progenitor
cells,19 and at least one Notch1 ligand, Jagged1, is
expressed by marrow stromal cells.11 Activation of Notch1
has recently been shown to result in the inhibition of granulocytic
differentiation of murine cells.11,19
Normal cell differentiation is thought to primarily occur through cell
functions accomplished in the G0 /G1 phase
of the cell cycle.20-24 Conversely, disruption of cell
cycle checkpoints has been associated with perturbed proliferation and
differentiation, as shown by cancer cells. As constitutively active
forms of Notch1 have the capacity to inhibit differentiation and act as
oncoproteins,25,26 we hypothesized that these effects might
be mediated, at least in part, through altered regulation of the cell
cycle. In this study, we show that Notch1 activation by either
transduction of an activated form or through cognate ligand stimulation
has the ability to delay the differentiation of human HL-60 cells and primary CD34+ progenitor cells, and that this effect
correlates with shortening of the G1 phase of the cell cycle.
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MATERIALS AND METHODS |
cDNA expression constructs.
cDNAs encoding a constitutively active form on Notch1 consisting of the
intracellular domain (base pairs [bp] 5308-7665; amino acids [aa]
1770-2555; ICN) or an inactive form on ICN with a deletion removing the coding sequence of the ankyrin repeat region (deletion bp
5570-6618; aa 1859-2207; ICN AR)27 were subcloned into
the multicloning site of the retroviral vector MSCV-GFP28
(Fig 1A). As the result of an internal
ribosomal entry sequence and a cDNA encoding enhanced green fluorescent
protein (GFP; Clontech Laboratories, Palo Alto, CA) that lie 3'
of the multicloning site, the 5' long terminal repeat promoter of
this vector drives the expression of a single biscistronic mRNA
encoding both Notch1 polypeptides and GFP.
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| Fig 1.
Retroviral constructs and transduction. (A) Structure of
Notch1 and ICN and of the retroviral vector MSCV 2.2. ( ),
hydrophobic leader; ( ), EGF-like repeats; ( ), LNR repeats;
(||), transmembrane domain;
( ), ankyrin repeats; Q, glutamine-rich region; P, PEST sequence. The
MSCV LTR, IRES motif, and GFP gene of the MSCV 2.2 vector are
indicated. (B) Flow cytometric analysis for GFP in HL-60-transduced
cells. Contour plots represent fluorescence intensity for GFP on the
x-axis and cell forward scatter on the y-axis. (C) Flow cytometric
analysis for Notch1 on gated HL-60 and CD34+ GFP-positive
cells. The polyclonal antibody directed against the intracytoplasmatic
region of Notch1, T3, was used as described. Histograms are expressed
as fluorescence intensity for Notch1 on the x-axis (log scale) and cell
count on the y-axis. Superimposed are fluorograms with anti-GST control
antibody on MSCV-GFP cells (left, filled curve), anti-T3 antibody on
MSCV-GFP cells (middle curve, solid line), and anti-T3 antibody on
MSCV-ICN/GFP cells (right curve, dotted line). Fluorograms with anti-T3
antibody on MSCV-GFP cells and uninfected cells were superimposable.
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Retroviral transduction.
MSCV-GFP vectors were cotransfected into 293T cells using a calcium
phosphate precipitation method25,29 with pKat (kindly provided by M. Finer, Cell Genesys Inc, Foster City, CA), an
amphotropic packaging plasmid, and pCMV-VSV-G, a plasmid encoding the
vesicular stomatitis virus G-glycoprotein (kindly provided by T. Friedmann, University of San Diego, CA). Supernatants containing
pseudo-typed retrovirus were collected at 48 and 72 hours and were used
to infect primary CD34+ cells and the HL-60 cell line.
CD34+ progenitors were cultured in Iscove's modified
Dulbecco's medium (IMDM) (Mediatech Inc, Harndon, VA) containing 10%
fetal calf serum (FCS; Sigma, St Louis, MO) (IMDM 10) supplemented with Kit ligand (KL; 50 ng/mL), thrombopoietin (TPO; 20 ng/mL), FLT3 ligand
(FLT-3 L; 50 ng/mL), and interleukin-3 (IL-3; 50 ng/mL) (R & D,
Minneapolis, MN) for 48 hours before infection, to induce cells to
enter into cell cycle. Stimulated CD34+ cells or HL-60
cells were washed in phosphate-buffered saline (PBS; Sigma) and
resuspended at the concentration of 1 to 2 × 105/mL
in 1 mL of 80% retroviral supernatant and 20% fresh complete medium
plus Polybrene (final concentration, 8 µg/mL; Sigma). Cells in
suspension were placed in a 48-well plate (Becton Dickinson Inc,
Franklin Lakes, NJ), spinoculated at 1,700 revolutions per minute for
50 minutes, then incubated at 37°C 5% CO2 for an
additional 6 to 8 hours, washed, and resuspended in fresh medium
overnight. A second and a third infection were conducted on the
following days using an identical procedure. Infection efficiency was
evaluated by GFP expression 4 days after the last infection. ICN and
ICNAR overexpression was also determined using the T3 antibody
against Notch1.30 In some experiments, GFP-positive cell
enrichment was performed by fluorescence-activated cell sorting (FACS
Vantage; Becton Dickinson, San Jose, CA).
Cells and cell culture.
Cord blood samples were obtained from the Pediatric Research Institute,
University of St Louis, MO, according to the guidelines established by
the Human Investigation Committee. Mononuclear cells were separated by
Ficoll-Hypaque (Pharmacia-Biotech, Uppsala, Sweden)
density gradient centrifugation, and CD34+ cells purified
by immunomagnetic bead selection (Milteny Biotec, Auburn, CA),
according to the manufacturer's instructions. Transduced CD34+ cells were grown in IMDM 10 supplemented with KL (20 ng/mL), FLT-3 L (20 ng/mL), and IL-3 (50 ng/mL; R & D) or PIXY
321 (50 ng/mL; Immunex, Seattle, WA). Cells were initially
cultured in 0.2 mL final volume at the density of 0.5 to 1 × 106/mL in 96-well plate wells. Fresh media and
cytokines were added every 3 to 4 days and cells were subsequently
transferred to 24-well plates (Becton Dickinson Labware) to maintain
the initial density.
The promyelocytic cell line HL-60 was obtained from American Type
Culture Collection (Rockville, MD). HL-60 and
HL-60-transduced cell lines were maintained in RPMI medium
supplemented with 10% FCS (R10). To induce HL-60 cells to
differentiate, 1 to 2 µmol/L all-trans-retinoic acid (ATRA; Sigma) or
2 to 4 nmol/L tetrahydro-phorbol-ester acid (TPA; Sigma) were added to
R10. In these experiments, cells were cultured in 1 mL of R10 with or
without inducing agents in 24-well plate wells; an equal volume of
fresh medium was added at day 3 of culture.
Human embryonic kidney-derived 293T cells and murine NIH-3T3 cells
were grown in Dulbecco's modified Eagle's medium supplemented with
10% FCS (D10). CD34+ cells, HL-60 cells, and NIH3T3 cells
were cultured at 37°C under 5% CO2, whereas 293T cells
were cultured at 33°C under 5% CO2. For coculture
experiments, 3T3pBABE and 3T3pBABE-Jagged2 cells10 were
cultured in 24-well plate wells to approximately 70% of confluence. Medium was then aspirated and the monolayers rinsed gently with PBS
before adding 2 mL of HL-60 or CD34+ cell suspension at the
density of 0.1 × 106/mL. ATRA, TPA, or medium was
added to the 3T3/HL-60 cocultures while 3T3/CD34+
cocultures were supplemented with FLT-3 L (20 ng/mL) and PIXY (20 ng/mL). Half of the medium volume was replaced every 3 days with fresh
medium. During 3T3/CD34+ cell cocultures, primary cells
were split and transferred from their 3T3 monolayers onto fresh
monolayers (70% confluent) every 5 to 6 days. Before analysis, HL-60
or CD34+ cells were detached from the feeder layer either
by vigorous pipetting or by EDTA treatment (no differences were noted
when the two methods were compared) and subsequently washed in R10 medium.
Clonogenic progenitor assays were performed with cells obtained from
the liquid cultures or the 3T3 cocultures. Harvested cells were
cultured in methylcellulose (Stem Cell Technologies, Vancouver, British
Columbia, Canada) supplemented with IL-3 (50 ng/mL), KL (50 ng/mL), and
erythropoietin (EPO; 2 U/mL; Amgen, Thousand Oaks, CA) according to the
manufacturer's recommendations. Colonies were evaluated by phase
microscopy and scored according to standard morphologic criteria after
14 days.
Antibodies.
Production, purification, and characterization of polyclonal rabbit
antibodies against a portion of the intracellular domain of Notch1
defined as T3 (aa 1763-1877), the Notch ligand Jagged2 (J2), and
glutathione S-transferase (GST) have been previously described.10,30 Intracellular staining was performed using fixing and permeabilization solutions (Fix and Perm) from Caltag (Burlingame, CA), according to manufacturer's instructions. Antibodies against T3, J2, and GST were added to permeabilized cells at the concentration of 5 µg/mL for 30 minutes at room temperature. Cells were than washed twice and incubated with the monoclonal antibody (MoAb) goat anti-rabbit conjugated to phycoerythrin (PE; Sigma) (1 µg/mL).
Fluoroisothiocyanate (FITC) and PE-conjugated MoAbs directed against
CD11b, CD14, CD15, and CD34 (Becton Dickinson) were used to analyze
differentiation. Cells were incubated with 0.5% human IgG (Sigma) in
PBS for 20 minutes at 4°C to block the Fc receptor before staining
with antigen-specific antibodies. Flow cytometric analysis was
performed using the FACScalibur instrument (Becton Dickinson).
Cell cycle analysis.
Cell cycle analysis was performed using the DNA binding dye TOPRO-3
(Molecular Probes, Eugene, OR). The emission spectrum of TOPRO-3
permits its use in conjunction with other fluorochromes. Three-color
analysis of GFP, TOPRO-3, and PE-conjugated antibodies was performed
simultaneously by flow cytometry. Cells stained with PE-conjugate
antibodies were fixed in PBS 1% formaldehyde for 30 minutes,
permeabilized in 0.1% Triton X-100 (Sigma) for 30 minutes at room
temperature, washed, and resuspended in a PBS 1 mg/mL RNAse (Sigma) and
1 µg/mL TOPRO-3 for a minimum of 30 minutes at 4°C before FACS analysis.
Cell cycle synchronization experiments were conducted on
HL-60-transduced cells enriched for GFP (to 95% purity) by cell
sorting. Cells were incubated with hydroxyurea (10 mmol/L) or nocodazol (200 ng/mL; Calbiochem Inc, La Jolla, CA) for 22 hours in R10. Then,
cells were washed twice with PBS and incubated in the fresh medium
without hydroxyurea or nocodazol. Cell cycle profiles were analyzed by
bromodeoxyuridine (BrdU; Sigma) incorporation every 6 hours for the
following 24 hours. BrdU-pulsed cells were fixed in 70% ethanol at
 20°C overnight and denatured by 2 N HCl, 0.5% Triton X-100 for 30 minutes at room temperature, followed by
neutralization with borate buffer (pH 8.5). Treated cells were
subjected to dual-color staining with anti-BrdU MoAb (Becton Dickinson)
followed by goat anti-mouse FITC-conjugated and 5 µg/mL
propidium iodide. Analyses were performed using the FACSCalibur flow
cytometer, and Cell Quest and Modefit software (Becton Dickinson).
Statistical analysis.
Equality of distributions for matched pairs of observations were tested
using the Wilcoxon matched-pairs signed ranks test. The analyses were
performed using the software STATA (r) (Stata Corporation, College
Station, TX).
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RESULTS |
The following two approaches were used to evaluate the effects of human
Notch1 on human hematopoiesis: (1) ligand-independent activation using
retroviral constructs driving the expression of a constitutively active
form of Notch1, termed ICN, and (2) ligand-dependent activation of
endogenous Notch1 using NIH 3T3 feeder cells engineered to express the
physiological ligand, Jagged2.10 To facilitate the former
approach, we used the retroviral vector MSCV-GFP, which is known to
drive high-level expression in hematopoietic progenitor cells and has
an internal ribosomal entry sequence that permits expression of cDNAs
of interest and a marker protein, GFP, from a single bicistronic mRNA.
To begin to investigate the effects of Notch1 activation on the
differentiation of human hematopoietic cells, we used the myeloid cell
line HL-60, which is capable of both granulocytic or monocytic
differentiation,31 as well as CD34+ primary
cells isolated from human cord blood. Differentiation toward the
myeloid lineage was evaluated by determining CD11b and CD14 expression,
surface markers that correlate with granulocytic and monocytic
differentiation.32-34
Expression of endogenous Notch1 and activated Notch1 in HL-60 cells
and CD34+ progenitor cells.
The efficiency of infection of HL-60 cells and CD34+
progenitor cells by MSCV-ICN/GFP or empty MSCV-GFP retroviruses (Fig
1A) was assessed by flow cytometric analysis; results of a
representative experiment are shown in Fig 1B. Independent experiments
(n = 4) showed a mean infection efficiency of 42% and 39% with
MSCV-GFP or MSCV-ICN/GFP retroviruses, respectively, for HL-60 cells.
Infection efficiency was more variable in CD34+ cells,
ranging from 5% to more than 30% of cells stably expressing GFP. To
assess expression of endogenous Notch1 and transduced ICN, flow
cytometric analysis was also performed after staining with
affinity-purified anti-Notch1 (Fig 1C). Both HL-60 cells and
CD34+ progenitor cells infected with MSCV-GFP expressed
endogenous Notch1, as shown by a shift in mean fluorescent staining
intensity with anti-Notch1 as compared with anti-GST control polyclonal IgG. Overexpression of ICN by MSCV-ICN/GFP-transduced cells was shown
by a further increase in the mean intensity of fluorescence (MIF).
Constitutively active Notch1 inhibits differentiation of HL-60 cells
and CD34+ progenitor cells.
Evaluation of the effect of constitutively active ICN was first studied
in HL-60 cells by following the induction of CD11b expression in the
presence of the granulocytic differentiation agent ATRA or in the
presence of the monocytic differentiation agent, TPA. HL-60 cells
transduced with MSCV-GFP or MSCV-ICN/GFP were plated at equal density
and induced with ATRA or TPA. A consistent delay in CD11b expression
acquisition was evident in the MSCV-ICN/GFP-transduced cells compared
with MSCV-GFP in multiple independent experiments (n = 4), as shown in
Fig 2. In a representative experiment, on ATRA stimulation (Fig 2A), at day 3 and 4 of culture only 12% and 30%
of HL-60 MSCV-ICN/GFP cells expressed CD11b, compared with 57% and
74% of HL-60 MSCV-GFP cells. By day 5, the HL-60 MSCV-CN/GFP cells
expressing CD11b increased in number and resembled that of control
cultures (92% v 98%). Figure 2C summarizes the results of
four experiments in the presence of ATRA. Similar results were observed
when cells were cultured in the presence of TPA. Upon TPA stimulation
(Fig 2B), at day 1 and 2 of culture, 8% and 28% of HL-60 MSCV-ICN/GFP
cells expressed CD11b, compared with 27% and 78% of HL-60 MSCV-GFP
control. By day 3, most of the HL-60 MSCV-ICN/GFP cells also expressed
CD11b. Figure 2D summarizes the results of two experiments. It should
be noted that in each experiment, the differentiation of
MSCV-ICN/GFP-infected cells lagged behind that of nontransduced
GFP-negative cells in the same culture, which constitutes an internal
control. In contrast to the results observed with ICN, introduction of
an inactive form of Notch1 bearing a deletion that spans the ankyrin
repeats, ICNAR, had no effect on differentiation in response to
either agent (data not shown).
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| Fig 2.
Effects of activated Notch1 on the differentiation of
HL-60 cells. Two-color flow cytometric analysis for GFP and CD11b on
transduced HL-60 cells in the absence (day 0) and in the presence of 1 µm ATRA (A and C) or 4 nm TPA (B and D). At each time point, cells
were harvested, washed, and labeled with anti-CD11b PE-conjugated MoAb
and analyzed by FACS. Contour plots (A and B) represent fluorescence
intensity for CD11b on the x-axis and for GFP on the y-axis. Values in
the line graphics (C and D) represent the mean percentage of
CD11b-expressing cells in the GFP-positive population in four (C) and
two (D) independent experiments. Bars represent standard error.
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To extend these observations to primary marrow progenitor cells, the
effect of constitutively activated Notch1 on the differentiation of
CD34+ cells was studied in liquid culture.
CD34+ cells infected with MSCV-GFP and MSCV-ICN/GFP grown
in the presence of 10% FCS and the cytokines KL, FLT-3 L, and IL-3 or
PIXY were harvested at different time points and stained with MoAbs
directed against CD34, CD11b, and CD14. CD34+ cells
infected with MSCV-ICN/GFP showed consistently lower expression of
CD11b and CD14 markers compared with CD34+ population
infected with the control vector MSCV-GFP.
Table 1 shows the mean percentage of CD11b
and CD14 expression in cultured cells from six experiments: at day 18, 23% of MSCV-ICN/GFP-positive cells expressed CD11b and 19% expressed
CD14, whereas 49% of the MSCV-GFP-positive cells expressed CD11b and
33% expressed CD14 (P = .04 and P = .06, respectively)
(Table 1). Expression of CD11b and CD14 in the GFP-negative cells in
both types of cultures did not differ from that observed in the
MSCV-GFP-transduced control cells (not shown), indicating that the
observed difference in expression of differentiation markers was
confined to the MSCV-GFP/ICN-transduced cell population.
To further assess the effect of ICN on the phenotype of progenitor
cells, we performed methylcellulose colony-forming assays as a measure
of a functional level of differentiation. We evaluated the
colony-forming ability of the cultured cells in three samples that
showed a high efficiency of transduction (approximately 30%). After 2 weeks of liquid culture, cells were replated at 10,000 cells/mL in
methylcellulose. CD34+ cells infected by MSCV-ICN/GFP gave
rise to a significantly higher number of colony-forming cells (CFC)
compared with CD34+ cells infected with MSCV-GFP (P = .03). CFC included colony-forming unit-granulocyte/macrophage
(CFU-GM), CFU-G, and CFU-M; CD34+ MSCV-ICN/GFP also
generated burst-forming units-erythroid (BFU-E) in some but not all
experiments. Figure 3A shows average colony number in three independent experiments. After 2 weeks in culture, progenitors infected with MSCV-ICN/GFP gave rise to an approximately twofold increase in CFC compared with controls (mean, 26 SEM ± 5 v mean, 14 SEM ± 4/10,000 cells). Thus, Notch1 activation
decreased the fraction of differentiated cells as measured by
immunophenotype and function, preserving progenitor cells in a state of
higher potentiality.
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| Fig 3.
Effects of Notch activation on colony-forming ability of
primary hematopoietic progenitors. (A) CD34+ MSCV-GFP and
MSCV-ICN/GFP cells were grown in liquid culture in presence of
cytokines. After 2 weeks, cells were obtained and plated in triplicate,
at a density of 10,000 cells/mL in methylcellulose supplemented with
IL-3, KL, and EPO. Columns represent the average of three experiments.
CFC are expressed per 10,000 cells. Error bars represent standard
deviation. Difference between the two populations is statistically
significant (P = .033). (B) CD34+
cocultivated with 3T3pBABE or 3T3pBABE-J2 were obtained and plated in
triplicate at the density of 1,000 cells/mL in methylcellulose, as
described above. Columns represent fold increase in colony formation by
progenitors exposed to Jagged2 versus control in two experiments.
CFU-mix included at least two myeloid lineages and erythroid cells as
assessed by phase microscopy. Error bars represent standard deviation.
*, P = .046; **, P < .03.
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Activation of endogenous Notch1 with the ligand Jagged2 inhibits
differentiation of HL-60 cells and CD34+ primary
cells.
One potential criticism of effects observed with constitutively active
forms of Notch1, such as ICN, is that their physiological relevance is
uncertain. We thus next assessed the ability of NIH 3T3 feeder lines
expressing Jagged2, a ligand for Notch1, to inhibit the differentiation
of HL-60 cells and CD34+ progenitor cells through
activation of endogenous Notch1. HL-60 cells cocultivated with NIH3T3
pBABE-Jagged2 cells showed a marked and consistent inhibition of
myeloid differentiation in response to ATRA and TPA as measured by
expression of CD11b and CD14 (Fig 4). In a
representative experiment (Fig 4A), 74% and 65% of the HL-60 cells
expressed at least one of these markers after 4 days of treatment with
ATRA or TPA, respectively, when cocultivated with NIH3T3 pBABE control
cells. In contrast, only 47% and 40% of HL-60 cells expressed at
least one of these markers after 4 days of treatment with ATRA or TPA,
respectively, when cocultivated with NIH3T3 pBABE-Jagged2 cells. In
addition, even among cells expressing CD14 and CD11b, there was a
decreased MIF. Figure 4B shows the kinetic of CD11b expression in HL-60
cells on stimulation with ATRA or TPA in three independent experiments.
The mean percentage of HL-60 cells expressing CD11b was 41% versus
27% at day 2, 49% versus 38% at day 3, and 60% versus 38% at day 4 when HL-60 cells were cocultured with NIH3T3 pBABE and NIH3T3 pBABE-J2,
respectively.
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| Fig 4.
Effects of Notch ligand-dependent activation on HL-60
differentiation. HL-60 cells were washed and seeded on NIH3T3pBABE and
NIH3T3pBABE-J2 cell monolayers at the density of 0.2 × 106/mL when in the absence and at the density of 0.4 × 106/mL when in the presence of ATRA or TPA. At days 2, 3, and 4 of coculture, HL-60 cells were detached from the 3T3 layer,
washed, and labeled with anti-CD11b PE-conjugated and anti-CD14
FITC-conjugated MoAbs and subsequently analyzed by two-color flow
cytometric analysis. (A) Two-color flow cytometric analysis at day 4 of
differentiation in the absence and in the presence of ATRA or TPA.
Contour plots represent fluorescence intensity for CD11b on the x-axis
and for CD14 on the y-axis. Contaminating NIH3T3 cells were excluded
from the analysis based on their different forward scatter (FSC) and
side scatter (SSC) and their negativity to CD15 (positive
100% on HL-60 cells). (B) Values in the line graphic represent the
mean percentage of HL-60 cells expressing CD11b during coculture with
NIH3T3pBABE and NIH3T3pBABE-J2 at days 0, 2, 3, and 4 after induction
of differentiation. Bars represent standard error.
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We next evaluated the effects of Notch ligand-dependent stimulation on
primary cells. Purified CD34+ were seeded on NIH 3T3-BABE
and NIH 3T3-pBABE-Jagged2 monolayers and analyzed after 7, 14, and 21 days for CD34, CD11b, and CD14 expression and for colony-forming
ability. Equivalent cell numbers were observed in the two culture
conditions at each time point. The absolute number of CD34+
cells and the intensity of CD34 expression decreased with time in both
culture systems. However, input CD34+ cells cocultivated
with NIH3T3 pBABE-Jagged2 showed a consistent delay in the decline of
CD34 expression, compared with CD34+ cocultivated with
NIH3T3 pBABE, in four independent experiments. The mean fraction of
cells expressing CD34 in the two populations was, respectively, 74%
versus 62% at day 7, 42% versus 36% at day 14, and 24% versus 19%
at day 21 (P < .01) (Table 2).
Conversely, expression of the myeloid differentiation markers CD11b and
CD14 increased with time in both cocultures. However, input
CD34+ cells exposed to Jagged2 showed consistently lower
percentages of cells expressing CD11b and CD14, with the largest
difference being observed at day 14. As shown in Table 2, the mean
percentage of cells expressing CD11b at day 14 was 22% in the
population cultured with Jagged2 versus 32% in the control population.
Similarly, the mean percentage of cells expressing CD14 was 17% versus
10%. These differences in expression of CD34, CD11b, and CD14 were statistically significant (P < .01). A representative
experiment is shown in Fig 5 in which at
day 7, 71% of the initial CD34+ population exposed to
Jagged2 ligand expressed CD34, as compared with 52% of control cells.
Conversely, at day 14 only 20% and 11% of the cells exposed to
Jagged2 ligand expressed CD11b and CD14, respectively, as compared with
32% and 21% of control cells.
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| Fig 5.
Effects of Notch ligand-dependent activation on
differentiation of primary hematopoietic progenitors. Purified
CD34+ were seeded on NIH3T3pBABE and NIH3T3pBABE-J2 at a
density of 0.1 × 106/mL in IMDM supplemented with 10%
FCS and FLT-3 L. At different time points cells were detached from the
3T3 layer, washed, and labeled with FITC- and PE-conjugated MoAbs
directed to CD34, CD11b, or CD14. The top panel of contour plots shows
fluorescence intensity of the cultured CD34+ cells
stained with control antibodies (IgG1-FITC and IgG1-PE) at day 14. The
middle panel shows fluorescence intensity of the cultured
CD34+ cells stained with anti-CD34 antibody at day 7;
CD34 intensity of fluorescence is expressed on the x-axis, and forward
scatter on the y-axis. The bottom panel shows two-color flow cytometric
analysis of cultured CD34+ cells at day 14; anti-CD11b
antibody fluorescence intensity is shown on the x-axis, and anti-CD14
antibody fluorescence intensity on the y-axis.
|
|
The observation that myeloid differentiation of CD34+ cells
was inhibited by exposure to Jagged2 suggested that a higher fraction of cells were being maintained in a relatively undifferentiated state.
This possibility was supported by the results of colony-forming assays
performed after 7 and 14 days of coculture. At all time points, cells
cocultivated with Jagged2-expressing feeder cells generated an
approximately twofold higher number of CFC colonies than control cells
(mean, 180 SEM ± 4.8 v mean, 125 SEM ± 4.8/1,000 cells
at day 7; and mean, 95 SEM ± 9 v mean, 54 SEM ± 11/10,000 cells at day 14, respectively). Most strikingly,
CD34+ progenitor cells exposed to Jagged2 generated
fourfold greater numbers of CFU-mix colonies compared with controls,
(mean, 15 SEM ± 5 v mean, 5 SEM ± 1.7/1,000 cells at
day 7; and mean, 4 SEM ± 1.3 v mean, 1 SEM ± 0.6/10,000
cells at day 14, respectively), consistent with the increased
maintenance of multipotent progenitors. Figure 3B represents fold
increase in CFU-C, CFU-GM, and CFU-mix formation by progenitors exposed
to Jagged2 relative to control. These data indicate that
stimulation of endogenous Notch1 by Jagged2 is capable of retaining
CD34+ progenitor cells in a more immature state, suggesting
a physiological role for Notch1 signaling in the regulation of
hematopoietic progenitor cell differentiation.
Constitutively active Notch1 alters cell cycle kinetics.
The effect of Notch1 activation on differentiation may be based on
multiple possible mechanisms. Since in most of developing systems,
withdraw from cell cycle is an event strictly correlated with ability
of cells to complete their differentiation program, we investigated
whether Notch1 activation alters cell cycle kinetics. Expression of ICN
did not affect either cell viability or cell proliferation, as
indicated by superimposable cell proliferation curves for HL-60
MSCV-GFP and HL-60 MSCV-ICN/GFP (Fig 6),
confirmed in multiple independent experiments. In multiple independent
observations, HL-60 MSCV-ICN/GFP cells consistently showed a lower
fraction of cells in G0 /G1 phase and a
higher fraction of cells in G2/M compared with HL-60
MSCV-GFP cells. Representative cell cycle profiles of infected HL-60
cells are shown in Fig 7A. Under conditions of exponential growth, nonsynchronized (untreated) HL-60 cells transduced with MSCV-ICN/GFP showed a lower percentage of cells in
G0 /G1 phase (41%) compared with control
cells HL-60 MSCV-GFP (52%) and HL-60 MSCV-ICNAR/GFP (49%) and a
higher proportion of cells in G2/M phase (26%) compared with controls
(12% and 14%, respectively). To determine the kinetics of the
observed altered cell cycle distribution, cell cycle profiles were
analyzed by propidium iodide and BrdU incorporation following
G1 synchronization after release from hydroxyurea or
nocodazol block. In five independent experiments, HL-60 cells
expressing activated Notch1 at 6 hours from G1 synchronization showed a
mean 15% higher level of BrdU incorporation compared with controls
(P = .04), indicating a more rapid transit from G1
to S-phase (Fig 7B). Overall cycle length was unaffected as determined
by sequential analysis of synchronized cells. Next, we analyzed cell
cycle kinetics under conditions of differentiation. The effect of
Notch1 activation on cell cycle distribution was observed, and was
consistently more marked when cells were induced to differentiate,
either with ATRA or TPA. To exclude the possibility that the
differences observed in the cell cycle profiles were merely reflecting
the various proportions of terminally differentiated cells rather than
the intrinsic cell properties, we focused the cell cycle analysis on
the fraction of cells that remained undifferentiated at each time point
(Fig 8, top panel, CD11b-negative gated
population). In response to ATRA treatment, HL-60 MSCV-ICN/GFP
CD11b-negative cells maintained a lower proportion of cells in
G1 compared with controls (Fig 8). After 48 hours of ATRA
treatment, only 40% of HL-60 MSCV-ICN/GFP CD11b-negative cells were in
G1 compared with 60% and 57% of HL-60 MSCV-GFP and HL-60
MSCV-ICNAR/GFP CD11b-negative fractions, respectively. With time,
HL-60 MSCV-ICN/GFP CD11b-negative cells also started to accumulate in
G1, but in a smaller proportion compared with controls:
55% versus 67% and 71% at day 3, and 64% versus 72% and 76% at
day 4 (data not shown). By day 5, most of the cells in both populations
were differentiated and blocked in G1. Of note, the delay
in G1 arrest in HL-60 MSCV-ICN/GFP-transduced cells
preceded the differentiation delay (Fig 2).
View larger version (16K):
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[in a new window]
| Fig 6.
Effects of activated Notch on HL-60 cell proliferation.
Cells were plated at density of 0.1 × 106/mL in RPMI
supplemented with 10% FCS (A) and at density of 0.2 × 106/mL in RPMI supplemented with 10% FCS and 1 µmol/L
ATRA (B). Each day, cells were obtained, counted, and the percentage of
GFP-positive cells determined by flow cytometry. Numbers express the
mean of three experiments and represent the absolute number of HL-60
MSCV-GFP- and MSCV-ICN/GFP-positive cells/mL. Error bars represent
standard deviation.
|
|
View larger version (25K):
[in this window]
[in a new window]
| Fig 7.
Effects of activated Notch1 on HL-60 cell cycle kinetics.
(A) HL-60 MSCV-GFP, HL-60 MSCV-ICN/GFP, and MSCV-ICNAR/GFP cells
cultured in growth media were obtained and stained with the DNA dye
TOPRO-3. Cell cycle distribution was analyzed gating on GFP-positive
cells. (B) Cells were synchronized as described, and aliquots were
obtained every 6 hours after 30 minutes of incubation with BrdU.
Samples were stained with anti-BrdU MoAb and propidium iodide before
FACS analysis. Columns represent the average of five experiments and
express the percentage of cells that were in S-phase by their
positivity to BrdU staining. Bars represent standard error. The
difference between HL-60 MSCV-GFP and HL-60 MSCV-ICN/GFP populations is
statistically significant (P = .04).
|
|
View larger version (48K):
[in this window]
[in a new window]
| Fig 8.
Effects of activated Notch1 on cell cycle distribution
during HL-60 cell differentiation. Transduced HL-60 induced to
differentiate in presence of ATRA were analyzed every 24 hours for DNA
content and CD11b expression. Cells were harvested, labeled with
anti-CD11b-PE MoAb and than stained with the DNA dye TOPRO-3 as
described. Top panel: contour plots represent fluorescence intensity
for CD11b on the x-axis and for GFP on the y-axis; gates show the
populations, GFP-positive and CD11b-negative, selected for DNA
analysis. Central panel: cell cycle distribution analyzed on gated
populations at 48 and 72 hours after ATRA treatment. Bottom panel:
values represent the average of 10 independent observations at days 2, 3, and 4 of ATRA treatment and express the percentage of cells in
G0/G1, S-phase and G2/M
analyzed as described above.
|
|
These results were consistent in multiple experiments as summarized in
the table in Fig 8 (bottom). In 10 independent observations at days 2, 3, and 4, the average proportion of cells in G1 was 52% in
HL-60 MSCV-ICN/GFP cells compared with 64% in HL-60 MSCV-GFP cells
(P = .005); conversely, a higher proportion of cells in S-phase
and G2/M was noted. On the basis of these observations, we
hypothesize that Notch may inhibit differentiation by altering cell
cycle kinetics.
| |
DISCUSSION |
Notch family members have been shown to mediate cell specification in
many developmental and differentiation systems18,35-38 and
several studies have demonstrated a role for Notch in maintaining an
immature precursor phenotype.7,8,17,39 Observations showing
that Notch activation blocks granulocytic differentiation in the murine
multipotential cell line 32D11,40 suggested a possible role
for the Notch pathway in the regulation of hematopoiesis. The results
reported here show for the first time the role of Notch1 in the
regulation of human hematopoietic cell differentiation. Further, the
data link the inhibitory effect of Notch1 on cell differentiation with
altered cell cycle kinetics and suggest that the preservation of a
primitive phenotype may be mediated through enhanced cell cycle
progression through G1.
These studies show that Notch1 activation inhibits monocytic as well
granulocytic differentiation in human HL-60 cells in the presence of
prodifferentiative stimuli, such as retinoic acid and TPA. The effects
of Notch1 on myeloid differentiation were corroborated in primary human
hematopoietic CD34+ progenitors, where Notch1 activation,
by either transduction or stimulation by its physiological ligand
Jagged2, was associated with a delay in the acquisition of mature
surface markers. Notably, preservation of functional immature
characteristics were also shown by threefold to fivefold increases in
the formation of colonies of primitive phenotype (CFU-mix). Consistent
with these findings, a recent report noted a significant preservation
of immature precursors capable of generating high proliferative
potential-mix (HPP-mix) colonies when mouse progenitor
cells were exposed to the Notch ligand Jagged141; however,
in that report, no changes in the number of mature myeloid cells were
noted a distinct difference with our results that may reflect
species-specific or ligand-specific effects of Notch1 activation.
These observations suggest a potential physiological role for Notch
signaling in the maintenance of multipotency in vivo. A physiological
role for Notch signaling in regulation of human hematopoiesis is
further supported by the expression of endogenous Notch1 protein on
primary CD34+ progenitor cells and the cognate ligands
Jagged1 and Jagged2 on primary bone marrow stroma11 (N.C.,
unpublished observations, April 1997). It is interesting
that the effect of Notch activation on HL-60 and CD34+
progenitor cell differentiation is limited, resulting in a delay in
differentiation rather than an absolute block. It may be that countervailing regulators are present in human cells that mitigate the
effect of Notch1 activation and permit differentiation even in the face
of high Notch signaling. Our results suggest that activated Notch1 acts
in vitro as a brake on myeloid differentiation; however, the
possibility that it may also induce hematopoietic cells to take
alternative fates in vivo cannot be excluded.
The potential for complexity in vivo is made likely by the presence in
mammals of three other Notch genes,13-15,38 at least one of
which (Notch2) is also expressed in CD34+ cells, and of
three different genes for Notch ligands,10,11,38,42 two of
which (Jagged1 and Jagged2) are expressed by bone marrow stroma. As
more is learned it may be possible to exploit these regulatory pathways
to alter differentiation of hematopoietic cells in the context of ex
vivo stem cell expansion strategies, potentially using their effects to
retain a multipotential phenotype.
Mechanisms by which Notch1 mediates its effects have been delineated in
Drosophila and Caenorhabditis elegans,
where the transcription factors Suppressor of Hairless (Su[H]) and
lag-1 were identified, respectively, as important downstream
factors.43,44 In Drosophila, activated Notch alters the
expression of other transcription regulators (such as the Enhancer of
Split [E(spl)] complex) affecting neuronal fate.45 In
mammalian systems, Notch1 signaling has been shown to be mediated by
the transcription factor RBP-J /CBF,27,46 which is
homologous to Su(H) and lag. Activation of RBP-J by Notch1 has been
shown to lead to altered expression of the E(spl) homologue, Hairy
Enhancer of Split,44,47,48 indicating that essential
components of the Notch signaling pathway outlined in invertebrates are
conserved in vertebrates. Altering the pattern of transcription factor
interactions with lineage-specific genes may account for the ability of
Notch1 to impede differentiation.
We explored an alternative mechanism of action for Notch1 based on cell
cycle regulation, reasoning that because transition through the
G0 /G1checkpoint and differentiation are
highly coordinated events, Notch1 activation might affect
G1 progression. The evolutionary conservation of the Notch
family influencing differentiation of simple as well as highly complex
organisms raised the possibility of a mechanism interacting with other
highly conserved regulatory pathways, such as cell cycle control.
Analysis of cell cycle kinetics in the context of Notch1 activation
showed that the overall cell proliferative rates and cell cycle length
were unaffected, as determined after release from cycle blockade
induced by hydoxyurea or nocodazole. However, the relative proportion
of cells in specific phases of the cell cycle were consistently altered
in the presence of activated Notch1. The decline in the fraction of
cells in G1 is consistent with a shortening of
G1, a supposition confirmed by BrdU labeling. Precedent for
the effects of G1 shortening on differentiation can be
found in models assessing the impact of D-type cyclins on cell cycle
control. Of greatest relevance, overexpression of cyclin D2 and D3 in
murine 32D cells results in a delay of granulocytic differentiation49 similar to that produced by activated
Notch1.40 Importantly, cyclin D overexpression in that
study was accompanied by a decreased proportion of cells in
G1 similar in magnitude to what we observed in human HL-60
cells expressing activated Notch1. In our study, activated Notch1
enhanced G1 progression in the absence as well in the
presence of prodifferentiative stimuli, consistent with a direct effect
of Notch signaling on cell cycle kinetics.
These data indicate a potential link of Notch effects on
differentiation with its effects on cell cycle regulation. Accelerated G1 progression may modulate the ability of cells to respond
to prodifferentiative signals. Definition of the relationships of Notch
to checkpoint regulators will further clarify the mechanisms involved
and may provide insight into both the physiological and pathophysiological effects of Notch activation. Whereas Notch may
affect G1 to perturb differentiation kinetics, alterations in other regulators of G1 progression, such as cyclin D1,
Rb, and p16, are prominently associated with malignancy, raising the possibility that Notch1 oncogenicity may also be mediated, at least in
part, through effects on cell cycle progression. Examination of Notch
interaction with cell cycle machinery may prove fruitful in defining
the relationship of Notch to cell fate determination and malignant
transformation. The ability of Notch to dissociate differentiation from
proliferation may be a mechanism for expanding precursors cells without
inducing terminal differentiation.
| |
ACKNOWLEDGMENT |
We gratefully acknowledge the contributions of Cory Johnson and Dr
Robert Fallon in providing umbilical cord blood specimens, Drs Daniel
Haber and Nicholas Dyson in thoughtful review of the manuscript, Dr
Fred Preffer and David Dombrowski in guidance with flow cytometry
analyses, and Ellen Kornell in manuscript preparation.
| |
FOOTNOTES |
Submitted June 22, 1998; accepted September 29, 1998.
Supported in part by DK 50234 and HL 55718 of the National Institutes
of Health (D.T.S.), Defense Advanced Research Projects Agency and the
Saltonstall Charitable Trust (D.T.S.), by 5R29 CA 66849 (J.C.A.) and by
5RO1 CA62450 (J.S.).
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 David T. Scadden, MD, 149 13th St, 5212D,
Boston, MA 02129; e-mail: scadden.david{at}mgh.harvard.edu.
| |
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E. Fung, S.-M. T. Tang, J. P. Canner, K. Morishige, J. F. Arboleda-Velasquez, A. A. Cardoso, N. Carlesso, J. C. Aster, and M. Aikawa
Delta-Like 4 Induces Notch Signaling in Macrophages: Implications for Inflammation
Circulation,
June 12, 2007;
115(23):
2948 - 2956.
[Abstract]
[Full Text]
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M. Magnusson, A. C. M. Brun, N. Miyake, J. Larsson, M. Ehinger, J. M. Bjornsson, A. Wutz, M. Sigvardsson, and S. Karlsson
HOXA10 is a critical regulator for hematopoietic stem cells and erythroid/megakaryocyte development
Blood,
May 1, 2007;
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[Abstract]
[Full Text]
[PDF]
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R. F. de Pooter, T. M. Schmitt, J. L. de la Pompa, Y. Fujiwara, S. H. Orkin, and J. C. Zuniga-Pflucker
Notch Signaling Requires GATA-2 to Inhibit Myelopoiesis from Embryonic Stem Cells and Primary Hemopoietic Progenitors
J. Immunol.,
May 1, 2006;
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[Abstract]
[Full Text]
[PDF]
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C. Hu, A. Dievart, M. Lupien, E. Calvo, G. Tremblay, and P. Jolicoeur
Overexpression of Activated Murine Notch1 and Notch3 in Transgenic Mice Blocks Mammary Gland Development and Induces Mammary Tumors
Am. J. Pathol.,
March 1, 2006;
168(3):
973 - 990.
[Abstract]
[Full Text]
[PDF]
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F. Vianello, N. Papeta, T. Chen, P. Kraft, N. White, W. K. Hart, M. F. Kircher, E. Swart, S. Rhee, G. Palu, et al.
Murine B16 Melanomas Expressing High Levels of the Chemokine Stromal-Derived Factor-1/CXCL12 Induce Tumor-Specific T Cell Chemorepulsion and Escape from Immune Control.
J. Immunol.,
March 1, 2006;
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[Abstract]
[Full Text]
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C. Delaney, B. Varnum-Finney, K. Aoyama, C. Brashem-Stein, and I. D. Bernstein
Dose-dependent effects of the Notch ligand Delta1 on ex vivo differentiation and in vivo marrow repopulating ability of cord blood cells
Blood,
October 15, 2005;
106(8):
2693 - 2699.
[Abstract]
[Full Text]
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J. Zhu, Y. Zhang, G. J. Joe, R. Pompetti, and S. G. Emerson
NF-Ya activates multiple hematopoietic stem cell (HSC) regulatory genes and promotes HSC self-renewal
PNAS,
August 16, 2005;
102(33):
11728 - 11733.
[Abstract]
[Full Text]
[PDF]
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L. M. Sarmento, H. Huang, A. Limon, W. Gordon, J. Fernandes, M. J. Tavares, L. Miele, A. A. Cardoso, M. Classon, and N. Carlesso
Notch1 modulates timing of G1-S progression by inducing SKP2 transcription and p27Kip1 degradation
J. Exp. Med.,
July 5, 2005;
202(1):
157 - 168.
[Abstract]
[Full Text]
[PDF]
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S. J. C. Mancini, N. Mantei, A. Dumortier, U. Suter, H. R. MacDonald, and F. Radtke
Jagged1-dependent Notch signaling is dispensable for hematopoietic stem cell self-renewal and differentiation
Blood,
March 15, 2005;
105(6):
2340 - 2342.
[Abstract]
[Full Text]
[PDF]
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R. N. La Motte-Mohs, E. Herer, and J. C. Zuniga-Pflucker
Induction of T-cell development from human cord blood hematopoietic stem cells by Delta-like 1 in vitro
Blood,
February 15, 2005;
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1431 - 1439.
[Abstract]
[Full Text]
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E. Ishiko, I. Matsumura, S. Ezoe, K. Gale, J. Ishiko, Y. Satoh, H. Tanaka, H. Shibayama, M. Mizuki, T. Era, et al.
Notch Signals Inhibit the Development of Erythroid/Megakaryocytic Cells by Suppressing GATA-1 Activity through the Induction of HES1
J. Biol. Chem.,
February 11, 2005;
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[Abstract]
[Full Text]
[PDF]
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B. K. Hadland, S. S. Huppert, J. Kanungo, Y. Xue, R. Jiang, T. Gridley, R. A. Conlon, A. M. Cheng, R. Kopan, and G. D. Longmore
A requirement for Notch1 distinguishes 2 phases of definitive hematopoiesis during development
Blood,
November 15, 2004;
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[Abstract]
[Full Text]
[PDF]
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M. Noseda, L. Chang, G. McLean, J. E. Grim, B. E. Clurman, L. L. Smith, and A. Karsan
Notch Activation Induces Endothelial Cell Cycle Arrest and Participates in Contact Inhibition: Role of p21Cip1 Repression
Mol. Cell. Biol.,
October 15, 2004;
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[Abstract]
[Full Text]
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S. M. Vercauteren and H. J. Sutherland
Constitutively active Notch4 promotes early human hematopoietic progenitor cell maintenance while inhibiting differentiation and causes lymphoid abnormalities in vivo
Blood,
October 15, 2004;
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2315 - 2322.
[Abstract]
[Full Text]
[PDF]
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N. Tanimizu and A. Miyajima
Notch signaling controls hepatoblast differentiation by altering the expression of liver-enriched transcription factors
J. Cell Sci.,
July 1, 2004;
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3165 - 3174.
[Abstract]
[Full Text]
[PDF]
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Y. Satoh, I. Matsumura, H. Tanaka, S. Ezoe, H. Sugahara, M. Mizuki, H. Shibayama, E. Ishiko, J. Ishiko, K. Nakajima, et al.
Roles for c-Myc in Self-renewal of Hematopoietic Stem Cells
J. Biol. Chem.,
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[Abstract]
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Y. Nefedova, P. Cheng, M. Alsina, W. S. Dalton, and D. I. Gabrilovich
Involvement of Notch-1 signaling in bone marrow stroma-mediated de novo drug resistance of myeloma and other malignant lymphoid cell lines
Blood,
May 1, 2004;
103(9):
3503 - 3510.
[Abstract]
[Full Text]
[PDF]
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R. Qi, H. An, Y. Yu, M. Zhang, S. Liu, H. Xu, Z. Guo, T. Cheng, and X. Cao
Notch1 Signaling Inhibits Growth of Human Hepatocellular Carcinoma through Induction of Cell Cycle Arrest and Apoptosis
Cancer Res.,
December 1, 2003;
63(23):
8323 - 8329.
[Abstract]
[Full Text]
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G. F. Hoyne
Notch signaling in the immune system
J. Leukoc. Biol.,
December 1, 2003;
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[Abstract]
[Full Text]
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B.-C. Lee, T. Cheng, G. B. Adams, E. C. Attar, N. Miura, S. B. Lee, Y. Saito, I. Olszak, D. Dombkowski, D. P. Olson, et al.
P2Y-like receptor, GPR105 (P2Y14), identifies and mediates chemotaxis of bone-marrowhematopoietic stem cells
Genes & Dev.,
July 1, 2003;
17(13):
1592 - 1604.
[Abstract]
[Full Text]
[PDF]
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T. Schroeder, H. Kohlhof, N. Rieber, and U. Just
Notch Signaling Induces Multilineage Myeloid Differentiation and Up-Regulates PU.1 Expression
J. Immunol.,
June 1, 2003;
170(11):
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[Abstract]
[Full Text]
[PDF]
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B. Varnum-Finney, C. Brashem-Stein, and I. D. Bernstein
Combined effects of Notch signaling and cytokines induce a multiple log increase in precursors with lymphoid and myeloid reconstituting ability
Blood,
March 1, 2003;
101(5):
1784 - 1789.
[Abstract]
[Full Text]
[PDF]
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L. Espinosa, J. Ingles-Esteve, A. Robert-Moreno, and A. Bigas
Ikappa Balpha and p65 Regulate the Cytoplasmic Shuttling of Nuclear Corepressors: Cross-talk between Notch and NFkappa B Pathways
Mol. Biol. Cell,
February 1, 2003;
14(2):
491 - 502.
[Abstract]
[Full Text]
[PDF]
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A. P. Weng, Y. Nam, M. S. Wolfe, W. S. Pear, J. D. Griffin, S. C. Blacklow, and J. C. Aster
Growth Suppression of Pre-T Acute Lymphoblastic Leukemia Cells by Inhibition of Notch Signaling
Mol. Cell. Biol.,
January 15, 2003;
23(2):
655 - 664.
[Abstract]
[Full Text]
[PDF]
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S. Weijzen, M. P. Velders, A. G. Elmishad, P. E. Bacon, J. R. Panella, B. J. Nickoloff, L. Miele, and W. M. Kast
The Notch Ligand Jagged-1 Is Able to Induce Maturation of Monocyte-Derived Human Dendritic Cells
J. Immunol.,
October 15, 2002;
169(8):
4273 - 4278.
[Abstract]
[Full Text]
[PDF]
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M. De Smedt, K. Reynvoet, T. Kerre, T. Taghon, B. Verhasselt, B. Vandekerckhove, G. Leclercq, and J. Plum
Active Form of Notch Imposes T Cell Fate in Human Progenitor Cells
J. Immunol.,
September 15, 2002;
169(6):
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[Abstract]
[Full Text]
[PDF]
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M. Dorsch, G. Zheng, D. Yowe, P. Rao, Y. Wang, Q. Shen, C. Murphy, X. Xiong, Q. Shi, J.-C. Gutierrez-Ramos, et al.
Ectopic expression of Delta4 impairs hematopoietic development and leads to lymphoproliferative disease
Blood,
August 28, 2002;
100(6):
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[Abstract]
[Full Text]
[PDF]
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S. Jeffries, D. J. Robbins, and A. J. Capobianco
Characterization of a High-Molecular-Weight Notch Complex in the Nucleus of Notchic-Transformed RKE Cells and in a Human T-Cell Leukemia Cell Line
Mol. Cell. Biol.,
June 1, 2002;
22(11):
3927 - 3941.
[Abstract]
[Full Text]
[PDF]
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S. Stier, T. Cheng, D. Dombkowski, N. Carlesso, and D. T. Scadden
Notch1 activation increases hematopoietic stem cell self-renewal in vivo and favors lymphoid over myeloid lineage outcome
Blood,
April 1, 2002;
99(7):
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[Abstract]
[Full Text]
[PDF]
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L. Espinosa, S. Santos, J. Ingles-Esteve, P. Munoz-Canoves, and A. Bigas
p65-NF{kappa}B synergizes with Notch to activate transcription by triggering cytoplasmic translocation of the nuclear receptor corepressor N-CoR
J. Cell Sci.,
March 15, 2002;
115(6):
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[Abstract]
[Full Text]
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K. Kumano, S. Chiba, K. Shimizu, T. Yamagata, N. Hosoya, T. Saito, T. Takahashi, Y. Hamada, and H. Hirai
Notch1 inhibits differentiation of hematopoietic cells by sustaining GATA-2 expression
Blood,
December 1, 2001;
98(12):
3283 - 3289.
[Abstract]
[Full Text]
[PDF]
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P. Cheng, A. Zlobin, V. Volgina, S. Gottipati, B. Osborne, E. J. Simel, L. Miele, and D. I. Gabrilovich
Notch-1 Regulates NF-{kappa}B Activity in Hemopoietic Progenitor Cells
J. Immunol.,
October 15, 2001;
167(8):
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[Abstract]
[Full Text]
[PDF]
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A. C. Jaleco, H. Neves, E. Hooijberg, P. Gameiro, N. Clode, M. Haury, D. Henrique, and L. Parreira
Differential Effects of Notch Ligands Delta-1 and Jagged-1 in Human Lymphoid Differentiation
J. Exp. Med.,
October 1, 2001;
194(7):
991 - 1002.
[Abstract]
[Full Text]
[PDF]
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J. Wang, L. Shelly, L. Miele, R. Boykins, M. A. Norcross, and E. Guan
Human Notch-1 Inhibits NF-{{kappa}}B Activity in the Nucleus Through a Direct Interaction Involving a Novel Domain
J. Immunol.,
July 1, 2001;
167(1):
289 - 295.
[Abstract]
[Full Text]
[PDF]
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B. K. Hadland, N. R. Manley, D.-m. Su, G. D. Longmore, C. L. Moore, M. S. Wolfe, E. H. Schroeter, and R. Kopan
gamma -Secretase inhibitors repress thymocyte development
PNAS,
June 19, 2001;
98(13):
7487 - 7491.
[Abstract]
[Full Text]
[PDF]
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T. Imai, A. Tokunaga, T. Yoshida, M. Hashimoto, K. Mikoshiba, G. Weinmaster, M. Nakafuku, and H. Okano
The Neural RNA-Binding Protein Musashi1 Translationally Regulates Mammalian numb Gene Expression by Interacting with Its mRNA
Mol. Cell. Biol.,
June 15, 2001;
21(12):
3888 - 3900.
[Abstract]
[Full Text]
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D. R. Sherwood and D. R. McClay
LvNotch signaling plays a dual role in regulating the position of the ectoderm-endoderm boundary in the sea urchin embryo
Development,
June 15, 2001;
128(12):
2221 - 2232.
[Abstract]
[Full Text]
[PDF]
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V. Sriuranpong, M. W. Borges, R. K. Ravi, D. R. Arnold, B. D. Nelkin, S. B. Baylin, and D. W. Ball
Notch Signaling Induces Cell Cycle Arrest in Small Cell Lung Cancer Cells
Cancer Res.,
April 1, 2001;
61(7):
3200 - 3205.
[Abstract]
[Full Text]
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F. N. Karanu, B. Murdoch, T. Miyabayashi, M. Ohno, M. Koremoto, L. Gallacher, D. Wu, A. Itoh, S. Sakano, and M. Bhatia
Human homologues of Delta-1 and Delta-4 function as mitogenic regulators of primitive human hematopoietic cells
Blood,
April 1, 2001;
97(7):
1960 - 1967.
[Abstract]
[Full Text]
[PDF]
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J. Zhang, H. Chen, G. Weinmaster, and S. D. Hayward
Epstein-Barr Virus BamHI-A Rightward Transcript-Encoded RPMS Protein Interacts with the CBF1-Associated Corepressor CIR To Negatively Regulate the Activity of EBNA2 and NotchIC
J. Virol.,
March 15, 2001;
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[Abstract]
[Full Text]
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H. Höfelmayr, L. J. Strobl, G. Marschall, G. W. Bornkamm, and U. Zimber-Strobl
Activated Notch1 Can Transiently Substitute for EBNA2 in the Maintenance of Proliferation of LMP1-Expressing Immortalized B Cells
J. Virol.,
March 1, 2001;
75(5):
2033 - 2040.
[Abstract]
[Full Text]
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T. Morimura, S. Miyatani, D. Kitamura, and R. Goitsuka
Notch Signaling Suppresses IgH Gene Expression in Chicken B Cells: Implication in Spatially Restricted Expression of Serrate2/Notch1 in the Bursa of Fabricius
J. Immunol.,
March 1, 2001;
166(5):
3277 - 3283.
[Abstract]
[Full Text]
[PDF]
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F. N. Karanu, B. Murdoch, L. Gallacher, D. M. Wu, M. Koremoto, S. Sakano, and M. Bhatia
The Notch Ligand Jagged-1 Represents a Novel Growth Factor of Human Hematopoietic Stem Cells
J. Exp. Med.,
November 6, 2000;
192(9):
1365 - 1372.
[Abstract]
[Full Text]
[PDF]
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H. T. Tan-Pertel, L. Walker, D. Browning, A. Miyamoto, G. Weinmaster, and J. C. Gasson
Notch Signaling Enhances Survival and Alters Differentiation of 32D Myeloblasts
J. Immunol.,
October 15, 2000;
165(8):
4428 - 4436.
[Abstract]
[Full Text]
[PDF]
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S. Tsai, J. Fero, and S. Bartelmez
Mouse Jagged2 is differentially expressed in hematopoietic progenitors and endothelial cells and promotes the survival and proliferation of hematopoietic progenitors by direct cell-to-cell contact
Blood,
August 1, 2000;
96(3):
950 - 957.
[Abstract]
[Full Text]
[PDF]
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S. Jeffries and A. J. Capobianco
Neoplastic Transformation by Notch Requires Nuclear Localization
Mol. Cell. Biol.,
June 1, 2000;
20(11):
3928 - 3941.
[Abstract]
[Full Text]
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F. Radtke, I. Ferrero, A. Wilson, R. Lees, M. Aguet, and H. R. MacDonald
Notch1 Deficiency Dissociates the Intrathymic Development of Dendritic Cells and T Cells
J. Exp. Med.,
April 3, 2000;
191(7):
1085 - 1094.
[Abstract]
[Full Text]
[PDF]
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H. Harada, P. Kettunen, H.-S. Jung, T. Mustonen, Y. A. Wang, and I. Thesleff
Localization of Putative Stem Cells in Dental Epithelium and Their Association with Notch and Fgf Signaling
J. Cell Biol.,
October 4, 1999;
147(1):
105 - 120.
[Abstract]
[Full Text]
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J. M. Verdi, A. Bashirullah, D. E. Goldhawk, C. J. Kubu, M. Jamali, S. O. Meakin, and H. D. Lipshitz
Distinct human NUMB isoforms regulate differentiation vs. proliferation in the neuronal lineage
PNAS,
August 31, 1999;
96(18):
10472 - 10476.
[Abstract]
[Full Text]
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J. Ingles-Esteve, L. Espinosa, L. A. Milner, C. Caelles, and A. Bigas
Phosphorylation of Ser2078 Modulates the Notch2 Function in 32D Cell Differentiation
J. Biol. Chem.,
November 21, 2001;
276(48):
44873 - 44880.
[Abstract]
[Full Text]
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Top
Abstract
Introduction