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*
Division of Hematology-Oncology, Department of Medicine, University of California at Los Angeles School of Medicine and Jonsson Comprehensive Cancer Center, Los Angeles, CA 90095; and
Department of Biological Chemistry, University of California at Los Angeles Molecular Biology Institute, Los Angeles, CA 90095
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The Notch family of receptors (Notch1–4) consists of large transmembrane proteins. The extracellular domain of Notch is composed of 36 epidermal growth factor (EGF)3-like repeats and three Lin-12/Notch repeats; the intracellular domain consists of six ankyrin-like repeats with two flanking nuclear localization sequences, a core binding factor 1 (CBF1) interaction region (RAM domain), and a proline-glutamate-serine-threonine-rich (PEST) motif. In Drosophila and mammalian systems, constructs expressing an isolated intracellular domain of Notch result in a constitutively active, gain-of-function phenotype (13, 14, 15). Genetic and biochemical studies suggest that signaling by this activated form of Notch involves the transcriptional regulator, CBF1 (the mammalian homolog of Drosophila suppressor of hairless) (16, 17, 18). Further downstream, CBF1 up-regulates the expression of a basic helix-loop-helix (bHLH) transcription factor, HES1 (the mammalian homolog of the Drosophila hairy enhancer of split) (19, 20) to repress lineage-specific genes such as achaete-scute in neurons (21). Recently, this simple concept of Notch signal transduction has been challenged in an in vitro myoblast system in which Notch signaling inhibits muscle cell differentiation through both CBF1-dependent and -independent pathways (22). Nofziger et al. (23) suggested that the CBF1-independent pathway provides a general block in cell differentiation that is reinforced by cell type-specific, CBF1-dependent signals.
The hematopoietic system provides an excellent model for studying the
function of Notch in cell fate specification, because hematopoiesis
requires continuous progenitor cell proliferation, lineage commitment,
differentiation, and maturation. Considerable evidence suggests that
Notch plays critical roles in several steps of hematopoiesis. The best
characterized examples are in lymphoid development. The
TAN-1/Notch1 translocation (7, 9) (q34;q34.3)
results in an active form of Notch1 mobilized to the TCR
locus, and is responsible for the formation of a subset of acute
T-lymphoblastic leukemias in humans (24). Transplantation
of retrovirally transduced bone marrow cells expressing activated forms
of Notch induces T cell leukemia and influences B- vs T-lineage
determination in mice (25, 26). Studies of transgenic mice
over-expressing an activated form of Notch1 in developing T cells
suggest that Notch1 participates in the CD4 vs CD8 and the 
ß vs
decisions (27, 28). A conditional knockout of
Notch1 in newborn mice results in a severe deficiency in
thymocyte development (29). Furthermore, activation of
the Notch signaling pathway confers resistance to TCR-mediated
(9) and glucocorticoid-induced (8) apoptosis
in CD4+/CD8+ thymocytes and
up-regulates cellular markers correlating with maturation
(8).
In myeloid development, Notch signaling has been reported to inhibit or delay differentiation of the 32D myeloblast cell lines (7, 30, 31) and HL-60 (32), and we and others have shown it to increase the formation of primitive precursor cell populations from humans and mice (33, 34). The conditional knockout of Notch1 did not yield obvious anomalies in myeloid development (29), perhaps because of the functional redundancy between Notch1 and Notch2, or because Notch2 may be the predominant form in myeloid progenitor cells, as is suggested by its prevalence in 32D myeloid progenitor cells (35). Furthermore, transplantation of bone marrow cells overexpressing active forms of Notch1 did not yield an apparent granulocytic phenotype, as assessed by FACS analysis of Mac1+/Gr1+ cells (26), probably because such cells represent a pool of myeloid cells without distinguishing stages of differentiation during myelopoiesis. Clearly, a more detailed analysis of Notch function in the progression of myeloid differentiation is needed.
To gain further insight into the function of Notch signaling during hematopoiesis in general, and specifically in myelopoiesis, we began to study the function of Notch in 32D (clone 3) myeloblast differentiation. In addition, we examined the Notch/CBF1/HES1 signaling pathway in these cells. We chose to use the well-characterized cell line, 32D (36), because these cells closely resemble bipotent granulocyte-macrophage progenitors in bone marrow. Originally derived from normal murine bone marrow, 32D cells are diploid and not leukemic in syngeneic murine recipients. Similar to CFU-GM, these cells have both granulocytic and monocytic potential (37). The cytokines, GM-CSF and IL-3, support their proliferation, whereas granulocyte CSF (G-CSF) induces granulopoiesis and maturation, followed by cell cycle arrest. We generated 32D cell stable populations expressing truncated forms of Notch1 or Notch2 that can or cannot signal through CBF1 to examine the effect of Notch activation on 32D cell survival and differentiation. Our results suggest that Notch1 and Notch2 enhance 32D cell survival, promote initial granulopoiesis, and inhibit postmitotic differentiation through a CBF1-dependent pathway.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Total cellular RNA was isolated from 32D cells (clone 3), a factor-dependent murine myeloid cell line (generously provided by Dr. Joel Greenberger, University of Pittsburgh, Pittsburgh, PA) using RNAzol reagents (Tel-Test, Friendswood, TX) according to the manufacturer’s protocol. PCR primers were designed to be specific to nonhomologous regions of Notch1 and Notch2 extracellular domains. The primer sequences for Notch1 were 5'-gccacagattgaggaggcct and 3'-accattggtgccaggaagca; and the primers for Notch2 were 5'-accagcacccctcctgctac and 3'-tcctgttcctgctcatcagg. RT-PCR was performed using Titan RT-PCR reagents (Roche, Indianapolis, IN). To eliminate genomic DNA contamination, total RNA was treated with DNase I (Life Technologies, Grand Island, NY). Reactions lacking reverse transcriptase were used as negative controls. Reverse transcription was conducted at 65°C for 30 min. PCRs were conducted in a thermal cycler (PTC100; MJ Research, San Francisco, CA) for 36 cycles at 94°C for 1 min, 55°C for 1 min, and 72°C for 1 min.
Flow cytometry for detecting Notch1 and Notch2 expression
32D cells were washed in PBS, fixed in 0.5% paraformaldehyde for 20 min at room temperature, then permeabilized in 70% EtOH for 10 min at 4°C. After two PBS washes, cells were resuspended in PBS, pH 7.2, containing 5 mM EDTA and 0.5% BSA (Sigma, St. Louis, MO) and allowed to rehydrate for 30 min at 37°C. Cells (106) were then blocked with 1 µg anti-CD32 mAb (PharMingen, San Diego, CA) for 10 min at room temperature, then incubated for 30 min with 0.4 µg of affinity-purified rabbit antiserum to either the intracellular domain of rat Notch1 (93-4) or Notch2 (93-7) or with an equivalent amount of normal rabbit IgG (Sigma). After PBS washes, cells were stained for 15 min with a 1:100 dilution of human-adsorbed FITC-conjugated goat anti-rabbit IgG (Caltag, Burlingame, CA). Cells were washed again and analyzed immediately on a FACScan flow cytometer, using Cell Quest software (Becton Dickinson, Mountain View, CA).
cDNA expression constructs
The truncated forms of Notch used in our work have been
described previously (22, 23). Briefly, deletion mutants
of Notch1 and Notch2 cDNA (see Fig. 2
A) were cloned in a
retroviral vector, SRMSVtKNeo (generously provided by Dr.
Owen Witte, University of California at Los Angeles, Los Angeles, CA),
to encode the following amino acids: ZEDN1, 1–23 plus 1712–2531;
OEDN1, 1–23 plus 24–60 and 1712–2531; CDN1, 1848–2531; and CDN2,
1786–2472.
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Retroviral transduction and transient transfections
Infectious retroviral supernatants were produced by
simultaneously transfecting (CalPhos Maximizer Transfection Kit;
Clontech Laboratories) 293T cells with two plasmids, one encoding a
deletion form of Notch in the SR retroviral vector, and
the other encoding the ecotropic helper virus, 
E. 32D cells
(105/ml) were then incubated in 1 ml of
retroviral supernatant, 2 ml of growth medium (see Cell
culture), and 2 µg/ml polybrene (hexadimethrine bromide; Sigma)
at 37°C for 24 h. Cell populations overexpressing HES1 were
generated by electroporating 30 µg of the pBos-EF12-HES1
plasmid (provided by Dr. Ryoichiro Kageyama of Kyoto University, Kyoto,
Japan) with 3 µg of the pCDNA3 plasmid (Invitrogen, San
Diego, CA) at 250 V, 960 µF, using a Gene Pulser (Bio-Rad, Richmond,
CA). This strategy was used to maximize the level of HES1 expression.
Twenty-four hours after transduction or electroporation, cells
populations stably expressing the desired proteins were selected and
maintained in 1.2 mg/ml of geneticin (G418; Life Technologies).
For subcellular localization studies, 3 µg of pEGFP-ZEDN1, pEGFP-OEDN1, pEGFP-CDN1, pEGFP-CDN2, or pEGFP vector control was transfected into NIH 3T3 cells (no. 1658-CRL; American Type Culture Collection, Manassas, VA) using CalPhos Maximizer Transfection reagents (Clontech). Photographs were taken at x400 magnification, using an Olympus (New Hyde Park, NY) fluorescence microscope (model IX50) at 24 h posttransfection.
Cell culture
32D cells were maintained in IMDM supplemented with 10% FBS (Omega Scientific, Tarzana, CA), 0.4 mM L-glutamine, 100 U/ml penicillin, 0.1 mg/ml streptomycin, and 200 pM murine GM-CSF (a gift from Amgen, Thousand Oaks, CA). When growing in GM-CSF culture, cells were split 1:100 every 3 days. To differentiate cells, 32D stable cell populations (1.6 x 104 cells/ml) were washed three times in medium to remove residual GM-CSF, then cultured in medium supplemented with 50 ng/ml of human G-CSF (provided by Amgen). After a time course of 4 days, the number of viable cells, the percentage of dead cells, and the percentage of differentiation were evaluated. The numbers of viable cells were evaluated by trypan blue exclusion using a hemacytometer. Equal culture volumes were centrifuged onto individual slides using a Cytospin 3 (Shandon, Pittsburgh, PA). Cytospin slides were stained with a Diff Quick Set (Dade Diagnostics, Aguada, PR) and evaluated in a blinded fashion for granulocytic differentiation. The criteria for differentiation included nuclear segmentation, cytoplasm-nucleus ratio, and granularity. The differential counts were performed on 300 cells per slide for blasts, promyelocytes, myelocytes, metamyelocytes, mature granulocytes, and dead cells.
Western blot analysis
Cells (3 x 105 per lane) were lysed
and electrophoresed on a 7.5 or 10% SDS-PAGE mini-gel. Separated
proteins were transferred to a Hybond nitrocellulose membrane
(Amersham, Arlington Heights, IL) and blocked with 5% nonfat dry milk
in TBS containing 1% Tween 20 for 2 h at room temperature.
Membranes were then incubated with one of the three polyclonal rabbit
antisera for 2 h at room temperature. The 93–4 antiserum
(22) against the intracellular domain of rat Notch1 was
used at a dilution of 1:4000, the 93–7 antiserum (38)
against the intracellular domain of rat Notch2 was affinity-purified
and used at 
32 µg/ml, and the IgG fraction of the HES1 antiserum
(a gift from Dr. Tetsuo Sudo, Toray Industries, Osaka, Japan) was used
at 3.6 µg/ml. Immunoreactivity was detected using a biotinylated
donkey anti-rabbit IgG Ab (Amersham), HRP-conjugated streptavidin
(Amersham), and enhanced chemiluminescence Western blot reagents
(Amersham).
TUNEL assay
Twenty-four hours after G-CSF induction (see Cell culture), 32D cell populations were washed twice with PBS containing 1% BSA and fixed for 1 h with 4% paraformaldehyde in PBS, pH 7.4. After washing with PBS twice, cells were permeabilized for 2 min with 0.1% Triton X-100 in 0.1% sodium citrate. Cells were then washed in PBS and resuspended in TUNEL reaction mixture for 1 h at 37°C according to the manufacturer’s protocol. After two washes in PBS, cells were analyzed by flow cytometry.
CBF1 trans-activation assay
32D cells (107) were electroporated with
30 µg of the indicated Notch deletion construct (see Fig. 2
A), along with 10 µg of
4xCBF1Luc (16) and 1 µg of the
ß-galactosidase plasmid pCH110 (Pharmacia, Piscataway,
NJ). Twenty-four hours postelectroporation, cells were evaluated for
both luciferase and ß-galactosidase activity according to the
manufacturer’s instructions (Promega, Madison, WI). ß-galactosidase
expression was used to control for differences in transfection
efficiencies. CBF1 activity is reported as fold increase in normalized
luciferase values for each Notch construct relative to vector control
values.
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Results |
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To confirm a previous report of endogenous Notch expression in 32D
cells (7), we used RT-PCR and flow cytometric analysis to
detect both Notch1 and Notch2. Because Notch1 and Notch2 are highly
homologous, PCR primers were designed against unique regions of either
Notch1 or Notch2, and primer specificity was verified using
Notch1 or Notch2 plasmid cDNAs. Fig. 1A shows that the
Notch1 primers only amplified Notch1 cDNA
(lane 2) and not Notch2 cDNA
(lane 3). Conversely, the Notch2 primers
were specific to Notch2 cDNA (lane 5) and
did not amplify Notch1 cDNA (lane 4). In
RT-PCR analysis, endogenous RNA species for Notch1 and
Notch2 were detected in lanes 7 and 9,
respectively. No bands were detected in the absence of RT enzyme
(lanes 6 and 8), indicating that there was
no genomic DNA contamination. Fig. 1B shows flow cytometric
analysis of 32D cells stained with antiserum against intracellular
Notch1 or Notch2, indicating the presence of endogenous Notch1 and
Notch2 protein isoforms.
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To identify structural motifs important for Notch function in 32D
cells, we used deletion constructs of Notch1
(OEDN1, ZEDN1, CDN1) and
Notch2 (CDN2) (see Fig. 2A).
OEDN1 and ZEDN1 encode the signal peptide, the
transmembrane domain, and the entire intracellular domain of Notch1. In
addition, OEDN1 encodes one and a half extracellular EGF
repeats. Both ZEDN1 and OEDN1 contain the entire 117 amino acids of the
RAM domain, which is located between the transmembrane domain and the
ankyrin repeats, and is required for CBF1 binding. CDN1, which
initiates 17 amino acids upstream of the ankyrin repeats, lacks most of
the RAM sequences necessary for productive CBF1 interactions. CDN2,
which initiates 36 amino acids upstream of the ankyrin repeats of
Notch2, encodes the RAM 1b domain (a weak CBF1 interaction domain), but
not the stronger affinity 1a domain (16, 38).
Protein expression was confirmed by Western analysis of 32D-SR,
32D-ZEDN1, 32D-OEDN1, 32D-CDN1, and 32D-CDN2 stable populations, using
antisera raised against the cytoplasmic sequences of Notch1 or Notch2.
The CDN1 and CDN2 proteins (lanes 2 and
6), with approximate molecular masses of 75 kDa, as well as
OEDN1 (lane 3) and ZEDN1 (lane 4)
proteins, with approximate molecular masses of 97 and 93 kDa,
respectively, are indicated by arrows in Fig. 2B. The level
of ZEDN1 expression was consistently lower than that of OEDN1, CDN1,
and CDN2.
Subcellular localization of ZEDN1, OEDN1, CDN1, and CDN2
The only difference between constructs ZEDN1 and OEDN1 is the presence of a sequence encoding one and a half extracellular EGF repeats in OEDN1. It is known that full-length Notch must be proteolytically processed at two extracellular sites and one transmembrane/intracellular site before full activation and nuclear localization (39, 40, 41, 42). We hypothesized that the one and a half extra EGF-like repeats in the extracellular domain of OEDN1 might affect OEDN1 processing and subcellular localization. For this reason, we examined the subcellular localization of all the Notch constructs to be characterized in the biological assay.
To elucidate the subcellular localization of ZEDN1, OEDN1, CDN1, and
CDN2, we created the EGFP-fusion constructs, and transfected
NIH 3T3 cells either with the EGFP vector or with sequences
encoding fusion constructs. When the pEGFP vector was
transfected, the green fluorescence was distributed throughout the
entire cell (Fig. 3b). The EGFP-ZEDN1 protein was
predominantly localized in the cell nucleus (Fig. 3d), with
aggregates possibly located in the endoplasmic reticulum and the Golgi
complex. In contrast, Fig. 3f shows that the EGFP-OEDN1
protein did not localize to the cell nucleus, suggesting that the
processing of OEDN1 was abnormal, preventing its nuclear import. Fig. 3, h and j, shows that CDN1-EGFP and CDN2-EGFP
were found in both the cytoplasm and the nucleus.
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To investigate the effects of overexpression by various forms of
Notch in granulopoiesis, 32D-SR, 32D-ZEDN1, 32D-OEDN1, 32D-CDN1, and
32D-CDN2 cell populations were cultured in growth or differentiation
conditions, and the numbers of viable and dead cells, as well as the
apoptosis profile, were evaluated. As shown in Fig. 4A, when the cells were
cultured in growth medium supplemented with the cytokine GM-CSF, all
five cell populations proliferated at a similar rate, indicating that
overexpression of the various Notch1 and Notch2 proteins did not alter
their growth properties in GM-CSF.
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B
demonstrates that the cells overexpressing ZEDN1 and CDN2 maintained
higher numbers of viable cells compared with the 32D-SR control on
days 1–4 (mean ± SEM of triplicates). ZEDN1 exhibited the
strongest phenotype, although it was expressed at a lower level than
the other Notch constructs, suggesting that the observed phenotype did
not result directly from variations in protein expression levels. Only
a modest increase in cell numbers was observed in the 32D-OEDN1
population. 32D-CDN1 did not show a significant difference in total
viable cell numbers compared with the SR control. Fig. 4C
shows the cytospin preparations of G-CSF cultures on day 2. The
cytopreps were conducted with an equal volume of culture from each
group rather than with an equivalent cell number. Therefore, the cell
number differences seen on the slides reflect the cell densities of the
cultures. Again we observed that the numbers of viable cells in G-CSF
culture were highest among the ZEDN1 (Fig. 4C, c)
and CDN2 (Fig. 4C, f) populations.
To determine whether the observed increases of viable cells in
32D-ZEDN1 and 32D-CDN2 were due to enhanced survival of these cells in
the presence of G-CSF, we calculated the number of dead cells as a
percentage of total cells in all five groups. Fig. 4D shows
that 32D-ZEDN1 and 32D-CDN2 stable populations had a significantly
lower percentage of cell death compared with the 32D-SR control on
days 1–3. The 32D-OEDN1 population had a slight decrease in cell death
on days 1 and 2, whereas the 32D-CDN1 population did not differ from
the vector control.
Fig. 4E shows a TUNEL analysis assessing apoptosis in the
stable cell populations at day 1 in G-CSF culture. The 32D-ZEDN1
population had less than half the levels of apoptosis (24%) compared
with the vector control (53%). 32D-CDN2 cells also displayed lower
levels of apoptosis (37%), whereas 32D-OEDN1 and 32D-CDN1 cells
exhibited minimal alterations in apoptosis. These trends were seen in
two separate experiments and were confirmed by Annexin V staining (data
not shown).
From these results, we concluded that active forms of Notch containing the entire RAM domain (ZEDN1) or a partial RAM domain (CDN2) enhanced cell survival and inhibited apoptosis of 32D myeloblasts in G-CSF culture.
Dual effects of Notch signaling on 32D differentiation
The course of granulopoiesis in 32D cells progresses sequentially
from undifferentiated blasts to promyelocytes, myelocytes, then
metamyelocytes, and finally mature granulocytes, followed by cell cycle
arrest. To examine the role of enforced Notch expression in the
progression of 32D cell differentiation, we analyzed the percentage of
differentiation for each stage of granulopoiesis. The cell populations
(shown in Fig. 4C) were grouped into three categories:
undifferentiated blasts, intermediate cells (including promyelocytes,
myelocytes, and metamyelocytes), and mature granulocytes. The
percentage of undifferentiated blasts remaining during G-CSF induction
correlates inversely with the ability of the population to commit to
granulopoiesis and differentiation. The percentage of mature
granulocytes reflects the levels of postmitotic differentiation, the
final step in differentiation of 32D cells.
Fig. 5A shows the percentage
of undifferentiated blasts in G-CSF cultures. 32D-ZEDN1 and 32D-CDN2
cells displayed a lower percentage of blasts than the SR control,
whereas 32D-OEDN1 and 32D-CDN1 cells were not different from the
control. We then examined the percentage of mature granulocytes in the
culture. Fig. 5C demonstrates that 32D-ZEDN1 cells had a
lower percentage of mature granulocytes than the SR control.
32D-CDN2 cells had a similar though less profound phenotype, as
compared with 32D-ZEDN1 cells. When compared with controls, more cells
in the 32D-ZEDN1 and 32D-CDN2 populations progressed to promyelocytes,
and a lower percentage of cells underwent terminal differentiation to
mature granulocytes. Consequently, most of the cells in these
populations accumulated in the early stages of differentiation as an
expanded pool of promyelocytes, myelocytes, and metamyelocytes (Fig. 5B). 32D-OEDN1 and 32D-CDN1 cells had percentages of
intermediate cells similar to the vector control.
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Notch deletion forms that contain the RAM domain or partial RAM domain can trans-activate endogenous CBF1
It has been reported that the RAM domain and the ankyrin repeats are required for CBF1 binding (43). Therefore, Notch constructs that lack almost the entire RAM domain, such as CDN1, should not activate CBF1. The interaction of endogenous CBF1 with Notch RAM domain sequences has not been tested in 32D cells. In addition, whether a construct such as OEDN1, which contains the entire RAM domain but is primarily localized outside the nucleus, can activate CBF1, has not been tested. To investigate the relationship of the Notch-induced survival and differentiation phenotype with CBF1 activity in 32D cells, we analyzed CBF1 trans-activation by ZEDN1, OEDN1, CDN1, and CDN2 in a CBF1-luciferase reporter system. When overexpressed in host cells, active forms of Notch that interact with endogenous CBF1 transform CBF1 from a transcriptional repressor to a transcriptional activator and drive the expression of the luciferase reporter gene.
Fig. 6 demonstrates that ZEDN1 strongly
trans-activates CBF1 (20-fold) in 32D cells, and CDN2
trans-activates CBF1 to a modest extent (5-fold). Neither
OEDN1 nor CDN1 show significant CBF1 trans-activation. The
CBF1 trans-activation results described here correlate well
with the cell survival and differentiation results described in the
previous section, suggesting that Notch acts via a CBF1-dependent
pathway in these cells.
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Genetic and biochemical studies suggest that Notch activation of
CBF1 leads to up-regulation of a bHLH transcription factor, HES1
(19, 20). 32D cells express low levels of HES1, as seen by
Northern analysis (data not shown). We postulated that if, in 32D
cells, Notch signals via trans-activation of CBF1, which in
turn up-regulates HES1, HES1-overexpressing cells should yield a
phenotype similar to Notch-overexpressing cells. Therefore, we created
32D-HES1 stable cell populations and examined the differentiation
properties of these cells. Fig. 7A shows that HES1 expression
in 32D-HES1 cells was readily detectable, and Fig. 7B
indicates the number of viable cells in 32D-HES1 and 32D-pBos cells on
day 2 of G-CSF culture. The 32D-HES1 cells maintained higher numbers of
viable cells in the presence of G-CSF than the vector control, similar
to what was observed in 32D-ZEDN1 and 32D-CDN2 cells, suggesting that
HES1 and Notch may function in the same pathway, and that HES1 may be a
downstream effector of Notch in 32D cells. The 32D-HES1 phenotype was
more subtle than that of 32D-ZEDN1 and 32D-CDN2, probably because HES1
is not the only effector of Notch. Therefore, we are probably only
seeing a partial response of Notch activation when we overexpress HES1.
For a subtle phenotype, such as that observed in 32D-HES1 cells, it was
difficult to observe significant differences in differential counts
(data not shown).
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Discussion |
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The dramatically higher numbers of viable cells and fewer dead cells in
the 32D-ZEDN1 and 32D-CDN2 populations during G-CSF culture indicated
that Notch signaling enhanced cell survival. The biphasic growth
properties of 32D cells displayed in Fig. 4B reflected a
window of adjustment for the 32D cells at days 1 and 2, before the
cells began to proliferate transiently at days 3 and 4. The dramatic
differences in viable cell numbers were largely due to differences in
viable cell numbers at days 1 and 2, when the cells were in the
nonproliferative period. 32D-ZEDN1 shortened the refractory period from
2 days to 1 day and, therefore, intensified the differences in viable
cell numbers. Further supporting the conclusion that Notch signaling
enhanced 32D cell survival was that TUNEL analysis demonstrated that
Notch signaling inhibited apoptosis during the initial 24 h of
G-CSF treatment.
We observed a dual effect of Notch signaling on 32D differentiation
(Fig. 5). In the presence of G-CSF, Notch signaling promoted
progression from myeloblasts to promyelocytes, but inhibited
postmitotic differentiation from metamyelocytes to mature granulocytes.
Our data suggest that Notch signaling might be potentiating the G-CSF
cytokine signal to initiate granulopoiesis. This action of Notch is
consistent with other systems in which Notch signaling modulates the
response of precursor cells to environmental signals
(1, 2, 3). Because 32D cells resemble bipotent CFU-GM, which
becomes committed to granulopoiesis when stimulated with G-CSF, it is
possible that the initial lineage specification requires Notch to
progress from myeloid blasts to promyelocytes.
Notch signaling inhibited postmitotic differentiation from
metamyelocytes to mature granulocytes, as shown in Fig. 5C.
This block to differentiation may contribute to fewer dead cells and
more viable cells. The block to differentiation is consistent with
Notch-mediated inhibition of granulocytic differentiation observed by
Milner et al. (7) and with the oncogenic property of TAN-1
(translocation-associated Notch homolog) in T lymphoblastic leukemia
(24, 25). However, we did not observe the expansion of
undifferentiated 32D cells (namely, myeloblasts) observed by Milner et
al. (7). In addition, these authors suggested that the
active form of Notch1 inhibited 32D differentiation by G-CSF but not
GM-CSF, whereas an active form of Notch2 inhibited differentiation
induced by GM-CSF but not G-CSF (31).
In contrast, we did not observe an expansion of undifferentiated cells,
but rather, the opposite; we observed that both Notch1 and Notch2
overexpression caused more cells to progress from myeloblasts to
promyelocytes, resulting in a decreased percentage of undifferentiated
cells (Fig. 5A). These conflicting observations could be due
to differences in the cell populations under study. The 32D cells used
in our investigation were regularly maintained in growth medium
supplemented with GM-CSF, whereas the cells used by Milner et al. were
regularly maintained in growth medium supplemented with
WEHI-conditioned medium as a source of IL-3. In addition, there is
inherent variability in the populations of 32D cells used in the two
studies. Furthermore, Milner et al. used clones for their study,
whereas we used freshly transduced cell populations. We also selected
clones overexpressing an active form of Notch, and conducted G-CSF
differentiation and cell survival experiments with similar results
(data not shown).
With regard to structure-function analysis of Notch signaling, the current model suggests that there are three functional domains for the Notch intracellular region: the RAM domain and the ankyrin repeats necessary for CBF-1 binding and corepressor displacement, and the C-terminal trans-activation domain (43). Our CBF1 trans-activation data are consistent with this model. The CBF1 trans-activation results also correlate well with G-CSF survival and differentiation. 32D-ZEDN1, which contains all three functional domains, exhibited the strongest phenotype of enhanced survival and differentiation, and could strongly trans-activate CBF1. CDN2, which contains a partial RAM domain and showed an intermediate phenotype, could weakly trans-activate CBF1. OEDN1, due to its impaired nuclear localization, did not strongly activate CBF1, nor did it strongly affect 32D survival and differentiation. CDN1 does not contain the RAM domain and did not activate CBF1, nor did it yield phenotypes in G-CSF culture. The correlation of CBF1 trans-activation with enhanced survival and altered differentiation in 32D cells suggests that Notch functions through a CBF1-dependent pathway in these cells.
It is known that proteolytic release of the Notch intracellular domain is required for its function. One of the cleavages that results in the release of the Notch1 intracellular domain occurs within the transmembrane domain or immediately adjacent to it. There are recent reports demonstrating that presenilin is physically associated with Notch1 and may be involved in proteolytic cleavage (44, 45, 46, 47). Our observation that the truncated Notch1 protein, OEDN1, was not localized to the nucleus suggests that OEDN1 was not processed correctly. The extra EGF repeats may affect the conformation of OEDN1, such that proteolytic enzymes such as presenilin cannot access the cleavage site.
Tomita et al. did not conclude that the HES1 knockout mice showed a difference in Mac1+/Gr-1+ cells in fetal liver (11.1 vs 8.7%, and 12.5 vs 8.8%, respectively) (48). As is the case for Notch1 knockout mice, overlapping and redundant signaling pathways may preclude the generation of a robust myeloid phenotype in HES1-/- mice. In our experiments, overexpression of HES1 in 32D cells caused a phenotype similar to that caused by activated forms of Notch is consistent with the hypothesis that HES1 is a downstream target of Notch in these cells. Because HES1 is a bHLH protein, which usually interacts with other factors to influence transcription of downstream lineage-specific genes, we speculate that in 32D cells, HES1 may affect the activity of other transcriptional regulators to control the expression of myeloid-specific genes. Now that components of the Notch signaling pathway have begun to be characterized in hematopoietic cells, the next challenge is to identify downstream myeloid-specific genes regulated by the Notch/CBF1/HES1-dependent pathway.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Judith C. Gasson, Jonsson Comprehensive Cancer Center, 8-684 Factor, Box 951781, University of California, Los Angeles, CA 90095-1781.
3 Abbreviations used in this paper: EGF, epidermal growth factor; CBF1, core binding factor 1; G-CSF, granulocyte CSF; bHLH, basic helix-loop-helix protein; EGFP, enhanced green fluorescent protein.
Received for publication May 3, 2000. Accepted for publication July 24, 2000.
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References |
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ß versus 
T cell lineage decision. Cell 88:833.[Medline]
-secretase-like protease mediates release of Notch intracellular domain. Nature 398:518.[Medline]
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