 |
INTRODUCTION |
Receptor-coupled heterotrimeric G proteins are known to mediate
signals that modulate growth factor-dependent cellular
proliferation (1). G protein-linked second messengers, such as
Ca2+ or cAMP, as well as G protein 
-subunits released
upon G protein activation, may be involved in this modulation.
Ultimately, these signaling elements are thought to act on
mitogen-activated protein kinase or on c-Jun amino-terminal kinase
modules, which serve as central regulators for cellular growth and
differentiation (2, 3). In some endocrine tumors, mutations in
clearly defined codons for conserved amino acids result in
GTPase-deficient (i.e. GTP-bound, constitutively active) G
protein
-subunits of Gs and Gi2, known as
the products of the gsp and gip oncogenes (4, 5).
In transfected cell lines, GTPase-deficient mutants of G
s and G
i2, as well as corresponding
mutants of G
12/13 and G
q, stimulate
mitogenic responses. The mechanisms for mitogenic stimulation may
involve elevated levels of cAMP (G
s) or activation of
phospholipase C
(G
q). However, in many cases, the
signaling pathways involved are not clear.
Paradoxically, G protein-mediated signaling may also be associated with
growth inhibition and cellular differentiation. Overexpression of
GTPase-deficient G
q in rat pheochromocytoma cells
induces neurite outgrowth (6). Constitutively active forms of
G
13 promote differentiation of P19 mouse embryonal
carcinoma cells into an endodermal phenotype (7). Both appear to
involve stimulation of the c-Jun amino-terminal kinase pathway. In
mouse embryonic stem cells, expression of G
i2 or its
GTPase-deficient mutant results in adipogenic differentiation (8).
Furthermore, expression levels of members of all classes of G protein
-subunits are found to be regulated during cellular differentiation
(9-14). These observations indicate a potentially widespread role of G
protein
-subunits in differentiation programs. However, conclusive
evidence that G protein regulation is the initiating event of the
differentiation process is still lacking in most cases.
The molecular cloning of G
16, a novel member of the
Gq family of G proteins, and the analysis of its tissue
distribution revealed that it is uniquely expressed in normal and in
malignant hematopoietic cells. Its expression is confined to
hematopoietic cell lines that were derived from early stages of
differentiation and is absent or strongly down-regulated in
differentiated normal cells or in leukemia cell lines after induction
of differentiation (12, 13, 15-17). In peripheral blood T-lymphocytes,
G
16 expression is transiently up-regulated after
lymphocyte activation, whereas the expression of G
i2 and
of G
q remains unchanged (18). In order to determine
whether G
16 modulates T-cell activation, regulated
expression of G
16 was disrupted by stably overexpressing
G
16 or G
16 antisense RNA in Jurkat T-cells, a human T lymphoma cell line. Activation of
G
16-deregulated Jurkat T-cells was inhibited as
demonstrated by a reduced up-regulation of interleukin-2 and of the
activation-specific surface antigen CD69 (18). These results suggest a
critical role of tightly regulated G
16 expression in
lymphocyte activation.
Further information about potential roles of G
16
accumulated mostly from experiments in nonhematopoietic cells by
overexpression studies. In Swiss 3T3 fibroblasts, constitutive
activation of Gq-dependent pathways by
overexpression of GTPase-deficient mutants of G
q and
G
16 results in growth arrest or in reduced growth in
response to platelet-derived growth factor or serum, respectively (19).
In small cell lung carcinoma, overexpression of a GTPase-deficient
mutant of G
16 inhibits growth (20), and in rat
pheochromocytoma cells, it induces differentiation (6). Surprisingly,
G
16 has also been reported to be involved in growth
stimulatory events. Activation of the receptor for complement fragment
C5a in human embryonic kidney cells leads to a pronounced activation of
mitogen-activated protein kinase when coexpressed with
G
16 (21). Taken together, these findings imply that G
16-dependent signaling may modulate
cellular proliferation or differentiation, depending on the
specific cellular environment.
Although G16 may interact with a broad spectrum of
receptors in some overexpressing systems (22), selective coupling of receptors to G16, but not to Gq, was observed
for the C-X-C chemokine interleukin-8, as well as for complement
fragment C5a, and for the chemotactic peptide
formyl-methionyl-leucyl-phenylalanine (23-25). In the human
erythroleukemia (HEL)1 cell
line, G
16 specifically couples to the P2Y2
(P2U) purinoceptor (26), suggesting that G
16
might assume specific roles in individual cells or cell lines. However,
its role in hematopoietic cells is still poorly defined in view of its
lineage-independent but differentiation stage-dependent expression.
In the present study, we examined the role of G
16 in
growth and differentiation of erythroleukemic cells. A reporter gene
assay was established, to detect entry into erythroid differentiation in transiently transfected cells. The results indicate that changes in
the expression level and functional activity of G
16 lead to the induction of differentiation in the factor-dependent
erythroleukemia cell line MB-02. The data suggest a new role of
G
16-dependent signaling in the decision
between cellular proliferation and differentiation.
 |
EXPERIMENTAL PROCEDURES |
Reagents--
Cell culture media and medium supplements were
purchased from Life Technologies, Inc. Healthy donors who had received
no recent medications were the source of human serum, which was
heat-inactivated at 56 °C for 30 min and stored at
20 °C for up
to 6 months. Granulocyte-macrophage colony-stimulating factor (GM-CSF),
erythropoietin (Epo), and stem cell factor (SCF) were generous gifts
from Werthenstein-Chemie (Schachen, Switzerland), Cilag AG
(Schaffhausen, Switzerland), and Dr. E. K. Thomas (Immunex Corp.,
Seattle, WA), respectively. Unless otherwise mentioned, all chemicals
(analytical grade), were from Sigma Chemicals (Buchs, Switzerland) or
from Merck AG (Dietikon, Switzerland).
Cell Culture and Induction of Differentiation--
MB-02 cells
(27) were maintained in basal medium (RPMI 1640 medium, 10% human
serum, 2 mM Glutamax I (Life Technologies, Inc.), 1 mM sodium pyruvate, 50 units/ml penicillin, 50 µg/ml streptomycin), supplemented with 5 ng/ml GM-CSF and kept at 37 °C in
a humidified atmosphere of 95% O2 and 5% CO2.
Cells were passaged every third day and replated at a density of 4 × 105/ml. The protocol for the induction of
differentiation was adapted from Broudy et al. (28).
Briefly, cells were washed once with phosphate-buffered saline and
plated at a density of 106 cells/ml in fresh basal medium
supplemented with 25 ng/ml SCF and 4 units/ml Epo and left undisturbed
for a week prior to further routine passaging for another 7-10 days.
Hemoglobin-producing cells were detected by benzidine staining of
cellular suspensions (29).
The HEL cell lines, 3D4 and 1E3, with suppressed G
16
expression were generated by stable transfection of parental HEL cells
derived from clone 92.1 (American Type Culture Collection, Manassas,
VA) with an antisense plasmid to G
16 and have been described previously (26). G418 sulfate-resistant cells that expressed
wild-type levels of G
16 (9G10 and 7H6) were used as
controls. All cell lines were maintained in the presence of G418
sulfate (400 µg/ml) in RPMI 1640 medium, which was supplemented with
2% fetal bovine serum, 2 mM Glutamax I, 1 mM
sodium pyruvate, 50 units/ml penicillin, and 50 µg/ml streptomycin.
For growth studies, cultures were counted every 48 h using a
Coulter Counter (Coulter Electronics, Ltd.)
COS-1 cells (American Type Culture Collection) were maintained in
Dulbecco's modified Eagle's medium supplemented with 10% fetal
bovine serum and 2 mM Glutamax I, 1 mM sodium
pyruvate, 50 units/ml penicillin, 50 µg/ml streptomycin (complete
Dulbecco's modified Eagle's medium).
Plasmid Construction--
The reporter plasmid piGFP was
assembled as follows: the full-length sequence encoding a highly
fluorescent version of green fluorescent protein (GFP) was excised from
phGFP-S65T (CLONTECH) using the SacI and
XbaI sites and inserted into the cloning vector pUC19 (New
England Biolabs). After EcoRI and SalI double
digestion, the fragment including GFP was inserted into the similarly
digested expression vector pEV3 (generously provided by Dr. H. Weir
Zeneca Pharmaceuticals, Cheshire; United Kingdom). As a positive
control, a plasmid constitutively expressing GFP (pcGFP) was generated by cloning the cytomegalovirus (CMV) promoter into piGFP upstream of
GFP. The full-length sequence encoding CMV was cut out from the
expression vector pcDNA3 (Invitrogen, Inc.) by digestion with EcoRI and MunI and then ligated into the unique
EcoRI site of piGFP. As a nonfluorescent control, pclacZ was
constructed replacing GFP in pcGFP with lacZ from the expression vector
pCMV-lacZ (a gift from Dr. S. Rusconi, University of Fribourg, Switzerland).
For cotransfection experiments, the different G protein genes were
cloned into the expression vector pcDNA3. The plasmids pG16AS and
pG16WT, harboring a full-length copy of the human G
16 cDNA in antisense or sense direction, respectively, were described previously (26). In order to generate pG16RC, the GTPase-deficient, constitutively active mutant of G
16,
G
16R186C, was recovered from
pVL1393G
16R186C (a gift from Drs. A. Dietrich and P. Gierschik, University of Ulm, Germany) and cloned into the unique XbaI site of pcDNA3. A plasmid expressing the
corresponding GTPase-deficient mutant of G
q,
pCISG
qR182C, was a gift of Dr. M. I. Simon (California Institute of Technology, Pasadena, CA). A plasmid encoding
the GTPase-deficient mutant of G
i2,
G
i2Q205L, was constructed by excising the coding region
from pCW1G
i2Q205L (30) and inserting it into pcDNA3 at
the HindIII site. pGi3AS was assembled using the
BamHI fragment of pCISG
i3 (also from Dr.
M. I. Simon), which was inserted in antisense orientation into pcDNA3.
Transient Transfection--
MB-02 cells were washed with
phosphate-buffered saline and resuspended at a density of 2 × 106/ml in phosphate-buffered sucrose (sucrose, 272 mM; MgCl2, 1 mM; NaH2PO4, 7 mM; glucose, 20 mM; KCl, 1 mM; pH 7.4). In transfection experiments, 5 µg of piGFP in combination with 5 µg of one of the G
protein-containing plasmids were used, or 5 µg of pcGFP and 5 µg of
pclacZ (positive control), or 10 µg pclacZ (nonfluorescent control).
106 cells and the DNA were mixed in the electroporator
cuvette (gap width, 4 mm) and equilibrated on ice for 5 min. Cells were
then electroporated using a Gene Pulser unit (Bio-Rad) that was set to
350 V, 100 µF (exponential decay), and immediately plated into 1 ml
of prewarmed (37 °C) complete RPMI medium with human serum and
GM-CSF or Epo plus SCF (for noninduced or induced cells, respectively). For the transfection of COS-1 cells, 105 cells in complete
Dulbecco's modified Eagle's medium were seeded into 35-mm dishes.
After 24 h, the culture medium was replaced with Opti-MEM (Life
Technologies, Inc.), and cells were transfected with a total of 1 µg
of plasmid DNA per dish using the
N-[1-(2,3-dioleoyloxy)propyl]-N,N,N-trimethylammoniummethyl sulfate (Boehringer Mannheim) reagent according to the manufacturer's instructions. Cells were harvested after 48 h, and the membrane fraction was analyzed by protein immunoblotting as described previously (26).
Flow Cytometry--
MB-02 cells were washed three times and
resuspended in phosphate-buffered saline (1 ml) supplemented with 22 mM glucose. Cells were analyzed using a FACScan flow
cytometer (Becton Dickinson, Mountain View, CA). Laser excitation was
at 488 nm (argon laser) for GFP, and emission was measured at 515-545
nm. The single color analysis was gated on forward scatter and side
scatter. This gate contained all viable cells and excluded cell debris
and cellular aggregates. A threshold for intensity of fluorescence was
set high enough to exclude autofluorescence and was determined by running samples of untransfected and lacZ-transfected cells prior to
the analysis. A total of 10,000 events was counted and analyzed for
each sample using the CellQuest software (Becton Dickinson). To correct
for variability in transfection efficiencies from experiment to
experiment, inducible GFP expression was normalized in each experiment
to the average of fluorescent cells observed upon expression of GFP
under the constitutively active CMV promoter, i.e. after transfection with pcGFP (three to four independent transfections per experiment).
Data Analysis--
Statistical analyses were performed using the
software packages StatView, version 4.02 for Macintosh (Abacus
Concepts, Inc., Berkeley, CA) or GraphPad Prism, version 2.0 (GraphPad
Software, Inc., San Diego, CA).
 |
RESULTS AND DISCUSSION |
Down-regulation of G
16 in HEL Cells Impairs
Cell Growth--
We recently established several sublines of HEL cells
that showed reduced expression of endogenous G
16 protein
after transfection with a plasmid harboring a full-length copy of
G
16 in antisense orientation (26). In these sublines,
mobilization of intracellular Ca2+ through activation of
the P2Y2 (P2U) purinoceptor by UTP is impaired, whereas Ca2+-mobilization via other receptors is not or is
only partially affected, demonstrating a specific functional defect
(26). Compared to controls with normal levels of G
16,
cellular growth was impaired in the G
16-deficient
sublines (Fig. 1). Population doubling
times were significantly higher in the G
16-suppressed cell lines 3D4 and 1E3 (41.4 ± 1.4 and 41.2 ± 1.6 h,
respectively) than in the cell lines 9G10 and 7H6 expressing normal
levels of G
16 (34.8 ± 1.0, 33.2 ± 0.6 h, respectively) (mean ± S.E.). The results suggested that
G
16-mediated cellular signaling may be involved in the
regulation of cellular proliferation. However, from these experiments,
it could not be established whether G
16-mediated inhibition of proliferation was associated with erythroid
differentiation in hematopoietic cells, because the factor-independent
HEL cells only partially differentiate in response to various inducers
(31, 32). To address this question, a factor-dependent cell
line, MB-02, was chosen that can be readily differentiated along the erythroid pathway (27, 28).

View larger version (14K):
[in this window]
[in a new window]
|
Fig. 1.
Expression of antisense RNA to
G 16 increases population doubling
times of HEL cells. Stably transfected HEL cells with reduced
levels of G 16 expression (G 16-suppressed
cell lines 3D4 and 1E3) or controls with unaltered G 16
expression (G 16-normal cell lines 7D6 and 9G10) were
seeded at a density of 2 × 105 cells/ml. Forty-eight
hours later, total numbers of viable cells were counted, and population
doubling times were calculated. The box plot shows the
summary of multiple measurements. The 10th, 25th, 50th (median), 75th,
and 90th percentiles of the variables are indicated by vertical
bars. One-way analysis of variance indicated significant
differences between the means (po < 0.0001).
Post hoc analysis by the method of Bonferroni-Dunn showed that
differences between controls and G 16-suppressed cell
lines are highly significant (po 0.0014).
|
|
Differentiation of MB-02 Cells--
MB-02 cells maintained in the
presence of GM-CSF differentiate along the erythroid pathway upon
withdrawal of GM-CSF and subsequent treatment with SCF and Epo (28).
Under our experimental conditions, the properties of the cell line
corresponded to the ones described in the original literature:
treatment for 10 days or longer with SCF/Epo resulted in a substantial
proportion of cells that expressed hemoglobin, which was detected by
benzidine staining (Fig. 2A). An average of 45 ± 5% of cells expressed hemoglobin upon
induction, whereas noninduced cells expressed detectable levels only in
6 ± 2% of the population (Fig. 2B). MB-02 cells
showed rapid growth in the presence of GM-CSF, whereas induction of
differentiation with SCF/Epo led to a reduction in the population
growth rate (Fig. 2C). Withdrawal of GM-CSF resulted in
massive cell death within 24 h, confirming their factor dependence
(not shown), whereas withdrawal of SCF during the induction process,
i.e. continuation of induction in the presence of Epo alone,
resulted in a population that was almost stationary in its number of
viable cells (Fig. 2C). Under the latter condition, a higher
proportion of cells expressed hemoglobin, but the overall viability of
the population decreased sharply (not shown). Therefore, in our
experiments, induction of cells was always performed in the presence of
Epo and SCF.

View larger version (50K):
[in this window]
[in a new window]
|
Fig. 2.
Hemoglobin expression and growth
characteristics of MB-02 cells upon induction of differentiation by
SCF/Epo. A, benzidine staining (dark gray)
of hemoglobin in noninduced (maintained in GM-CSF) and induced (SCF/Epo
for 14-16 days) MB-02 cells. B, cells treated as in
A were counted under the microscope, and the percentage of
benzidine positive cells from noninduced and induced cultures was
calculated and plotted (mean ± S.E., n = 6).
C, MB-02 cells were cultured in the presence of GM-CSF,
SCF/Epo, or Epo alone during successive days and counted. The
cumulative fold increase in cell numbers is plotted.
|
|
Assembly and Functional Evaluation of the Reporter System--
In
order to study the effects of G protein overexpression in MB-02 cells,
we used a transient transfection system to acutely manipulate G protein
levels. As the transfection efficiency is low in hematopoietic cells,
potential effects of such treatments could not be studied by
biochemical or immunological methods. Thus, a reporter gene assay was
established based on a plasmid that was co-transfected with the
gene of interest. As a reporter for erythroid differentiation, the
nucleotide sequence encoding GFP was placed under the control of the
-globin promoter in the original plasmid pEV3 (33). In this reporter
plasmid (piGFP), the
-globin promoter is located downstream of locus
control region sequences derived from the human
-globin gene cluster
(Fig. 3A). In constructs
bearing the locus control region-
-globin promoter arrangement,
strong inducible expression from the
-globin promoter has been shown
upon induction of differentiation in erythroid cells (33). In addition,
a plasmid (pcGFP) with constitutive expression of GFP under the control
of the CMV promoter was constructed by splicing in CMV promoter
sequences upstream of the GFP sequence (Fig. 3B).

View larger version (12K):
[in this window]
[in a new window]
|
Fig. 3.
Schematic representations of expression
plasmids piGFP and pcGFP. A, reporter plasmid
piGFP, which expresses GFP upon activation of the -globin promoter.
B, control plasmid pcGFP, which constitutively expresses GFP
under the control of the CMV promoter. The construction of the plasmids
is described in detail under "Experimental Procedures."
|
|
In order to test whether the reporter plasmid was indeed capable of
detecting erythroid differentiation, cells treated with GM-CSF
(noninducing conditions) or with SCF/Epo (inducing conditions) were
transiently transfected with piGFP or with the control plasmid pcGFP.
Expression of GFP was then detected by flow cytometry. As shown in Fig.
4A, the relative number of
GFP-expressing cells was 2.7-fold higher in SCF/Epo-treated cells than
in GM-CSF-treated controls when normalized to GFP expression under the
constitutively active CMV promoter. The relative number of
differentiating cells as measured by the reporter assay reached a
maximum after 11-14 days of induction (Fig. 4B), consistent
with previously published data on SCF/Epo-induced expression of globin
proteins in this cell line (28). In order to quantify the extent of
-globin gene induction in differentiating cells, their fluorescence
intensities resulting from the expression of the reporter gene
construct were compared with that of noninduced cells. The histogram in
Fig. 4C shows an analysis in bins of 0.5 log units over the
4 log units of fluorescence intensities recorded by the FACScan. The
first log unit of fluorescence (Fig. 4C, bins 1 and
2) represents autofluorescence as demonstrated by cells that
were transfected with lacZ instead of the reporter gene construct.
Overall, SCF/Epo-treated cells that were transfected with the reporter
plasmid showed a 2.9-fold increase in their number of fluorescent cells
contained in bins 3-8 when compared with noninduced cells (averages of
95.2 and 32.5 cells, respectively). The corresponding total
fluorescence of induced cells was 4.3-fold higher than in noninduced
cells (Fig. 4C). Thus, SCF/Epo treatment resulted in a
substantial increase in the number of fluorescent cells in the gated
window (encompassing bins 3-8) but only in a slight increase of
average fluorescence per cell (+48%), which is reflected by higher
ratios of fluorescent cells appearing in bins 5-8 than in bins 3 and 4 (3.7, 4.5, 6.5, and 3.3 versus 2.7 and 2.1, respectively).
Apparently, cells that undergo spontaneous differentiation under
noninducing conditions (see also Fig. 2) show
-globin promoter
activities almost equal to those of cells that were treated with
SCF/Epo. Consequently, the number of fluorescent cells rather than gene
expression levels on a per cell basis in the gated window was taken as
a measure for erythroid induction.

View larger version (35K):
[in this window]
[in a new window]
|
Fig. 4.
Detection of erythroid differentiation by the
GFP-reporter assay in induced MB-02 cells. A, induced
and noninduced MB-02 cells were transfected with either pcGFP or piGFP
and analyzed by flow cytometry after 24 h. Numbers of fluorescent
cells observed in piGFP transfectants were normalized to the numbers of
fluorescent cells in pcGFP transfectants to account for different
transfection efficiencies of noninduced and induced (14 days) cells
(4.54 and 1.64% of total numbers of viable cells, respectively).
Induced cells (n = 12) showed a 2.7-fold increase in
normalized numbers of fluorescent cells as compared with noninduced
cells (n = 11). The difference between noninduced and
induced cells is highly significant (po < 0.0002, unpaired t test). B, time course of the
appearance of differentiated MB-02 cells during SCF/Epo treatment.
Cells were transfected with 5 µg of either pcGFP or piGFP and
analyzed by flow cytometry 24 h later at the different time points
indicated in the figure. Cells that were transfected with pclacZ (5 µg) were used as a control to size the acquisition window. Inducible
GFP fluorescence from the reporter plasmid was normalized to
constitutive GFP fluorescence (resulting from the expression of pcGFP),
and the results were plotted. The means (± S.E.) of four different
transfections are shown. C, frequency histogram of
fluorescence intensities of control (pclacZ)- and piGFP-transfected
MB-02 cells (data from B). Fluorescence intensities
(logarithmic scale) were collected in 1024 channels. For the frequency
histograms, the entire range of fluorescence was divided into eight
equal gates. Thus, data from 128 channels each (representing 0.5 log
units of fluorescence) were binned and plotted (bins 1-8, bin 1 representing the lowest and 8 the highest fluorescence intensities).
The mean of four transfections for noninduced (day 0) and six
transfections for induced (day 14) cells are shown (±S.E.). A total of
10,000 cells were counted, and the fluorescence intensities were
analyzed. In this experiment, transfection efficiencies in noninduced
and induced cells matched closely (1.15 and 1.09%, respectively),
allowing for direct comparison of numbers of fluorescent cells (for
details, see under "Experimental Procedures").
|
|
The results shown in Fig. 4A indicate a ratio of
differentiated cells of 46 ± 6% in SCF/Epo-treated cells. The
reporter gene assay closely reflects the relative number of
differentiated MB-02 cells, as detected by benzidine staining of cells
after treatment with SCF/Epo (Fig. 2B), which resulted in
induction rates of 45 ± 5% of the population. However, it
appears that the reporter assay is more sensitive than benzidine
staining at lower levels of
-globin promoter activity: in noninduced
cultures, 6 ± 2% of the cells were benzidine-positive,
whereas 17 ± 1% of the cells showed increased fluorescence in
the reporter assay (compare Figs. 2B and 4A). In
COS-1 cells, transient transfection of pcGFP resulted in strong
fluorescence, whereas no fluorescence was observed in cells transfected
with piGFP (not shown). These results rule out the possibility that low
levels of the reporter gene might have been expressed in the absence of
activators of the
-globin promoter. Thus, the reporter system
provides a valid and sensitive assay for monitoring erythroid
differentiation in MB-02 cells.
Induction of Differentiation in MB-02 Cells upon Changes in
G
16 Expression--
Using the reporter assay, we then
examined the effect of G protein expression on
-globin promoter
activation in nondifferentiated cells. Cells were co-transfected with
the inducible reporter plasmid together with a plasmid encoding either
wild-type G
16 or the GTPase-deficient mutant of
G
16 (G
16R186C). Cultures transfected with
wild-type G
16 showed a significant increase of
GFP-expressing cells to 154% of levels observed in control cells
transfected with pclacZ (Fig.
5A). Expression of the
GTPase-deficient mutant of G
16 led to an even higher
proportion (195% of control) of transfected cells that showed
-globin promoter activity, which was also significantly higher than
for the wild-type
-subunit. In order to test whether this effect was
specific for G
16 or resulted from the mere
overexpression of G protein
-subunits, a GTPase-deficient mutant of
G
q, G
qR182C, was also expressed. G
q represents the phylogenetically closest relative of
G
16, which also belongs to subfamily I of
G
q proteins (34). In contrast to
G
16R186C, no induction was observed when
G
qR182C was expressed (Fig. 5B). Expression
of a GTPase-deficient member of the more distant G
i
family, G
i2Q205L, resulted in a slight increase to
127 ± 19% (mean ± S.E.) of control values, which, however,
was not significantly different from the control (not shown). Although
from the same subfamily of G proteins, G
q apparently is
not able to functionally substitute for G
16 in inducing
differentiation of MB-02 cells. We previously observed that in HEL
cells the P2U purinoceptor specifically couples to G
16, leading to similarly exclusive
G
16-dependent signaling (26). These
experiments demonstrate that the specificity of G
16-dependent signal transduction is not
limited to specificity in the receptor-G protein coupling but may also
result from the interaction of G
16 with downstream
effector systems. The inducing effect of G
16 appears to
depend on functional activity of G
16, because the
GTPase-deficient mutant showed a significantly stronger induction than
the wild-type form. The inability of G
i2 and of
G
q to effectively induce
-globin promoter activity indicates that differentiation strictly depends on G
16
rather than being a general effect of G protein
-subunit
overexpression.

View larger version (16K):
[in this window]
[in a new window]
|
Fig. 5.
Alterations of
G 16 activity trigger entry into
erythroid differentiation in MB-02 cells. Noninduced MB-02 cells
that were maintained in GM-CSF were transiently transfected
(electroporation) with piGFP and various G protein constructs or lacZ
as indicated. After 72 h, cells were analyzed by flow cytometry
for induced fluorescence. Inducible GFP expression as detected in piGFP
transfected cells was then normalized to the total number of
transfected cells as measured in parallel for each experiment by
transfection with pcGFP. The percentages of fluorescent cells 72 h
after transfection of cultures with pcGFP were in the range of
0.41-1.95% of gated, viable cells. A, the ratio of cells
showing inducible fluorescence as a percentage of transfected cells is
shown; the figure represents 16 (lacZ, control), 21 (G16RC), 16 (G16AS), and 15 (G16WT)
independent transfections and flow cytometric analyses, which were
performed in 5-8 different experiments. One-way analysis of variance
of the experimental data indicated that the observed means are not
equal (po < 0.0015). Post hoc analysis by the
method of Dunnett (comparing the control value against all other
values) indicated significant differences of the means against the
control at the level of po < 0.01 (*) or
po < 0.05 (#). ¥ indicates significant
differences between G16RC and G16WT and between G16RC and G16AS when
tested by paired t test for each individual experiment
(po < 0.05). B, MB-02 cells were
transfected with G16RC or a GTPase-deficient, constitutively active
form of G q, G qR182C (GqRC).
Each column represents the mean ratio (± S.E.) of
fluorescent cells observed from eight transfections. One way analysis
of variances indicated that the observed means were not equal
(p < 0.0001), and post hoc analysis by Dunnett's
method indicated a statistically significant difference between the
control and G16RC (p < 0.01 (§)), but no difference
between GqRC and control (p > 0.05 (¶)).
|
|
As HEL cells with suppressed G
16 showed reduced growth
rates (Fig. 1) but were not well suited to examine
G
16-dependent effects on cellular
differentiation, MB-02 cells were also transfected with a
G
16 antisense-plasmid to down-regulate endogenous G
16. Interestingly, transcription of antisense-RNA also
significantly increased the number of GFP-positive cells to 156% of
control values (Fig. 5A). Since differentiation is
associated with reduced rates of proliferation, this inducing effect of
G
16-down-regulation in MB-02 cells is consistent with
its effect on proliferation that was detected in HEL cells (Fig. 1).
However, the observation that either reducing or up-regulating the
levels of G
16 protein expression triggers
differentiation may not easily be reconciled. Although this phenomenon
has also been observed for acquiring functional competence of Jurkat T
cells (18), the underlying mechanism cannot be deduced from the
available data and warrants further investigation. A cell-specific,
optimal level of G
16 activity seems to be required for
proliferation, and any deviation from it will decrease growth rates and
result in increased ratios of differentiating cells.
Expression of Antisense RNA Inhibits G
16 Expression
in Transiently Transfected COS-1 Cells--
Due to low transfection
efficiencies, expression of G
16 protein or
antisense-dependent down-regulation of endogenous G
16 could not be verified directly in MB-02 cells.
Therefore, expression of G
16 protein and its
down-regulation through antisense RNA was verified by protein
immunoblotting of solubilized membranes of transiently transfected
COS-1 cells. Transfection with G
16R186C or with
wild-type G
16 resulted in a single band with a relative
mobility identical to the single immunoreactive band observed in
membranes of MB-02 cells (Fig. 6). As
expected, in COS-1 cells that were transfected with the control plasmid pclacZ, no immunoreactivity was detected, as these cells do not express
endogenous G
16. Transfection of smaller amounts of the
G
16 plasmid resulted in lower expression levels,
indicating that under these conditions, plasmid-dependent
protein expression was in a dynamic range, thus allowing for the
detection of copy number-dependent alterations of
expression. As shown in Fig. 6, co-transfection of G
16
with the G
16 antisense construct resulted in a marked
down-regulation of G
16 expression when compared with
controls that were transfected with G
16 alone. A plasmid
leading to a G
i3 antisense RNA transcript was not capable of reducing the expression of G
16,
indicating specific interaction of the G
16-transcript
with G
16 antisense sequences (Fig. 6, compare
lanes G16WT+lacZ and G16WT+Gi3AS).

View larger version (13K):
[in this window]
[in a new window]
|
Fig. 6.
G 16
antisense expression down-regulates
G 16 expression in COS-1
cells. COS-1 cells were transfected with plasmids encoding
G 16R186C (G16RC), lacZ (lacZ), or
wild-type G 16 (G16WT) or with
G 16 (G16AS) or G i3
(Gi3AS) antisense plasmids as indicated in the figure. A
total of 1 µg of DNA was used in each transfection. Co-transfections
were done with pG16WT (0.15 µg) plus either pclacZ (0.85 µg),
pG16AS (0.85 µg), or pGi3AS (0.85 µg). Forty-eight hours after
transfection, membrane fractions were prepared, and the proteins (15 µg of total protein per lane) were separated by 10%
SDS-polyacrylamide gel electrophoresis and immunoblotted for
G 16 using the antiserum AS 339 (35) as described earlier
(26). The membrane fraction (15 µg of protein) of MB-02 cells was
also analyzed, showing expression of endogenous G 16 in
this cell line. The figure shows the immunoblot, which was developed
using the enhanced chemiluminescence protocol (Amersham Pharmacia
Biotech).
|
|
The results suggest a causative role for a
G
16-dependent signaling pathway in the
induction of differentiation in MB-02 cells. The identification of
constitutively active, GTP-bound G
16 as the active
molecular entity suggests that in a natural environment, activation of
G
16 in response to receptor activation may indeed induce
differentiation. This conclusion is supported by the results from the
experimental up- or down-regulation of wild-type G
16,
which is expected to translate into changes in basal levels of
GTP-bound G
16, and consequently into changes of
G
16-dependent signaling activity. It is not
known which receptors may engage G
16 in MB-02 cells.
Therefore, an assessment of the role of agonists that activate
G
16-coupled receptors is not yet possible. The
observation that the GTPase-deficient G
16 shows a higher
efficacy than the wild-type form also excludes the possibility that
G
16 may act solely as a receptor-independent regulator
of G protein signaling, e.g. by capturing 
-subunits that might have been liberated by activation of other G proteins.
Transfection of MB-02 cells with the G
16R186C mutant
resulted in a 1.9-fold increase of induced cells as compared with the
number observed in control-transfected cells (Fig. 5A). This increase is substantial in view of the 2.7-fold increase observed when
cells were induced to differentiate by SCF/Epo (Fig. 4A). Importantly, cells induced with SCF/Epo received the differentiating stimulus for 14 days, whereas transfected cells had to be scored after
3 days due to the transient nature of transfection. It was not possible
to subject cells to transfection after 3 days of differentiation by
SCF/Epo due to the fragile nature of the cultures at this time point.
However, as demonstrated in Fig. 4B, SCF/Epo treatment
required several days to fully differentiate MB-02 cells. Thus,
G
16R186C
and deregulation of G
16
activity in general
may be considered as potent inducers of differentiation.
In this context, it should be noted that our attempts to generate
stable HEL cell lines transfected with G
16R186C were not
successful. It appeared that expression of constitutively active
G
16 inhibits proliferation in these cells to an extent that precluded the isolation of cell clones. Expression of the functionally similar GTPase-deficient mutant G
16Q212L in nonhematopoietic cells apparently led to growth retardation, but stable
cell lines could still be established (19). G
16 may thus
act as a much stronger (negative) regulator of proliferation in
hematopoietic cells than in other cell lines. Whether this could be
caused by a unique coupling of G
16-dependent
signaling to downstream effector systems or by stronger coupling to
effector systems that are used by other members of the
G
q-family is not known.
In conclusion, in hematopoietic cells an increase of G
16
function or its down-regulation have profound effects on proliferation and may cause erythroid differentiation in specific cell lines. It
remains to be established whether differentiation along the erythroid
pathway is specifically determined by G
16 or whether G
16 preconditions hematopoietic cells for
differentiation independently of lineage determination. Deviations from
optimal levels of G
16 activity seem to be associated
with acquiring functional competence by inducing either differentiation
or activation of cellular proliferation, as seen in T-lymphocytes (18).
One might speculate that agonists employing G
16-coupled
receptors directly regulate the proportion of functionally competent
cells. With the reporter assay based on transient transfection of cells and subsequent analysis by flow cytometry, it should now be possible to
identify the G
16-dependent signaling
pathways involved in these processes. Furthermore, these studies could
be extended to primary hematopoietic progenitor cells, in which G
protein expression cannot be easily manipulated by other techniques.