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INTRODUCTION |
Opioid receptors are a typical seven-transmembrane domain receptor
family that signal through inhibitory G proteins to a multitude of
second messengers and cellular effectors, including adenylyl cyclase,
voltage-operated calcium channels, and inwardly rectifying K+ channels (1); intracellular calcium stores (2); and the extracellular signal-regulated kinase
(ERK)1 mitogen-activated
protein kinase (MAPK) pathway (3, 4). There are three principal types
of opioid receptors, µ,
, and
, with ~60% homology. However,
the µ-opioid receptor has generated the most interest as the
principal site of action for clinical analgesics and abused opiate
drugs. The µ-opioid receptor can couple to all members of the
G
i/o family, with little selectivity for particular G
subunits (5). Selectivity of intracellular µ-opioid signaling would
therefore appear to depend on cell-specific expression of G protein
subunits coupled with the selectivity of G proteins to interact with
particular effectors. However, it has been suggested that other factors
besides G protein and effectors may contribute to signaling specificity
(6-8).
Agonist activation of G protein-coupled receptors leads to exchange of
GDP for GTP on G
and dissociation of the G
-GTP and G
subunits. Deactivation is brought about by the intrinsic GTPase activity of G
, causing GTP to be hydrolyzed to GDP and the
subsequent reassociation of the G
-GDP and G
subunits. G
protein signaling in this fashion is negatively controlled by a family
of proteins known as RGS (regulators of G
protein signaling) proteins (6). These proteins act as
GTPase-activating proteins (GAPs) for G
and speed up the hydrolysis
of the G
-bound GTP, thus reducing the steady-state levels of
G
-GTP and inhibiting signaling. Therefore, it has been suggested
that, as with other G protein-coupled receptors, RGS proteins act to
inhibit µ-opioid signaling and may play a controlling role in the
effectiveness of opioid receptor ligands. In support of this idea,
overexpression of RGS2 shifts the concentration effect curve for
morphine-stimulated pigment aggregation to the right to a small
(2-fold) degree in cultured dermal melanophores from Xenopus
laevis transiently expressing a murine µ-opioid receptor (9). Furthermore, a reduction in RGS9 levels in mice using antisense
oligonucleotide leads to an increase in the anti-nociceptive potency of
morphine (10). Although these changes are small, they are suggestive of
a role for RGS proteins in opioid coupling efficiency.
An important question is whether RGS proteins alter the efficiency of
all intracellular signaling pathways equally or whether there is a
variable effect that would provide for selectivity. Selectivity for
particular pathways may be obtained by several mechanisms.
RGS-containing proteins have a wide variety of non-RGS domains (11-13)
that, when RGS protein binds to G
, can link other proteins and
signaling pathways to provide for diversity of signaling. In addition,
the interaction of RGS protein with receptors may contribute to
selectivity; for example, RGS12 binds to the carboxyl terminus of the
interleukin-8 receptor (14), and inhibition of Ca2+
signaling in rat pancreatic acinar cells by RGS4 is selective for
muscarinic receptors relative to bombesin and cholecystokinin receptors
(15) possibly through interaction of the N-terminal domain of RGS4 with
the receptors (16). Recently, Wang et al. (17) demonstrated,
using ribozyme technology, that RGS3 is a negative modulator of m3
muscarinic receptor signaling, whereas RGS5 is a negative modulator of
angiotensin type 1a receptor signaling through
Gq/11.
In addition to RGS proteins selectively modulating the coupling of
different receptors to a single effector, it is possible that RGS
proteins could selectively modulate the coupling of a single receptor
to different effectors. Indeed, we have recently proposed a "kinetic
scaffolding" model for G protein signaling in which RGS proteins
confer selectivity for signaling pathways by their ability to shorten
the lifetime of G
-GTP (18). In this model, RGS protein
accelerates hydrolysis of the G
-bound GTP, permitting recombination
of G
-GDP and G
and recoupling of the heterotrimer and receptor
and allowing rapid reactivation by agonist-bound receptors. This
maintains active G
-GTP and G
proteins in the close vicinity of
the receptor, but spillover of G
-GTP and G
to more distant
effectors is prevented by the GAP activity of RGS. This effect can be
mediated by the RGS domain alone and does not depend on other protein
interaction modules.
Here we test the hypothesis that RGS proteins differentially regulate
µ-opioid receptor coupling to signaling pathways, thus contributing
to selectivity of receptor activation of second messenger pathways.
Because 30 mammalian proteins with RGS activity have been identified to
date (12, 13), the choice of which RGS protein to study is a difficult
one. We have therefore made use of a point mutation in
G
o (G184S) that is known to block interaction with all
members of the RGS family without affecting GTPase activity (RGS-insensitive) (19), together with a mutation (C351G) that confers pertussis toxin (PTX) insensitivity (PTXi) (20). In this way,
when the RGS- and PTX-resistant G
o mutant
(G
oRGS/PTXi) is expressed in a cellular system, coupling
to endogenously expressed G proteins can be inactivated by PTX
treatment, and the system must then signal through the expressed
G
o mutant (21).
Our findings demonstrate that the µ-opioid agonists DAMGO and
morphine showed increased potency and/or efficacy of signaling to
adenylyl cyclase in cells expressing RGS-insensitive G
o
compared with those expressing RGS-sensitive G
o.
Signaling through the MAPK pathway also showed an increased potency
with the full agonist DAMGO, but not an increased maximal effect,
although the maximal effect of the partial agonist morphine was
significantly enhanced. In contrast, the ability of DAMGO or morphine
to stimulate the release of calcium from intracellular stores was
altered to a much lesser extent in cells expressing RGS-insensitive
G
o compared with those expressing RGS-sensitive
G
o. These results confirm that RGS proteins can modulate
effector signaling by a single G protein and may play an important role
in directing effector responses to µ-opioid receptor signaling.
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EXPERIMENTAL PROCEDURES |
Materials--
[3H]DAMGO,
[
-32P]GTP, and [35S]GTP
S were from
PerkinElmer Life Sciences, and cAMP radioimmunoassay kits were from
Diagnostic Products Corp. (Los Angeles, CA). Tissue culture medium,
LipofectAMINE Plus reagent, Geneticin, Zeocin, fetal bovine serum, and
trypsin were from Invitrogen. Morphine and naloxone were obtained
through the Opioid Basic Research Center at the University of Michigan (Ann Arbor, MI), and DAMGO was obtained from Sigma. Trizma (Tris base),
GDP, ATP, and other biochemicals were purchased from Sigma and were
analytical grade. Anti-phospho-p44/42 MAPK (ERK1/2) antibody and
anti-p44/42 MAPK (ERK1/2) antibody were from Cell Signaling Technology,
Inc. (Beverly, MA); anti-G
o antibody (K-20) and
secondary antibodies were from Santa Cruz Biotechnology, Inc. (Santa
Cruz, CA); and SuperSignal West Pico chemiluminescent substrate was from Pierce. PTX-insensitive G
o (G
oPTXi,
C351G) and RGS- and PTX-insensitive G
o
(G
oRGS/PTXi, G184S/C351G) DNAs in the pCI vector were
obtained from Dr. Stephen Ikeda (Guthrie Research Institute, Sayre,
PA). GST fusion protein containing RGS8 (GST-RGS8) and
His6-tagged G
o were prepared as previously
described (22).
Expression of G
oPTXi or
G
oRGS/PTXi in C6µ Cells and Cell
Culture--
G
oPTXi or G
oRGS/PTXi DNA
was excised from the plasmid vector pCI with NotI and
NheI restriction enzymes and inserted into the Zeocin
resistance vector pcDNA3.1zeo
. Plasmid DNA was
transfected into C6 glioma cells stably expressing the rat µ-opioid
receptor (C6µ cells) (23) using LipofectAMINE Plus reagent. Colonies
were isolated from transfected cells grown in Dulbecco's modified
Eagle's medium containing 10% fetal bovine serum under 5%
CO2 in the presence of 0.25 mg/ml Geneticin (to maintain
expression of the µ-opioid receptor in a Geneticin-resistant plasmid)
and 0.4 mg/ml Zeocin. Clones were maintained under the same conditions
and typically subcultured at a ratio of 1:20 to 1:30, with partial
replacement of the medium on Day 4 and the day before subculturing or
harvesting at Day 7.
Membrane Preparation--
Cells were treated with or without PTX
(100 ng/ml) overnight before collection. Cells were washed twice with
ice-cold phosphate-buffered saline (0.9% NaCl, 0.61 mM
Na2HPO4, and 0.38 mM
KH2PO4, pH 7.4), detached from the plates by
incubation in harvesting buffer (20 mM HEPES, pH 7.4, 150 mM NaCl, and 0.68 mM EDTA) at room temperature, and pelleted by centrifugation at 200 × g for 3 min.
The cell pellet was suspended in ice-cold 50 mM Tris-HCl
buffer, pH 7.4, and homogenized with a Tissue Tearor (Biospec Products,
Inc.) for 20 s at setting 4. The homogenate was centrifuged at
20,000 × g for 20 min at 4 °C, and the pellet was
resuspended in 50 mM Tris-HCl, pH 7.4, with a Tissue Tearor
for 10 s at setting 2, followed by recentrifugation. The final
pellet was resuspended in 50 mM Tris-HCl, pH 7.4, to
0.5-1.0 mg/ml protein and frozen in aliquots at
80 °C. To
determine protein concentration, membrane samples were dissolved with 1 N NaOH for 30 min at room temperature, neutralized with 1 M acetic acid, and assayed by the method of Bradford (42)
using bovine serum albumin as the standard.
Determination of G
o Expression--
Membrane
proteins (20 µg) or His6-G
o standards
(10-40 ng) were separated by SDS-PAGE on 12% gels (Protogel, National
Diagnostics, Inc., Atlanta, GA). Proteins were transferred to a
nitrocellulose membrane (45 µm; Osmonics, Inc., Minnetonka, MN),
probed with a 1:200 dilution of anti-G
o antibody,
treated with horseradish peroxidase-conjugated goat anti-rabbit IgG,
and visualized by enhanced chemiluminescence. Quantification was done
using a Eastman Kodak Image Station 440.
Receptor Binding Assay--
Membranes (10-20 µg) were
incubated in 50 mM Tris-HCl, pH 7.4, with 0.2-28
nM [3H]DAMGO with or without 50 µM naloxone (to determine nonspecific binding) in a total
volume of 0.2 ml for 60 min in a shaking water bath at 25 °C.
Samples were filtered through glass-fiber filters (No. 32; Schleicher & Schüll) mounted in a Brandel cell harvester and rinsed three
times with ice-cold 50 mM Tris-HCl, pH 7.4. Radioactivity retained on the filters was counted by liquid scintillation counting in
4 ml of EcoLume scintillation mixture (ICN, Aurora, OH).
[35S]GTP
S Binding Assay--
Membranes (14-20
µg) were incubated for 60 min in a shaking water bath at 25 °C
with 20 mM Tris-HCl, pH 7.4, 5 mM
MgCl2, 100 mM NaCl, 0.1 mM
dithiothreitol (DTT; freshly prepared), 30 µM GDP, 0.1 nM [35S]GTP
S, and 0.01-10
µM DAMGO, 0.01-10 µM morphine, or
dH2O. Samples were filtered through the glass-fiber filters
mounted in a Brandel cell harvester and rinsed three times with
ice-cold 50 mM Tris-HCl, pH 7.4, 5 mM
MgCl2, and 100 mM NaCl. Radioactivity retained
was determined as described above. For kinetic studies, membranes (~20 µg) were incubated for 10 min at 25 °C in 20 mM
HEPES, 10 mM MgCl2, 100 mM NaCl,
0.1 mM DTT, and 1 mM EDTA, pH 7.4, containing 30 µM GDP. DAMGO (10 µM) was added, and the
mixture was further incubated for 10 min before the addition of 0.1 nM [35S]GTP
S to start the reaction. At
various times (3 min to 2 h), bound and free radioactivities were
separated and quantified as described above.
GTPase Assay--
Membranes (14-20 µg) were prewarmed for
5-20 min at 30 °C with 10 mM Tris, pH 7.6, 2 mM MgCl2, 20 mM NaCl, 0.2 mM EDTA, 0.1 mM DTT (freshly prepared),
an ATP-regenerating system (0.2 mM ATP, 0.2 mM
AppNHp, 50 units/ml creatine phosphokinase, and 5 mM phosphocreatine), and 0.01-10 µM DAMGO,
10 µM morphine, or dH2O with or without 1 µM GST-RGS8. The reaction was initiated by the addition
of 0.1 µM [
-32P]GTP (prewarmed to
30 °C) to a final volume of 0.1 ml. The reaction was stopped after
15-120 s by the addition of ice-cold 15% charcoal with 20 mM phosphoric acid in 0.1% gelatin. After at least 30 min
on ice, samples were centrifuged at 4000 × g for 20 min at 4 °C, and 0.3 ml was taken from the supernatant for liquid
scintillation counting with 4 ml of EcoLume scintillation mixture.
Blank values for each time point (without membranes) were subtracted
from each value.
Inhibition of cAMP Accumulation--
Cells were plated to
confluency in 24-well plates the day before the assay and treated
overnight with 100 ng/ml PTX. To start the assay, the cells were rinsed
with serum-free medium and then incubated with serum-free medium
containing 30 µM forskolin, 1 mM
3-isobutyl-1-methylxanthine, and 0.001-10 µM DAMGO,
0.01-10 µM morphine, or dH2O for 30 min at
37 °C. The reaction was stopped by replacing the medium with
ice-cold 3% perchloric acid. After at least 30 min at 4 °C, 0.4 ml
was removed from each sample, neutralized with 0.08 ml of 2.5 M KHCO3, vortexed, and centrifuged at
15,000 × g for 1 min. A radioimmunoassay kit was used
to quantify accumulated cAMP in a 10-µl aliquot of the supernatant
from each sample. Inhibition of cAMP formation was determined as a
percentage of forskolin-stimulated cAMP accumulation in the absence of
opioid agonist.
Stimulation of p44/42 MAPK Phosphorylation--
Cells
were plated in six-well plates the day before the assay to reach
70-90% confluency on the day of the assay and treated overnight with
100 ng/ml PTX. The medium was replaced with serum-free medium for
2 h before the addition of 0.001-10 µM DAMGO, 10 µM morphine, or dH2O. The assay was stopped
after 1-20 min by rinsing the cells twice with ice-cold
phosphate-buffered saline and adding 0.1 ml of ice-cold SDS sample
buffer (62.5 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol,
50 mM DTT, and 0.01% bromphenol blue). Samples were
removed from the wells; sonicated for 10-15 s; boiled for 5 min; and
then subjected (120 µg) to 12% SDS-PAGE, followed by transfer to
45-µm nitrocellulose membranes for Western blotting. The blot was
probed with a 1:2000 dilution of anti-phospho-p44/42 MAPK (ERK1/2)
antibody and visualized using horseradish peroxidase-conjugated anti-mouse IgG, followed by enhanced chemiluminescence detection and
quantification using the Image Station 440. To assure equal loading,
membranes were stripped and reblotted with a 1:1000 dilution of
anti-p44/42 MAPK (ERK1/2) antibody to measure total ERK levels.
Release of Intracellular Calcium--
After overnight treatment
with 100 ng/ml PTX and 5 µM forskolin, confluent cells
were harvested with 10 mM HEPES-buffered 0.9% saline
containing 0.05% EDTA, pH 7.4, and washed twice with and then
resuspended in Krebs-HEPES buffer of the following composition: 143.3 mM NaCl, 4.7 mM KCl, 2.5 mM
CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 11.7 mM
glucose, and 10 mM HEPES, pH 7.4, with 10 M
NaOH. Cell suspensions were loaded with 3 µM
fura-2/acetoxymethyl ester for 30 min at 37 °C, washed, incubated at
20 °C for 20 min, and then rewashed. Intracellular calcium
concentrations were measured in 1-ml volumes at 37 °C using a
Shimazdu RF5000 spectrofluorophotometer at 340/380 nm excitation and
510 nm emission. In certain experiments, nominally
Ca2+-free buffer containing 0.1 mM EGTA was
used and was included in the final resuspension only. Data are
presented as the
340/380 nm ratio (mean ± S.E.).
Data Analysis--
Concentration-effect data from GTPase,
[35S]GTP
S binding, adenylyl cyclase, MAPK
phosphorylation, and [Ca2+]i assays were fitted
to sigmoidal concentration-effect curves using GraphPAD Prism to
determine EC50 values and maximal effects. Specific binding
data were fitted to a one-site binding hyperbola using GraphPAD Prism
to determine KD and Bmax values. Data are presented as means ± S.E. from at least three separate experiments and are compared using two-tailed Student's t test unless stated otherwise.
 |
RESULTS |
Expression of the G
oPTXi and
G
oRGS/PTXi Mutants in C6µ Glioma
Cells--
C6µ cells were stably transfected with
G
oPTXi (C6µ-G
oPTXi cells) or
G
oRGS/PTXi (C6µ-G
oRGS/PTXi cells). A
C6µ-G
oPTXi clone (C1) and a
C6µ-G
oRGS/PTXi clone (M1) chosen as expressing similar levels of G
o (Fig.
1A) were used for most
studies. In membrane preparations, the binding affinity of
[3H]DAMGO in Tris-HCl buffer was similar, with values of
2.0 ± 0.1 nM for C6µ-G
oPTXi cells
(clone C1) and 3.5 ± 0.9 nM for
C6µ-G
oRGS/PTXi cells (clone M1). The
Bmax values for [3H]DAMGO were
7.6 ± 1.3 pmol/mg in C6µ-G
oPTXi cells (clone C1) and 14.4 ± 2.0 pmol/mg in C6µ-G
oRGS/PTXi cells
(clone M1). PTX treatment (100 ng/ml overnight) of wild-type C6µ
cells abolished opioid agonist-mediated signaling, as assessed by
inhibition of cAMP accumulation, stimulation of
[35S]GTP
S binding, stimulation of MAPK
phosphorylation, and increases in [Ca2+]i through
endogenous G proteins (data not shown). PTX treatment of C6µ cells
expressing G
oPTXi or G
oRGS/PTXi allowed signaling through the transfected PTX-resistant Go proteins
to be measured.

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Fig. 1.
Expression and activation of
G oPTXi or
G oRGS/PTXi in
C6µ cells. A, membranes were
prepared from wild-type (wt) C6µ cells or from C6µ cells
stably transfected with G oPTXi (PTXi) or
G oRGS/PTXi (RGSi) as described under
"Experimental Procedures" and subjected to SDS-PAGE (20 µg of
membranes or 20 ng of G o standard (Go)).
Proteins were transferred to a nitrocellulose membrane, incubated with
anti-G o antibody followed by horseradish
peroxidase-conjugated anti-rabbit IgG, and visualized by
chemiluminescence as described under "Experimental Procedures."
Shown is a representative blot from three separate blots. B,
membranes (14-20 µg) from PTX-treated C6µ-G oPTXi
(closed symbols) or C6µ-G oRGS/PTXi
(open symbols) cells were incubated for 60 min at 25 °C
with 50 mM Tris-HCl, pH 7.4, 5 mM
MgCl2, 100 mM NaCl, 1 mM EDTA, 1 mM DTT, 30 µM GDP, 0.1 nM
[35S]GTP S, and DAMGO (squares), morphine
(circles), or dH2O. Data are derived from four
assays, each carried out in duplicate, and are expressed as a
percentage of basal binding. C, membranes (14-20 µg) from
PTX-treated C6µ-G oPTXi ( ) or
C6µ-G oRGS/PTXi ( ) cells were incubated for 10 min
at 25 °C in 20 mM HEPES, 10 mM
MgCl2, 100 mM NaCl, 0.1 mM DTT, and
1 mM EDTA, pH 7.4, containing 30 µM GDP,
followed by a 10-min incubation with or without 10 µM
DAMGO before the addition of 0.1 nM
[35S]GTP S to start the assay, which was allowed to
proceed for 3-100 min. Samples were then harvested and counted as
described under "Experimental Procedures." Data are presented as a
percentage of the mean maximal DAMGO response from five assays, each
performed in duplicate.
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Stimulation of [35S]GTP
S Binding--
Basal
levels of [35S]GTP
S binding were not different in
membranes from PTX-treated G
oPTXi-expressing (0.026 ± 0.003 pmol/mg) and G
oRGS/PTXi-expressing (0.031 ± 0.002 pmol/mg) cells. The maximal stimulation of
[35S]GTP
S binding produced by DAMGO over basal binding
in membranes (Fig. 1B) was lower in
C6µ-G
oRGS/PTXi membranes (157 ± 26% over control) versus C6µ-G
oPTXi membranes
(250 ± 67% over control), but the difference was not
statistically significant. A similar pattern was seen for maximal
stimulation by morphine (C6µ-G
oRGS/PTXi membranes,
48 ± 7% over control; and C6µ-G
oPTXi membranes,
66 ± 17% over control). The potencies of the two µ-agonists
for stimulation of [35S]GTP
S binding was also the same
(DAMGO, EC50 = 296 ± 30 nM in C6µ-G
oPTXi cells and EC50 = 316 ± 30 nM in C6µ-G
oRGS/PTXi cells; and morphine,
EC50 = 124 ± 24 nM in
C6µ-G
oPTXi cells and EC50 = 159 ± 41 nM in C6µ-G
oRGS/PTXi cells). The rate of
stimulation of [35S]GTP
S binding by 10 µM DAMGO (Fig. 1C) was the same in
C6µ-G
oPTXi membranes (k = 0.17 ± 0.05 h
1) and C6µ-G
oRGS/PTXi membranes
(k = 0.17 ± 0.09 h
1).
Stimulation of GTPase Activity--
The rates of basal GTP
hydrolysis were 10.3 ± 0.3 and 13.2 ± 0.3 pmol/mg/min in membranes from PTX-treated C6µ-G
oPTXi
and C6µ-G
oRGS/PTXi cells, respectively. DAMGO
stimulated GTP hydrolysis in membranes from both PTX-treated
C6µ-G
oPTXi and C6µ-G
oRGS/PTXi cells
(Fig. 2A). The DAMGO
stimulation of GTP hydrolysis in C6µ-G
oPTXi cells
(5.70 ± 0.50 pmol/mg/min) was greater (p < 0.05)
than in C6µ-G
oRGS/PTXi cells (2.64 ± 0.75 pmol/mg/min), consistent with reduced GTPase activity. Furthermore, the
addition of 1 µM GST-RGS8 increased the DAMGO stimulation
of GTP hydrolysis by a maximal concentration of DAMGO (1 µM) in C6µ-G
oPTXi (but not
C6µ-G
oRGS/PTXi) membranes, indicating the
effectiveness of the RGS-insensitive mutation in preventing the GAP
activity of RGS8. GTPase stimulation in C6µ-G
oPTXi
membranes by DAMGO at 2 min was concentration-dependent (Fig. 2B). GST-RGS8 increased the maximal stimulation over
basal levels by DAMGO from 60 ± 7 to 151 ± 19%
(p < 0.05), with a shift in the EC50 value
from 34 ± 12 to 92 ± 31 nM, although this did not reach significance (p = 0.16). The maximal
stimulation of GTP hydrolysis by 10 µM morphine at 2 min
was 45 ± 5% in the C6µ-G
oPTXi membranes and
increased significantly (p < 0.05) to 94 ± 17%
in the presence of 1 µM GST-RGS8, although relative to
DAMGO, morphine was significantly less efficacious (p < 0.05) in the presence of GST-RGS8 (61 ± 3%) than in its
absence (75 ± 1%). RGS8 was chosen for these studies because it
is structurally a simpler RGS protein and is known to be a GAP for
G
o (22).

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Fig. 2.
A, stimulation of GTPase activity by 10 µM DAMGO in C6µ-G oPTXi or
C6µ-G oRGS/PTXi cells in the presence or absence of
RGS8. Membranes (14-20 µg) from PTX-treated
C6µ-G oPTXi (closed symbols) or
C6µ-G oRGS/PTXi (open symbols) cells were
prewarmed for 5 min at 30 °C with 10 µM DAMGO in the
absence (squares) or presence (triangles) of 1 µM RGS8 in a buffer system of 50 mM Tris, pH
7.4, 1 mM EDTA, 10 mM MgCl2, 100 mM NaCl, 1 mM DTT, and an ATP-regenerating
system. The reaction was initiated by the addition of prewarmed
[32P]GTP to a final concentration of 0.1 µM
and stopped at varying times between 15 and 120 s by the addition
of ice-cold 15% charcoal in 20 mM
H3PO4 plus 0.1% gelatin. Data are expressed as
stimulation of Pi released by DAMGO after subtraction of
basal release. Shown are the combined data from three assays.
B, concentration-effect curve for stimulation of
GTPase activity by DAMGO in C6µ-G oPTXi cells in the
presence and absence of RGS8. Membranes (14-20 µg) from PTX-treated
C6µ-G oPTXi cells were prewarmed for 5 min at 30 °C
with varying concentrations of DAMGO in the absence ( ) or presence
( ) of 1 µM RGS8 in a buffer system of 50 mM Tris, pH 7.4, 1 mM EDTA, 10 mM
MgCl2, 100 mM NaCl, 1 mM DTT, and
an ATP-regenerating system. The reaction was initiated by the addition
of prewarmed [32P]GTP to a final concentration of 0.1 µM and stopped after 120 s by the addition of
ice-cold 15% charcoal in 20 mM
H3PO4 plus 0.1% gelatin. Data are given as
pmol/Pi released/mg/min and are the combined data from
three assays.
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Inhibition of Adenylyl Cyclase--
Adenylyl cyclase activity was
measured by the accumulation of cAMP stimulated by forskolin in the
presence of the phosphodiesterase inhibitor 3-isobutyl-1-methylxanthine
in PTX-treated whole cells. The level of accumulated
forskolin-stimulated cAMP was the same in both the
G
oPTXi- and G
oRGS/PTXi-expressing cells.
The maximal inhibition of cAMP accumulation by DAMGO was significantly
greater (p < 0.05) in C6µ-G
oRGS/PTXi
cells (58 ± 5% inhibition) than in C6µ-G
oPTXi
cells (35 ± 6% inhibition) (Fig.
3). More impressive was the fact that
DAMGO was ~35-fold more potent (p < 0.05) in C6µ-G
oRGS/PTXi cells (EC50 = 12 ± 1 nM) than in C6µ-G
oPTXi cells (EC50 = 404 ± 112 nM). Morphine
inhibition of forskolin-stimulated cAMP accumulation (Fig. 3) increased
significantly (p < 0.01) from a maximum of 10 ± 5% in C6µ-G
oPTXi cells to 54 ± 7% in
C6µ-G
oRGS/PTXi cells and showed an 8-fold increase in
potency (C6µ-G
oRGS/PTXi cells, EC50 = 21.7 ± 11.2 nM; and C6µ-G
oPTXi
cells, EC50 = 170 ± 53 nM), although this
did not quite reach significance at the 0.05 level (p = 0.053). To ensure that the striking difference between the
G
oPTXi- and
G
oRGS/PTXi-expressing cells was not caused
by differences in receptor and G
o expression levels,
inhibition of cAMP accumulation was measured in two additional
C6µ-G
oPTXi clones (C2 and C3) and an additional
C6µ-G
oRGS/PTXi clone (M2). DAMGO and morphine were
consistently more potent and gave higher maximal effects in the
C6µ-G
oRGS/PTXi clones (Table
I).

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Fig. 3.
Inhibition of cAMP accumulation. Cells
were plated to confluency in 24-well plates the day before the assay
and treated overnight with 100 ng/ml PTX. C6µ-G oPTXi
(closed symbols) or C6µ-G oRGS/PTXi
(open symbols) cells were rinsed with serum-free medium and
then incubated with serum-free medium containing 30 µM
forskolin, 1 mM 3-isobutyl-1-methylxanthine, and DAMGO
(squares), morphine (circles), or
dH2O for 30 min at 37 °C. The reaction was stopped by
replacing the medium with ice-cold 3% perchloric acid. After keeping
the samples at 4 °C for at least 30 min, the samples were
neutralized, and cAMP was quantified using a radioligand binding assay
kit as described under "Experimental Procedures." Values are
expressed as a percentage of the values with forskolin only (in the
absence of ligand), which were the same for C6µ-G oPTXi
(6.6 ± 0.5 pmol/mg) and C6µ-G oRGS/PTXi
(6.8 ± 0.9 pmol/mg) cells. Shown are the combined data from three
assays, each measured in duplicate.
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Table I
Inhibition of cAMP accumulation in C6µ-G oPTXi and
C6µ-G oRGS/PTXi cells by DAMGO and morphine
after treatment with PTX
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In wild-type C6µ, C6µ-G
oRGS/PTXi (clone M1), and
C6µ-G
oPTXi (clone C1) cells not treated overnight with
PTX, DAMGO and morphine robustly inhibited cAMP accumulation with
similar EC50 values and maximal effects (Table II).
DAMGO had somewhat increased potency in
the G
oRGS/PTXi-expressing cells compared with the
G
oPTXi-expressing cells and the wild-type C6µ cells.
Morphine was more potent in the cells expressing
G
oRGS/PTXi. When cells were treated with PTX overnight,
the effect of DAMGO and morphine was completely lost in the C6µ cells
expressing wild-type G
proteins; however, inhibition of cAMP
accumulation was retained in the cells expressing G
oPTXi.
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Table II
Inhibition of cAMP accumulation in wild-type C6µ,
C6µ-G oPTXi, and C6µ-G oRGS/PTXi cells by
DAMGO and morphine without PTX treatment
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Stimulation of p44/42 MAPK
Phosphorylation--
To determine whether MAPK regulation was also
under the control of endogenous RGS proteins, stimulation of ERK
phosphorylation by DAMGO (100 nM) was measured in
PTX-treated C6µ cells expressing either G
oPTXi or
G
oRGS/PTXi at intervals from 0 to 20 min (Fig. 4, A and B).
Stimulation of phosphorylation by 100 nM DAMGO followed a
similar time course in both cell clones, but the percent stimulation over basal levels was consistently higher in the
C6µ-G
oRGS/PTXi cells. To determine whether the
increased phosphorylation by 100 nM DAMGO in the
G
oRGS/PTXi-expressing cells was due to a change in
potency or a change in maximal effect, a concentration-effect curve for DAMGO was determined at the 5-min time point. DAMGO stimulated the phosphorylation of p44/42 MAPK (Fig.
5, A and B) with an
18-fold greater potency (p < 0.05) in
C6µ-G
oRGS/PTXi cells (clone C1, EC50 = 48 ± 11 nM) than in C6µ-G
oPTXi cells (clone M1, EC50 = 839 ± 187 nM). Although
the basal phosphorylation level was consistently lower in
C6µ-G
oRGS/PTXi cells, the maximal percent increase
over basal levels was similar (C6µ-G
oPTXi cells, 530 ± 115% stimulation; and C6µ-G
oRGS/PTXi
cells, 590 ± 104% stimulation). Thus, the enhanced ERK
phosphorylation seen in the initial time course study was due to a
change in the EC50 for DAMGO and not in the maximal
response. In contrast, even at 10 µM, morphine stimulated
a low level of p44/42 MAPK phosphorylation in
C6µ-G
oPTXi cells, representing just 14% of the
stimulation seen with DAMGO. However, this concentration of morphine
was significantly (p < 0.05) more efficacious in
C6µ-G
oRGS/PTXi cells (527 ± 174% stimulation
over basal levels) than in C6µ-G
oPTXi cells (68 ± 41% stimulation over basal levels) (Fig. 5C).

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Fig. 4.
Time dependence of DAMGO-stimulated p44/42
MAPK phosphorylation. C6µ-G oPTXi ( ) or
C6µ-G oRGS/PTXi ( ) cells were treated overnight with
100 ng/ml PTX. The medium was replaced with serum-free medium for
2 h. The assay was started by the addition of 100 nM
DAMGO and stopped after 0-20 min, and Western blotting was performed
as described under "Experimental Procedures." Shown in A
is a representative blot of phosphorylated and total MAPKs. Bands were
quantitated as sum intensity (pixels) and plotted as a percentage of
basal levels (without ligand). Shown in B are the combined
data from three to four assays.
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Fig. 5.
Dose dependence of agonist-stimulated p44/42
MAPK phosphorylation. C6µ-G oPTXi ( ) or
C6µ-G oRGS/PTXi ( ) cells were treated overnight with
100 ng/ml PTX. The medium was replaced with serum-free medium for
2 h. The assay started by the addition of 0-10 µM
DAMGO (B) or 10 µM morphine (C) and
stopped after 5 min, and Western blotting was performed as described
under "Experimental Procedures." Shown in A is a
representative blot of phosphorylated and total MAPKs. Bands were
quantified as sum intensity (pixels) and plotted as a percentage of
basal levels (without ligand). The basal sum intensity was lower in
C6µ-G oPTXi cells (157,000 ± 17,000 pixels) than
in C6µ-G oRGS/PTXi cells (67,000 ± 12,000 pixels). Shown in B are the combined data from three to four
assays.
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Increases in Intracellular Calcium--
To obtain a measurable
increase in the intracellular calcium signal in response to opioid
agonists, cells were grown for 24 h in the presence of 5 µM forskolin (24, 25). In these cells, DAMGO stimulated
[Ca2+]i in PTX-treated
C6µ-G
oRGS/PTXi (EC50 = 80 ± 34 nM) and C6µ-G
oPTXi (EC50 = 89 ± 17 nM) cells. There was no significant difference in the EC50 values or maximal stimulation (Fig.
6A). Compared with DAMGO, the
relative maximal effect of morphine was somewhat higher in
C6µ-G
oRGS/PTXi cells (71.9 ± 9.1%) than in C6µ-G
oPTXi cells (35.3 ± 14.5%) (Fig.
6B), but this did not reach statistical significance
(p = 0.11, Wilcoxon matched pairs). At 1 µM DAMGO, the induced rise in
[Ca2+]i was from intracellular stores because the
increase was not significantly different (p > 0.05, Wilcoxon matched pairs) in the presence of extracellular
Ca2+ (
340/380 nm ratios of 0.11 ± 0.02 in
C6µ-G
oRGS/PTXi cells and 0.06 ± 0.01 in
C6µ-G
oPTXi cells) or in its absence (
340/380 ratios
of 0.07 ± 0.01 in C6µ-G
oRGS/PTXi cells and
0.07 ± 0.01 in C6µ-G
oPTXi cells).

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Fig. 6.
Agonist-stimulated increase in
[Ca2+]i. [Ca2+]i was
measured in the presence of 0-10 µM DAMGO (A)
or 10 µM morphine (B) in fura-2-loaded whole
cell suspensions from C6µ-G oPTXi ( ) and
C6µ-G oRGS/PTXi ( ) cells treated for 24 h with
forskolin and overnight with PTX as described under "Experimental
Procedures." Morphine stimulation is expressed as a percentage of the
10 µM DAMGO response. Shown are the combined data from
six to seven assays.
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To confirm that the G
oPTXi- and
G
oRGS/PTXi-expressing cells treated overnight with
forskolin still showed different sensitivities to µ-opioid agonist
inhibition of adenylyl cyclase, cells were examined for DAMGO
inhibition of cAMP accumulation. Forskolin treatment increased the
maximal effect and potency of DAMGO in the
G
oPTXi-expressing cells. However, the differential
response between the G
oPTXi- and
G
oRGS/PTXi-expressing cells was retained. The degree of
maximal inhibition of cAMP accumulation was increased in the presence
of the RGS-insensitive G
o mutant (G
oPTXi,
68.5 ± 5.4%; and G
oRGS/PTXi, 83.8 ± 1.2%;
p = 0.05), as was the enhanced potency of DAMGO
(G
oRGS/PTXi, EC50 = 4.24 ± 1.44 nM; and G
oPTXi, EC50 = 157 ± 35 nM; p < 0.05).
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DISCUSSION |
In this study, we have expressed G
o that is
insensitive to RGS protein action in C6 cells expressing a µ-opioid
receptor to demonstrate a role for endogenous RGS proteins in the
control of opioid receptor signaling. Two main conclusions can be drawn from this study. First, endogenous RGS proteins reduce the
effectiveness of G
o signaling to adenylyl cyclase and
MAPK pathways, suggesting that endogenous RGS protein action
has substantial regulatory effects on agonist potency and maximal
response. Second, because only minor differences were seen between the
RGS-insensitive G
o mutant compared with its
RGS-sensitive counterpart in coupling to intracellular calcium release,
but significant differences were measured with the adenylyl cyclase and
MAPK pathways, we conclude that endogenous RGS proteins contribute to
the control of effector selectivity of G
o signaling.
One of the most direct measures of receptor activation is stimulation
of [35S]GTP
S binding seen upon the addition of
agonist. The potency and maximal effect of DAMGO and morphine in
stimulating [35S]GTP
S binding were similar in
membranes from C6µ cells expressing G
oPTXi and
G
oRGS/PTXi or, if anything, were greater for
RGS-sensitive G
o. One would not expect the GAP activity
of RGS proteins to play a role in the binding of the GTP analog to the
G
o subunit because the assay is done in the presence of
a large excess of GDP and the rate-limiting step is the dissociation of
GDP from the G
subunit (26).
In contrast to agonist-stimulated [35S]GTP
S binding,
DAMGO-stimulated GTPase activity in membranes from cells expressing
G
oRGS/PTXi was significantly less than in membranes from
cells expressing G
o/PTXi. This indicates that endogenous
RGS protein GAP activity in C6µ cell membranes is significant, but
this cannot function to stimulate GTP hydrolysis by the
G
oRGS/PTXi mutant. Furthermore, DAMGO-stimulated GTP
hydrolysis in membranes from the G
oPTXi-expressing cells
was markedly enhanced in the presence of added RGS8, whereas no
stimulation by exogenously added RGS8 was observed in membranes from
the G
oRGS/PTXi-expressing cells, confirming the
insensitivity of RGS-insensitive G
o. As a
percentage of the maximal DAMGO stimulation, morphine produced a
significantly smaller increase in GTPase activity in the presence of
GST-RGS8 than in its absence, demonstrating that RGS protein is able to
produce a greater enhancement of steady-state GTPase in the presence of
a full agonist; this could relate to the greater rate of GDP release
caused by the full agonist (26).
The potency and maximal effect of DAMGO and morphine in inhibiting
adenylyl cyclase through activation of the transfected RGS-sensitive
G
oPTXi mutant in PTX-treated cells were poor compared with both wild-type C6µ cells and cells expressing
G
oPTXi before treatment with PTX. C6 cells do not
endogenously express G
o, and G
i2 is the
predominant G
subunit expressed (27). The robust inhibition of
adenylyl cyclase in C6µ cells expressing exogenous G
o
before PTX treatment confirms that endogenous G
i2
couples very efficiently to adenylyl cyclase. In contrast, it is
probable that the transfected G
oPTXi mutant, although
activated efficiently by the µ-opioid receptor, couples poorly to
adenylyl cyclase such that this response can be dramatically improved
by inhibition of RGS activity. In support of this, G
o
does not couple well to adenylyl cyclase in NG108-15 cells or
HEK293 cells compared with G
i2 (28, 29), although
it is reported to have a role in this regard in SH-G
-interacting
protein SY5Y cells (30). The inhibition of cAMP accumulation was very
much improved in the G
oRGS/PTXi-expressing cells, giving
a large increase in both the maximal inhibition and agonist potency in
the different clones examined. It was also noticeable that, without PTX
treatment, cells expressing the G
oRGS/PTXi mutant
were more efficiently inhibited by morphine and DAMGO than cells
expressing only G
oPTXi. In a series of experiments using
the opposite approach, expression of RGS4 or G
-interacting
protein in HEK293 cells reduced the level of somatostatin
receptor-induced inhibition of cAMP accumulation (31), demonstrating
the ability of RGS proteins to control the magnitude of inhibitory G
protein signaling. Our data extend this by demonstrating a role for
endogenous RGS proteins in this effect.
This effect of the RGS-insensitive G
o mutant on
agonist-mediated inhibition of adenylyl cyclase was particularly marked
for the partial µ-agonist morphine, which became almost (90%) as
efficacious as the full agonist DAMGO in the
C6µ-G
oRGS/PTXi cells. This indicates that RGS proteins
may be more effective when the receptor/G protein/effector system is
signaling at submaximal levels. In agreement with this, several authors
have shown that, at high agonist concentrations, RGS proteins are less
effective (32, 33). In addition, RGS5-mediated reduction in
intracellular calcium release by the angiotensin type 1a receptor
(Gq-linked) is less effective when receptors are expressed
at high levels (34). Because the relative efficacy of an agonist is
tissue-specific, it may be possible that differential expression of RGS
proteins in tissues is one factor in determining agonist efficacy.
µ-Opioids strongly activate phosphorylation of p44/42 ERK in
C6µ cells (3). This effect was retained in C6µ cells expressing either G
oRGS/PTXi or G
oPTXi after PTX
treatment. However, similar to the µ-agonist effect on the cAMP
system, µ-agonist activation of the MAPK pathway was increased in
C6µ cells expressing the G
oRGS/PTXi mutant compared
with the RGS-sensitive G
oPTXi mutant. Thus, when
the µ-receptor was coupled through G
oRGS/PTXi, a
>10-fold increase in DAMGO potency and an increase in the maximal
effect of morphine were observed compared with coupling through
G
oPTXi. These results are consistent with
findings that 5-hydroxytryptamine 1
receptor activation of ERK
(p44/42 MAPK) is reduced by overexpression of RGS4 in
neuroblastoma cells (32), as is stimulation of MAPK through
interleukin-8 (35) and dopamine D2 (33) receptor
activation. In contrast to the effects on adenylyl cyclase and MAPK,
the µ-opioid-mediated increase in [Ca2+]i
showed a much smaller increase in effect in the
G
oRGS/PTXi-expressing cells such that the
difference between these and the
G
oPTXi-expressing cells did not quite reach
significance. This differential effect causes a shift in the most
potent response to the µ-opioid agonist DAMGO. Thus, in the presence
of endogenous RGS activity, the rise in intracellular calcium is the
most potent response (EC50 = 89 nM), and the
inhibition of adenylyl cyclase is the weakest response (EC50 = 400 nM); but in the absence of
endogenous RGS activity, the order is reversed, and the inhibition of
adenylyl cyclase is the most potent response (EC50 = 12 nM), whereas the rise in intracellular calcium is the
weakest response (EC50 = 80 nM).
The present data show that endogenous RGS proteins may differentially
affect signaling by a single G protein depending on the effector
pathway to which the G protein couples. Several mechanisms could
account for this specificity. One is that the GAP activity of
endogenous RGS proteins controls signaling by a kinetic scaffolding mechanism (18). The kinetic scaffolding model predicts that RGS action
reduces depletion of local G
-GTP levels and so permits rapid
recycling of G protein and rapid recoupling of the receptor and
maintains local G protein activation. The adenylyl cyclase and MAPK
pathways are poorly signaled to in the presence of RGS activity; but
when this activity is blocked, as in cells expressing the
RGS-insensitive G
o mutant, then signaling can occur
because spatial control is lost, allowing spillover of G
-GTP and
G
subunits to more distant effectors. In contrast, coupling to
intracellular calcium stores is more similar in cells expressing
G
oPTXi and G
oRGS/PTXi. Thus, for the
kinetic scaffolding model to account for this effect, the G proteins
involved in coupling to this pathway must be organized closely with
receptor and effector such that they show a reduced
RGS-dependent effect.
The differential effect of RGS on the three pathways examined is
consistent with this theory, but does not provide direct proof. Other
mechanisms may explain the findings. The increased opioid effect
at adenylyl cyclase and MAPK may simply be due to an increased lifetime
of G
-GTP in the absence of RGS protein GAP activity; but if so, then
the question arises as to why the intracellular calcium signal is not
enhanced to a similar extent. There may be differential
localization or compartmentalization of effectors within the
cell (36) such that the intracellular calcium signaling complex is
protected from RGS action. One possibility is that RGS proteins provide
a protein scaffold that allows signaling to the intracellular calcium
pathway; but in the absence of this restraining scaffold, other
pathways become available to the G
-GTP or G
subunits.
Certainly, C6 cells contain message for a variety of RGS proteins
(RGS2, RGS3, RGS8, RGS10, RGS12, and RGS14) (37), several of which have
regions outside of the RGS box that could be involved in
protein-protein interactions. Alternatively, RGS proteins may not have
a controlling function in modulation of the intracellular calcium
signal; the rate of GTP hydrolysis and the lifetime of G
-GTP
may not be the rate-limiting step in this signaling pathway. The
increase in [Ca2+]i response is transient in
nature. Opioid stimulation of intracellular concentration is thought to
be mediated through G
subunit activation of phospholipase
C
1 to break down phosphatidylinositol 4,5-bisphosphate and to provide inositol 1,4,5-trisphosphate, which binds to the inositol 1,4,5-triphosphate receptor on the intracellular store, causing an increase in
[Ca2+]i (23). The nature of the intracellular
calcium signal may be controlled by other factors such as inositol
1,4,5-triphosphate receptor desensitization and the fullness of the
Ca2+ store; indeed, there may not be a direct temporal
relationship between inositol 1,4,5-triphosphate and Ca2+
signaling (38).
In summary, we have shown, in a transfected C6 cell line, that RGS
proteins differentially regulate µ-opioid receptor-mediated signaling
to different effectors through G
o, consistent with a
kinetic scaffolding mechanism. Coupling to adenylyl cyclase and the
MAPK pathway appears to be efficiently limited by endogenous RGS
proteins, whereas coupling to intracellular calcium stores is less
susceptible to RGS protein action. Because the potency and maximal
effect of agonist are altered, it is possible that differential
expression of RGS proteins in tissues plays a role in tissue-specific
differences in agonist selectivity and efficiency. Finally, because
cAMP (39) and MAPK (40, 41) have been implicated in contributing to the
tolerance associated with long-term opioid administration, the effect
of endogenous RGS proteins on these cellular adaptations merits further investigation.