(Received for publication, December 4, 1995; and in revised form, January 23, 1996)
From the
The phospholipase C-1 (PLC-
1) signaling pathway was
reconstituted by addition of purified PLC to phospholipid vesicles that
contained purified recombinant m1 muscarinic cholinergic receptor,
G
, and 2-4 mol %
[
H]phosphatidylinositol 4,5-bisphosphate. In this
system, the muscarinic agonist carbachol stimulated steady-state PLC
activity up to 90-fold in the presence of GTP. Both GTP and agonist
were required for PLC activation, which was observed at physiological
levels of Ca
(10-100 nM). PLC-
1
is also a GTPase-activating protein for G
. It accelerated
steady-state GTPase activity up to 60-fold in the presence of
carbachol, which alone stimulated activity 6-10-fold, and
increased the rate of hydrolysis of G
-bound GTP by at least
100-fold. Despite this rapid hydrolysis of G
-bound GTP, the
receptor maintained >10% of the total G
in the active
GTP-bound form by catalyzing GTP binding at a rate of at least
20-25 min
,
10-fold faster than previously
described. These and other kinetic data indicate that the receptor and
PLC-
1 coordinately regulate the amplitude of the PLC signal and
the rates of signal initiation and termination. They also suggest a
mechanism in which the receptor, G
, and PLC form a
three-protein complex in the presence of agonist and GTP (stable over
multiple GTPase cycles) that is responsible for PLC signaling.
Heterotrimeric G proteins transmit signals from cell-surface
receptors to intracellular effectors (such as PLC- (
)enzymes, adenylyl cyclase, or ion channels) by transiting
a controlled cycle of GTP binding and hydrolysis. GTP binding to the
subunit activates G proteins and allows them, in turn, to
activate effectors. They are deactivated when they hydrolyze bound GTP
to GDP. Receptors increase the rate at which G proteins bind GTP.
Agonist-bound receptor thus determines the rate of G protein
activation, and hydrolysis of bound GTP determines the rate of
deactivation. The relative rates of activation and deactivation
determine the steady-state concentration of the activated G protein
and, therefore, the amplitude of the signal transmitted through the
receptor-G protein-effector pathway (see (1, 2, 3) for reviews).
Until recently,
the rate of GTP hydrolysis by G protein subunits was not known to
be regulated directly, although there was evidence that GTP hydrolysis
in cells may occur faster than had been described for purified G
proteins(1) . A model for such regulation was the small
monomeric GTP-binding proteins, such as p21
. For
these proteins, GTP hydrolysis is negligible in the absence of
GTPase-activating proteins (GAPs), which accelerate hydrolysis
dramatically. For example, the rate of hydrolysis of GTP bound to
p21
is increased by Ras GAP
10
-fold (from 1.2
10
s
to 19 s
)(4) .
Numerous GAPs specific for one or more small GTP-binding proteins have
now been identified. They regulate signal duration and/or amplitude,
but apparently do not directly transmit signal(5) .
Only two
GAPs for heterotrimeric G proteins have been described so far, one each
for G and G
. The G
GAP is
PLC-
, the principal effector regulated by
G
(6) . When purified G
and m1AChR
were co-reconstituted into phospholipid vesicles, addition of purified
recombinant PLC-
1 stimulated steady-state GTPase activity
>20-fold when the muscarinic agonist carbachol was added to
stimulate GTP binding(6) . Steady-state kinetic arguments and
preliminary measurements of the rate of hydrolysis of bound GTP in this
system suggested that PLC-
1 stimulates hydrolysis of
G
-bound GTP >50-fold, from
0.8 min
to at least 40 min
(6) .
The G GAP activity described for PLC-
1 is reciprocally specific
for the PLC-
family and the G
family. PLC-
1,
PLC-
2, and PLC-
3 are all G
GAPs, (
)and
preliminary data suggest that the PLC-
encoded by the Drosophila NorpA gene is a GAP for Drosophila visual
G
(7) . In contrast, neither PLC-
(not G
protein-regulated) nor adenylyl cyclase (regulated by G
,
but not by G
) has G
GAP activity.
In addition, while PLC-
1 is a GAP for the G
family members G
, G
, and G
,
it is not a GAP for G
, G
, G
, or
G
(6, 7, 8) .
The
G GAP also involves the G
effector protein,
cyclic GMP phosphodiesterase. In this case, however, both the
subunit of the phosphodiesterase and a second, unidentified membrane
protein are required to stimulate GTP hydrolysis by
G
(9, 10, 11, 12, 13) .
Phosphodiesterase
and the membrane-bound factor together can
increase the hydrolysis rate constant of G
from 0.05
s
to >1 s
, approaching the
physiological deactivation rate of photoreceptor
signaling(13) .
Not all G protein-regulated effectors are
GAPs. We have been unable to detect G GAP activity in
several preparations of adenylyl cyclase(14) . However, the
existence of other G protein GAPs is strongly predicted because the
rates of deactivation of many G protein-mediated signaling processes
are significantly faster than the rates of hydrolysis of bound GTP by
isolated G proteins (reviewed in (1) ).
The finding that
PLC-1 is a GAP for G
raises the question of how a
signal can be generated when an effector that is a GAP terminates its
own stimulus. Indeed, the discovery of PLC-
1 as a G
GAP was sparked by our failure to detect GTP-supported
stimulation of PLC activity by muscarinic agonists when m1AChR,
G
, and PLC-
1 were co-reconstituted into phospholipid
vesicles(15) . Although our measurements of GTP binding, GTP
hydrolysis, and accumulation of activated G
-bound GTP
suggested that added PLC-
1 should be substantially activated, the
G
GAP activity of PLC-
1 accelerated GTP hydrolysis,
and no phospholipase stimulation was observed. More recently, Nakamura et al.(8) succeeded in demonstrating stimulation of
PLC activity by carbachol plus GTP in a similarly reconstituted system,
suggesting that the three proteins are sufficient to generate signal in
response to agonist in the presence of GTP. However, activation was
modest (2.2-fold), leaving open the question of how receptor-promoted
GTP binding can keep up with GAP-stimulated hydrolysis.
For this
study, we have modified our previous reconstitution protocol (6, 15) to develop a system that generates a large
IP signal in response to GTP and carbachol
(50-100-fold stimulation). This system provides an experimental
model in which to study PLC-
1 activity in terms of the regulation
of the G
GTPase cycle. The data reported here reconcile the
rates of receptor-stimulated GTP binding by G
and of
GAP-stimulated GTP hydrolysis with the steady-state concentration of
G
-bound GTP and activated PLC-
1.
The extraction
supernatant (100 ml) was diluted 5-fold into buffer G1 and mixed with
12 ml of NTA-Ni-agarose. The mixture was stirred for
1 h, and the NTA-Ni
-agarose was collected in a
chromatography column. The resin was washed with buffer G1, buffer G1
plus 0.5 M NaCl, buffer G1 plus 0.5 M NaCl and 10
mM imidazole, buffer G1 plus 1 M NaCl, and buffer G1.
Each wash step was deemed complete when the absorbance base line (280
nm) remained flat for at least 24 ml. The column was then moved to room
temperature and washed with buffer G2 (100 ml) to remove the endogenous
Sf9
-like protein and with buffer G3 (80 ml) to remove
residual GTP
S and Lubrol. G
was separated from
the G
His
G
His
dimer at room temperature in buffer G4 (100 ml), and the eluate
was collected at 0 °C. The Al
/F
concentration in the eluate was reduced at least 1000-fold by
repeated concentration (Amicon PM-10 membrane) and dilution in buffer
G5. G
was prepared by mixing G
and
G
in a 1:2 molar ratio. The
concentration of active G
was determined according to
bound GDP(28) , and the G
concentration was based on total protein(29) .
The
estimated purity of G was >95%, and that of
G
was >85% (Fig. 1). No Sf9
cell G
was detectable in the G
preparation by silver staining or by immunoblotting (P-960 antibody) (30) . We were also unable to detect contamination with Sf9
cell G
by comparing the bands detected on a single
immunoblot that was sequentially probed with an antibody that
recognizes Sf9 cell G
(Z811) (31) and with a
second antibody that recognizes only recombinant G
.
The yield of G
was 130-150 µg/liter of Sf9
cells; the yield of G
was 340
µg/liter of Sf9 cells.
Figure 1:
SDS-polyacrylamide gel electrophoresis
of purified proteins. Each protein, purified as described under
``Experimental Procedures,'' was electrophoresed and
silver-stained. Two samples of each protein (nanograms applied) are
shown, one of which is 10-fold overloaded to highlight impurities. Arrowheads indicate the major protein bands. The lower
arrowhead for the G gel indicates G
, which stains
poorly but is visible on the original gel.
Figure 2:
Reconstitution of G-mediated
activation of PLC-
1 by m1AChR. m1AChR and G
were
co-reconstituted with [
H]PIP
as
described under ``Experimental Procedures.'' The activity of
added PLC-
1 was measured in the presence of guanine nucleotide
and/or muscarinic ligands. A, GTP-dependent PLC activity; B, GTP
S-stimulated PLC activity. Conditions were as
follows: no addition(-), 10 µM GTP (GTP), 1
mM carbachol (Cch), carbachol plus 10 µM atropine (C+A), and 100 nM GTP
S (
S). All samples contained 10 nM free
Ca
, 2.4 nM G
, 0.33 nM m1AChR, 1 nM PLC-
1, and 0.56 µM accessible PIP
. A and B show data
(mean ± S.D.) from the same experiment on different
scales.
Regulation of PLC-1 by
reconstituted m1AChR and G
displayed the dependence on the
concentrations of carbachol, GTP, Ca
,
PIP
, and PLC-
1 that is typical of intact biological
systems (Fig. 3Fig. 4Fig. 5). Carbachol plus GTP
markedly stimulated the activity of PIP
even at very low
Ca
concentrations (Fig. 3). Stimulation was
100-fold at 10 nM free Ca
, and this
concentration was used in all other experiments shown here. Stimulation
in the physiological range of 10-100 nM Ca
was appreciable. Ca
also stimulated PLC
activity directly, and stimulation by Ca
and that by
G
were strongly synergistic. This overall pattern of
regulation is observed for PLC-
in native membranes (see (38) for a review or (39) for an example). We have not
explored the interaction of muscarinic stimulation and Ca
further, however, because agonist-stimulated PLC activities at
micromolar Ca
concentrations were very high and
became nonlinear after a few seconds.
Figure 3:
Effect of Ca on
G
-stimulated PLC-
1 activity. m1AChR and G
were co-reconstituted with
[
H]PIP
, and the activity of added
PLC-
1 was measured at various free Ca
concentrations. The data plotted are the initial rates determined
from time courses of PLC activity conducted at each Ca
concentration in the presence of 10 µM GTP and
either 1 mM carbachol (filled circles) or 10
µM atropine (empty circles). Reactions contained
0.35 nM m1AChR, 3.0 nM G
, 0.33 µM accessible PIP
, and 1.0 nM PLC-
1.
Figure 4:
Dependence of carbachol-stimulated PLC
activity on concentration of PIP. m1AChR and G
were co-reconstituted in vesicles that contained various
concentrations of PIP
. PLC activity was measured in the
presence of 10 nM free Ca
, 10 µM GTP, 1 nM PLC-
1, and either 1 mM carbachol
or 10 µM atropine, Each data point represents the initial
rate of carbachol-stimulated PIP
hydrolysis derived from a
time course performed with a different batch of vesicles. The abscissa is the concentration of PIP
accessible to
PLC-
1, as described under ``Experimental Procedures.''
The concentration of G
in the 12 batches of vesicles varied
between 0.6 and 0.8 nM. Although other experiments in this
study were frequently performed at molar concentrations of PIP
higher than those shown here, the mole fraction of PIP
in the vesicles was maintained in the range shown here
(1.1-5.5 mol %). The discrepancy reflects experimental scale. The
data come from multiple small-scale reconstitutions wherein
lipid/protein ratios were decreased by half to conserve materials and
to maximize yield during reconstitution.
Figure 5:
GTPase and PLC activities at increasing
concentrations of PLC-1. m1AChR-G
vesicles were mixed
with increasing concentrations of PLC-
1 and assayed for both
GTPase (filled circles) and PLC activity (empty
circles) under identical conditions. Assays contained 0.27 nM m1AChR, 1.7 nM G
, 0.44 µM accessible PIP
, 10 µM GTP, 10 nM Ca
, and either 1 mM carbachol or 10
µM atropine. Data show increases in activity caused by
carbachol. Assay times were adjusted to ensure linear
reactions.
Carbachol-stimulated PLC
activity increased linearly with the concentration of PIP in the vesicles up to
6 mol % of total phospholipid (Fig. 4). The routine concentration of PIP
was held
at
3 mol %, which is not unreasonable with respect to the
concentrations of PIP
in the inner leaflet of animal plasma
membranes(40, 41) . Much higher concentrations of
PIP
decreased recovery of m1AChR during reconstitution and
may inhibit m1AChR-G
coupling (as measured by
agonist-stimulated GTP
S binding) (data not shown). PLC activity
also increased linearly with increasing amounts of total vesicles added
to the assays (data not shown). Added PLC-
1 presumably has access
only to PIP
in the outer leaflet of the vesicle bilayer,
and prolonged incubation of the vesicles with 100 nM GTP
S, 1 mM carbachol, and 1 µM Ca
resulted in hydrolysis of only about half of
the PIP
. We have therefore defined PLC-accessible PIP
as that which can be hydrolyzed in the presence of these
activators after 40 min at 30 °C. PLC assays were linear with time
so long as <15% of the accessible PIP
was hydrolyzed.
Note that carbachol-stimulated GTPase activity, measured in either
the presence or absence of PLC-1, was not altered by the presence
of Ca
or by the concentration of PIP
in
the vesicles (data not shown).
In addition to its activity as a
phospholipase, PLC-1 stimulates the GTPase activity of
G
(6) . Maximal stimulation at steady state can
exceed 60-fold in the reconstituted vesicles when carbachol is added to
promote GDP/GTP exchange (35-fold in Fig. 5). The G
GAP activity of PLC-
1 is saturable, with an EC
of 1-2 nM(6) (1.1 nM in Fig. 5). Carbachol-stimulated PLC activity also reached a
plateau in this same range of PLC-
1 concentrations, with
half-maximal activity at 0.5 nM PLC-
1. This behavior was
not caused by substrate depletion at high enzyme concentrations; <7%
of the accessible PIP
was hydrolyzed in the assays shown in Fig. 5. The plateau in activity at high nanomolar concentrations
of PLC-
1 apparently reflects the limiting steady-state
concentration of GTP-activated G
present during the assay
(data not shown).
Figure 6:
Dependence of steady-state GTPase and PLC
activities on concentrations of GTP and carbachol. GTPase and PLC
activities were measured under identical conditions using the same
batch of vesicles for all assays shown in each panel (except filled
squares). The concentration of PLC-1 was 10 nM in
the PLC assays (filled triangles) and, where added, in the
GTPase assays (filled squares and circles). GTPase
was also assayed without added PLC (empty circles).
Carbachol-stimulated activities are shown. A, activities were
assayed in the presence of 10 µM GTP with 0.19 nM m1AChR, 2.0 nM G
, and 0.52 µM accessible PIP
. Maximum activities were 93.6 ±
3.2 fmol of P
/min without PLC, 976 ± 39 fmol of
P
/min with PLC, and 8.58 ± 0.37 pmol of
IP
/min. B, activities were assayed in the presence
of 1 mM carbachol with 0.37 nM m1AChR, 2.3 nM G
, and 0.45 µM accessible
PIP
. Maximum activities were 40.5 ± 5.5 fmol of
P
/min without PLC, 2610 ± 170 fmol of
P
/min with PLC, and 4.50 ± 0.31 pmol of
IP
/min. Filled squares and circles in B are from separate experiments.
As shown in Fig. 6B, addition of PLC-1 also increased the K
for the agonist-stimulated GTPase reaction from
0.25 µM to 5.1 µM GTP. Such an increase is
predicted by classical enzyme kinetic theory for a GAP that increases
the rate of hydrolysis of bound substrate without altering affinity for
substrate. (
)The GTP concentration curve for the
agonist-stimulated PLC reaction fell about midway between the two
GTPase curves in several independent experiments (EC
= 1.0 µM in Fig. 6B) (data
not shown). This behavior is also qualitatively consistent with a
simple analysis of the GTPase reaction, but why the EC
is
lower than the K
for GAP-stimulated GTP hydrolysis
is not clear.
Figure 7:
Kinetics of PLC-1 activation and
deactivation. Vesicles that contained m1AChR, G
, and
[
H]PIP
were incubated initially in
buffer that contained 1 nM PLC-
1, 10 nM free
Ca
, and 1 mM carbachol (A), 10
µM GTP (B), or 0.3 mM carbachol plus 10
µM GTP (C). At the times shown by the arrows, 10 µM GTP (A), 1 mM carbachol (Cch; B), or 50 µM atropine (C) was added, and the assay was allowed to
continue (filled circles). The linear extrapolations of the
initial rates, shown in B and C, are validated by
data from at least three other experiments in which reactions were
continued both with and without the second addition. The dotted
line in B is an extrapolation of the final stimulated
rate. Key concentrations were 0.3 (A), 0.6 (B), and
0.2 (C) nM m1AChR; 1.6 (A), 1.2 (B), and 3.3 (C) nM G
; and 0.35 (A), 0.58 (B), and 0.42 (C) µM accessible PIP
.
The deactivation of PLC-1 upon addition of a
muscarinic antagonist is also quite rapid. Carbachol is a low affinity
agonist (K
0.1 mM) (25) and
dissociates rapidly, such that addition of excess antagonist (50
µM atropine) should block receptors in <1 ms. As shown
in Fig. 7C, addition of atropine during
agonist-stimulated steady-state PIP
hydrolysis inhibited
PLC activity with no perceptible lag. Based on the temporal resolution
of the earliest data points, we estimate from multiple experiments that
the deactivation rate constant in such an experiment is >25
min
. Assuming that deactivation requires hydrolysis
of G
-bound GTP, 25 min
is thus also a
lower limit on the GAP-stimulated k
for
G
. This rate is 30-fold higher than the GTPase k
displayed by isolated
G
(15) .
Figure 8:
Carbachol-stimulated binding of GTPS (A) and GTP (B) to m1AChR-G
vesicles.
Reactions were initiated by adding vesicles to prewarmed reaction
buffers that contained either 1 mM carbachol (filled
circles) or 10 µM atropine (empty circles)
and either 100 nM [
S]GTP
S (A) or 500 nM [
-
P]GTP (B). The dashed lines are the differences between
binding measured in the presence of carbachol and atropine. Reactions
shown in A and B contained 0.15 or 0.10 nM m1AChR and 2.1 or 0.70 nM G
,
respectively.
Since the
relatively slow nucleotide binding reactions described above might
reflect an initial delay, we measured agonist-stimulated GDP/GDP
exchange under equilibrium conditions in which there was no hydrolysis
and slow association of proteins had had time to occur. For this
experiment, vesicles were first incubated for 10 min in the presence of
both agonist and unlabeled GDP. The rate of GDP binding was then
monitored at equilibrium by addition of a tracer amount of
[-
P]GDP. Subsequently, excess unlabeled GDP
was added to measure the rate of dissociation of labeled nucleotide
(still at equilibrium). As shown in Fig. 9, both the binding and
dissociation of GDP were very rapid under these conditions. Although
the association and dissociation time courses were best fit by biphasic
functions, the major kinetic components for both the binding and
release of GDP displayed rate constants of
20
min
. These rates are probably underestimates because
they are calculated according to only a few data points at the limits
of resolution of the manual binding assay. Because the rate of binding
of GDP is well below the rate predicted by diffusion-controlled
association and because the rate of GDP binding equals the rate of GDP
release, dissociation of bound GDP is probably rate-limiting for
association of a second GDP molecule. According to these data, the
reconstituted m1AChR-G
vesicle system can achieve
agonist-stimulated guanine nucleotide exchange rates commensurate with
the high steady-state GTPase rates observed in the presence of agonist
and PLC-
1. Data from preliminary experiments similar to those
shown in Fig. 9indicate that PLC-
1 does not alter the
kinetics of agonist-stimulated GDP/GDP exchange.
Figure 9:
Receptor-stimulated equilibrium GDP/GDP
exchange. Binding and release of GDP were measured at a saturating
concentration of free nucleotide. A, a suspension of
m1AChR-G vesicles was incubated for 10 min at 30 °C in
1.8 ml of binding assay buffer that contained 500 nM unlabeled
GDP and 0.3 mM carbachol. At time 0, 5.4
10
cpm [
-
P]GDP (36 µl) was added,
and 50-µl aliquots were withdrawn at the indicated times for
measurement of bound radiolabeled GDP. These data were best fit by the
sum of two exponential binding functions, one with a rate constant of
22 ± 11 min
that accounted for 44% of the
total (31 fmol) and a second with a rate constant of 2.1 ± 0.5
min
that accounted for the remainder. B, at
12.25 min, concentrated unlabeled GDP (8.5 µl) was added to the
mixture, and bound radiolabeled GDP was measured at the indicated times
as described for A. GDP dissociation was also biphasic; 67% of
the total (46 fmol) dissociated rapidly with a rate constant of 20
± 7 min
, and the remainder dissociated more
slowly with a rate constant of 1.4 ± 0.3 min
.
The data shown in A and B were derived from the same
binding assay mixture; each 50-µl aliquot contained 0.35 nM (17.5 fmol) m1AChR.
To correlate the
high equilibrium nucleotide exchange rates shown in Fig. 9with
the slow initial onset of agonist-stimulated GTP binding (Fig. 8) and PLC activation (Fig. 7B), we
measured the rate of formation of the presumed complex of agonist,
m1AChR, and G that accounts for the faster rates. Vesicles
were first incubated with carbachol for increasing times, and GTP
binding was then initiated by simultaneously adding both
[
-
P]GTP and atropine. The reaction was then
allowed to proceed for 60 s to selectively monitor the rapid component
of GTP binding. As shown in Fig. 10, initial exposure of the
vesicles to carbachol created a species that bound GTP during the
subsequent 60-s incubation. Formation of this putative
carbachol-m1AChR-G
complex occurred with a rate constant of
1.6 min
, somewhat slower than the rate of
stimulation of PLC by added carbachol (Fig. 7B) and
faster than the initial nucleotide exchange rate measured by exposing
m1AChR-G
vesicles to agonist (Fig. 8). The
relationship of these rates is discussed below.
Figure 10:
Formation of agonist-m1AChR-G
complex that rapidly binds GTP. A, m1AChR-G
vesicles were pre-equilibrated for 5 min in binding assay buffer
at 30 °C and then exposed to 0.3 mM carbachol for the
times shown. At each time point, 300 nM [
-
P]GTP and 50 µM atropine were added, and the incubation was continued for 60 s
before quenching for measurement of bound nucleotide. The zero time
point was determined in vesicles that were equilibrated, exposed to
atropine and [
-
P]GTP for 60 s, and then
quenched before carbachol was added. The concentrations of m1AChR and
G
were 0.2 and 2.2 nM, respectively. The
experimental background (no added vesicles) was subtracted from all
data.
We have not studied
the stability of the m1AChR-G complex in detail, but
initial evidence suggests that it is not stable over long periods of
time. Although
50% of the total G
was able to exchange
GDP rapidly in experiments of the sort shown in Fig. 9, only
10% of the total G
could be trapped as a rapidly
binding species in experiments of the sort shown in Fig. 10. For
the complex that does remain stable in this experiment, however, the
observed decay rate after addition of antagonist was only
1.3
min
(data not shown), suggesting that the average
lifetime of the complex is equivalent to >30 rounds of the GTPase
cycle. The lifetime of the complex could of course be considerably
longer in the continued presence of agonist, and we do not know the
molecular event(s) that initiates its decay.
In the G-mediated regulation of PLC-
,
receptor-promoted activation of G
is opposed by the GAP
activity of the PLC-
effector. Although this acceleration of GTP
hydrolysis allows rapid termination of signaling upon removal of
agonist, accelerated deactivation must be balanced by rapid catalysis
of GDP/GTP exchange by receptor in order to generate a substantial
signal. In general, a G protein must transit the GTPase cycle with
markedly different kinetics if its effector protein is or is not a GAP.
To address the mechanism of G
-mediated signaling, we
developed a purified and reconstituted assay system in which the
individual steps of activation and deactivation of G
and
PLC-
1 can be studied in parallel, either immediately upon
activation by agonist or while at steady state.
The reconstituted
system described here duplicates several important aspects of natural
G-mediated signaling: 1) low basal synthesis of IP
in the presence of either agonist or GTP alone, 2) massive
activation (50-100-fold) in the presence of both ligands, and 3)
rapid onset and termination of signaling. The vesicle system also
substantially amplifies its signal, although
``amplification'' is subject to several definitions and is
difficult to evaluate in intact cells. In the m1AChR-G
vesicles, stimulation by agonist results in synthesis of
200-600 molecules of IP
/min/receptor under our
standard assay conditions. Such amplification is comparable to that
observed in preparations of plasma membranes (see (31) and (42) for examples), and PLC activity in the vesicles can be
further increased 3-6-fold by increasing the concentration of
Ca
within the physiological range (Fig. 3) or
by altering phospholipid concentrations (Fig. 4).
The
m1AChR-G vesicles are obviously not an exact duplicate of
any specific plasma membrane, and plasma membranes vary enormously
among different cells. However, the composition and behavior of the
vesicles fall within a well described biological range. The
phospholipid composition of the vesicles mimics the inner monolayer of
the plasma membrane only approximately, but the concentration of
PIP
(2-4 mol %) is reasonable, and the 10 nM Ca
concentration is well within the range
described for the cytoplasm of resting cells. The ratio of m1AChR to
G
, usually 0.1-0.5, is also within the likely
physiological range. We varied the concentration of PLC-
1 broadly
to ask mechanistic questions, and the actual cytoplasmic concentration
of PLC-
near the plasma membrane is unknown. Most important,
100-fold GTP-dependent muscarinic stimulation of PLC activity indicates
that the basic signaling pathway has been efficiently reconstituted.
Thus, the m1AChR-G
vesicles appear to be a valid model
system in which to study the mechanism of G protein signaling when an
effector is a G protein GAP. Since the system is active and well
regulated in the absence of other proteins suspected of involvement in
IP
signaling (phosphatidylinositol exchange protein (43, 44) or actin- and PIP
-binding
proteins(45) ), it should also allow the evaluation of how
these factors modulate receptor-G
-PLC-
signaling.
Rigorous interpretation of results from this system depends on the
assumption that the PLC activity of PLC-1 is strictly a reflection
of the concentration of GTP-activated G
and of the affinity
of the two proteins for each other. This assumption appears to be
valid. The concentration ranges over which GTP
S-bound G
activates PLC-
1 and PLC-
1 stimulates the GTPase
activity of G
are identical, with EC
values of
1-3 nM(6) . This value, an apparent K
for the binding of activated G
to
PLC-
1, is consistent with the data of Fig. 5(although we
have not been able to test the relationship over a wide range of
vesicle concentrations). The similar dependence of GTPase and PLC
activities on agonist concentration (Fig. 6) also argues for
this assumption. Qualitatively, these findings suggest strongly that
the complex of proteins that displays the high GTPase activity is also
responsible for the stimulated PLC activity.
In contrast to our
previous effort to reconstitute GTP-supported stimulation of PLC-1
by m1AChR(15) , this study demonstrates that the combination of
receptor and G
in phospholipid vesicles that contain
PIP
allows both efficient stimulation of PLC and the rapid
initiation and termination of signaling. The most interesting and
informative outcome, however, is how the rate of agonist-stimulated
binding of GTP (or GTP
S) to G
, which is relatively
slow under usual in vitro assay conditions (Fig. 8)(6, 8, 15) , can balance the
rapid hydrolysis of G
-bound GTP to account for muscarinic
activation of PLC-
1 in the presence of GTP.
The data of Fig. 8Fig. 9Fig. 10indicate that initiation of
receptor-stimulated nucleotide exchange by G is indeed
slow, but that a more rapidly exchanging species forms over
2 min.
This species, apparently a complex of receptor and G
,
displays a nucleotide exchange rate of
20-25 min
(Fig. 9) that can balance rapid GTP hydrolysis and thereby
sustain a small but adequate fraction of G
in the active
GTP-bound conformation at steady state (Table 1). This mechanism,
described by the inner cycle in Fig. 11, contrasts with more
widely accepted schemes for the action of G protein-coupled receptors
in which receptor is assumed to dissociate from GTP-activated G protein
during each cycle. In the more traditional scheme (outer cycle in Fig. 11), association of agonist-bound receptor with G
is rate-limiting for the slow activation step ( Fig. 8and Fig. 10A) and is followed by the rapid binding of GTP.
This scheme is essentially that proposed for the G
-mediated
activation of adenylyl cyclase by Levitzki(46) , in which the
initial encounter of receptor and G protein in the plasma membrane was
suggested to be the actual slow step in the action of agonist. The
feature that distinguishes the behavior of m1AChR and G
from that described by Levitzki for G
is that the
complex of receptor and G
, once formed, is sufficiently
stable to remain associated over multiple GTPase cycles (inner cycle in Fig. 11).
Figure 11:
Alternative GTPase cycles for stable or
unstable association of receptor (R) and G protein (G). Both cycles assume saturating agonist, such that R is
always agonist-bound. Activated species are shown with asterisks. In the slow outer cycle, where the effector has no
GAP activity and GGTP has a lifetime
10 s,
association of R with G
GDP is rate-limiting for regeneration of
active G
GTP (see (46) ). In the rapid inner
cycle, proposed for the agonist-stimulated GTPase reaction catalyzed by
G
under the influence of a GAP such as PLC-
1, a
complex of R, G, and effector (E) remains intact throughout
the cycle. In the rapid cycle, 1) GAP-stimulated hydrolysis is fast
under the influence of the GAP activity of E, such that R does not
dissociate; 2) GDP dissociation is fast since R is already associated
with G; and 3) GTP association is very fast, either diffusion-limited (28, 48) or nearly so(50) , such that E does
not dissociate significantly before reactivation of G by GTP. Although
steady-state GTPase activity is high, enough GTP-activated G is
maintained to mediate signal transduction.
The GAP activity of PLC-1 is central to the
stability of the receptor-G
-PLC complex and its ability to
rapidly traverse the inner cycle of Fig. 11. Although m1AChR
might be expected to dissociate from the activated G
-PLC
complex, rapid GAP-stimulated hydrolysis allows the receptor to cause
the dissociation of GDP instead. (Receptors displace GDP more
efficiently than GTP (47) (confirmed by us for several G
proteins(25) .) Since the receptor then maintains the
nucleotide-binding site on G
in the ``open''
conformation, GTP binding is rapid, and another round of activation is
completed before PLC itself would dissociate. We cannot determine
whether GTP binding to the carbachol-m1AChR-G
complex is
actually diffusion-controlled (28, 48) because GDP
dissociation rather than GTP binding appears to be rate-limiting (Fig. 9, compare A and B). Regardless, binding
is fast.
Two sets of data support this general model. First,
although binding of either GTP or GTPS is relatively slow when
initiated by addition of agonist (Fig. 8), the rate is
10
times faster after a few minutes of incubation with agonist ( Fig. 9and Fig. 10). A similar but smaller difference was
seen in the PLC reaction, which was stimulated only after a substantial
delay upon addition of agonist, but which was stimulated at the
earliest observable time point when GTP was added after preincubation
with agonist. Thus, an initial slow accumulation of a coupled
agonist-receptor-G
species is followed by the relatively
faster binding of GTP, and the complex is then sustained over multiple
GTP binding and hydrolytic events. Second, the initial rate of
nucleotide binding upon addition of agonist is simply too slow to
account for the steady-state turnover of the GTPase reaction. Typical
GTPase turnover numbers in this system are usually
10
min
in the presence of agonist and PLC-
1 (20
min
has been observed). This is at least 5-fold
faster than the observed initial rates of nucleotide binding when the
reaction is initiated with agonist. The more rapid nucleotide exchange
rate during steady-state hydrolysis is, however, congruent with the
binding reactions described in Fig. 9.
We were able to
estimate the rate of GAP-stimulated hydrolysis of G-bound
GTP from steady-state nucleotide binding data (Table 1) using the
rate of receptor-catalyzed GTP binding during steady-state GTP
hydrolysis. This calculation is constrained by experimental limitations
on the data, but all of the data point to a catalytic rate constant of
100-200 min
(1.5-3.0
s
), an increase of 150-300-fold in hydrolysis
rate over that observed with G
alone (0.7
min
)(6) . First, this is an enormous
increase in k
by an allosteric effector,
although smaller than the effects of GAPs for small monomeric
GTP-binding proteins such as p21
(4) . Second,
since this rate constant remains 4-12-fold higher than the
exchange rate constant, receptor-catalyzed GTP binding remains the slow
step in the overall GTPase cycle, as indicated by the dependence on
agonist concentration shown in Fig. 6.
Given the scale of the
reaction rates and the extent of stimulation of PLC activity observed
in the reconstituted system described here, the
receptor-G-PLC-
1 system has the capacity to generate
the speed and net activity to account for virtually any described
physiological IP
release without invoking the need for
other modulatory or accessory proteins. The potential function of such
proteins is not, of course, ruled out by these data, and this system
seems well suited for the study of any such ancillary factors. Because
the activation and deactivation rates described here are adequate to
account for the fast G protein-mediated responses that have been
described electrophysiologically, the search for the G protein GAPs
that act in these pathways takes on new interest.