From the Institut für Medizinische Physik und Biophysik, Humboldt-Universität zu Berlin, Universitätsklinikum Charité, Schumannstrasse 20-21, 10098 Berlin, Germany
Received for publication, October 17, 2000, and in revised form, December 4, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Despite the growing structural information on
receptors and G proteins, the information on affinities and kinetics of
protein-protein and protein-nucleotide interactions is still not
complete. In this study on photoactivated rhodopsin (R*) and the rod G
protein, Gt, we have used kinetic light scattering,
backed by direct biochemical assays, to follow G protein activation.
Our protocol includes the following: (i) to measure initial rates on
the background of rapid depletion of the GtGDP substrate;
(ii) to titrate GtGDP, GTP, and GDP; and (iii) to apply a
double displacement reaction scheme to describe the results. All data
are simultaneously fitted by one and the same set of parameters. We
obtain values of Km = 2200 Gt/µm2 for GtGDP and
Km = 230 µM for GTP; dissociation
constants are Kd = 530 Gt/µm2 for R*-GtGDP dissociation
and Kd = 270 µM for GDP release from
R*GtGDP, once formed. Maximal catalytic rates per
photoexcited rhodopsin are 600 Gt/s at 22 °C and 1300 Gt/s at 34 °C. The analysis provides a tool to allocate
and quantify better the effects of chemical or mutational protein
modifications to individual steps in signal transduction.
In retinal rod cells, absorption of a photon by the visual pigment
rhodopsin initiates a cascade of biochemical reactions that eventually
generates an electrical signal (1). Much of the tremendous overall gain
(105-106) of the visual cascade depends on two
enzymatic amplification stages, namely the receptor-catalyzed
nucleotide exchange in the rod G protein, transducin, and the
hydrolysis of the second messenger cGMP by the effector, cGMP phosphodiesterase.
Rhodopsin (R) and transducin
(Gt) display fundamental similarities to other receptors
and G proteins. However, the function of the rod as a single photon
sensory cell requires both low basal activities of R and Gt
and high speed of catalytic nucleotide exchange in the G protein. Any
catalytic activity of rhodopsin is effectively blocked in the dark by
the covalently bound inverse agonist 11-cis-retinal.
Although a mammalian rod cell contains ~107
rhodopsin molecules, thus providing the necessary target for efficient
light absorption, spontaneous single photon-like activity originates
only every 100 s from one of the many dormant receptors (2). On
absorption of a photon, light-induced isomerization converts the
chromophore to the agonist all-trans-retinal, thereby triggering conformational changes in the receptor protein that result
within milliseconds in the formation of the enzymatically active
intermediate metarhodopsin II (for review, see Refs. 3 and 4). Once
activated, rhodopsin finds by diffusion its substrate Gt.
In its inactive, GDP-bound state (GtGDP), the
heterotrimeric Gt holoprotein is peripherally bound to the
disc membrane by weak hydrophobic and ionic interactions (5-7). The
activation of the G protein proceeds through a sequence of two mutual
displacements (R* for GDP and GTP for R*; so-called double displacement
mechanism). Collisional interaction between light-activated rhodopsin
(R*) and GtGDP (Fig.
1A, step 1) triggers a
conformational change that opens the Gt
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
APPENDIX
REFERENCES
nucleotide-binding site. Upon GDP release, a stable R*Gt
complex with an empty nucleotide-binding site on the
Gt
-subunit is formed (step 2). Binding of GTP
to the Gt
-subunit within the R*Gt complex
enables a second conformational change (step 3) that eventually induces the dissociation of active GtGTP
(Gt*) from the receptor (step 4) and the
(simultaneous or unmeasurably delayed) separation of the
- and
-subunits (Gt
GTP and Gt
). At
least in vitro, activation is further accompanied by
immediate (delay <1 ms (8)) dissociation of both Gt
GTP
and Gt
from the disc membrane. The high rate of
R*-catalyzed nucleotide exchange leads to the rapid (transient)
accumulation of Gt*. Active Gt
GTP in turn
binds to the cGMP phosphodiesterase (PDE) within less than 5 ms (8).
The noncatalytic, stoichiometric interaction keeps the PDE active, and
hydrolysis of cGMP leads to the closure of cGMP-dependent
ion channels in the plasma membrane of the rod outer segment (for
review, see Refs. 9 and 10).
View larger version (14K):
[in a new window]
Fig. 1.
Reaction schemes of Gt activation
by R*. A, R*-catalyzed nucleotide exchange on
Gt includes (at least) four microsteps as follows. The
formation of the R*GtGDP complex by successful collision of
R* and GtGDP (step 1). Release of GDP and
formation of the stable R*Gt complex with empty
nucleotide-binding site (step 2). GTP uptake forms the
transient R*GtGTP complex (step 3), which in
turn dissociates into R* and active GtGTP (step
4). B, R*-catalyzed nucleotide exchange on
Gt, using the Cleland notation (horizontal line
symbolizes the reaction coordinate (33)). The double displacement
mechanism centers around the stable R*Gt complex with empty
nucleotide-binding site and involves two successive mutual
displacements on the G protein, namely R* for GDP and GTP for R*. The
initially formed encounter complex (R*·GtGDP) converts
into a complex with open Gt nucleotide-binding site
(R*Gt·GDP). GTP uptake leads to the release of active
Gt via analogous intermediates.
Despite the detailed knowledge of the reaction mechanisms and despite the growing information about the underlying structures (11, 12), sufficiently accurate estimates for the affinities and rates of the protein-protein and protein-nucleotide interactions are still not available. In this study we focus on the crucial activation reaction of the G protein, its rate of catalytic activation, and its dependence on GTP and GDP. Together with the kinetic parameters of PDE activation and cGMP turnover, this is the key parameter for any quantification of the gain of phototransduction in the rod cell (see Refs. 13 and 14). Efforts to measure the actual rate of the catalytic power of rhodopsin date back to the late 70s. To account for the rapid cGMP hydrolysis, Liebman and co-workers (15) concluded that light activation of a single molecule of rhodopsin results in the activation of several hundred molecules of PDE. After identification of transducin (16-18), direct GDP release and GTP binding studies yielded lower Gt activation rates (see Refs. 13 and 14). However, data obtained with low time resolution underestimate the actual rate, when not accounting for rapid depletion of the substrate, and are contaminated by both the onset of deactivation reactions and by slow activation of soluble Gt.
Complementary to the biochemical assays are the light-scattering (LS)
techniques, which follow flash-induced changes in the scattering of
near infrared light as an endogenous probe of specific molecular
changes (see Ref. 19). The LS monitor allows one to measure
Gt activation continuously and in real time, so that the decisive 500 ms after flash excitation are obtained. Following the
classical approach of Kühn and co-workers (5, 20), we combine it
with biochemical tools to quantify the amount of membrane-bound Gt and to calibrate the LS signals. The experimental data
can then be used to extract initial rates of Gt activation,
thereby separating them from slower, associated reactions, such as
membrane interaction, rhodopsin kinase interference (21), and the decay of the active receptor. To account for the influence of nucleotide concentration, we titrate GtGDP, GDP, and GTP. The double
displacement reaction scheme that takes into account of all these
components (22) is then applied to the experimental data.
![]() |
EXPERIMENTAL PROCEDURES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Membrane and Transducin Preparations--
Rod outer segments
were prepared from frozen bovine retinas using a sucrose gradient
procedure as described (23). Hypotonically stripped disc membranes were
prepared from rod outer segments either by two consecutive extractions
with low salt buffer as described (24) or by the Ficoll floating
procedure similar to the procedure described (25) except that 2% w/v
Ficoll instead of 5% was used. Both methods yield osmotically intact
disc vesicles with a vesicle size >400 nm. No significant difference
was found between the two types of preparations. Contamination by
vesicle aggregates was removed by a 2-µm filter (Nucleopore).
Membranes were kept on ice and used within 4 days without any loss of
activity. Rhodopsin concentration was determined from its absorption
spectrum using 500 = 40,000 M
1
cm
1.
Transducin was purified as described (8). Subunits were further
purified on Blue-Sepharose (1 ml of HiTrap Blue, Amersham Pharmacia
Biotech) at a flow rate of 1.2 ml/h. Proteins eluted with starting
buffer (20 mM BTP, pH 7.5, 1 mM
MgCl2, 2 mM dithiothreitol) contain inactive
Gt (26). Active Gt
was eluted with
a linear gradient of 0-0.3 M NaCl (15 ml).
Gt
was eluted with 1 M NaCl. The subunits
were dialyzed against measuring buffer (20 mM BTP, pH 7.4, 130 mM NaCl, 5 mM MgCl2 and 2 mM dithiothreitol), concentrated (Amicon, YM-10), and
stored at
40 °C. Gt
concentration was determined
by the method of Bradford (27) using bovine serum albumin as the
standard. The amount of intact, activable Gt
was determined precisely by fluorometric titration with GTP
S (28).
Nucleotides--
GTP (>98%, Fluka) was used without further
purification. GDP (80%, Fluka) was further purified by two successive
ion exchange chromatographies on QAE-Sepharose (Amersham Pharmacia
Biotech) using a linear gradient of 0.2-1 M
triethylammonium hydrogen carbonate buffer, pH 7.7. The procedure
yields GDP virtually free of contaminating GTP. Concentration of the
nucleotides was determined spectroscopically using 252 = 13,700 M
1
cm
1.
Because it is known that GDP is slowly converted to GTP by guanylate kinase present in preparations of disc membranes (29, 30), the incubation time of GDP prior to the recording of the dissociation signal was minimized to less than 3 min.
Centrifugation Assay-- The relative amount of soluble and membranous Gt was determined using the centrifugation assay (18, 19, 21). Aliquots (100 µl) of the samples used for recording of LS signals were pelleted by centrifugation (2 min, 21,000 × g, 22 °C). After removal of the supernatant, the pellet was washed once (without resuspending, to avoid loss membrane-bound Gt) with buffer and then resuspended in 100 µl of buffer. The amount of Gt either bound to the membrane pellet or present in the supernatant was analyzed by densitometry on Coomassie Blue-stained sodium dodecyl sulfate-polyacrylamide gel electrophoresis.
All samples were heated to 95 °C for 10 min in the presence of SDS,
to aggregate most of rhodopsin, which would otherwise mask the
Gt-subunit. Background staining by residual rhodopsin was subtracted using a sample without Gt (see Fig.
2A, lanes 1 and
2). To ensure reproducible quantification, arrestin was
added to the sample buffer as an internal standard.
|
Kinetic Light Scattering-- Light-scattering changes were measured in a set-up described in detail in Heck et al. (19). All measurements were performed in 10-mm path cuvettes with 300-µl volumes in isotonic buffer (20 mM BTP, pH 7.4, 130 mM NaCl and 5 mM MgCl2) and at 22 °C unless specified otherwise. Reactions are triggered by flash photolysis of rhodopsin with a green (500 ± 20 nm) flash, attenuated by appropriate neutral density filters. The flash intensity is quantified photometrically by the amount of rhodopsin bleached and expressed either in terms of the mole fraction of photoexcited rhodopsin (R*/R) or in the surface density of R* (R*/µm2).
Dissociation signals were recorded with a 0.5-5-ms dwell time of the
A/D converter (Nicolet 400, Madison, WI). To suppress base-line
activation, NH2OH was added to the membrane stock at a
concentration of 5 mM. The final concentration of
NH2OH in the samples never exceeded 300 µM
NH2OH, to keep the decay of the flash-induced R* small. For
calibration of the LS monitor, dissociation signals were induced by
saturating flashes (R*/R = 0.5%). For the kinetic steady state
analysis dissociation signals were routinely measured at R*/R = 2.3·104 (5.7 R*/µm2),
i.e. in the linear range of the light titration curve (Fig. 3). Binding signals (R*/R = 32%)
were corrected by a reference signal (N signal) measured on a sample
without added Gt as described (19). All data were taken at
pH 7.4, i.e. in the maximum of the bell-shaped pH/rate
profile (31); at this pH, membrane binding of GtGDP (in the
dark) is also near its
maximum.2
|
Amplitudes of the signals are expressed as relative scattering
intensity changes (I/I, where I represents the
intensity measured before the flash). Calibration of the amplitudes was
routinely performed prior to each set of experiments by measuring
dissociation and binding signals induced by saturating flashes on
aliquots of disc membranes (3 µM R) supplemented with 0.5 µM Gt (see Fig. 2B). As described
below (see under "Results"), the dissociation signal exclusively
monitors activation of the membrane-bound pool of GtGDP
(Gtmb), whereas the binding signal exclusively
measures transition of the soluble fraction of GtGDP
(Gtsol) to the membranes. Thus, the (absolute
values of the) maximum scattering change of the signals are
proportional to the concentration of Gtmb and Gtsol, respectively. Together with the known
amount of total Gt added to the samples, the amplitude of
the signals can be converted to concentration units by the
scaling factor (F) as shown in Equations 1-3.
![]() |
(Eq. 1) |
![]() |
(Eq. 2) |
![]() |
(Eq. 3) |
![]() |
(Eq. 4) |
Reaction Scheme and Mathematical Analysis--
The R*-catalyzed
steady state Gtmb activation rate depends on the
Gtmb surface concentration as well as on the
volume concentrations of both GDP and GTP (Fig.
4C). The multifactorial
dependence is well described by the double displacement scheme
(formally equivalent to the ping-pong scheme; see Fig. 1B
and see Refs. 22 and 33).
|
By using the steady state approach, the following rate Equation 5 can
be derived for the initial rate of Gtmb
activation (vRG) in absence of the product
GtGTP (see e.g. Ref. 34),
![]() |
(Eq. 5) |
Each set of experiments comprises the titration of the dissociation signal with exogenous Gt in the presence of fixed concentrations of GTP and GDP. Both the initial activation rate of Gtmb (vRG) and the initial Gtmb surface concentration were determined from the maximum slope and the maximum amplitude of the dissociation signal, respectively (see above and "Results"). The Gt titrations were repeated (i) at different concentrations of GTP (10-3000 µM) and (ii) at 200 µM GTP with different concentrations of GDP (75-2000 µM). In this way the dependence of vRG on the three variables (Gtmb, GTP, and GDP) was obtained (Fig. 4).
The data points of 23 independent sets of titration experiments (232 data points overall) were numerically fitted with Equation 5, using a multiple least squares fit, i.e. the simultaneous fit to all 23 titrations using one and the same set of the parameters Km(G), Km(GTP), Kd(GDP), Kd(G), and Vmax. In the fit procedure (Scientist Software, MicroMath), the concentrations of GTP and GDP were fixed for each individual titration (i.e. for each pair of [GTP] and [GDP]), and each of the five kinetic parameters was allowed to vary. The turnover number (Vmax/R*) was then calculated with the known surface density of R* (5.7 R*/µm2).
Note that Equation 5 is converted into a simple Michaelis-Menten type
of hyperbolic function for any given (fixed) set of nucleotide
concentrations to yield Equation 6,
![]() |
(Eq. 6) |
The validity of the steady state approach is essentially based on the
following: (i) the initial Gtmb concentration is
high as compared with R* even at the lowest Gtmb
concentrations investigated, and (ii) at the time the maximum slope was
obtained, less than 5% of Gtmb is activated,
thus rendering depletion of the substrate negligible. We have also
found that neither lowering the membrane concentration (1.8 µM instead of 3 µM R) nor addition of
excess Gt leads to a significant change of the
observations (a putative persistent Gt
-receptor
interaction may be integrated in the reaction scheme and the respective
rate equations derived (22)). Note that both
Km(GTP) and
Kd(GDP) do not depend on the correct
determination of Gtmb (in contrast to
Vmax, Kd(G), and
Km(G), which linearly depend on any
error in the Gtmb concentration).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Quantification of Membrane-bound Transducin--
Fig.
2A shows the typical, variable membrane binding of
Gt and its dependence on light and nucleotides as analyzed
by the centrifugation assay (18, 19, 21) (see "Experimental
Procedures"). Upon dilution of the membranes, GtGDP
(unlike other G proteins) is in equilibrium between a membrane-bound
(Gtmb) and a soluble form
(Gtsol) in the dark (Fig. 2A, lanes 3 and 4). The extent of this "dark binding" depends on
several factors, including overall protein and membrane concentration,
temperature, pH, ionic strength, and divalent cations (see Refs. 32 and
35). Photoactivation of excess rhodopsin in the absence of GTP or GDP
locks Gt in its nucleotide-free, receptor-bound
conformation (R*Gt), which is seen in a complete shift of
Gt to the membrane (Fig. 2A, lanes 5 and 6). Conversely, light activation of rhodopsin in the
presence of GTP leads to a complete dissociation of Gt from
the membranes (Fig. 2A, lanes 7 and 8). Due to
the low membrane concentration used in the experiments (equivalent to 3 µM R), both Gt- and Gt
-subunits almost quantitatively dissociate from the
membrane upon activation (Fig. 2A), even at high
Gt concentration (data not shown).
The gain of mass of a membrane, when Gt is bound from the solution, and the loss of mass with its dissociation produces large and readily measurable changes in near-infrared light scattering (LS signals; see Ref. 19). Representative light-induced LS signals (measured on the samples used for the centrifugation assay shown in Fig. 2A) are shown in Fig. 2B. Transition of all Gtsol to the membrane induced by bleaching excess rhodopsin in the absence of GTP gives rise to an increase of the scattered light ("binding signal" (20, 36); Fig. 2B, trace a). Accordingly, dissociation of Gtmb from the membrane upon activation is seen as decrease of LS ("dissociation signal" (20, 36); Fig. 2B, trace b). Since the dissociation step itself is not rate-limiting (see below), the rising phase of the dissociation signal is a real time monitor of Gt activation. Notably, binding signals are generally slow as compared with dissociation signals. Previous work resolved this apparent conflict by the finding that, although binding of Gtmb to R* is fast, interaction of Gtsol with the membrane is slow (36). Consequently, the activation rate of Gtsol is limited by its membrane binding, thereby leading to artificially slowed activation rates when total active Gt* formation is assayed under conditions where a significant fraction of GtGDP is solubilized.
As indicated in Fig. 2B, the dissociation signal exclusively monitors fast activation of Gtmb. Its maximum amplitude is not contaminated by a contribution of Gtsol, since the slow mass gain of the membrane upon its binding is just canceled by the fast subsequent dissociation of Gt*. Accordingly, the maximum amplitude of the dissociation signal is proportional to the Gtmb pool. On the other hand, the maximum amplitude of binding signals induced by saturating flashes is proportional to Gtsol, since binding of Gtmb to R* does not lead to any LS changes of the membrane suspension (note that this is in contrast to LS measurement on rod outer segment preparations (37, 38)). Consequently, comparison of the amplitudes of the dissociation and binding signals allows determination of the sizes of the Gtmb and Gtsol pools (Fig. 2, B and C). The relative fraction of Gtmb varied with different membrane and Gt preparations between 40 and 60% of the total Gt added. Densitometry on gels as shown in Fig. 2A was used to independently quantify dark binding of Gt under the conditions of the LS experiments. Comparison of the results obtained by the two methods (Fig. 2C) shows good agreement, which further confirms the interpretation of the LS signals.
The sum of the maximum amplitudes (absolute values) of the dissociation and binding signals is proportional to the known amount of Gt added (i.e. Gtmb + Gtsol), thereby allowing the calibration of the LS monitor. A calibration factor (F) was calculated for every set of experiments, which relates the relative scattering intensity change to Gt concentration units (see Equations 1-3). The calibration factor varied only little between different preparations (7.0 < F < 8.3 µM). The scaling factor was used to transform the maximum amplitude of individual dissociation signals to yield the respective initial (i.e. at time of the flash) volume concentration of Gtmb. Based on the known surface density of rhodopsin in the disc membrane (25,000 R/µm2; see Ref. 32), the Gtmb volume concentration is then easily transformed to surface concentration units (Equations 4 and 7; see Fig. 2D).
Titration of the amplitude of the dissociation signal with
Gt yields the dependence of Gtmb on
added Gt (Fig. 2D). The slight but reproducible
sigmoidal dependence of Gtmb on added
Gt may be explained by a relatively weak
Gt-Gt
-subunit interaction in solution
and a negligible interaction (relative to the
Gt-holoprotein) of the isolated subunits with the membrane,
which is in agreement with gel filtration and centrifugation
experiments.2
Dependence of Gtmb Activation Rate on the Concentration
of R*--
Gtmb activation is accelerated with
increasing concentration of R*. Accordingly, the rise time of the
dissociation signal depends on the mole fraction of photoexcited
rhodopsin (Fig. 3). The light titration curve saturates at high
bleaching levels due to the local depletion of Gt on the
disc membrane (and possibly rate limitation by dissociation of
Gt* from the membranes). At low bleaching levels (R*/R < 103) the maximum slope of the dissociation
signal depend linear on the R* concentration (dashed line in
Fig. 3), which proves that the dissociation of Gt* from the
membrane is not rate-limiting under these conditions and establishes
the rising phase of the dissociation signal as a real time monitor of
Gt* formation.
Furthermore, the linearity is an important criterion for the analysis
of the activation rates since it allows us to normalize the activation
rates to R*, i.e. to calculate the turnover number (Vmax/R*) of R*-catalyzed Gt
activation. Consequently, we routinely measured the dissociation signal
at R*/R = 2.3·104 (5.7 R*/µm2).
With decreasing R* the amplitude of the dissociation signal decreases
(Fig. 3, inset), since it is proportional to the number of
individual vesicles hit by at least one photon (the slow activation of
Gtmb initially bound to vesicles that do not
contain an R* is not detected on the time scale of the experiments). It
is seen that under the experimental standard conditions (R*/R = 2.3·104) more than 80% of the maximum
amplitude is evoked. Fit of the data points to a Poisson sum
(solid line in inset of Fig. 3) yield 15.000 R
per domain (i.e. vesicle). With the known rhodopsin surface density of 25,000/µm2, a vesicle size of 440 nm
(diameter) is calculated (see Ref. 39 for details), which is in good
agreement with the size as estimated from electron micrographs of our
preparations (not shown). The slight systematic deviation of the data
points from the best fitting Poisson sum is most likely due to slightly
nonuniform vesicle size (and contamination by slow GTPase reaction,
significant at very low R*).
Steady State Analysis of Gt Activation Rate--
Fig.
4A shows a typical set of dissociation signals, each evoked
by a small flash producing 5.7 R*/µm2 (i.e. in
average about 3.4 R* per disc membrane vesicle) with increasing amounts
of purified Gt added. As expected, both the maximum
amplitude (Fig. 2D) and the maximum slope of the signal (Figs. 4 and 5) increase with
Gt concentration. Expansion of the scales (Fig.
4B) shows that the maximum slope is not immediately reached
after light activation of the receptor but is delayed by a short,
R*-depending period (8). It is the time it takes all the individual
reactions, R* formation, nucleotide exchange, and dissociation of
Gt* from the membrane, to reach a steady state. The maximal
rate of Gt* production is reached 40-100 ms after the
flash under the condition of the experiments; despite of this delay, we
will refer to it as "initial" rate of Gt activation. After the short linear period, the slope of the dissociation signal slows due to depletion of Gtmb and increasing
activation of Gtsol. Because membrane binding of
Gtsol is slow, distortion of the maximum slope
by this effect was neglected. Rapid substrate depletion is a
consequence of the membrane-bound state of the proteins, which limits
the "reservoir" of the substrate as compared with reactions in
solution. As calculated from the dependence of
[Gtmb] on [Gt] added (Fig.
2D), Gt membrane binding saturates at about 7500 Gt/µm2 (0.3 Gtmb per
R), which is in good agreement with previous estimates (40, 41).
|
The maximum slope of the signals (dashed lines in Fig. 4B) was taken from the linear period, which lasts 100-300 ms, depending on Gt concentration (Fig. 4B). It was then converted to the steady state Gt* formation rate by the calibration procedure described above (see Equations 1-4). Importantly, the initial Gtmb concentration was calculated from the maximum amplitude of each signal (see above and under "Experimental Procedures"), thus avoiding errors inferred from variations of total Gt added (see Fig. 2D).
The titration of the dissociation signal with exogenous Gt (Fig. 4A) was repeated (i) with [GDP] = 0 and different concentrations of GTP (10-3000 µM; Fig. 4C, upper panel) and (ii) with 200 µM GTP and different concentrations of GDP (75-2000 µM, Fig. 4C, lower panel). The resulting dependence of the initial rate of Gtmb activation (vRG) on the three variables (Gtmb, GTP, and GDP) was then fitted by a simultaneous fit to the data, using Equation 5 and 23 sets of dissociation signals; different colors in Fig. 4C identify different concentrations of nucleotide, and different symbols identify different preparations. The kinetic parameters thereby obtained are summarized in Table I.
|
Dependence of Gtmb Activation Rate on
Temperature--
The temperature dependence of
Gtmb activation was studied by varying
Gtmb at each temperature in the presence of 3 mM GTP (no GDP added; Fig. 5A). Because the
experiments were done at a single, fixed nucleotide concentration, the
data were fit to a simple hyperbolic function (Equation 6). The
apparent turnover number of R*-catalyzed Gtmb
activation (V
|
The temperature dependence of the apparent turnover number of R*
catalyzed Gt activation
(V
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We have studied the receptor-catalyzed nucleotide exchange of a heterotrimeric G protein with the aim to quantify both the maximal catalytic activity of the activated receptor (turnover number) and the influence of GTP and GDP on the velocity of G protein activation. For such analysis, the visual system is well suited because (i) neither the receptor nor the G protein show any measurable basal activity; (ii) the G protein (Gt) is easily isolated and available in the quantities required for titration; and (iii) the light trigger allows us to load the system with defined doses of active receptor.
Initial Rate Analysis--
Enzyme-catalyzed reactions are commonly
assayed by steady state product formation with a high initial substrate
concentration (S0 Et). In the case
of G protein-coupled systems, the membrane localization of the
substrate (the G protein) limits its initial concentration. Bovine rod
disc membranes operate at a Gt/R*
(S0/Et) ratio of 3000:1 (single photon
detection). The low amount of membrane-bound GtGDP
(Gtmb) leads to rapid depletion of the substrate
and causes a very short steady state period. It is obvious that the
rapid depletion of the G protein substrate cannot be overcome by
overloading the membranes with G protein. Even if, as in the visual
system, the G protein is in vitro in equilibrium between a
membrane-bound and soluble form, the problem is not solved by a large
excess of G protein in solution. The transition of soluble
GtGDP (Gtsol) to the membrane is far
too slow to maintain a high concentration of membrane-bound
GtGDP. In other words, the rate of activation of soluble
GtGDP is limited by the slow membrane binding step, which
is particularly evident when Gt activation is assayed at
low membrane concentrations. As a consequence, true activation rates
are only obtained when the initial rate of Gt activation is
assayed in the presence of sufficiently high membrane concentrations.
The initial rate approach kinetically separates the rapid activation of
membrane-bound GtGDP from the slower activation of the
soluble GtGDP pool (and from other slow reactions that
influence the rate, such as R* deactivation and GTPase reaction). The
keys to reliable activation rates are thus a millisecond time
resolution and an accurate quantification of the membrane-bound
fraction of the G protein.
Unfortunately, biochemical assays so far applied to this system lack the necessary time resolution. Stopped-flow techniques, which were successfully applied to the study of nucleotide uptake or GTPase reaction of various systems, including small G proteins (see e.g. Refs. 42 and 43) and EF-Tu (see e.g. Ref. 44), are hard to apply to the visual system with its light sensitivity. We thus used the kinetic light-scattering technique to obtain initial rates of Gtmb activation. Previous work established the dissociation signal as a real time monitor of the fast activation of the membrane-bound fraction of GtGDP (20, 36, 37). A precise calibration can be obtained from the dependence on [R*] (37, 38) and/or from classical biochemical assays (this work). Both the initial surface concentration of Gtmb and the initial Gtmb activation rate can be obtained (Figs. 2 and 4).
Double Displacement Scheme--
The dependence of the
Gt activation rate on nucleotide or initial
GtGDP concentration is commonly analyzed employing a
Michaelis-Menten type of hyperbolic function (e.g. Equation 6). For example, each titration with Gt (Figs. 4 and 5) can
be fitted with Equation 6. The resulting parameters
(K'm and
V'max), however, are only apparent
values when multiple substrates are involved. The
K
|
We have used the classical double displacement reaction scheme to describe adequately the receptor-catalyzed activation of a G protein (Fig. 1 (22)). By using the steady state approach, rate equations can be derived (e.g. Equation 5) that explicitly account for the concentrations of all components involved (R*, GtGDP, GTP, GDP) and allow us to extract the true kinetic parameters for the individual steps.
We note that the inclusion of the transitory complex (R*GtGDP) in the reaction scheme is justified by the experimental data: as seen in Fig. 4C, Vmax is not approached in the presence of GDP even at infinite GtGDP concentrations. This shows that GDP is not a competitive inhibitor implying that binding of GtGDP to R* and the release of GDP are separated by a transitory R*GtGDP complex with finite lifetime.
Affinity of GtGDP for Photoactivated
Rhodopsin--
The affinity of GtGDP for R* depends on the
equilibrium of the active receptor conformation with its tautomeric
forms (see under "Appendix"). For the following discussion, we use
the corrected parameters Km(G) and
Kd(G) to characterize the interaction of
Gtmb with R* (Table I). As compared with the
native surface concentration of Gt (3000 µm2) the value of
Kd(G) (530 ± 260 µm
2) reflects a surprisingly weak
interaction of the proteins. However, this does not contradict the
experimentally confirmed stability of the R*Gt complex
(i.e. the complex without bound nucleotide (45, 46)),
because formation of the latter is composed of two
reactions, namely the initial interaction of GtGDP with R* (step 1 in Fig. 1A) and the succeeding GDP
release (step 2 in Fig. 1A). Consequently, the
formation of the complex depends on both
Kd(G) and
Kd(GDP) (see under "Appendix"). Despite the low affinity of GtGDP to R* (high value of
Kd(G)), the reaction is almost
quantitatively shifted toward formation of the R*Gt complex
under typical experimental conditions (i.e. low membrane
concentration, no added GDP) because the concentration of the
endogenous GDP released is too low to dissociate the complex (Kd(GDP) = 270 µM; see
Table I). At high membrane concentrations, however, the endogenous GDP
can significantly affect R*Gt formation.
When volume concentrations are used and/or the influence of the endogenous GDP is omitted (see e.g. Refs. 47 and 48), the resulting Gt-R* affinity depends on all the concentrations used in the experiment. Thus the proper definition and separation of the individual reaction steps is not an academic problem but has an immediate impact on the reaction mechanism.
Dependence on Nucleotides--
An important result of this study
is the high Kd (low affinity) of GDP to
R*Gt, once formed (Kd(GDP)),
and the high Km of GTP
(Km(GTP)). This may seem surprising in
view of the higher apparent affinities of the nucleotides obtained 1)
by Gt activation assays performed at low concentration of
Gt and/or membrane, or 2) under equilibrium conditions.
Reasons for these apparent discrepancies are as follows. 1) As
described above, low [Gt] (as compared with
Km(G)) necessarily leads to lower
apparent K
The low affinity of GDP to R*Gt obtained in this study (Kd(GDP) = 270 µM) is necessary to explain the essentially quantitative formation of the R*Gt complex at low overall concentrations and in the absence of added GDP (see above). The low Michaelis constant for GTP (Km(GTP) = 230 µM) is in agreement with conclusions drawn from independent analyses of GTP-induced dissociation of the R*Gt complex (50).
Gt Activation Rate-- Published estimates of the Gt activation rate vary from about 10 to >3000 Gt*/s per R* formed (for review see Ref. 13). The extreme variation is at least partially due to differences with respect to the method, preparation, and measuring conditions used. For example, when aliquots for a filter assay of GTP uptake are taken in seconds intervals (see e.g. Ref. 51), not only the depletion of membrane-bound GtGDP but also slow reactions may severely affect the results. Activation of soluble GtGDP will be the predominant artifact in broken rod outer segment preparations or reconstituted systems, whereas deactivation of R* may interfere in more intact systems. Evidently, measurements of Gt activation on rod outer segments require a sufficiently fast assay that measures the rising phase of Gt activation before these influences cut in (<100 ms for mammalian rods). With such approaches, initial rates in the order of 1000 Gt*/R*s are obtained (38, 52, 53).
In the present study on isolated membranes, measuring initial rates of Gtmb activation with known Gtmb surface concentrations, the steady state approach yields a turnover number of 600 and 1300 Gt*/R*s at 22 and 34 °C, respectively. With the kinetic parameters obtained at 22 °C (Table I), the activation rate can be plotted as a function of GTP and GDP (Fig. 6).
The maximal rate is in good agreement with a previous study
on whole rod outer segments (800 Gt*/R*s at 21 °C (38)).
However, there is a discrepancy because with the isolated
membranes and the assumed native Gt concentration,
Vmax, is not approached (see Fig.
6A). This is due to an increased
Km(G) and
Kd(G) for R*-GtGDP
interaction in the reconstituted preparation. The intact membranes may
reserve more quantitative formation of active metarhodopsin II and/or
more efficient collisional coupling. The latter is likely to depend on
the reversible carboxymethylation of the G-subunit (26).
What Is the Rate-limiting Step?-- Given the complexity of the reaction scheme (Fig. 1), the question arises whether the maximum rate of receptor-catalyzed G protein activation is limited by the rate of diffusion of the G protein or GTP or by protein conformation changes.
In the Arrhenius representation, the dependence of
V'max/R* on temperature (Fig.
5B) yields a low temperature (<22 °C) branch with an
activation energy of 111 kJ/mol. This value is significantly lower than
the one determined for Gt activation between 2 and 12 °C and at very low [GTP] (175 kJ/mol (50)). Although the reasons for the difference remain unknown, the high activation energy
indicates that a protein conformational change within the reaction
sequence is likely to be rate-limiting in this range (54). At
sufficiently high temperature, a process with smaller activation energy
(Ea <30 kJ/mol) takes over, possibly a diffusion
limited process (54). However, it is difficult to extract an individual
step when the kinetic parameters are composed of various individual
rate constants (see Table I), each of which may have its own
temperature dependence.
The lower limit of the R*-GtGDP encounter rate obtained
from our analysis (see under "Appendix") is 820 s1 at 22 °C, i.e. well below
its theoretical limit of 7000 s
1 (13). The
lower limit for the bimolecular rate constant of formation of the
encounter complex R*GtGTP is 2.6·106
M
1 s
1
(see under "Appendix"), which is much smaller than the diffusional limit of about 108-109
M
1 s
1.
Thus, at least in the reconstituted system, neither binding of the G
protein nor GTP uptake is diffusion-controlled.
Application to Other Receptors--
We have shown that the
four-step analysis adequately describes the experimental data obtained
with the visual system. Modifications of the analysis, to account for
ligand binding (22), should make it applicable to other receptor
systems, thus providing a basis for the assignment and quantification
of chemical and mutational probing.
![]() |
ACKNOWLEDGEMENTS |
---|
We are indebted to Ulrike Laitko for helpful suggestions. We thank Ingrid Semjonow for excellent technical assistance.
![]() |
FOOTNOTES |
---|
* This work was supported by Grant SFB 366 from the Deutsche Forschungsgemeinschaft (to K. P. H.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence may be addressed: Institut für
Medizinische Physik und Biophysik, Humboldt-Universität zu
Berlin, Universitätsklinikum Charité, Schumannstrasse
20-21, 10098 Berlin, Germany. Tel.: 49-30-450-524111; Fax:
49-30-450-524952; E-mail: martin.heck@charite.de.
§ To whom correspondence may be addressed: Institut für Medizinische Physik und Biophysik, Humboldt-Universität zu Berlin, Universitätsklinikum Charité, Schumannstrasse 20-21, 10098 Berlin, Germany. Tel.: 49-30-450-524111; Fax: 49-30-450-524952; E-mail: kph@charite.de.
Published, JBC Papers in Press, December 14, 2000, DOI 10.1074/jbc.M009475200
2 M. Heck and K. P. Hofmann, unpublished observations.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
R, rhodopsin;
R*, light-activated rhodopsin;
Gt, G protein of the rod,
transducin;
GtGDP, inactive, GDP-bound transducin;
GtGTP or Gt*, active, GTP-bound transducin;
Gt,
-subunit of transducin;
Gt
,
-subunit of transducin;
Gtmb, membrane-bound transducin;
Gtsol, soluble
transducin;
LS, light-scattering;
PDE, phosphodiesterase;
BTP, 1,3-bis[tris(hydroxymethyl)methylamino]propane;
GTP
S, guanosine
5'-3-O-(thio)triphosphate.
![]() |
APPENDIX |
---|
Membrane Localization of the Proteins--
It is obvious that the
density of rhodopsin in the disc membrane does not change upon dilution
of the disc membrane suspension. As a consequence, neither the
equilibrium of the R*-GtGDP interaction nor the rate of the
R*GtGDP complex formation depends on the dilution of the
membranes for a given amount of membrane-bound GtGDP
(Gtmb). Hence, both the concentrations and the
corresponding kinetic parameters are only properly specified in
two-dimensional terms. On the other hand, any surface density,
[X]2D (including e.g. the values of
Km(G),
Kd(G), and microscopic rate constants),
is easily transformed to a volume concentration,
[X]3D (or vice versa), by the relation shown
in Equation 7.
![]() |
(Eq. 7) |
The problem becomes even more evident when reactions are investigated
that depend on both soluble components (such as GDP and GTP) and
membrane-bound proteins. For instance, in the coupled equilibrium (see
Equation 8)
![]() |
(Eq. 8) |
![]() |
(Eq. 9) |
Influence of the Relative Fraction of Active Receptor on the Kinetic Parameters-- A general characteristic of G protein-coupled receptors is the equilibrium of inactive and active receptor conformations. As a consequence, the actual amount of active receptor may be lower than the total amount of "activated" receptor (R*). The receptor equilibrium does not affect the interaction of the nucleotides with the R*Gt complex once formed nor the turnover number Vmax/R* (due to the stabilizing effect of the G protein on the active receptor conformation). However, it strongly influences the interaction between R* and the G protein; the more the equilibrium is shifted to the inactive receptor conformation, the higher the observed values of Km(G) and Kd(G). The true values of Km(G) and Kd(G) are obtained by multiplying the observed values by the fraction (fA) of the active receptor (A) (measured in the absence of G protein).
The active rhodopsin conformation can be identified with metarhodopsin
II (MII), which is in a pH and temperature dependent equilibrium with
its precursor metarhodopsin I (MI; for review see Ref. 55) (Equation 10):
![]() |
(Eq. 10) |
![]() |
(Eq. 11) |
Lower Limit of the Rate of G Protein and GTP Binding-- It is important to note that the steady state treatment does not yield values of microscopic rate constants. However, lower limits of k1 and k3 (Fig. 1) can be calculated from kcat/Km (see Ref. 34). The minimum second-order rate constant of the formation of the encounter complex R*GtGDP (k1) is thus given by Equation 12,
![]() |
(Eq. 12) |
Analogously, a lower limit for the second-order rate constant of the
formation of the encounter complex R*GtGTP
(k3) is given by
![]() |
(Eq. 13) |
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Baylor, D. A., Lamb, T. D., and Yau, K. W. (1979) J. Physiol. (Lond.) 288, 613-634[Abstract] |
2. | Lamb, T. D. (1987) J. Opt. Soc. Am. 4, 2295-2300 |
3. | Hofmann, K. P. (1999) in Rhodopsins and Phototransduction, Novartis Foundation Symposium 224 (Takeuchi, I. , Bock, G. , and Goode, J. A., eds) , pp. 158-175, John Wiley & Sons Ltd., Chichester, UK |
4. | Hofmann, K. P. (2000) in Handbook of Biological Physics (Stavenga, D. G. , de Grip, W. J. , and Pugh, E. N., Jr., eds), Vol. 3 , pp. 91-141, Elsevier Science Publishers B.V., Amsterdam |
5. | Kühn, H. (1981) Curr. Top. Membr. Transp. 15, 171-201 |
6. | Seitz, H. R., Heck, M., Hofmann, K. P., Alt, T., Pellaud, J., and Seelig, A. (1999) Biochemistry 38, 7950-7960[CrossRef][Medline] [Order article via Infotrieve] |
7. |
Melia, T. J.,
Malinski, J. A.,
He, F.,
and Wensel, T. G.
(2000)
J. Biol. Chem.
275,
3535-3542 |
8. | Heck, M., and Hofmann, K. P. (1993) Biochemistry 32, 8220-8227[Medline] [Order article via Infotrieve] |
9. | Chabre, M., and Deterre, P. (1989) Eur. J. Biochem. 179, 255-266[Medline] [Order article via Infotrieve] |
10. | Helmreich, E. J., and Hofmann, K. P. (1996) Biochim. Biophys. Acta 1286, 285-322[Medline] [Order article via Infotrieve] |
11. | Lambright, D. G., Sondek, J., Bohm, A., Skiba, N. P., Hamm, H. E., and Sigler, P. B. (1996) Nature 379, 311-319[CrossRef][Medline] [Order article via Infotrieve] |
12. |
Palczewski, K.,
Kumasaka, T.,
Hori, T.,
Behnke, C. A.,
Motoshima, H.,
Fox, B. A.,
Le Trong, I.,
Teller, D. C.,
Okada, T.,
Stenkamp, R. E.,
Yamamoto, M.,
and Miyano, M.
(2000)
Science
289,
739-745 |
13. | Pugh, E. N., Jr., and Lamb, T. D. (1993) Biochim. Biophys. Acta 1141, 111-149[Medline] [Order article via Infotrieve] |
14. | Pugh, E. N., Jr., and Lamb, T. D. (2000) in Handbook of Biological Physics (Stavenga, D. G. , De Grip, W. J. , and Pugh, E. N., Jr., eds), Vol. 3 , pp. 183-255, Elsevier Science Publishers B.V., Amsterdam |
15. | Yee, R., and Liebman, P. A. (1978) J. Biol. Chem. 253, 8902-8909[Medline] [Order article via Infotrieve] |
16. | Godchaux, W., III, and Zimmerman, W. F. (1979) J. Biol. Chem. 254, 7874-7884[Abstract] |
17. | Fung, K. K. B., and Stryer, L. (1980) Proc. Natl. Acad. Sci. U. S. A. 77, 2500-2504[Abstract] |
18. | Kühn, H. (1980) Nature 283, 587-589[Medline] [Order article via Infotrieve] |
19. | Heck, M., Pulvermüller, A., and Hofmann, K. P. (2000) Methods Enzymol. 315, 329-347[CrossRef][Medline] [Order article via Infotrieve] |
20. | Kühn, H., Bennett, N., Michel Villaz, M., and Chabre, M. (1981) Proc. Natl. Acad. Sci. U. S. A. 78, 6873-6877[Abstract] |
21. | Pulvermüller, A., Palczewski, K., and Hofmann, K. P. (1993) Biochemistry 32, 14082-14088[Medline] [Order article via Infotrieve] |
22. | Waelbroeck, M., Boufrahi, L., and Swillens, S. (1997) J. Theor. Biol. 187, 15-37[CrossRef][Medline] [Order article via Infotrieve] |
23. | Papermaster, D. S. (1982) Methods Enzymol. 81, 48-52[Medline] [Order article via Infotrieve] |
24. | Bauer, P. J. (1988) J. Physiol. (Lond.) 401, 309-327[Abstract] |
25. | Smith, H. G., Jr., Stubbs, G. W., and Litman, B. J. (1975) Exp. Eye Res. 20, 211-217[Medline] [Order article via Infotrieve] |
26. | Parish, C. A., and Rando, R. R. (1994) Biochemistry 33, 9986-9991[Medline] [Order article via Infotrieve] |
27. | Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve] |
28. | Fahmy, K., and Sakmar, T. P. (1993) Biochemistry 32, 7229-7236[Medline] [Order article via Infotrieve] |
29. |
Panico, J.,
Parkes, J. H.,
and Liebman, P. A.
(1990)
J. Biol. Chem.
265,
18922-18927 |
30. |
Zera, E. M.,
Molloy, D. P.,
Angleson, J. K.,
Lamture, J. B.,
Wensel, T. G.,
and Malinski, J. A.
(1996)
J. Biol. Chem.
271,
12925-12931 |
31. |
Kisselev, O. G.,
Meyer, C. K.,
Heck, M.,
Ernst, O. P.,
and Hofmann, K. P.
(1999)
Proc. Natl. Acad. Sci. U. S. A.
96,
4898-4903 |
32. | Liebman, P. A., Parker, K. R., and Dratz, E. A. (1987) Annu. Rev. Physiol. 49, 765-791[CrossRef][Medline] [Order article via Infotrieve] |
33. | Cleland, W. W. (1963) Biochim. Biophys. Acta 67, 104-137[CrossRef] |
34. | Segel, I. H. (1975) Enzyme Kinetics , pp. 606-623, John Wiley & Sons, Inc., New York |
35. | Hofmann, K. P., and Heck, M. (1996) in Biomembranes II (Lee, A. G., ed) , pp. 141-197, Jai Press Inc., Greenwich, CT |
36. | Schleicher, A., and Hofmann, K. P. (1987) J. Membr. Biol. 95, 271-281[Medline] [Order article via Infotrieve] |
37. | Bruckert, F., Vuong, T. M., and Chabre, M. (1988) Eur. Biophys. J. 16, 207-218[Medline] [Order article via Infotrieve] |
38. | Bruckert, F., Chabre, M., and Vuong, T. M. (1992) Biophys. J. 63, 616-629[Abstract] |
39. | Liebman, P. A., and Pugh, E. N., Jr. (1979) Vision Res. 19, 375-380[Medline] [Order article via Infotrieve] |
40. |
Baehr, W.,
Morita, E. A.,
Swanson, R. J.,
and Applebury, M. L.
(1982)
J. Biol. Chem.
257,
6452-6460 |
41. | Liebman, P. A., and Sitaramayya, A. (1984) Adv. Cyclic Nucleotide Protein Phosphorylation Res. 17, 215-224[Medline] [Order article via Infotrieve] |
42. | Klebe, C., Prinz, H., Wittinghofer, A., and Goody, R. S. (1995) Biochemistry 34, 12543-12552[Medline] [Order article via Infotrieve] |
43. | Rensland, H., John, J., Linke, R., Simon, I., Schlichting, I., Wittinghofer, A., and Goody, R. S. (1995) Biochemistry 34, 593-599[Medline] [Order article via Infotrieve] |
44. | Rodnina, M. V., Savelsbergh, A., Katunin, V. I., and Wintermeyer, W. (1997) Nature 385, 37-41[CrossRef][Medline] [Order article via Infotrieve] |
45. | Emeis, D., Kühn, H., Reichert, J., and Hofmann, K. P. (1982) FEBS Lett. 143, 29-34[CrossRef][Medline] [Order article via Infotrieve] |
46. | Bornancin, F., Pfister, C., and Chabre, M. (1989) Eur. J. Biochem. 184, 687-698[Abstract] |
47. | Bennett, N., and Dupont, Y. (1985) J. Biol. Chem. 260, 4156-4168[Abstract] |
48. | Parkes, J. H., Gibson, S. K., and Liebman, P. A. (1999) Biochemistry 38, 6862-6878[CrossRef][Medline] [Order article via Infotrieve] |
49. | Ernst, O. P., Bieri, C., Vogel, H., and Hofmann, K. P. (2000) Methods Enzymol. 315, 471-489[Medline] [Order article via Infotrieve] |
50. | Kohl, B., and Hofmann, K. P. (1987) Biophys. J. 52, 271-277[Abstract] |
51. | Leskov, I. B., Klenchin, V. A., Handy, J. W., Whitlock, G. G., Govardovskii, V. I., Bownds, M. D., Lamb, T. D., Pugh, E. N., Jr., and Arshavsky, V. Y. (2000) Neuron 27, 525-537[Medline] [Order article via Infotrieve] |
52. | Vuong, T. M., Chabre, M., and Stryer, L. (1984) Nature 311, 659-661[Medline] [Order article via Infotrieve] |
53. | Kahlert, M., and Hofmann, K. P. (1991) Biophys. J. 59, 375-386[Abstract] |
54. | Gutfreund, H. (1995) Kinetics for the Life Sciences , p. 237, Cambridge University Press, Cambridge, UK |
55. | Hofmann, K. P. (1986) Photobiochem. Photobiophys. 13, 309-327 |
56. | Parkes, J. H., and Liebman, P. A. (1984) Biochemistry 23, 5054-5061[Medline] [Order article via Infotrieve] |