Rapid Kinetics of Regulator of G-protein Signaling (RGS)-mediated Gαi and Gαo Deactivation

Regulator of G-protein signaling (RGS) proteins accelerate GTP hydrolysis by Gα subunits speeding deactivation. Gα deactivation kinetics mediated by RGS are too fast to be directly studied using conventional radiochemical methods. We describe a stopped-flow spectroscopic approach to visualize these rapid kinetics by measuring the intrinsic tryptophan fluorescence decrease of Gα accompanying GTP hydrolysis and Gα deactivation on the millisecond time scale. Basal k cat values for Gαo, Gαi1, and Gαi2 at 20 °C were similar (0.025–0.033 s−1). GlutathioneS-transferase fusion proteins containing RGS4 and an RGS7 box domain (amino acids 305–453) enhanced the rate of Gα deactivation in a manner linear with RGS concentration. RGS4-stimulated rates could be measured up to 5 s−1 at 3 μm, giving a catalytic efficiency of 1.7–2.8 × 106 m −1 s−1 for all three Gα subunits. In contrast, RGS7 showed catalytic efficiencies of 0.44, 0.10, and 0.02 × 106 m −1s−1 toward Gαo, Gαi2, and Gαi1, respectively. Thus RGS7 is a weaker GTPase activating protein than RGS4 toward all Gα subunits tested, but it is specific for Gαo over Gαi1 or Gαi2. Furthermore, the specificity of RGS7 for Gαo does not depend on N- or C-terminal extensions or a Gβ5 subunit but resides in the RGS domain itself.

G-protein 1 -coupled receptor-mediated signal transduction governs many important physiological functions. Upon binding of a ligand, such as light, neurotransmitter, hormone, chemokine, etc., to heptahelical receptors, the heterotrimeric G-proteins composed of ␣, ␤, and ␥ subunits are stimulated to release GDP and bind GTP. In the GTP-bound form, G␣ dissociates from G␤␥ and both interact with downstream effectors. This pathway is terminated when G␣ hydrolyzes the bound GTP, thereby promoting reassociation of G␣ and G␤␥ and returning the system to inactive state (reviewed in Refs. [1][2][3]. Members of the recently described family of proteins, Regulators of G-protein signaling (RGS), act as negative regulators of G-protein function by enhancing GTP hydrolysis by G␣ (4 -6) or by functioning as effector antagonists (7). Although the first RGS protein, Sst2p, was identified in yeast (8,9), more than 20 variants have been found in mammalian species, all being characterized by a conserved RGS domain of approximately 120 amino acids (reviewed in Refs. 10 -12). The GTPase-activating property (GAP) of RGS proteins has been demonstrated by direct in vitro biochemical studies (13)(14)(15). Further studies using GDP-AlF 4 Ϫ suggest that RGS proteins enhance GTP hydrolysis by stabilizing the transition state conformation of G␣ (16), which leads to the crystal structure of the RGS4⅐G␣ i1 complex (17). Mutagenesis analyses have also illustrated the importance of specific residues on the interface between G␣ and RGS proteins (18 -21). Besides being GAPs, many RGS proteins have also been found to serve as links to other cellular signaling pathways through non-RGS domains such as GGL, DEP, DH/PH, and PDZ domains (reviewed in Refs. 11 and 12). The wide expression of RGS in various tissue and cell types (see Ref. 12 for review) thus indicate that RGS proteins may play important physiological roles directly or indirectly.
The specificity of RGS proteins for interactions with G␣ subtypes (G␣ s , G␣ i /G␣ o , G␣ q , and G␣ 12 ) is now being vigorously explored. The majority of the RGS proteins are GAPs for G␣ i and G␣ q but not for G␣ s subunit (reviewed in Refs. 10,[22][23]. Numerous studies on the specificity between RGS proteins and G␣ subunits have been reported using biochemical, immunochemical, and functional methods. RGS4 is one of the most extensively studied RGS protein and is a highly effective GAP for all G␣ i /G␣ o family members (reviewed in Refs. 10 -11, 22, 24 -26). In contrast, limited and apparently inconsistent data on RGS7 have been reported. The RGS domain of RGS7 has been shown to have effective GAP activity toward G␣ i1 and G␣ o (21,27), whereas it preferentially bound G␣ o , G␣ i3 and G␣z, but not G␣ i1 or G␣ i2 (28). However, accurate quantitation of GAP activity is difficult, because rates of GTP hydrolysis are too fast to measure with standard assays (see below). Furthermore, full-length RGS7 was recently found to form a complex with G␤ 5 (29,30), and this complex was shown to have moderate but significant GAP activity toward G␣ o but not G␣ i1 or G␣ i2 (31), despite the fact that it did not seem to favor the physical binding of RGS7 to G␣ o (32). Therefore, it is not clear whether the specificity of full-length RGS7 for G␣ o over G␣ i1 and G␣ i2 is encoded in the RGS domain or it depends on G␤ 5 .
Direct measurement of the kinetics of GTP hydrolysis and deactivation of G␣ mediated by RGS would be the most straightforward approach to determining specificity. The current challenge in this regard is that the conventional biochem-ical assays can only delineate kinetics as fast as t1 ⁄2 ϳ 10 s, whereas the RGS-mediated G␣ deactivation, as well as the turn-off of many physiological processes, takes place on the subsecond scale. One recent study in a m1AchR-Gq vesicle system using a quench-flow method has reported more than a 1000-fold enhancement of GTP hydrolysis rate mediated by RGS4 with t1 ⁄2 ϳ 50 ms, indicating that data acquisition on the millisecond time scale is required to study the effect of RGS on G␣ deactivation kinetics under physiological conditions (33).
The present study aimed to develop a stopped-flow spectroscopic approach to visualize these rapid kinetics and to quantitatively assess RGS⅐G␣ specificity. The rapid-mix stoppedflow florescence spectroscopy utilizes the intrinsic tryptophan fluorescence of G␣. Intrinsic tryptophan fluorescence is high in GTP-bound, active G␣ subunits and low for GDP-bound G␣ (34 -37). The intrinsic fluorescence change is due to alterations in the environment of the ␣2 helix tryptophan residue, Trp 212 in G␣ o (38), and Trp 207 in transducin (39). With purified G␣ o , G␣ i1 , and G␣ i2 proteins in solution, we successfully employed this stopped-flow method to quantitate catalytic efficiencies of RGS4 and RGS7 and obtained new evidence on the specificity of RGS4 and RGS7 (box domain) toward G␣ subunits.
Purification of His 6 -tagged G␣ o and G␣ i1 and Myristoylated G␣ i1 and G␣ i2 Proteins-His 6 G␣ o and His 6 G␣ i1 were expressed from the vector, pQE60, in Escherichia coli strain BL21/DE3 and purified as described previously (21). His 6 G␣ o and His 6 G␣ i1 eluted from nickel resin columns (Qiagen, Santa Clara, CA) were approximately 80% pure by Coomassie Blue staining of SDS gels. Myristoylated G␣ i1 and G␣ i2 were expressed from the pQE6 expression vector cotransformed in the E. coli strain BL21/DE3 with the N-myristoyl transferase vector based on the protocol of Mumby and Linder (40). The specific activities of the G␣ proteins determined by [ 35 S]GTP␥S binding assay (41) ranged from 6 to 18 pmol/g.
Purification of GST-RGS Fusion Proteins-The GST-RGS fusion proteins were expressed and purified using PGEX expression vectors in E. coli strain BL21/DE3 as described previously (21). The bacterial supernatant was incubated with glutathione-agarose beads (Amersham Pharmacia Biotech) at 4°C overnight. After washing with ice-cold PBS buffer (pH 7.3), the GST-RGS fusion proteins were eluted with 15 mM glutathione in PBS and dialyzed against PBS buffer. Where indicated, the fusion proteins were cleaved by incubation overnight at 4°C with 10 units of thrombin/mg of fusion protein followed by incubation with glutathione-agarose to remove GST and any uncleaved GST-RGS proteins. Cleavage of the GST to yield RGS4 alone did not alter the rate constants for G␣ deactivation (data not shown). RGS proteins were approximately 60 -90% pure by Coomassie Blue staining of SDS gels.
Slow Time-based Fluorescence Measurements-Slow time-based fluorescence measurements were determined using a PTI Alphascan fluorometer (Photon Technology, Monmouth Junction, NJ) with a watercooled 150-watt xenon arc lamp as described (42). Stopped-flow G␣ Deactivation Kinetics-The rapid kinetics of G␣ deactivation in the presence or absence of RGS proteins was measured in an Applied Photophysics DX-17MV stopped-flow spectrofluorometer.
The intrinsic fluorescence change of G␣ proteins was measured at an excitation wavelength of 290 nm with 2.3-nm slits. Emission light was detected with a photomultiplier tube behind a WG320 band pass filter (Corion, Holliston, MA). G␣ o (400 nM), G␣ i1 (600 nM), or G␣ i2 (600 nM) proteins were preloaded with 2 M GTP in the absence of magnesium in HED buffer (50 mM Hepes, 5 mM EDTA, 2 mM DTT, pH 8.0). Incubations were for 20 min at 20°C for G␣ o and for 15 min at 30°C for G␣ i1 and G␣ i2 . Samples were then stored on ice until use (Ͻ2 h). GTP-loaded G-protein was equilibrated in the instrument at 20°C for at least 3 min, then samples were mixed 1:1 with magnesium-containing HEDM buffer (50 mM Hepes, 5 mM EDTA, 2 mM DTT, 30 mM MgSO 4 , pH 8.0) in the absence or presence of RGS proteins as indicated. Tryptophan fluorescence was recorded for at least 100 s in the absence of RGS and for 20 or 50 s in the presence of RGS, depending on the rates achieved. Data from four to six shots were averaged and fit to a one-phase exponential decay equation using Prism v.3.0 for Windows (GraphPad Software, San Diego, CA).
Determination of Single Turnover k cat for Hydrolysis of [␥-32 P]GTP-Single turnover [␥-32 P]GTPase assays at low temperature (4°C) were performed as described previously (21). Single turnover [␥-32 P]GTP hydrolysis was also determined at room temperature (24°C) as below. His 6 G␣ o or His 6 G␣ i1 (10 nM) was preloaded in magnesium-free HEDL buffer (50 mM Hepes, 1 mM EDTA, 1 mM DTT, 20 ppm deionized Lubrol, pH 8.0) with 1 M [␥-32 P]GTP (7500 cpm/pmol). The preloading incubations were for 20 min at room temperature for G␣ o and for 15 min at 30°C for G␣ i1 . After equilibration at room temperature, the single turnover hydrolysis reaction was then started by addition of MgSO 4 and GTP␥S to final concentrations of 20 mM and 200 M, respectively. MgSO 4 activates His 6 G␣ o and triggers catalysis, while GTP␥S prevents [␥-32 P]GTP from rebinding to the G-protein. Aliquots (50 l) were taken and diluted in 1 ml of 15% (w/v) charcoal solution (50 mM NaH 2 PO 4 , pH 2.3, 0°C) at the indicated time points. Background hydrolysis was determined in the absence of protein and represents less than 10% of total [␥-32 P]P i release. The amount of [␥-32 P]P i released at each time point was fit to an exponential function, cpm(t) ϭ cpm o ϩ ⌬cpm ϫ (1 Ϫ e Ϫkt ).

Slow Time-based Fluorescence
Measurements-To determine the feasibility of using fluorescence spectroscopy to study RGS function, we first examined the intrinsic fluorescence changes of G␣ o upon addition of GTP. The time course of intrinsic fluorescence changes of G␣ o (500 nM) was monitored using a slow time-based fluorometer before and after addition of 2 M GTP in the presence of 10 mM Mg 2ϩ (Fig. 1A). Similar to the findings reported previously (34,36), GTP and Mg 2ϩ caused a rapid increase in the fluorescence intensity followed by a slow decrease in fluorescence. Fluorescence approached the baseline level at approximately 30 min (1800 s). The fluorescence increase represents GTP binding to G␣ o to induce an active conformation. As the bound GTP is hydrolyzed and GDP produced, an increasing percentage of G␣ o becomes GDP-bound, which displays a lower intrinsic fluorescence. When a small amount of RGS4 (3 nM) was added with GTP and Mg 2ϩ , the peak of intrinsic fluorescence decreased and the time for the fluorescence to return to baseline was shortened. A more dramatic effect was observed when a higher concentration of RGS4 (30 nM) was added.
Because our stopped-flow approach required preloading of G␣ with GTP, we also tested the effect of adding RGS4 at the peak of the intrinsic fluorescence increase (Fig. 1B). RGS4 markedly accelerated the rate of fluorescence decrease.
Stopped-flow Measurements of G␣ Deactivation Kinetics-Most previous studies of RGS-stimulated GTP hydrolysis by G-proteins have been done at 0 -4°C to slow the single turnover kinetics (13,15,43). Although low temperature allows the measurement of the k cat of basal GTPase, which had rates of 0.002 and 0.004 s Ϫ1 for G␣ o and G␣ i1 , respectively, the RGS4enhanced k cat was still too fast to be measured (21). The intrinsic GTPase activity of His 6 G␣ o ( Fig. 2A) and His 6 G␣ i1 (not shown), as determined by [␥-32 P]GTP single turnover assays at room temperature, were 0.019 Ϯ 0.002 s Ϫ1 (n ϭ 5) and 0.028 Ϯ 0.002 s Ϫ1 (n ϭ 5), respectively, corresponding to half-times of 36 and 25 s. This is almost as fast as can be measured by manual methods, making the kinetics accelerated by RGS virtually inaccessible.
We therefore tested the feasibility of using stopped-flow methods to measure the intrinsic fluorescence decrease of GTPpreloaded G␣ upon addition of Mg 2ϩ . This depends on GTP hydrolysis by the G␣ subunit (34). Both His 6 G␣ o (Fig. 2B) and His 6 G␣ i1 (Fig. 2C) showed rapid Mg 2ϩ -induced fluorescence decreases. The rate constants were 0.022 s Ϫ1 for His 6 G␣ o , and 0.027 s Ϫ1 for His 6 G␣ i1 , corresponding to half-times (t1 ⁄2 ) of 32 and 25 s, respectively. These rates of fluorescence decrease were in excellent agreement with the rates of GTP hydrolysis (k cat ) as determined by single turnover [ 32 P]GTPase assays described above. Similar results were obtained for myrG␣ o , myrG␣ i1 , and myrG␣ i2 (graphs not shown), and the rates are summarized in Table I.
We further tested the effect of RGS on the rate of fluorescence decrease. In the presence of 100 nM RGS4, the rate of G␣ i1 deactivation increased nearly 10-fold to 0.26 Ϯ 0.03 s Ϫ1 (t1 ⁄2 , 2.7 s), whereas the magnitude of the fluorescence change was not affected (Fig. 3A). To be certain that the fluorescence decrease depended on the GTP hydrolysis and G␣ deactivation rather than mere binding of RGS4 to G␣, the following controls were performed. His 6 G␣ i1 was preincubated with GDP, GDP-AlF 4 Ϫ , or Gpp(NH)p, a non-hydrolyzable GTP analog, in the absence of Mg 2ϩ followed by addition of Mg 2ϩ with or without 100 nM RGS4. None of these reactions led to detectable fluorescence changes (Fig. 3A). Thus it was not simply binding of RGS4 to either the active form (Gpp(NH)p) or the transition state (GDP-AlF 4 Ϫ ) of G␣ that produced the fluorescence signal.  Furthermore, the RGS-insensitive mutant G183S G␣ i1 (21) was utilized to test whether or not the effect of RGS4 depended on G␣/RGS4 interactions seen in biochemical studies. As expected, RGS4 did not increase the rate of G␣ deactivation for G183S G␣ i1 (without RGS4, 0.027 Ϯ 0.00, n ϭ 2; with 100 nM RGS4, 0.013 Ϯ 0.001, n ϭ 2) compared with that for wild type His 6 G␣ (Fig. 3B). This result is in agreement with our previous biochemical measurements at 4°C (21). Therefore, the intrinsic fluorescence changes were dependent on and reflected the kinetics of GTP hydrolysis and G␣ deactivation. To simplify discussion, we will use the designation k deact throughout the text to represent the rate of intrinsic fluorescence change that represents the rate of GTP hydrolysis and G␣ deactivation kinetics.
To try to determine the maximal rate V max and the K m value of the RGS-facilitated G␣ deactivation, we used increasing concentrations of RGS4 (Fig. 4A). The rate of G␣ deactivation increased linearly with the concentration of RGS4 up to 3 M, as shown in Fig. 4C. Due to the deteriorating signal-to-noise ratio contributed by intrinsic fluorescence of the high concentration of RGS4, 3 M was the highest concentration of RGS4 for which acceptable data could be obtained. Similarly, the linear relationship between k deact and [RGS] was also found for RGS7 (Fig. 4, B and D) and RGS8 (Fig. 4C) as well as for the three RGS proteins toward His 6 G␣ i1 (Fig. 4, E and F).
Specificity of RGS4 and RGS7 toward G␣ i/o -Given that we cannot separately define a V max and a K m for the RGS-mediated enhancement of GTP hydrolysis by G␣, we used the slope of the plot of k deact versus [RGS] as a measure of the "catalytic efficiency" of that RGS toward particular G␣ subunits. This is equivalent to a k cat /K m in classical enzyme mechanisms. The catalytic efficiencies of RGS4 and RGS8 toward His 6 G␣ o and His 6 G␣ i1 were similar, being ϳ2 ϫ 10 6 M Ϫ1 s Ϫ1 (Fig. 4 and Table II). In contrast, the catalytic efficiency of the RGS7 box construct toward His 6 G␣ o was approximately eight times lower than those of RGS4 and RGS8, being 0.27 Ϯ 0.01 ϫ 10 6 M Ϫ1 s Ϫ1 (Fig. 4D, note scale change, and Table II). More surprisingly, very little effect was seen for RGS7 toward His 6 G␣ i1 , yet we were still able to detect a linear relationship of rate versus RGS7 concentration (Fig. 4F). The catalytic efficiency of RGS7 toward His 6 G␣ i1 was 0.03 Ϯ 0.01 ϫ10 6 M Ϫ1 s Ϫ1 , nine times lower than that toward His 6 G␣ o and nearly 60 times lower than that of RGS4 toward His 6 G␣ i1 (Table II). This clarifies the confusion in the literature and demonstrates that the G␣ specificity of RGS7 resides in the RGS box itself and doesn't depend on complex formation with G␤ 5 . Furthermore, we examined whether myristoylation of the G␣ subunits altered the kinetics or specificity of the RGS interactions. Comparable catalytic efficiencies were obtained for both RGS4 and RGS7 toward myrG␣ compared with the results for His 6 G␣ (Fig. 5 and Table The rate of intrinsic fluorescence decrease (or G␣ deactivation) was obtained by fitting the data to a single exponential decay curve. A and B are each from a single experiment representing a total of four and seven determinations, respectively. C-F, plots of rates as determined in A and B versus RGS concentrations. Linearity was shown for RGS4 up to 3 M and similarly for RGS7 and RGS8 up to 1 M for both His 6 G␣ o (C, D) and His 6 G␣ i1 (E, F). All lines shown are the linear fits of averaged data points at various RGS concentrations from at least three experiments (except for RGS8, n ϭ 2). Error bars representing S.E. are too small to be visible. The slopes of all lines are summarized in Table II. II). Thus the myristoylation of G␣ did not affect the activity of RGS. A similar but less pronounced difference in catalytic activity was also seen between RGS4 and RGS7 toward myrG␣ i2 . To summarize these results, RGS4 was essentially equally active on all G␣ subunits tested with a catalytic efficiency of ϳ2 ϫ 10 6 M Ϫ1 s Ϫ1 . RGS7 had lower activity overall, however, it was specific for G␣ o , being ϳ10 -30 times more active on G␣ o than on G␣ i1 and six times more active on G␣ o than on G␣ i2 (Fig. 5B and Table II).
To ensure that the specificity seen above by stopped-flow was not an artifact of the spectroscopic method, we used the standard single turnover [ 32 P]GTPase assay to check RGS4 and RGS7 (27,31). As shown in Fig. 6, His 6 G␣ o and His 6 G␣ i1 alone hydrolyzed GTP at similar rates at 4°C (k cat 0.004 Ϯ 0.0002 s Ϫ1 and 0.007 Ϯ 0.0008 s Ϫ1 , respectively, n ϭ 3). RGS4 at a concentration of 50 nM increased these rates about 10-fold (k cat 0.08 Ϯ 0.01 s Ϫ1 and 0.07 Ϯ 0.01 s Ϫ1 , respectively, n ϭ 3). RGS7, at a concentration of 100 nM, produced more modest increases (k cat 0.017 Ϯ 0.003 s Ϫ1 and 0.010 Ϯ 0.003 s Ϫ1 , n ϭ 3). We made qualitative estimates of catalytic efficiency from these single point data, and, as summarized in Table II, the results are consistent with the more extensive and quantitative data obtained by stopped-flow fluorimetry.

DISCUSSION
The recently identified RGS proteins are responsible for the fast physiological turnoff of G-protein-mediated signaling by accelerating the GTPase activity of G␣ (10,23,44,45). Conventional biochemical assays to determine single turnover rates are not sufficiently fast to study the subsecond kinetics of GTP hydrolysis in the presence of RGS proteins. We describe here a novel approach, stopped-flow fluorimetry, that permits the quantitative study of the effects of RGS proteins on G␣ on the millisecond time scale.
We were able to quantify the rates of G␣ i1 and G␣ o deacti-vation up to 5 s Ϫ1 (t1 ⁄2 140 ms) in the presence of micromolar concentrations of RGS4. This is approximately five times faster than the deactivation of muscarinic receptor-activated potassium channels in the heart (46). To put these results in context, Popov et al. (43) also found a linear relationship between the k cat of G␣ i1 and the concentration of RGS using single turnover [ 32 P]GTPase assay at 0°C but only studied the kinetics up to 60 nM RGS4. Their calculated catalytic efficiency of 0.9 ϫ 10 6  6. GAP effect of RGS4 and RGS7 as determined by single turnover [ 32 P]GTPase assays. Experiments were performed at 4°C using 100 nM His 6 G␣ o (A-C) or His 6 G␣ i1 (D-F) in the absence or presence of RGS4 and RGS7 as described previously (21). A and D, no RGS; B and E, 50 nM RGS4; C and F, 100 nM RGS7. Data were fit with one phase exponential curve, and the rates were used to calculate a 2-point slope k cat /[RGS] similar to those in Figs. 4 and 5. The slopes are reported in Table II. Data in each graph are single determinations from one experiment representative of three. M Ϫ1 s Ϫ1 was similar to our results (Table II). Because our rates of G␣ deactivation showed a linear relationship with the concentration of RGS4, RGS7, and RGS8 up to 1-3 M, we were unable to detect a K m or V max for G␣ deactivation with any of the RGS proteins. The RGS4 data were best, and their effects on G␣ o and G␣ i indicate that the K m and V max are even higher than those predicted by Berman et al. based on extrapolation of manual kinetics studies (2.5 M and 14 s Ϫ1 ) (16). Recently, Mukhopadhyay and Ross (33) used a quench-flow method to determine a rate constant of ϳ20 s Ϫ1 for RGS4-stimulated G␣ q single turnover GTP hydrolysis. Interestingly, they reported an EC 50 of 200 -300 nM for RGS4 enhancement of G␣ q -mediated steady-state GTP hydrolysis. They saw no further increase above 1 M RGS probably due to the rate-limiting release of GDP, whereas we saw clear increases up to 3 M.
Our data on the basal rates of fluorescence decrease (20°C) using the stopped-flow method are in excellent agreement with the rates of GTP hydrolysis (k cat ) as determined by single turnover [ 32 P]GTPase assays (24°C) (Fig. 2). It is thus tempting to compare the measurements of catalytic efficiency using the two different methods. The temperature dependence revealed in our results is quite interesting. Although the unstimulated GTPase rates of G␣ subunits are reduced approximately 10-fold upon reducing the temperature from 20°C to 4°C (0.02-0.03 s Ϫ1 to 0.002-0.004 s Ϫ1 ), there is little temperature dependence of the second order rate constant for RGS catalytic efficiency. For RGS4 we observed only a 15-25% lower slope of k deact /[RGS] at 20°C versus 4°C. Similarly, the value reported by Popov (0.9 ϫ10 6 M Ϫ1 s Ϫ1 ) at 0°C is within a factor of two of our results at 20°C. It has been proposed that RGS proteins enhance GTP hydrolysis by G␣ by stabilizing the transition state conformation of G␣ (16). The lack of temperature dependence indicated in our results supports the suggestion that there may be no large conformational change of the G-protein upon RGS binding and that RGS simply binds to those G-proteins already in the transition state conformation and stabilizes them. Therefore, the enhanced rates are largely dependent on the collision between RGS and G␣ in the transition state conformation, as suggested in the "asparagine knuckle" hypothesis (17). Thus diffusion might be the ratelimiting step, and a plateau of the G␣ deactivation rate may not be observed until extremely high concentrations of RGS protein are reached. This stopped-flow method, however, opens up a window of RGS-mediated fast G␣ deactivation kinetics beyond those available with the conventional approaches.
Structural Basis of Specificity of RGS4 and RGS7 toward G␣ o -The quantitative data obtained with this rapid mix method provides a more accurate picture of RGS⅐G␣ specificity. We previously reported that 100 nM RGS4 had k cat values of Ͼ5 min Ϫ1 toward G␣ i1 (21), which is consistent with our present results, shown in Table II with a k cat value of 14 min Ϫ1 . The catalytic efficiency of RGS4 for all three G␣ i/o subunits is quite comparable.
We have also demonstrated the specificity of the RGS7 box domain (amino acids 305-453) for G␣ o over G␣ i using both stopped-flow fluorimetry and single turnover [ 32 P]GTPase methods (Table II). These data on the specificity of RGS7 are largely consistent with and extend other studies previously reported. These include the preferential binding of an RGS7 box domain (amino acids 303-470) to G␣ o over G␣ i1 or G␣ i2 (28) and the moderate GAP activity of RGS7⅐G␤ 5 complex (compared with RGS4) toward G␣ o but not G␣ i1 , or G␣ i2 (31). Our results, in comparison, provide quantitative measurements on the specificity of RGS7 using accurate k cat values, especially for RGS at higher concentrations. In contrast, Shuey et al. (27), reported that the RGS7 box (amino acids 317-474) was active toward both G␣ o and G␣ i1 . This conclusion, however, was only qualitative, because they did see slower k cat for G␣ i1 than for G␣ o . The high concentration of RGS7 they used (800 nM) makes the estimation of rates in the manual assay very difficult. The apparent difference between the results of Shuey et al. and those reported here does not seem to be due to the different RGS7 region they expressed, because a shorter version of our RGS7 construct (amino acids 327-452) still retained specificity for G␣ o over G␣ i (data not shown). Thus the specificity of RGS7 GAP activity for G␣ o over G i1 and G i2 resides in the RGS box domain itself and does not require the full-length RGS7 or a complex with the ␤ 5 subunit.
In summary, we show results from a novel stopped-flow method to characterize the rapid G␣-RGS interactions. Our data are the first to report the effects of RGS proteins on G␣ o or G␣ i on the millisecond time scale. The rates of G␣ deactivation observed here are clearly fast enough to account for the physiological turning off of ion channels. Furthermore, using this approach, we quantitatively evaluate the specificity of G␣-RGS interactions. It is also clear that the mechanism of RGS-mediated G␣ GAP activity does not require substantial, slow conformation changes in the structure of the G␣ subunit.