Recipient of the American Heart Association Established Investigator award. To whom correspondence should be addressed: Pharmacology Dept., University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75390-9041. Tel.: 214-648-8581
* This work was supported in part by grants from the R. A. Welch Foundation and National Institutes of Health Grants GM31278 (to J. R. F.) and DK47890 (to T. M. W.).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.
Regulators of G protein signaling (RGS proteins) are GTPase-activating proteins (GAPs) for Gi and/or Gq class G protein α subunits. RGS GAP activity is inhibited by phosphatidylinositol 3,4,5-trisphosphate (PIP3) but not by other lipid phosphoinositides or diacylglycerol. Both the negatively charged head group and long chain fatty acids (C16) are required for binding and inhibition of GAP activity. Amino acid substitutions in helix 5 within the RGS domain of RGS4 reduce binding affinity and inhibition by PIP3 but do not affect inhibition of GAP activity by palmitoylation. Conversely, the GAP activity of a palmitoylation-resistant mutant RGS4 is inhibited by PIP3. Calmodulin binds all RGS proteins we tested in a Ca2+-dependent manner but does not directly affect GAP activity. Indeed, Ca2+/calmodulin binds a complex of RGS4 and a transition state analog of Gαi1-GDP-AlF4−. Ca2+/calmodulin reverses PIP3-mediated but not palmitoylation-mediated inhibition of GAP activity. Ca2+/calmodulin competition with PIP3 may provide an intracellular mechanism for feedback regulation of Ca2+ signaling evoked by G protein-coupled agonists.
Heterotrimeric G proteins link 7-transmembrane domain receptors to intracellular effector proteins in all mammalian cell types. These pathways are required for rapid responses to hormones and neurotransmitters. Signaling is initiated by agonist binding to receptors, which catalyze the exchange of GTP for GDP on the Gα subunit. Activated Gα-GTP and Gβγ subunits then regulate effector proteins that generate second messengers and subsequent downstream responses (reviewed in Ref.
). The intensity, duration, and specificity of G protein-mediated signaling depend on a newly identified class of proteins, termed regulators of G protein signaling (RGS1 proteins), which are GTPase-activating proteins (GAPs) for Gα subunits (Refs.
), could bind RGS4 and regulate its activity. We report that calmodulin binds in a Ca2+-dependent manner to all RGS proteins we tested, including RGS1, RGS2, RGS4, RGS10, RGS16, and GAIP. Surprisingly, Ca2+/calmodulin binding did not influence the GAP activity of RGS proteins.
Calmodulin regulates membrane association of many intracellular proteins (
) that could bind acidic lipids in the plasma membrane. Here we demonstrate that dipalmitoylphosphatidylinositol 3,4,5-trisphosphate (diC16-PIP3 or PIP3) binds RGS4 and inhibits its GAP activity in a concentration-dependent manner, in contrast to other phosphatidylinositol phosphates. PIP3inhibited GAP activity of other RGS proteins, including RGS1, RGS2, RGS10 and GAIP, but not RGS16. Amino acid substitutions in helix 5 of RGS4 reduced PIP3 binding (10-fold) and PIP3-dependent inhibition of GAP activity. A potential mechanism for feedback regulation of G protein-mediated Ca2+ signaling was suggested by the observation that PIP3-dependent inhibition of GAP activity was reversed by Ca2+/calmodulin. The concerted action of Ca2+/calmodulin and PIP3 may regulate RGS GAP activity to initiate [Ca2+]i oscillations evoked by G protein-coupled agonists.
MATERIALS AND METHODS
Calmodulin, Peptides, and Inositol Lipids
Bovine brain calmodulin and calmodulin covalently attached to agarose beads (CaM-agarose) were from Sigma. Peptide P1–33, MCKGLAGLPASCLRSAKDMKHRLGFLLQKSDSC, was described (
), and a scrambled sequence of P1–33, MLDQAPLKSKSACAGKLRMHGCLSFGLRLKCSD, ψP1–33, was synthesized by C. Slaughter (UT Southwestern). l-α-Phosphatidylcholine (PC),l-α-phosphatidylserine (PS),l-α-phosphatidyl-sn-glycerol from egg yolk lecithin (PG), l-α-phosphatidylinositol 4,5-bisphosphate (PIP2), l-α-phosphatidylinositol 4-phosphate (PIP), l-α-phosphatidylinositol (PI), all from bovine brain, d-myo-inositol 1,3,4,5-tetrakisphosphate (IP4), and 1-stearoyl-2-arachidonoyl-sn-glycerol (DAG) were purchased from Sigma.l-α-Phosphatidylethanolamine (PE) from bovine liver was purchased from Avanti Polar Lipids. Dihexadecanoylphosphatidylinositol 3,4,5-trisphosphate (diC16-PIP3), diC16-3,5-AP2dioctanoylphosphatidylinositol 3,4,5-trisphosphate (diC8-PIP3), dioctanoylphosphatidylinositol 3,4-bisphosphate (diC8–3,4-PIP2), dioctanoylphosphatidylinositol 3,5-bisphosphate (diC8–3,5-PIP2), and dioctanoylphosphatidylinositol 4,5-bisphosphate (diC8–4,5-PIP2) were synthesized as described (
). The fragment coding for mutant RGS4 K112E/K113E was generated by polymerase chain reaction using the oligonucleotide containing the above mutation as a primer and cloned into the pQE60 expression vector (Qiagen) as described for RGS4 (
). Concentration of RGS4 was measured using Bradford reagent from Bio-Rad. Band shift gel analysis of calmodulin binding to RGS4 and RGS2 in 4 m urea, in the presence of 0.1 mmCa2+ or 2 mm EDTA, was carried out as described (
). RGS binding to calmodulin-agarose was detected by SDS-PAGE of supernatants as follows: 15 μl of wet calmodulin-agarose beads (26 μg of CaM) washed with buffer (10 mm HEPES, pH 7.4, 0.1 mm CaCl2, 1 mm DTT) were pelleted; the supernatant was removed, and the beads were mixed with RGS4 (1.5 nmol) equal in amount to coupled calmodulin in a final volume 30 μl of buffer. The suspension was incubated at room temperature with constant agitation. After 30 min, the beads were washed twice with 500 μl of buffer. The suspension was brought to the original volume and incubated for 10 min. 500 mm EGTA and 5 m NaCl were added to final concentrations 2 and 50 mm, respectively, and the suspension was incubated for additional 10 min. Finally, 5 μl of the SDS-PAGE loading buffer was added to the beads and incubated another 10 min. Equal volume aliquots of supernatant were withdrawn after each incubation step, followed by separation by SDS-PAGE and Coomassie Blue staining.
Fluorescent detection of RGS4-calmodulin interaction was performed using Perkin-Elmer LS 50B and Hitachi F-2000 fluorescent spectrophotometers at 25 ± 0.1 °C. Both excitation and emission slit widths were 10 nm. For internal RGS4 tryptophan fluorescence measurements, the excitation and the emission wavelengths were 283 and 336 nm, respectively. Typically, aliquots of CaM solution were gradually added to the solution of RGS4 in the assay cuvette with constant agitation, and after each addition the mixture was equilibrated inside the instrument with the excitation beam shutter closed. The fluorescence measurements were made after 5–15 min. The excitation beam shutter was opened only long enough to get accurate readings. Measurements were repeated until reproducible readings were obtained. Data in Fig. 2 were corrected for dilution (which did not exceed 7%) and for fluorescence of calmodulin alone.
) and extensively dialyzed against 10 mm HEPES, pH 7.4. Fluorescence measurements of dansylated calmodulin (dansyl-CaM) were at 335 (excitation) and 500 nm (emission). All measurements were corrected for background fluorescence observed in control experiments. The Scatchard plot analysis of the fluorescence affinity measurement data indicated a single binding site interaction (or 2 sites with similar affinities) and fit well to linear approximations within the experimental error.
RGS GAP Assays
GAP assays were carried out in a soluble single turnover system with Gαi1 as described (
) with minor modifications. Briefly, 250–500 nmGαi1 in the assay buffer (10 mm HEPES, pH 8.0, 5 mm EDTA, 2 mm DTT) was loaded with [γ-32P]GTP at 30 °C for 20 min. The solution was placed on ice for 5 min, and all subsequent reactions were carried out on ice in a cold room. A drop of RGS protein (5–10 μl) was placed onto a wall of reaction tube next to a 5-μl drop of solution, 500 mm MgCl2, 5 mm cold GTP, and the GAP reaction was started by addition of 175 μl of Gαi1-GTP solution. The final concentration of free Mg2+ ions in the reaction mix was 3–4 mm. To study the influence of Ca2+/calmodulin on RGS4 GAP activity with Gαi1-GTP, RGS4 was preincubated with calmodulin in 10 mm HEPES, pH 8.0, 1 mm CaCl2, 2 mm DTT on ice for 20 min. Then 175 μl of Gαi1 loaded with [γ-32P]GTP was added with 5 μl of 500 mm GTP, 475 mmMgCl2, 25 mm CaCl2.
To prepare small unilamellar lipid vesicles (SUVs), chloroform-soluble lipids were evaporated under a stream of nitrogen into a dry film, and resuspended by vortexing in a sonication buffer, 10 mm HEPES, pH 8.0, 0.1 mm EDTA, 2 mm DTT, essentially as described (
). SUVs were generated by sonication of a lipid suspension (2 mg/ml total lipid) in a water bath at room temperature for 10–15 min. To prepare SUVs containing chloroform-insoluble diC16-PIP3, the suspension of diC16-PIP3 in the sonication buffer (10 mg/ml) was added to the dry film of other lipids, vortexed vigorously for 1 min, and sonicated after addition of the appropriate amount of the sonication buffer. Aggregated material was removed from the preparations of SUVs by centrifugation at 10,000 × g for 10 min. The micelles of diC16-PIP3 were obtained by its brief sonication in a buffer solution (
Surface plasmon resonance measurements were performed using BIAcore 1000 instrument (BIAcore, Inc.) at 25 °C. RGS protein was immobilized on the surface of the carboxymethylated dextran chip (CA-5) using standard carbodiimide chemistry in accordance with manufacturer's instructions. Lipid dissolved in the running buffer, 10 mm HEPES, pH 8.0, 150 mm NaCl, 3 mm EDTA, 0.005% surfactant NP20, was injected over the chip surface with a flow rate 5 μl/min. At least three different concentrations were used. Control injections were made with a blank chip without coupled RGS. The amount of coupled protein was 2500 (RGS4), 1580 (RGS4 K112E/K113E), and 2900 response units (RU) (RGS16). The regeneration of the chip after each binding experiment was achieved by injecting 0.01% SDS in running buffer. The data were analyzed using the BIAevaluation 2.1 software (BIAcore, Inc.). The binding curves showed a moderate heterogeneity of both association and dissociation phases. The minor components identified in the analysis of association and dissociation (less than 20% of total signal) were ignored. A relatively fast transition process evident at the beginning of the dissociation phase was not studied. The dissociation constant, Kd, was calculated for each sensorgram using kinetic constants ka andkd. The average values are shown in Table IV.
Table IVRGS-PIP3 kinetic and affinity constants
10 3 m −1 s −1
10 −4 s −1
12.1 ± 7.5
4.8 ± 1.6
44 ± 19 (n = 5)
3.2 ± 1.4
12.6 ± 1.1
430 ± 140 (n = 4)
5.2 ± 2.5
7.9 ± 2.7
163 ± 38 (n = 3)
Binding constants (± S.D.) were determined with the BIAcore at 25 °C.
To investigate the role of Ca2+ in feedback regulation of G protein signaling by RGS proteins, we characterized two potential calmodulin binding regions in RGS4 as follows: one in the N-terminal 33 amino acids and another between residues 99–113, in helixes 4 and 5 of the RGS domain (4Box). These regions contain amphiphatic sequences with bulky hydrophobic residues at certain positions and clusters of positively charged amino acids similar to calmodulin-binding sites in other proteins (Table I; see Ref.
). RGS4 binding to Ca2+/calmodulin was corroborated using calmodulin coupled to agarose beads. RGS4 binding to calmodulin-agarose beads required Ca2+ and was stable to buffer washes, but bound RGS4 could be eluted from beads either with EGTA or SDS (Fig.1C). We found that a previously characterized amphipathic calmodulin-binding peptide from CaM kinase II (
) competed with RGS4 binding to calmodulin-agarose beads (Fig. 1E, 4th and5th lanes). An RGS4 N-terminal 33 amino acid peptide (P1–33), which conveys high affinity and receptor-selective regulation of Gq signaling (
), bound Ca2+/calmodulin (Fig. 1D) and competed with RGS4 for binding to calmodulin-agarose beads (Fig. 1E, 6th and7th lanes). By contrast, a scrambled sequence composed of the same amino acids (ψP1–33) did not bind to calmodulin beads (data not shown) and did not compete with RGS4 binding (Fig. 1E, 8th and 9th lanes). Interestingly, RGS4 apparently formed a heterotrimeric complex with Gαi1-GDP-AlF4− and Ca2+/calmodulin (Fig. 1F), consistent with their predicted distinct binding sites on RGS4.
Table IPutative calmodulin binding regions in RGS domain proteins
(97) EYKKIKSPSKLSPKAKKI (114)
(125) ELKAEANQHVVDEKARLI (142)
(107) DYKKTES-DLLPCKAEEI (123)
(118) DFKKTKSPQKLSSKARKI (135)
(429) DFKKVKSQSKMASKAKKI (446)
(68) DFKKMQDKTQMQEKAKEI (85)
(99) EFKKIRSATKLASRAHHI (116)
The amino acid residues used are: −, negative; +, positive; h, hydrophobic; X, any; -, gap in alignment.
We used fluorescence spectroscopy to quantitate binding interactions between RGS4 and Ca2+/calmodulin. The fluorescence of two tryptophan residues within the RGS domain of RGS4 (4Box) was quenched by titration with Ca2+/calmodulin (which lacks tryptophan), indicative of RGS4-Ca2+/calmodulin binding in solution (Fig. 2). A sharp inflection in the titration curve indicates formation of a stable, equimolar complex between Ca2+/calmodulin and 4Box (20 mm NaCl, 10 mm HEPES, pH 7.4). Upon further addition of Ca2+/calmodulin to 4Box, the slope of the fluorescence titration curve paralleled that of unbound RGS4 and Ca2+/calmodulin (calculated as the sum of their fluorescence signals measured separately). This behavior is consistent with the prediction of a single Ca2+/calmodulin-binding site in the RGS domain. By contrast, in the absence of salt, the fluorescence intensity of RGS4 (Fig. 2) or 4Box (data not shown) did not change at higher molar ratios of Ca2+/calmodulin, suggesting additional, low affinity calmodulin-binding sites on RGS4. These low affinity interactions are probably electrostatic because they were not detected at higher ionic strength (Fig. 2, 4Box,and data not shown).
In addition to the RGS domain, Ca2+/calmodulin apparently binds to the N-terminal 33 amino acids of RGS4 because the peptide P1–33 binds in a Ca2+-dependent manner both to calmodulin beads (Fig. 1) and in solution, as detected by two-dimensional heteronuclear single quantum coherence NMR using [15N]calmodulin.
) which allowed us to compare the binding properties of P1–33 and RGS4 in similar conditions. The apparent Kd values (TableII) were calculated from Scatchard linear transformations of titration curves (Fig.3). In the presence of 100 mmNaCl, RGS4 and P1–33 bind to Ca2+/dansyl-CaM with similar affinities (Kd ≈5 μm). The affinity of P1–33 binding within the accuracy of measurements did not change with ionic strength, whereas RGS4 bound Ca2+/dansyl-CaM almost 5 times stronger than P1–33 in 20 mm NaCl, consistent with our observations that a decrease in salt concentration strengthened the interaction of RGS4 with Ca2+/calmodulin beads.
Table IIAffinity (μm) of RGS4-Ca 2+ /CaM interaction (25 °C)
Calmodulin binds to many RGS proteins in a Ca2+-dependent manner but with different salt dependences. Binding assays indicated that RGS4, RGS16, and GAIP had similar affinities toward calmodulin-agarose beads in 20 mmNaCl, whereas RGS10 interaction with Ca2+/calmodulin was relatively weak (even without salt). Only RGS1 and RGS2 bound to calmodulin-agarose beads in high salt (150 mm KCl). Stable interaction was corroborated by band shift analysis that revealed an RGS2-Ca2+/calmodulin complex in high ionic strength running buffer with 4 m urea, 275 mm Tris-HCl, pH 8.3 (data not shown).
Calmodulin Does Not Influence RGS4 GAP Activity
To test the biological relevance of Ca2+/calmodulin binding to RGS proteins, we studied its effect on RGS4 and RGS1 GAP activity. We found that neither 1 mm Ca2+ alone nor 1.6 μm Ca2+/calmodulin preincubated with either RGS protein altered their GAP activity toward Gαi1 in a single turnover assay (Fig. 4 and data not shown). Because the N terminus of RGS4 is not required for GAP activity (
) but binds calmodulin (Fig. 1, D andE), we also tested GAP activity of an N-terminal deletion mutant that lacked the first 57 residues of RGS4 but retained the calmodulin-binding site within the RGS domain (R4ΔN; Fig. 4B). The GAP activities of full-length RSG4 and R4ΔN were unaffected by Ca2+/calmodulin in the single turnover assay (Fig. 4). These results indicated that Ca2+/calmodulin binding neither sterically blocked Gαi1 binding nor irreversibly changed the conformation of the RGS4-Gαi1 interface. Considering the stability of RGS4 binding to Ca2+/calmodulin beads (Fig. 1), it seems improbable that GAP activity resulted from the rapid and transient displacement Ca2+/calmodulin from RGS4 by Gαi-GTP. Indeed, we found that Gαi1-GDP-AlF4−, which mimics a transition state of RGS4-catalyzed Gα-GTP hydrolysis (Kd = 0.6 nm at 25 °C, Ref.
), binds as a complex with RGS4 and Ca2+/calmodulin-agarose beads (Fig. 1E). This complex was stable to extensive washing with buffer, but RGS4-Gαi1-GDP-AlF4− was eluted from Ca2+/calmodulin beads by 1% SDS. Gαi1-GDP-AlF4− did not bind to Ca2+/calmodulin in the absence of RGS4. These results indicate that a heterodimeric complex of Ca2+/calmodulin-RGS4 retains GAP activity. We therefore favor a model in which calmodulin binds surface residues of the RGS domain without substantially altering the RGS4 conformation.
PIP3 Inhibits RGS GAP Activity
Previous studies indicated that brief dialysis of recombinant RGS4 into patch clamped pancreatic acinar cells potently inhibited Ca2+ signaling evoked by Gi- and Gq-coupled receptor agonists (
). This suggested the possibility that endogenous RGS proteins might be relatively inactive prior to agonist stimulation of Ca2+ signaling and that recombinant RGS proteins escaped this inhibition. To identify inhibitors of RGS4 GAP activity on Gαi1 in the single turnover assay, we tested various compounds related to and including either the substrate or products of PLCβ. We found that RGS4 GAP activity was inhibited by preincubation with an analog of PIP3, diC16-PIP3 (30 μm PIP3, at 0 °C for several minutes, Fig.5A), or phosphocholine vesicles containing 20% PIP3 (Fig. 5B). Inhibition of RGS4 GAP activity was dependent on the concentration of PIP3 (Fig. 5C). The kinetic curve of GTP hydrolysis in the presence of PIP3 closely approximated the basal activity of Gα without RGS4. This behavior indicated negligible dissociation of PIP3 from RGS4 during the assay (5 min) because GTP hydrolysis was initiated by a rapid 20-fold dilution of the RGS-PIP3 incubation mix into a solution containing Gαi1-GTP. In control experiments, the intrinsic GTPase activity of Gαi1 was unaffected by PIP3 (Fig.5D). The GAP activity of each RGS protein that was tested in the single turnover assay, except RGS16, was inhibited by PIP3 (Fig. 5D and data not shown).
By contrast to PIP3 inhibition of RGS GAP activity, no effect was observed following coincubation of RGS4 with 400 μm PIP2 from bovine brain (Fig.5B), and only 2–3-fold inhibition was observed following coincubation with PIP2 micelles (9 mm). We barely detected the inhibitory activity of 4-mono phosphorylated phosphatidylinositol phosphate, PIP (9 mm). As summarized in Table III, several synthetic PIP2 lipids were also without effect (assayed at 400 μm in phosphocholine vesicles), including dioctanoylphosphatidylinositol 3,4-bisphosphate (diC8–3,4-PIP2), dioctanoylphosphatidylinositol 3,5-bisphosphate (diC8–3,5-PIP2), and dioctanoylphosphatidylinositol 4,5-bisphosphate (diC8–4,5-PIP2). RGS4 GAP activity was also not affected by a lipid head group derivative, 1-stearoyl-2-arachidonoyl-sn-glycerol (diacylglycerol, DAG), which is one of the reaction products of PLCβ. No inhibitory activity was detected using PI or highly charged head group derivatives of PIP3 lacking the fatty acyl moieties, including inositol 1,3,4,5-tetrakisphosphate (IP4) and glycerophosphoinositol 3,4,5-trisphosphate (IP3, the other reaction product of PLCβ; data not shown). PIP3 binding to Rac1 and RhoA was shown to have similar requirements for both electrostatic and hydrophobic interactions (
). Surprisingly, we found that dioctanoylphosphatidylinositol 3,4,5-trisphosphate (diC8-PIP3), which only differed from PIP3(diC16-PIP3) in the length of the fatty acid chains, did not inhibit RGS4 GAP activity; nor did diC16-3,5-PIP2. Thus, diC16-PIP3 is the only phospholipid which inhibited RGS4 GAP activity, and its activity appears to require both the long chain fatty acid moiety and the highly charged head group.
Table IIIPIP3 charge density and fatty acid chain length required to inhibit RGS4 GAP activity
Protein-lipid binding affinities were estimated by surface plasmon resonance measurements on the BIAcore (Biacore, Inc). RGS4 coupled to the carboxymethylated dextran surface of the BIAcore chip bound diC16-PIP3 (Fig.6A) but not diC8-PIP3, PIP2, or other phosphoinositides, consistent with the observation that RGS GAP activity was most sensitive to inhibition by diC16-PIP3. The association phase of diC16-PIP3 binding was typically complete within several minutes, whereas dissociation was slow (Fig. 6A). No diC16-PIP3 binding was detected on a blank chip. TheKd value of diC16-PIP3 binding to RGS4 (44 ± 19 nm; Table IV) was calculated from the on and off rates extracted from the binding curves (Fig. 6A, and data not shown).
Ca2+/CaM Antagonizes the PIP3 Inhibition of RGS4 GAP Activity
The positively charged patch on the surface of helixes 4 and 5 in the RGS domain of RGS4 (residues 99–113; Ref.
) appeared to be a good candidate for binding not only to calmodulin but also to PIP3. We found that PIP3 inhibition of RGS4 GAP activity was reversed by coincubation of RGS4 and PIP3 (micelles or phosphocholine vesicles) with Ca2+/calmodulin (Fig. 7). Ca2+/calmodulin and PIP3 apparently compete for binding to helixes 4 and 5 in RGS4. To test this model further, we introduced amino acid substitutions of glutamate for lysine residues at positions 112 and 113 in helix 5 of RGS4. This mutant protein (RGS4 K112E/K113E) retained GAP activity and calmodulin binding, but it bound PIP3 with almost 10-fold lower affinity than did wild type RGS4 (Table IV). Similarly, low binding affinity was observed with RGS16. The comparatively weak binding of these proteins to PIP3 correlated with their relative insensitivity to PIP3 inhibition of GAP activity (Figs. 5D and6B). Helixes 4 and 5 are conserved in many RGS domains (Table I) and may provide an important regulatory feature of RGS proteins because Ca2+/calmodulin has been shown to serve as a molecular switch that regulates lipid-protein interactions (
PIP3 Inhibits GAP Activity of Palmitoylation-resistant RGS Proteins
We propose that PIP3-mediated inhibition of GAP activity acts by a concerted mechanism in which the highly charged head group interacts with the RGS domain residues in helixes 4 and/or 5 to position the palmitoyl moiety of PIP3 near its binding site. Palmitoylation of a nearby cysteine residue inhibited RGS4 and RGS10 GAP activity (
). Substitution of cysteine for valine (C95V) prevents covalent modification by palmitate at this position in RGS4. GAP activity of the C95V mutant protein is equivalent to wild type protein, but it is not inhibited by palmitoylation (
). By contrast, the GAP activity of RGS4 C95V is as sensitive to PIP3-mediated inhibition as is wild type protein (Fig. 8A). The GAP activities of RGS16 and the mutant RGS4 K112E/K113E, which were relatively insensitive to inhibition by PIP3, responded like wild type protein to inhibition by palmitoylation (Fig. 8,B and C). In contrast to RGS4 interaction with PIP3, inhibition of GAP activity by palmitoylation was not prevented by addition of Ca2+/calmodulin (Fig.8D). This is presumably because Ca2+/calmodulin competes with PIP3 binding to RGS4 but cannot displace the covalent modification of RGS4 by palmitate. The feedback mechanisms that regulate the palmitoylation of RGS proteins in vivo are unknown, but we propose that PIP3-mediated inhibition of RGS GAP activity may be reversed in a Ca2+-dependent manner through binding of Ca2+/calmodulin.
Feedback Regulation of RGS GAP Activity by Ca2+/Calmodulin and PIP3
Negative regulation of RGS GAP activity by PIP3 may be part of a reset mechanism that allows a new wave of G protein signaling in response to agonist. G protein-coupled receptors, such as formyl-methionyl-leucyl-phenylalanyl receptors in neutrophils, can elicit both Ca2+-release and a rapid and large accumulation of PIP3 by activating the effector proteins PLCβ and PI-3 kinase, respectively (reviewed in Ref.
). Ca2+ release from internal stores is one of the initial responses to PLCβ activation either by Gαq or Gβγ from Giclass G proteins. As the local concentration of Ca2+elevates in response to PLCβ activity, we postulate that Ca2+/calmodulin binding to RGS4 and other RGS proteins displaces PIP3 to restore GAP activity (modeled in Fig.9). Ca2+/calmodulin binding may also enhance GAP activity by relocating RGS proteins within the receptor signaling complex to be in proximity to their Gα-GTP substrates. Feedback inhibition of G protein-mediated PLCβ activation by RGS proteins would allow [Ca2+]i to decrease and promote the dissociation of calmodulin from RGS proteins. We hypothesize that PIP3 released after receptor stimulation could then bind RGS proteins to inhibit their GAP activity. If agonist stimulation persists, this may reactivate G protein signaling and allow another burst of Ca2+ release from internal stores. If agonist is no longer present, PIP3-mediated inhibition of RGS GAP activity may reset the signaling pathway to allow a robust cellular response to subsequent agonist stimulation. Feedback regulation of RGS GAP activity may provide an intracellular mechanism to initiate oscillations over a wide range of frequencies.
We thank Y. Tu for RGS4C95V protein; K. Chapman for [γ-32P]GTP; J. Rizo-Rey, K. Luby-Phelps, I. Fernandez, and C. Wigley for their contributions to NMR analysis of the Ca2+/calmodulin-RGS4 complex; C. Slaughter for peptide synthesis; and E. Ross, S. Muallem, and colleagues for discussions and comments on the manuscript.