A Model for Agonism and Antagonism in an Ancient and Ubiquitous cAMP-binding Domain
2006; Elsevier BV; Volume: 282; Issue: 1 Linguagem: Inglês
10.1074/jbc.m607706200
ISSN1083-351X
Autores Tópico(s)Protein Kinase Regulation and GTPase Signaling
ResumoThe cAMP-binding domain (CBD) is an ancient and conserved regulatory motif that allosterically modulates the function of a group of diverse proteins, thereby translating the cAMP signal into a controlled biological response. The main receptor for cAMP in mammals is the ubiquitous regulatory (R) subunit of protein kinase A. Despite the recognized significant potential for pharmacological applications of CBDs, currently only one group of competitive inhibitor antagonists is known: the (Rp)-cAMPS family of phosphorothioate cAMP analogs, in which the equatorial exocyclic oxygen of cAMP is replaced by sulfur. It is also known that the diastereoisomer (Sp)-cAMPS with opposite phosphorous chirality is a cAMP agonist, but the molecular mechanism of action of these analogs is currently not fully understood. Previous crystallographic and unfolding investigations point to the enhanced CBD dynamics as a key determinant of antagonism. Here, we investigate the (Rp)- and (Sp)-cAMPS-bound states of R(CBD-A) using a comparative NMR approach that reveals a clear chemical shift and dynamic NMR signature, differentiating the (Sp)-cAMPS agonist from the (Rp)-cAMPS antagonist. Based on these data, we have proposed a model for the (Rp/Sp)-cAMPS antagonism and agonism in terms of steric and electronic effects on two main allosteric relay sites, Ile163 and Asp170, respectively, affecting the stability of a ternary inhibitory complex formed by the effector ligand, the regulatory and the catalytic subunits of protein kinase A. The proposed model not only rationalizes the existing data on the phosphorothioate analogs, but it will also facilitate the design of novel cAMP antagonists and agonists. The cAMP-binding domain (CBD) is an ancient and conserved regulatory motif that allosterically modulates the function of a group of diverse proteins, thereby translating the cAMP signal into a controlled biological response. The main receptor for cAMP in mammals is the ubiquitous regulatory (R) subunit of protein kinase A. Despite the recognized significant potential for pharmacological applications of CBDs, currently only one group of competitive inhibitor antagonists is known: the (Rp)-cAMPS family of phosphorothioate cAMP analogs, in which the equatorial exocyclic oxygen of cAMP is replaced by sulfur. It is also known that the diastereoisomer (Sp)-cAMPS with opposite phosphorous chirality is a cAMP agonist, but the molecular mechanism of action of these analogs is currently not fully understood. Previous crystallographic and unfolding investigations point to the enhanced CBD dynamics as a key determinant of antagonism. Here, we investigate the (Rp)- and (Sp)-cAMPS-bound states of R(CBD-A) using a comparative NMR approach that reveals a clear chemical shift and dynamic NMR signature, differentiating the (Sp)-cAMPS agonist from the (Rp)-cAMPS antagonist. Based on these data, we have proposed a model for the (Rp/Sp)-cAMPS antagonism and agonism in terms of steric and electronic effects on two main allosteric relay sites, Ile163 and Asp170, respectively, affecting the stability of a ternary inhibitory complex formed by the effector ligand, the regulatory and the catalytic subunits of protein kinase A. The proposed model not only rationalizes the existing data on the phosphorothioate analogs, but it will also facilitate the design of novel cAMP antagonists and agonists. The cAMP-binding domain (CBD) 2The abbreviations used are: CBD, cAMP-binding domain; C, catalytic subunit of PKA; cAMP, 3′,5′-cyclic ester of adenosine monophosphate; cAMPS, phosphorothioate analog of cAMP; CPMG, Carr-Purcell-Meiboom-Gill pulse sequence; EPAC, guanine exchange factor for the Ras-like small GTPases Rap1 and Rap2; NOE, nuclear Overhauser effect; PBC, phosphate-binding cassette; PKA, cAMP-dependent protein kinase; R, regulatory subunit of PKA; RIα, isoform Iα of the R-subunit of PKA; R(CBD-A), CBD A of RIα; RC, relaxation-compensated; CT, constant time; MES, 4-morpholineethanesulfonic acid. 2The abbreviations used are: CBD, cAMP-binding domain; C, catalytic subunit of PKA; cAMP, 3′,5′-cyclic ester of adenosine monophosphate; cAMPS, phosphorothioate analog of cAMP; CPMG, Carr-Purcell-Meiboom-Gill pulse sequence; EPAC, guanine exchange factor for the Ras-like small GTPases Rap1 and Rap2; NOE, nuclear Overhauser effect; PBC, phosphate-binding cassette; PKA, cAMP-dependent protein kinase; R, regulatory subunit of PKA; RIα, isoform Iα of the R-subunit of PKA; R(CBD-A), CBD A of RIα; RC, relaxation-compensated; CT, constant time; MES, 4-morpholineethanesulfonic acid. represents a conserved regulatory motif that modulates the function of a diverse group of proteins, including protein kinase A (PKA) in eukaryotes (1Berman H.M. 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Sci. 2002; 968: 139-147Crossref PubMed Scopus (90) Google Scholar, 23Taylor S.S. Kim C. Vigil D. Haste N.M. Yang J. Wu J. Anand G.S. Biochim. Biophys. Acta. 2005; 175: 25-37Crossref Scopus (197) Google Scholar). For instance, for PKA, which is the major receptor of cAMP in vertebrates, over the past ∼40 years, hundreds of compounds were tested as competitive inhibitors of cAMP for therapeutic purposes. However, only one group of cAMP surrogates, the phosphorothioate (Rp)-cAMPS analogs (Fig. 1b), was found to have an antagonist function (24Van Haaster P.M. Van Driel R. Jastorff B. Baraniak J. Stec W.C. De Wit J.W. J. Biol. Chem. 1985; 259: 10020-10024Abstract Full Text PDF Google Scholar, 25Dostmann W.R.G. Taylor S.S. Genieser H.G. Jastorff B. Doskeland S.O. Ogreid D. J. Biol. Chem. 1990; 265: 10484-10491Abstract Full Text PDF PubMed Google Scholar, 26Dostmann W.R.G. Taylor S.S. Biochemistry. 1991; 30: 8710-8716Crossref PubMed Scopus (73) Google Scholar). In (Rp)-cAMPS, the equatorial exocyclic oxygen of cAMP is replaced by a sulfur atom, thus introducing chirality at the phosphorous position. Switching to the opposite chirality by placing the sulfur atom in the axial exocyclic position leads to the related diastereomeric cAMP analog: (Sp)-cAMPS (Fig. 1a). Unlike (Rp)-cAMPS, (Sp)-cAMPS behaves as a cAMP agonist, revealing the stringent stereo-specific requirements of antagonism. However, it is currently not fully understood in molecular terms why the (Rp)- and (Sp)-cAMPS analogs function as a cAMP antagonist and agonist, respectively. The crystal structures of the regulatory subunit of PKA (R) bound to both phosphorothioate analogs have been solved at 2.3 Å resolution and are overall very similar to each other (i.e. Cα root mean square deviation of 0.34 Å) and to the cAMP-bound state (i.e. Cα root mean square deviation of 0.5 Å) as well (27Wu J. Jones M.J. Xuong N-H. Eyck L.F.T. Taylor S.S. Biochemistry. 2004; 43: 6620-6629Crossref PubMed Scopus (68) Google Scholar). Despite the overall similarity of these three structures, subtle local conformational differences were found for the conserved arginines in the PBC (i.e. Arg209 for CBD-A and Arg333 for CBD-B). For example, the intermolecular distances indicate that the phosph(othio)ate-Arg209 guanidinium interaction is significantly tighter for (Rp)-cAMPS as compared with (Sp)-cAMPS and cAMP, although the affinity of the (Rp)-surrogate for this CBD is lower than that of the (Sp)-analog and of cAMP (27Wu J. Jones M.J. Xuong N-H. Eyck L.F.T. Taylor S.S. Biochemistry. 2004; 43: 6620-6629Crossref PubMed Scopus (68) Google Scholar). In addition, the crystallographic investigation revealed that the (Rp)-cAMPS bound state is probably more dynamic than the (Sp)-cAMPS- and cAMP-bound forms, based on its higher B-factors (27Wu J. Jones M.J. Xuong N-H. Eyck L.F.T. Taylor S.S. Biochemistry. 2004; 43: 6620-6629Crossref PubMed Scopus (68) Google Scholar). It has also been shown that the Gibbs free energy of urea unfolding of (Rp)-cAMPS-bound R is similar to that of cAMP-free R, whereas the thermodynamics of unfolding for (Sp)-cAMPS compares well with that of the more thermodynamically stable cAMP-bound R (28Dostmann W.R.G. FEBS Lett. 1995; 375: 231-234Crossref PubMed Scopus (63) Google Scholar). Here, we further investigate by NMR the effects of (Rp)- and (Sp)-cAMPS binding. For this purpose, we focus on the 119-244 RIα fragment, which maps to CBD-A of the R-subunit of PKA and represents its minimal central controlling unit (29Das R. Abu-Abed M. Melacini G. J. Am. Chem. Soc. 2006; 128: 8406-8407Crossref PubMed Scopus (36) Google Scholar). Using a comparative NMR strategy based on Nz exchange spectroscopy to assign different bound states, it was possible to unveil how the internal signaling pathways of CBD-A are differentially perturbed by (Rp)-cAMPS as compared with (Sp)-cAMPS and cAMP. In addition, 15N relaxation measurements (i.e. T1,2, HN NOE, and relaxation-compensated constant time CPMG NMR dispersion) combined with hydrodynamic simulations provide further insight on the sites and time scales of the enhanced dynamics caused by (Rp)-cAMPS. The interpretation of these results in the context of the allosteric networks of CBD-A and of the stereo-electronic effects caused by the oxygen-to-sulfur isolobal substitution has led to the proposition of a molecular model for the antagonism and agonism of the (Rp)- and (Sp)-cAMPS analogs, respectively. NMR Sample Preparation of RIα-(119-244) Bound to Different Ligands—Samples of uniformly 15N-labeled cAMP-bound RIα-(119-244) in MES buffer (50 mm MES, pH 6.5, 100 mm NaCl, and 0.02% NaN3) were prepared as previously discussed (29Das R. Abu-Abed M. Melacini G. J. Am. Chem. Soc. 2006; 128: 8406-8407Crossref PubMed Scopus (36) Google Scholar, 30Hamuro Y. Anand G. Kim J.S. Juliano C. Stranz D.D. Taylor S.S. Woods V.L. J. Mol. Biol. 2004; 304: 1185-1196Crossref Scopus (84) Google Scholar). The RIα-(119-244) samples with substoichiometric amounts of cAMP used for the Nz exchange experiments were obtained through a protocol of unfolding, partial dialyzing out of cAMP and refolding. Specifically, after adding 8 m urea to cAMP-bound RIα-(119-244), the sample was dialyzed for 18 h in the presence of 0.5 liters of MES buffer with 6 m urea and 1 mm dithiothreitol. After changing the dialysis buffer three times at regular intervals, the protein was then partially refolded by stepwise dilution of the dialysis solution to 0.5 m urea through the addition of MES buffer with 1 mm dithiothreitol. Essentially complete removal of urea was obtained by a final dialysis against the MES buffer, which resulted in a sample with NMR-detectable amounts of both cAMP-bound and free states of RIα-(119-244) at a total concentration of ∼0.1 mm. The addition of 50 μm (Sp)-cAMPS phosphorothioate cAMP analog (Sigma) to this sample resulted in another NMR sample with NMR-observable amounts of both cAMP-bound and (Sp)-cAMPS-bound RIα-(119-244). This sample was used to assign the (Sp)-cAMPS-bound RIα-(119-244) through Nz exchange spectroscopy. The sample with both cAMP-bound and (Rp)-cAMPS-bound (Sigma) RIα-(119-244) was prepared similarly. Samples containing only RIα-(119-244) bound to the (Sp)-cAMPS or (Rp)-cAMPS analogs were prepared by extensively rather than partially dialyzing the protein under denaturing conditions and then refolding. Specifically, we added urea to 6 ml of 0.15 mm RIα-(119-244) to a final urea concentration of 8 m. After keeping the resulting solution at room temperature for 30 min, it was dialyzed against 8 m urea buffer (50 mm MES, pH 6.5, 100 mm NaCl, and 1 mm dithiothreitol) for 16 h with three buffer changes. The protein solution was then dialyzed extensively with 6 m urea for 24 h to completely remove cAMP. After refolding the protein as indicated above, 1 mm excess (Sp)-cAMPS analog was added. The (Rp)-cAMPS-bound RIα-(119-244) sample was prepared likewise. NMR Spectroscopy—All NMR data were acquired using a Bruker AV 700-MHz spectrometer equipped with a TCI cryoprobe set at a temperature of 306 K. The calibration of the probe temperature was obtained using a thermocouple as well as an ethylene glycol sample. Unless otherwise specified, all spectral widths, number of digitization points, carrier frequencies, 15N GARP decoupling strength, and gradient shapes were set as previously discussed (29Das R. Abu-Abed M. Melacini G. J. Am. Chem. Soc. 2006; 128: 8406-8407Crossref PubMed Scopus (36) Google Scholar). Spectra were processed using the Xwinnmr (Bruker Inc.) or NMRPipe (31Delaglio F. Grzesiek S. Vuister G.W. Zhu G. Pfeifer J. Bax A. J. Biomol. NMR. 1991; 1: 439Crossref PubMed Scopus (96) Google Scholar) program based on previously published standard protocols (29Das R. Abu-Abed M. Melacini G. J. Am. Chem. Soc. 2006; 128: 8406-8407Crossref PubMed Scopus (36) Google Scholar, 30Hamuro Y. Anand G. Kim J.S. Juliano C. Stranz D.D. Taylor S.S. Woods V.L. J. Mol. Biol. 2004; 304: 1185-1196Crossref Scopus (84) Google Scholar, 31Delaglio F. Grzesiek S. Vuister G.W. Zhu G. Pfeifer J. 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NMR Spectrom. 1999; 34: 93-158Abstract Full Text Full Text PDF Scopus (1386) Google Scholar). Nz Exchange—The assignment of the backbone amides of RIα-(119-244) bound to (Sp)-cAMPS was obtained from that of cAMP-bound RIα-(119-244) through Nz exchange spectra (36Farrow N.A. Zhang O. Forman-Kay J.D. Kay L.E. J. Biomol. NMR. 1994; 4: 727-734Crossref PubMed Scopus (387) Google Scholar, 37Vialle-Printems C. Heijenoort C. Guittet E. J. Mag. Reson. 2000; 142: 276-279Crossref PubMed Scopus (9) Google Scholar, 38Rodriguez J.C. Jennings P.A. Melacini G. J. Biomol. NMR. 2004; 30: 155-161Crossref PubMed Scopus (12) Google Scholar) acquired for a sample with NMR-observable amounts of both cAMP-bound and (Sp)-cAMPS-bound RIα-(119-244). Similarly, the backbone amides of RIα (119-244) bound to (Rp)-cAMPS were assigned through Nz exchange spectra of a sample with NMR observable amounts of both cAMP-bound and (Rp)-cAMPS-bound RIα-(119-244). In both cases, the Nz mixing period was 230 ms, and the relaxation delay between subsequent scans was 2 s. After acquisition of 128 scans per t1 transient, the Nz exchange spectra were processed using linear prediction. The ( ((ΔδHN1)2+(ΔδNH15/6.5)2)) equation (39Mulder F.A. Schipper D. Bott R. Boelens R. J. Mol. Biol. 1999; 292: 111-123Crossref PubMed Scopus (220) Google Scholar) was used to compute the compounded 1H,15N chemical shift variations between the different states of RIα-(119-244) (i.e. X-bound versus free and X-bound versus cAMP-bound, where X = (Sp)- or (Rp)-cAMPS). Relaxation Dispersion NMR—A relaxation-compensated constant time (RC-CT) CPMG pulse sequence (40Tollinger M. Skrynnikov N.R. Mulder F.A.A. Forman-Kay J.D. Kay L.E. J. Am. Chem. Soc. 2001; 123: 11341-11352Crossref PubMed Scopus (416) Google Scholar, 41Mulder F.A.A. Skrynnikov N.R. Hon B. Dahlquist F.W. Kay L.E. J. Am. Chem. Soc. 2001; 123: 967-975Crossref PubMed Scopus (273) Google Scholar, 42Palmer III, A.G. 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In addition, considering that at 700 MHz and at the CPMG radio frequency strength employed (3.1 kHz), 15N pulses are affected by significant offset effects, all CT-RC-CPMG experiments were acquired with three different 15N carrier frequencies (110, 119, and 127 ppm) to span the 15N spectral width using three narrow bands. The NMR relaxation dispersion ΔR2eff was calculated as follows: ΔR2eff=R2eff(43Hz)-R2eff(472Hz). Since R2eff(νCPMG)=(-1/TCP)ln(IνCPMG/Io) (40Tollinger M. Skrynnikov N.R. Mulder F.A.A. Forman-Kay J.D. Kay L.E. J. Am. Chem. Soc. 2001; 123: 11341-11352Crossref PubMed Scopus (416) Google Scholar, 41Mulder F.A.A. Skrynnikov N.R. Hon B. Dahlquist F.W. Kay L.E. J. Am. Chem. Soc. 2001; 123: 967-975Crossref PubMed Scopus (273) Google Scholar), where IψCPMG and Io are the cross-peak intensities with and without the CPMG periods, respectively, the equation for ΔR2eff can be recast as follows: ΔR2eff=(1/Tcp)ln(I472Hz/I43Hz) (29Das R. Abu-Abed M. Melacini G. J. Am. Chem. 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The relaxation rates and their errors were computed through relaxation fitting simulations with Sparky 3.111 (50Ding Z. Lee G. Liang X. Gallazzi F. Arunima A. Van Doren S.R. Biochemistry. 2005; 44: 10119-10134Crossref PubMed Scopus (20) Google Scholar), using 1000 iterations with a Gaussian distributed random noise added. The uncertainty in the steady state NOE was measured based on the S.D. value of the distribution of the differences in fit heights between duplicate spectra (34Farrow N.A. Muhandiram R. Singer A.U. Pascal S.M. Kay C.M. Gish G. Shoelson S.E. Pawson T. Forman-Kay J.D. Kay L.E. Biochemistry. 1994; 33: 5984Crossref PubMed Scopus (2012) Google Scholar). All errors were treated as previously explained (51Kroenke C.D. Rance M. Palmer III, A.G. J. Am. Chem. Soc. 1999; 121: 10119-10125Crossref Scopus (126) Google Scholar). Selected cross-peaks could not be used in the relaxation analysis due to line broadening and/or overlap. 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Under this assumption, the J(ωN + ωH) and J(ωN) values are derived only from the measured 15NR1 and {1H}15N NOEs, whereas the J(0) values are computed using the measured 15NR2 rates as well (55Lefèvre J.F. Dayie K.T. Peng J.W. Wagner G. Biochemistry. 1996; 35: 2674-2686Crossref PubMed Scopus (233) Google Scholar). The calculated value of J(0) includes also chemical exchange contributions. Errors in the reduced spectral densities were assessed through propagation of the uncertainties in the experimentally determined 15N relaxation rates and NOEs. For reference purposes, the Lorentzian spectral density of an isotropically diffusing rigid molecule was computed as follows: J(ω) = (2/5)τc/(1 + (ωτc)2). Hydrodynamic Simulations—The contributions of the overall tumbling and the effect of diffusional anisotropy on the relaxation rates and the reduced spectral densities were modeled through bead method-based hydrodynamic simulations using the HYDRONMR program (56Bernado P. de la Torre J.G. Pons M. J. Biomol. NMR. 2002; 23: 139-150Crossref PubMed Scopus (76) Google Scholar, 57Garcìa de la Torre J. Biophys. Chem. 2001; 93: 159-170Crossref PubMed Scopus (79) Google Scholar). For this purpose, the coordinates for the 119-244 fragment of the 1RGS Protein Data Bank structure of RIα were employed with hydrogen atoms added through the program Molmol (58Koradi R. Billeter M. Wüthrich K. J. Mol. Graphics. 1996; 14: 51-55Crossref PubMed Scopus (6487) Google Scholar) and with an atomic element radius of 3.3 Å, which represents the optimal average value that has been previously found to best fit several hydrodynamic properties (i.e. translational diffusion, sedimentation coefficients, rotational diffusion, and intrinsic viscosity) for a set of model proteins (57Garcìa de la Torre J. Biophys. Chem. 2001; 93: 159-170Crossref PubMed Scopus (79) Google Scholar). An error of ±0.2 Å was considered for the atomic element radius to account for hydration layer variability. A temperature of 306 K was used for the HYDRONMR simulation, and the viscosity of water in centipoises at this temperature was computed as follows (57Garcìa de la Torre J. Biophys. Chem. 2001; 93: 159-170Crossref PubMed Scopus (79) Google Scholar): η = 1.7753-0.0565t + 1.0751 × 10-3 t2 - 9.2222 × 10-6 t3, where t is the temperature in Celsius. The 15N relaxation rates at a static field of 16.44 T computed by HYDRONMR for the rigid RIα-(119-244) assume an N-H distance of 1.02 Å and a chemical shift anisotropy of -160 ppm (56Bernado P. de la Torre J.G. Pons M. J. Biomol. NMR. 2002; 23: 139-150Crossref PubMed Scopus (76) Google Scholar, 57Garcìa de la Torre J. Biophys. Chem. 2001; 93: 159-170Crossref PubMed Scopus (79) Google Scholar). The Dpar/Dper ratio was computed as 2Dz/(Dx + Dy), where Dx and Dy are the pair of eigenvalues of the rotational diffusion matrix that are closest to each other, with Dx > Dy (56Bernado P. de la Torre J.G. Pons M. J. Biomol. 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