| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 279, Issue 46, 48024-48037, November 12, 2004
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
||||||||¶
From the
California Pacific Medical Center Research Institute, San Francisco, California 94115 and Kennesaw State University, Kennesaw, Georgia 30144
Received for publication, June 15, 2004 , and in revised form, August 19, 2004.
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
S) binding and GIRK1/4 channel current effects in Xenopus oocytes where the mutant proteins were expressed transiently. The F3.36(201)A mutation showed statistically significant increases in ligand-independent stimulation of GTPS binding versus wild type CB1, although basal levels for the W6.48(357)A mutant were not statistically different from wild type CB1. F3.36(201)A demonstrated a limited activation profile in the presence of multiple agonists. In contrast, enhanced agonist activation was produced by W6.48(357)A. These results suggest that a F3.36(201)/W6.48(357)-specific contact is an important constraint for the CB1-inactive state that may need to break during activation. Modeling studies suggest that the F3.36(201)/W6.48(357) contact can exist in the inactive state of CB1 and be broken in the activated state via a 
1 rotamer switch (F3.36(201) trans, W6.48(357) g+) 
(F3.36(201) g+, W6.48(357) trans). The F3.36(201)/W6.48(357) interaction therefore may represent a "toggle switch" for activation of CB1. |
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
-helical structure seen in Rho, thereby resulting in similar tertiary structures. In the work presented here, we provide evidence that "structural (functional) mimicry" by alternate microdomains may also support the core function of signaling activation through transmembrane helix conformational change in the class A family.
|
Khorana and co-workers (13) have reported that even in the dark (inactive) state of Rho, only some strong constraints exist, whereas the majority of the molecule experiences conformational flexibility. Therefore, light activation of Rho does not require the breaking and forming of thousands of specific contacts within nanoseconds, rather only a few specific contacts restricting the inactive state, including indole side chain contacts of tryptophan residues, need to break on activation. These changes can then be transmitted through the entire membrane protein because of its dynamic plasticity. One of the tryptophan residues that Khorana and co-workers (13) have reported to be restricted is W6.48(265). In the dark (inactive) state of Rho, the 
-ionone ring of 11-cis-retinal is close to W6.48(265) of the CWXP motif on TMH F and helps constrain it in a 1 = g+ conformation (3). In the light-activated state, the -ionone ring moves away from TMH F and toward TMH D where it resides close to A4.58(169) (14). This movement releases the constraint on W6.48(265), making it possible for W6.48(265) to undergo a conformational change. Lin and Sakmar (15) reported that perturbations in the environment of W6.48(265) of Rho occur during the conformational change concomitant with receptor activation. This suggests that the conformation of W6.48(265) when Rho is in its inactive/ground state (R; 1 = g+) changes during activation (i.e. W6.48(265) 1 g+ trans) (16).
In the class A cationic neurotransmitter receptors, a highly conserved cluster of aromatic amino acids is found on TMH6 that faces the binding site crevice bracketing W6.48 (F6.44, W6.48, F6.51, and F6.52) (16). Shi et al. (16) have proposed that an aromatic at 6.52 (F6.52) in the 2-adrenergic receptor may serve to constrain W6.48 in its inactive state (i.e. g+ 1) and is part of a rotamer toggle switch (C6.47 trans/W6.48 g+/F6.52 g+ C6.47 g+/W6.48 trans/F6.52 trans) for activation of this receptor.
Restriction of W6.48 by a TMH6 aromatic cluster is not possible in the cannabinoid receptors, as the CB1 receptor has leucines at 6.44, 6.51, and 6.52. Instead, the CB1 receptor contains a microdomain of aromatic residues that face into the ligand-binding pocket in the TMH3-4-5-6 region, including F3.25(190), F3.36(201), W4.64(256), Y5.39(276), W5.43(280), and W6.48(357) (Fig. 2). In work reported here, we suggest that the F3.36(201)/W6.48(357) interaction may act as a mimic of the 11-cis-retinal/W6.48 interaction in the Rho dark state and may serve as the "toggle switch" for CB1 activation, with F3.36(201) 1 trans/W6.48(357) 1 g+ representing the inactive (R) and F3.36(201) 1 g+/W6.48(357) 1 trans representing the active (R*) state of CB1. Modeling, mutation, and functional studies undertaken to test the importance of the TMH3-4-5-6 aromatic microdomain in ligand recognition and in the conformational changes that accompany activation of CB1 suggest that a F3.36(201)/W6.48(357)-specific contact is an important constraint for the CB1-inactive state.
|
|
EXPERIMENTAL PROCEDURES |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
Definition of Rotameric State of 1—Different nomenclatures have been used to define the rotameric state of side chain torsion angles. The nomenclature employed here for the 1 torsion angle is that described by Shi et al. (16). When the heavy atom at the position is opposite to the backbone nitrogen when viewed from the -carbon to the -carbon, the 1 is defined to be trans. When the heavy atom at the position is opposite to the backbone carbon when viewed from the -carbon to the -carbon, the 1 is defined to be gauche+ (g+). When the heavy atom at the position is opposite to the -hydrogen when viewed from the -carbon to the -carbon, the 1 is defined to be gauche- (g-). By using this nomenclature system, the side chain conformations discussed here are categorized into g- (0° < 1 < 120°), trans (120° < 1 < 240°), or g+ (240° < 1 < 360°).
Conformational Memories Studies of TMH6 and the W6.48A Mutant TMH6 —In order to explore the consequences of the W6.48A mutation upon the conformation of TMH6, we used the conformational memories (CM) method (18), a method that employs multiple Monte Carlo/simulated annealing random walks and the Amber* force field. Conformational memories has been shown to converge in a very practical number of steps and to be capable of overcoming energy barriers efficiently. By using CM, the conformational properties of a helix can be fully characterized by the free energy of each of the conformations that the helix can adopt, and this property includes not only the intrinsic energy of each conformational state but also the probability that the helix will adopt each particular conformation relative to all other ones accessible in an equilibrated thermodynamic ensemble.
The calculation is performed in two phases. In the first phase, repeated runs of Monte Carlo/simulated annealing are carried out to map the entire conformational space of the helix. In the second phase, new Monte Carlo/simulated annealing runs are performed only in the populated regions identified in the first phase of the calculation.
WT TMH6 Versus TMH6 W6.48A Mutant—The CB1 TMH6 (from residue 6.30 to residue 6.57, DIRLAKTLVLILVVLIICWGPLLAIMVY) and the TMH6 W6.48A mutant (DIRLAKTLVLILVVLIICAGPLLAIMVY) were built using MacroModel (19). In the Rho 2.8-Å crystal structure (3), 1 of W6.48 is g+. In order to be consistent with this result, 1 = g+, -60° was chosen as the starting conformation for W6.48(357) in the WT TMH6 run. Our previous CM studies of TMH6 have suggested that when C6.47(356) adopts a trans conformation (1 = 180°) and forms a hydrogen bond with the backbone of TMH6, TMH6 exhibits its greatest flexibility. In the studies reported here, the 1 of C6.47(356) was set to trans. The s for all other amino acids in TMH6 were left at the default values used by MacroModel when an -helix is built (19). The charges on all charged residues in each TMH6 studied were reduced to one-third of their values to prevent artifacts during the CM runs.
All calculations were performed using a distance-dependent dielectric. Each TMH6 was first minimized using the Amber* forcefield in MacroModel (19). For WT CB1 TMH6 73 torsion angles were allowed to vary during the CM runs, whereas 71 torsion angles were allowed to vary in the CB1 TMH6 W6.48(357)A mutant run. These included all helix backbone 
s and s, as well as amino acid side chain torsion angles for L6.39(348) to I6.54(363) (excluding 1 dihedrals on -branched residues and 2 on C6.47(356) in this span). The backbone s and s for I6.46(355) through P6.50(359) (i.e. the turn before P6.50(359)) were allowed to vary ±50° from their minimized values. All other backbone torsions were allowed to vary ± 10°. The 1 and 2 for W6.48(357) in WT TMH6 and the 1 for C6.47(356) (in both the WT and W6.48(357)A mutant runs) were allowed to vary ±60°. The 2 for C6.47(356) was not allowed to vary in either run. All other side chain torsion angles were allowed to vary ± 180° without constraints. The calculation was performed in two phases as indicated below.
Exploratory Phase—In the exploratory phase, a random walk was used to identify the region of conformational space that is populated for each torsion angle studied. Starting at a temperature of 2070 K, 20,000 steps were applied to the rotatable bonds with cooling in 18 steps to a final temperature of 310 K. Trial conformations were generated at each temperature by randomly picking three torsion angles from the set of 73 (71 in the W6.48A mutant) and changing each angle by a random value within the range set in the calculation (see above). After each step, the generated trial conformation was either accepted or rejected using the Metropolis criterion. This calculation was repeated for a total of 100 cycles. Accepted conformations were used to map the conformational space of TMH6 by creating "memories" of values for each torsion angle that were accepted.
Biased Annealing Phase—In the second phase of the CM calculation, the only torsion angle moves attempted were those that would keep the angle in the "populated conformational space" mapped in the exploratory phase. The biased annealing phase began at a temperature of 722 K cooling to 310 K in 8 steps. 100 structures were written out at 310 K.
Analysis of Output—Finally, the output of 100 structures at 310 K was clustered using X-Cluster in MacroModel (19). This program reorders the structures according to their root mean square deviation and groups the structures into families of similar conformers. The resulting 100 structures from CM were also analyzed using the program, ProKink (20). This program, which is embedded in the Simulaid Conversion program,2 was used to calculate the face shift, wobble, and bend angles of each helix. Statistically significant differences between the face shift, wobble, and bend angles of the R and R* CB1 WT versus W6.48A R and R* helix families were evaluated in the two-sample independent t test computed using OriginPro version 7 (Origin Lab Corp.).
Models of CB1 R and R* States—In the present study, the literature on GPCR activation discussed above was used to generate an R* CB1 TMH bundle from a model of the inactive (R) CB1 receptor based on the 2.8-Å crystal structure of rhodopsin (3). The creation of these two forms of CB1 is described briefly below.
Model of Inactive State (R) Form of CB1—A model of the R form of CB1 was created using the 2.8-Å crystal structure of bovine Rho (3). First, the sequence of the mouse CB1 receptor (22) (see Fig. 1) was aligned with the sequence of bovine Rho using the same highly conserved residues as alignment guides that were used initially to generate our first model of CB1 (23). TMH5 in CB1 lacks the highly conserved proline in TMH5 of Rho. Therefore, the sequence of CB1 in the TMH5 region was aligned with that of Rho as described previously using its hydrophobicity profile (23). The mouse CB1 sequence (22) is 97.7% identical to the human CB1 sequence (2) overall and 100% identical within the transmembrane regions. The mouse sequence is one residue longer (473 residues) than the human sequence (472 residues) due to an additional residue in the N terminus.
Initial helix ends for mouse CB1 were chosen in analogy with those of Rho (3). With the exception of TMH1, these helix ends were found to be within one turn of the helix ends originally calculated by us and reported in 1995 (23). Two changes dictated by the CB1 sequence were made in the helix ends. The shortness of the E1 loop region in CB1 necessitated starting TMH3 at 3.23 (N3.23(188) to R3.56(221)). The break in helicity caused by the GWNC sequence motif on the extracellular end of TMH4 necessitated that TMH4 end at 4.62 instead of 4.66 (as is found in Rho). Changes to the general Rho structure that were necessitated by sequence divergences included the absence of helixkinking proline residues in TMH1 and TMH5, the lack of a GG motif in TMH2, as well as the presence of extra flexibility in TMH6 (24). Our recent conformational memories study of TMH6 in CB1 revealed that TMH6 in CB1 has high flexibility due to the small size of residue 6.49 (a glycine) immediately preceding Pro 6.50. The conformer selected from our CM results for inclusion in the CB1 R bundle (Pro kink angle = 53.1°) was chosen so that R3.50(215) and D6.30(339) could form a salt bridge at the intracellular ends of TMH3 and -6 in the CB1 TMH bundle. An analogous salt bridge has been shown to be an important stabilizer of the inactive state of the 2-adrenergic receptor, the 5HT-2a receptor (10), and to be present in Rho (3). Because of the extreme flexibility of TMH6 in CB1, we have proposed that an additional TMH3–6 salt bridge, K3.28(193)–D6.58(367), stabilizes the inactive state on the extracellular side of the TMH bundle (25).
Model of Active (R*) Form of CB1—Based upon experimental results for rhodopsin and the 2-adrenergic receptor (6, 10, 15, 26), the R* (active) CB1 bundle was created from the inactive (R) model of CB1 by substituting a less kinked TMH6 (21.8° kink angle) from our CM results (25)for which the R3.50(215) and D6.30(339) salt bridge would be broken due to the movement of the intracellular end of TMH6 away from that of TMH3 and out into lipid (10). Rotations of both TMH3 and TMH6 were also central to the creation of the R* model. The details of these rotations are presented elsewhere (24).
Preparation of Helices—Each helix of the model was capped as the acetamide at its N terminus and as the N-methyl amide at its C terminus. Ionizable residues in the first turn of either end of the helix were neutralized, as were any lipid facing charged residues. Ionizable residues were considered charged if they appeared anywhere else in the helix.
Energy Minimization, Unoccupied Receptor States—The energy of the CB1 R or CB1 R* TMH bundle complex was minimized using the AMBER* united atom force field in Macromodel 6.5 (Schrödinger Inc., Portland, OR). A distance-dependent dielectric, 8.0-Å extended non-bonded cut-off (updated every 10 steps), 20.0-Å electrostatic cut-off, 4.0-Å hydrogen bond cut-off, and explicit hydrogens on sp2 carbons were used. The first stage of the calculation consisted of 2000 steps of Polak-Ribier conjugate gradient (CG) minimization in which a force constant of 225 kJ/mol was used on the helix backbone atoms in order to hold the TMH backbones fixed, while permitting the side chains to relax. The second stage of the calculation consisted of 100 steps of CG in which the force constant on the helix backbone atoms was reduced to 50 kJ/mol in order to allow the helix backbones to adjust. Stages one and two were repeated with the number of CG steps in stage two incremented from 100 to 500 steps until a gradient of 0.001 kJ/(mol·Å2) was reached. This same protocol was followed for the W6.48(357)A TMH bundle.
Assessment of Aromatic Stacking Interactions—Residues were designated here as participating in an aromatic stacking interaction if subject rings had centroid to centroid distances (d) between 4.5 and 7.0Å. These interactions were further classified as "tilted t" arrangements if 30° 
90° and as parallel arrangements for < 30° (where is the angle between normal vectors of interacting rings) (27). Parallel arrangements were considered favorable only if the interacting rings were offset from each other (28). All measurements were made using MacroModel 8.1.
Assessment of Cation-
Interactions—In their study of cation- interactions in the protein data bank, Gallivan and Dougherty (29) considered a cation- interaction to be present in structures when both r 10 Å and r' 10 Å, where r is the distance between the ammonium nitrogen (NZ) of the protonated residue and the aromatic ring centroid, and r' is the distance between CE of the protonated residue (in the case here, a Lys) and the aromatic ring centroid (29). However, these investigators reported that in 88% of the cation- interactions considered, r < 5 Å and r' < 5 Å. A cation- interaction was judged to be present in the work presented here if r < 5 Å and r' < 5 Å. The r and r' distances were measured here using MacroModel 8.1.
Ligand Conformations and Docking Positions—The binding site conformations and anchoring interactions inside the receptor used for each ligand discussed here are based on our recently published work (1).
Mutation Studies
Materials—SR141716A and CP55,940 were obtained from the National Institute on Drug Abuse. WIN 55,212-2 was purchased from RBI (Natick, MA), and anandamide was purchased from Tocris (Ellisville, MI).
Mutagenesis, Cell Culture, and Radioligand Binding—The cell lines used in this study have been described previously (1). Site-directed mutations were introduced into mouse CB1 in pcDNA3 at the designated sites using the QuikChange mutagenesis technique (Stratagene, La Jolla, CA). Stable cell lines were created by transfection of wild type or mutant CB1pcDNA3 into HEK 293 cells by the LipofectAMINE reagent (Invitrogen) and cultured as described previously (30). Cell lines containing moderate to high levels of receptor mRNA, assessed by Northern analysis, were tested for receptor binding and signal transduction properties. Receptor binding was determined as described previously (1). Cell lines with the most similar receptor expression profile, as ascertained by Bmax values, were chosen for further analysis (Table I).
|
S Binding Assay—The basal activity and the ability of various cannabinoids to stimulate the wild type CB1 receptor or the mutant receptors were tested with [35S]GTPS binding. Cells were harvested in phosphate-buffered saline containing 1 mM EDTA and centrifuged at 500 x g for 5 min. The cell pellet was homogenized and centrifuged at 50,000 x g for 10 min at 4 °C. The pellet was resuspended in buffer composed of (mM): Tris-HCl 50, EDTA 1, and MgCl2 3, pH 7.4, to yield protein concentration of 2–4 mg/ml. Membrane preparations were aliquoted and stored at -80 °C. Binding was initiated by the addition of 20 µg of membrane protein into glass tubes containing 0.1 nM [35S]GTPS, 10 µM GDP in GTPS binding buffer (50 mM Tris-HCl, 100 mM NaCl, 3 mM MgCl2, 0.2 mM EGTA, 0.1% bovine serum albumin, pH 7.4). Nonspecific binding was assessed in the presence of 20 µM unlabeled GTPS. Binding assays were performed for 90 min at 30 °C with various concentrations of WIN 55212-2, CP55,940, anandamide, and SR141716A in a total volume of 500 µl. Free and bound radioligands were separated by a rapid filtration through Whatman GF/C filters. Filters were shaken for 1 h in 6 ml of scintillation fluid (Fisher), and radioactivity was determined by a liquid scintillation counter.
Expression in Oocytes and Recordings—This technique was performed as described previously (31). Briefly, 0.013 pg of GIRK1 and GIRK4 and 25 ng of CB1 (wild type or mutant) cRNAs were co-injected using a micromanipulator (Drummond Scientific Co., Broomall, PA) into Xenopus laevis oocytes (Xenopus One, Dexter, MI). Recordings were performed after 7–9 days of incubation in 0.5x L-15 media (Sigma) supplemented with L-glutamine and antibiotics. For recordings, the eggs were placed in a chamber (total volume 200 ml) and perfused at 4 ml/min with LK (2 mM KCl, 96 mM NaCl, 2 mM CaCl2, 1.8 mM MgCl2, and 5 mM HEPES, pH 7.5), HK (96 mM KCl, 2 mM NaCl, 2 mM CaCl2, 1.8 mM MgCl2, and 5 mM HEPES, pH 7.5), or HK plus drug. Bovine serum albumin (3 µM) was added to all drug solutions to minimize absorption of cannabinoid compounds to the perfusion system. Oocytes were impaled with two microelectrodes filled with 3 M KCl and were voltage-clamped at reported voltages using an Axon GeneClamp amplifier (Axon Instruments, Foster City, CA). Currents were filtered at 10 Hz, collected, and analyzed using a Macintosh Centris 650 containing a 16-bit analog-digital interface board and voltage-clamp software running under the IGOR graphics environment (Wavemetrics, Lake Oswego, OR). Oocytes were voltage clamped at -80 mV and superfused in a low potassium (LK) solution containing 96 mM Na+ and 2 mM K+. When a high potassium (HK) solution containing 96 mM K+ and 2 mM Na+ was exchanged for LK, an inward current was produced and termed IHK. IHK represents the basal activity of GIRK1/4 channels. This current reached a plateau with 
30 s (31). At this point application of the cannabinoid agonists (WIN 55,212-2 or anandamide) in HK further enhanced the inward current. This agonistinduced current was named IAg. Concentration-response curves were generated by nonlinear regression of log percent concentration enhancement data (IAg/IHK x 100) with the use of the GraphPad Prism program (GraphPad, San Diego, CA).
Statistical Analyses—The EC50 and Emax values were calculated by unweighted least squares nonlinear regression of log concentration values versus percent effect. Significant differences were determined (GraphPad Prism) using analysis of variance or the unpaired Student's t test, where suitable. Bonferroni-Dunn post-hoc analyses were conducted when appropriate. p values <0.05 defined statistical significance.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
|---|
= 90°), whereas F3.36(201) stacks with W5.43(280) (d = 5.6 Å, = 40°). W5.43(280) also has an off-set parallel stack with Y5.39(276) (d = 5.9 Å, = 0°), whereas Y5.39(276) stacks with W4.64(256) (d = 6.5 Å, = 90°) (see "Experimental Procedures" for definitions of d and ).
Residue 3.25 (190) does not stack with the other aromatic residues in the TMH3-4-5-6 region but still appears to be an important residue. In the R bundle, F3.25(190) is located directly extracellular to K3.28 and is close enough to K3.28 to be able to form a cation- interaction with K3.28 (193) (NZ-centroid distance, r = 3.1 Å, CE-centroid distance, r' = 4.9 Å; see "Experimental Procedures" for definitions of distances). In the R state, the 1 for K3.28 (193) and for F3.25(190) is trans.
CB1 R* State—Fig. 2B illustrates key features of the CB1 TMH bundle model in the active (R*) state in the TMH3-4-5-6 region. The conformational changes that occur upon receptor activation result in rotations of TMH3 and -6 as well as a change in the conformation of TMH6 (by moderation of its proline kink angle) (6–8, 10). In our models, both W6.48(357) and F3.36(201) undergo a change in their 1 values from R to R*. 1 in W6.48(357) changes from g+ to trans and 1 of F3.36(201) changes from trans to g+ (see "Experimental Procedures" for definition of 1). In the R* TMH bundle, the K3.28(193) and D6.58(367) salt bridge is broken (N-O distance = 16.8 Å) because TMH3 and -6 rotate (counterclockwise from extracellular view) during the R to R* transition (i.e. activation) (26, 32). K3.28(193) has rotated away from D6.58(367) toward TMH2/TMH7, and D6.58(367) has rotated toward the TMH5–6 interface and is raised higher above the ligand-binding pocket due to the moderation of the TMH6 proline kink angle. The W6.48(357)/F3.36(201)/W5.43(280)/Y5.39(276)/W4.64(256) aromatic cluster present in the inactive state in the absence of ligand also undergoes rearrangement, with F3.36(201) no longer part of this cluster. In the TMH3-4-5-6 region of R* in the absence of ligand, W6.48(357) and W5.43(280) form an off-set parallel aromatic stacking interaction with each other (d = 4.9 Å, = 30°). W5.43(280) also stacks with Y5.39 (d = 6.6 Å, = 60°), whereasY5.39 stacks with W4.64 (d = 5.7 Å, = 90°). This series of aromatic stacking interactions results in a large aromatic stack in R* comprised of W6.48(357)/W5.43(280)/Y5.39(276)/W4.64(256). F3.36(201) (1 = g+) is not located near another TMH3-4-5-6 aromatic residue in the R* bundle, instead F3.36(201) is bounded by V3.40(205), V3.32(197), and L6.44(353).
As stated above, in the R* state, the rotation of TMH3 upon activation causes K3.28(193) to point toward TMH2/TMH7, and because F3.25(190) is one turn above K3.28(193), it also now faces the TMH2/TMH7 region. The 1 for both K3.28(193) and F3.25(190) is g+ in the R* bundle.
Ligand Binding
We have recently published mutation and modeling studies of the CB1 TMH3-4-5-6 aromatic microdomain in which binding sites for the inverse agonist SR141716A and the CB1 agonists WIN 55212-2 and anandamide were identified (1).
Functional Analysis of Mutant Receptors
We have shown previously that the aromatic residues F3.36(201), W5.43(280), and W6.48(357) form specific interaction sites in CB1 for aminoalkylindole agonist (WIN 55,212-2) and diaryl pyrazole inverse agonist/antagonist (SR141716A) ligands but not for endogenous (anandamide) and bicyclic cannabinoid (CP55,940) agonists (1). When the aromatic residues were individually mutated to alanine, a significant reduction in ligand binding affinity was only observed in the presence of WIN 55,212-2 and SR141716A but not CP55,940 and anandamide. The study presented here is a detailed functional analysis of these mutant receptors using two different cellular backgrounds. The mutants were studied in stable cell lines created in HEK cells or in oocytes where the mutant proteins were expressed transiently. The mutations were analyzed in two ways. Direct receptor-G protein stimulation in HEK cells was assessed using [35S]GTPS binding, and G protein stimulation downstream of the receptor was evaluated by measuring enhancement of GIRK channel activity in oocytes.
F3.36A(201)—The F3.36A(201) mutation produced a significant reduction in agonist-dependent activation produced by WIN 55,212-2 (Fig. 3, A and B, and Tables II and III). This could be observed at the level of direct receptor-G protein stimulation (GTPS), and this outcome was even more profound downstream at GIRK1/4 channels. The affinity of WIN 55,212-2 at F3.36A(201) was reduced 8.9-fold in HEK cells, whereas the affinity of CP55,940 and anandamide was unaffected (1). Even the addition of high concentrations of WIN 55,212-2 could not produce substantial receptor activation suggesting the loss of affinity for the ligand, produced by the mutation, was not solely responsible for decreased activation. To strengthen this conclusion further, CP55,940 and anandamide were also tested because the mutation to alanine at F3.36(201) had no effect on the affinity of these ligands for the receptor (Table I). Full receptor activation could also not be produced with CP55,940 (see Fig. 3D and Table II) or with anandamide (see Fig. 3C and Table III).
|
|
|
Agonist-independent activation of GPCRs can be assessed by measuring basal turnover of [35S]GTPS in transfected cells and by comparing the IHK of GIRK1/4 in injected oocytes (38, 39). Consistent with previous findings (39), we found a 56% increase (WT CB1 = 1055 ± 58 dpm, untransfected = 675* ± 73 dpm) in agonist-independent activation of [35S]GTPS in HEK cells expressing WT CB1 compared with untransfected cells (Fig. 7 and Table IV). When the agonist-independent GTPS activity of F3.36A(201) was compared with WT CB1, a 73% increase (F3.36(201)A = 1825 ± 138 dpm, WT CB1 = 1055 ± 58 dpm) was observed with the mutant (Fig. 7 and Table IV). A 62% increase was also observed when IHK or basal activity of GIRK1/4 was evaluated between F3.36A(201) and WT CB1 (Fig. 8 and Table IV). Although the amount of receptor expressed in oocytes used for the GIRK assay cannot be determined, the Bmax values for the WT and F3.36(201)A cell lines used for the GTPS assay were not statistically different from one another (WT CB1 Bmax = 4.4 (3.5–5.3) pmol/mg; F3.36(201)A-Bmax = 5.2 (3.6–6.9) pmol/mg; Table I). Thus, the 73% increase (relative to WT) in the level of GTPS binding seen in the HEK cells stably transfected with F3.36(201)A cannot be attributed to an overexpression of mutant receptor protein (Table I). These combined data strongly suggested the F3.36A(201) mutant receptor had even greater constitutive activity than WT CB1. To further support this hypothesis, the properties of the mutant receptor were evaluated using the CB1-selective inverse agonist SR141716A. An inverse agonist should reduce the basal activity of the constitutively activated mutant receptor because the inverse agonist will force the receptor to adopt an inactive conformation; this was the case (Figs. 3E and 7). Furthermore, compared with WT CB1 the inverse agonist response of SR141716A at F3.36A(201) was significantly increased (Fig. 3E). No inverse agonism was produced at WT CB1 except at the highest (5 µM) concentration (17 ± 2%, n = 3) (Fig. 3E). However, inhibition of GTPS binding occurred with F3.36(201)A in the presence of nanomolar concentrations of SR141716A with EC50 and Emax values of 0.6 (0.1–3.2) nM and -24 (-30 to -18) %, respectively.
|
|
|
W5.43(280)A—The EC50 values reported in Tables II and III show that the potency of WIN 55,212-2 at the W5.43(280)A mutant was reduced 245- and 16.8-fold when this receptor was studied by using GTPS binding or by measuring enhancement of GIRK channel activity, respectively (Fig. 4, A and B and Tables II and III).
|
W6.48A(357)—The EC50 values reported in Tables II and III show that the potency of WIN 55,212-2 at the W6.48(357)A mutant was reduced 12- and 1.8-fold when this receptor was studied using GTPS binding or by measuring enhancement of GIRK channel activity, respectively (Fig. 5, A and B, and Tables II and III). However, the loss of potency observed with GIRK channel activity was not significant. The effects on potency may be due to the decreased affinity of WIN 55,212-2 at W6.48(357)A. The affinity of WIN 55,212-2 at W6.48(357)A was reduced 3.8-fold in HEK cells, whereas the affinity of CP55,940 and anandamide was unaffected (Table I) (1). Because the affinity of CP55,940 and anandamide was unaffected at this mutation, the functional responses produced by these ligands were tested at W6.48A(357). As shown in Fig. 5, C and D, and Tables II and III, there was no significant reduction in the potency of either CP55,940 or anandamide at W6.48A(357). These data suggest the loss in potency observed for WIN 55,212-2 stimulation of GTPS binding at W6.48(357)A was the result of the selective loss of affinity for WIN 55,212-2 at this mutant receptor. It should be noted that a comparison of the shifts in potency for WIN 55,212-2 at both W6.48(357)A and W5.43(280)A demonstrate the GTPS assay is more sensitive to this observable effect compared with studying GIRK channel activity (Figs. 4, A and B and Fig. 5, A and B, and Tables II and III).
|
S activity produced by WIN 55,212-2 at W6.48(357)A was significantly enhanced compared with WT CB1 (Fig. 5A and Table II). To determine whether this effect was specific for aminoalkylindoles, the bicyclic cannabinoid CP55,940 was also tested at W6.48(357)A. As observed with WIN 55,212-2, CP55,940 also produced enhanced stimulation of GTPS activity at W6.48(357)A compared with WT CB1 (Fig. 5D). The enhanced activity of WIN 55,212-2 at W6.48(357)A was not observed when GIRK channel activity was measured (Fig. 5B). There were no significant changes in agonist-independent activation of W6.48(357)A in either cell system tested (Figs. 7 and 8 and Table IV). Consistent with unchanged constitutive activity, the response to the inverse agonist at W6.48(357)A was not different from the response of WT CB1 to the inverse agonist (Fig. 5E). Modeling studies also suggest that the W6.48(357)A mutant should not be constitutively active, if it is assumed that changes in the TMH3-4-5-6 aromatic cluster influence the state preference for the receptor. The W6.48A mutation will affect the TMH3-4-5-6 aromatic cluster in both the R and R* states. In the R state, the W6.48A mutation will result in the loss of one aromatic stacking interaction, i.e. the W6.48(357)/F3.36(201)/W5.43(280)/Y5.39(276)/W4.64(256) cluster in R will become a F3.36(201)/W5.43(280)/Y5.39(276)/W4.64(256) cluster in the mutant. In the R* state, the W6.48(357)/W5.43(280)/Y5.39(276)/W4.64(256) cluster present in the R* state will become a W5.43(280)/Y5.39(276)/W4.64(256) cluster in the mutant. Because aromatic stacking in both the R and R* states will be equally impacted, it is reasonable to expect that the W6.48A mutant will not produce a change in basal levels for the W6.48A mutant relative to WT CB1, and this is what was seen experimentally.
F3.25(190)A—We previously reported that F3.25(190) is not part of the binding site of WIN 55,212-2, SR141716A, or CP55,940 but is a part of the anandamide-binding pocket (1). When F3.25(190) was mutated to an alanine, a 6-fold reduction in binding affinity was observed in the presence of anandamide, but no changes were observed in the presence of WIN 55,212-2, SR141716A, and CP55,940 (Table I). When GIRK channel activity was measured there was a significant reduction in both the potency and efficacy of anandamide at F3.25(190)A but no change in the presence of WIN 55,212-2 (Fig. 6, A and B, and Table III).
|
S turnover, in HEK cells transfected with F3.25(190)A, could not be measured because the partial agonist nature of the endogenous ligand results in a nonsignificant amount of GTPS stimulation (41). Therefore, the findings with anandamide and the GIRK channel activity could not be compared with GTPS stimulation because anandamide did not produce a significant functional response in HEK cells. There was a 6.2-fold decrease in the potency of WIN 55,212-2 when receptor activation was assessed by measuring GTPS activity.
These agonists do not exhibit affinity changes from WT in the F3.25(190)A mutant. However, the effects observed on potency and/or efficacy reported here for WIN 55212-2 at the F3.25(190)A mutant may be related to the general role F3.25(190) plays in positioning K3.28 in the TMH bundle. As is clear from Fig. 2, F3.25(190) is just above K3.28 in the inactive state of CB1 and has a cation- interaction (r 5 Å) with this residue. Gallivan and Dougherty (29) have reported that 14.5% of all cation- interactions found in the "Protein Data Bank Select" list of Sander and co-workers (42, 43) represent such Lys-Phe interactions. Our modeling studies suggest that these residues move in concert with each other, as they both have trans 1s in the R state and g+ 1s in the R* state. F3.25(190) may act as a chaperone of K3.28 to position it in the correct location in R* for ligand interaction and to shield it from the extracellular milieu in the R state. Because residue positions in TMH3 R* may be changed in the absence of aromaticity at position 3.25, it is possible that ligands that do not bind to F3.25(190) can have their potency and/or efficacy affected by the F3.25(190)A mutation.
There were no significant changes in agonist-independent activation of F3.25(190)A in either cell system tested (Fig. 7 and Fig. 8 and Table IV). F3.25(190) is not part of the TMH3-4-5-6 aromatic cluster that characterizes the R and R* states of CB1 (see Fig. 2). Therefore, the mutation of this aromatic residue to a nonaromatic residue would not be expected to affect basal levels.
Toggle Switch Residues
Fig. 9 illustrates the relationship between F3.36(201) and W6.48(357)in the R and R* states of CB1. In the context of the inactive state model (see Fig. 9, left), F3.36(201) (1 = trans) is located opposite W6.48(357)(1 = g+) and has an aromatic stacking interaction with W6.48(357) that would prevent the 1 of W6.48(357) from changing from g+ to trans, thus stabilizing TMH6 in its inactive conformation. In the active state of CB1, F3.36(201) and W6.48(357) change conformations (F3.36(201)(1 = trans)/W6.48(357)(1 = g+) F3.36(201)(1 = g+)/W6.48(1 = trans)) in order to rotate past each other and F3.36(201) (1 = g+) and W6.48(357) (1 = trans) are located too far apart in R* to interact with each other (see Fig. 9, right).
|
-helix. Table V lists the results of ProKink analyses of WT TMH6 CM output and the W6.48A TMH6 CM output. This analysis yields values not only for the bend angle of this proline-containing helix but also the wobble angles and face shifts for these helices. The bend angle is the angle between the two parts when the helix is kinked along its axis. The wobble angle is the angle that defines the orientation of the post-proline helix in three-dimensional space, with respect to the pre-proline helix. The face shift measures the distortion that causes a twisting of the helix "face" in such a way that amino acids that used to be on the same side (face) of the helix are shifted and are on different sides of the helix as a result of the bend (20).
|
For WT CB1, cluster 1 contained 40 members with an average proline bend (kink) angle of 75.9 ± 1.0°, whereas cluster 2 contained 51 members with average bend angle of 33.7 ± 1.1°. The TMH6 used in our CB1 R model was selected from the more bent cluster, cluster 1, whereas the TMH6 used in our CB1 R* model was selected from the straighter cluster, Cluster 2 (see "Experimental Procedures"). For the TMH6 W6.48A mutant, cluster 1 contained 19 members with an average proline bend (kink) angle of 74.0 ± 1.0°, whereas cluster 2 contained 72 members with an average proline bend angle of 36.8 ± 1.2°.
At the 0.01 level, the difference of population means for the kink, wobble, and face shift angles for the W6.48A mutant reported in Table V were not significantly different from the corresponding measures in WT CB1, except for the R* wobble angle. Here the W6.48A R* wobble angle (-105.6 ± 3.4°) was found to be significantly different from the WT R* wobble angle (-120.5 ± 4.6°) at the 0.01 level. Fig. 10 illustrates the steric consequence of this 15° difference in wobble angle. In the R to R* transition, the salt bridge between R3.50 and D/E6.30 is thought to be broken via a conformational change in TMH6 mediated by the flexible hinge region (CWXP motif) of TMH6. Fig. 10 shows that D6.30 in the W6.48A mutant is capable of pulling further away from the intracellular end of TMH3 and R3.50 than is D6.30 in WT TMH6.
|
|
DISCUSSION |
|---|
|
TOPABSTRACT |
|---|
) and F3.36(201)A (
) receptor activation. WIN 55,212-2 (A), CP55,940 (D), and SR141716A (E) were used for concentration-response analysis of [35S]GTP
