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J. Biol. Chem., Vol. 282, Issue 34, 25100-25113, August 24, 2007

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Helix 8 Leu in the CB₁ Cannabinoid Receptor Contributes to Selective Signal Transduction Mechanisms^*

Sharon Anavi-Goffer, Daniel Fleischer, Dow P. Hurst, Diane L. Lynch, Judy Barnett-Norris, Shanping Shi^¶, Deborah L. Lewis^¶, Somnath Mukhopadhyay^||, Allyn C. Howlett^||1, Patricia H. Reggio, and Mary E. Abood²

From the California Pacific Medical Center Research Institute, San Francisco, California 94107, Center for Drug Discovery, Department of Chemistry and Biochemistry, University of North Carolina, Greensboro, North Carolina 27402, the ^¶Department of Pharmacology and Toxicology, Medical College of Georgia, Augusta, Georgia 30912, and the ^||Neuroscience of Drug Abuse Research Program, North Carolina Central University, Durham, North Carolina 27707

Received for publication, April 24, 2007 , and in revised form, June 19, 2007.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The intracellular C-terminal helix 8 (H8) of the CB₁ cannabinoid receptor deviates from the highly conserved NPXXY(X)_5,6F G-protein-coupled receptor motif, possessing a Leu instead of a Phe. We compared the signal transduction capabilities of CB₁ with those of an L7.60F mutation and an L7.60I mutation that mimics the CB₂ sequence. The two mutant receptors differed from wild type (WT) in their ability to regulate G-proteins in the [³⁵S]guanosine 5'-3-O-(thio)triphosphate binding assay. The L7.60F receptor exhibited attenuated stimulation by agonists WIN-55,212-2 and CP-55,940 but not HU-210, whereas the L7.60I receptor exhibited impaired stimulation by all agonists tested as well as by the inverse agonist rimonabant. The mutants internalized more rapidly than WT receptors but could equally sequester G-proteins from the somatostatin receptor. Both the time course and maximal N-type Ca²⁺ current inhibition by WIN-55,212-2 were reduced in the mutants. Reconstitution experiments with pertussis toxin-insensitive G-proteins revealed loss of coupling to G_i3 but not G_0A in the L7.60I mutant, whereas the reduction in the time course for the L7.60F mutant was governed by G_i3. Furthermore, G_i3 but not G_0A enhanced basal facilitation ratio, suggesting that G_i3 is responsible for CB₁ tonic activity. Co-immunoprecipitation studies revealed that both mutant receptors were associated with G_i1 or G_i2 but not with G_i3. Molecular dynamics simulations of WT CB₁ receptor and each mutant in a 1-palmitoyl-2-oleoylphosphatidylcholine bilayer suggested that the packing of H8 is different in each. The hydrogen bonding patterns along the helix backbones of each H8 also are different, as are the geometries of the elbow region of H8 (R7.56(400)-K7.58(402)). This study demonstrates that the evolutionary modification to NPXXY(X)_5,6L contributes to maximal activity of the CB₁ receptor and provides a molecular basis for the differential coupling observed with chemically different agonists.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
The CB₁ cannabinoid receptor is a member of the class A G-protein-coupled receptor (GPCR)³ superfamily (see Fig. 1). The CB₁ receptor is activated by endocannabinoid lipid mediators (anandamide, 2-arachidonoylglycerol), as well as cannabinoid (HU-210, CP-55940) and aminoalkylindole (WIN-55,212-2) agonists, leading to diverse signaling responses (1). CB₁ receptor agonists can selectively regulate G-protein coupling (2–5). Competitive antagonists, such as the arylpyrazole rimonabant (also known as SR141716), can inhibit CB₁ constitutive activity and thereby behave as inverse agonists (6–8). However, the protein conformation signals that underlie the ligand-directed recognition of the CB₁ receptor for different G-protein subtypes and their activation are not understood.

Crystal structures of bovine rhodopsin (9, 10), a prototype member of class A GPCRs, reveal an intracellular -helical extension of transmembrane helix (TMH) 7 (Lys³¹¹ to Cys³²³). This domain, referred to as helix 8 (H8), is positioned along the intracellular surface of TMH1 (9, 10). Photoactivation of rhodopsin has been associated with movement of H8 as detected in cysteine cross-linking studies and in nitroxide spin label studies (11). The highly conserved NPXXY(X)_5,6F motif of rhodopsin has been proposed to provide structural constraints via the aromatic interaction of Y7.53(306) in the NPXXY motif of TMH7 to F7.60(313) in H8, which rearrange in response to photo-isomerization (12). This same interaction has been shown to be important for the switching of the 5HT-2C receptor among multiple active and inactive conformations (13). Taken together, these studies suggest that the H8 domain plays a role in docking specific G-protein subunits and in the transition between conformational states.

The CB₁ receptor possesses a defined intracellular helical segment (H8 D7.59(403) to P7.69(413)), as predicted by Fourier transform analysis of the primary sequence of the CB₁ receptor (14). Significant helicity was demonstrated when the CB₁ 401–417 peptide was present in an anionic micelle environment (SDS or phosphatidic acid) but not in aqueous media (15). Recent NMR and circular dichroism studies also revealed significant -helical structure of synthetic peptides in dodecyl phosphocholine micelles (16, 17) or SDS (18). However, the CB₁ receptor differs from the NPXXY(X)_5,6F motif, as amino acid residue 7.60(404) is a Leu in CB₁.We explored the ramifications of this deviation from the highly conserved NPXXY(X)_5,6F motif, by comparing the signal transduction capabilities of the CB₁ wild-type with those of an L7.60F mutation which mimics the rhodopsin motif, and an L7.60I mutation which mimics the homologous CB₂ receptor sequence.

View larger version (60K): [in this window] [in a new window]   FIGURE 1. A helix net representation of the sequence of the human CB1 receptor. Residues Y7.53(397) of the TMH7 NPXXY motif and L7.60(404) of intracellular H8 are shown in boldface type.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Mutation Studies
Materials—[³H]CP-55,940 was purchased from PerkinElmer Life Sciences. WIN-55,212-2 was procured from Tocris. [³H]Rimonabant, rimonabant, and CP-55,940 were obtained from the National Institute on Drug Abuse. HU-210 (11-hydroxy-⁸-THC dimethylheptyl) was a gift from Prof. Mechoulam (Hebrew University, Jerusalem, Israel). D-Trp⁸ somatosatin-14 was from Bachem. Anti-G_i antibody was from Biomol. GDP and protease inhibitor mixture (4-(2-aminoethyl)benzenesulfonyl fluoride) were purchased from Sigma; GTPS was from Roche Applied Science, and [³⁵S]GTPS and ECL reagents were from Amersham Biosciences. All G-protein subunits were obtained from UMR cDNA Resource Center (Rolla, MO). Pertussis toxin was from List Biological Laboratories, Campbell, CA. For radioligand binding assays, all the drugs were dissolved in ethanol. For [³⁵S]GTPS assays, drugs were dissolved in Me₂SO.

Mutagenesis—Mutations were introduced into the hCB₁ receptor that had been subcloned into pcDNA3 vector (Invitrogen) with a QuikChange site-directed mutagenesis kit (Stratagene) as described previously (19). The hCB₁ L7.60I mutation was made with the mutagenic primer (forward) TCGCTGAGGAGTAAGGACATCCGACACGCTTTCCGGAGC. The hCB₁ L7.60F mutation was made with the mutagenic primer (forward) TCGCTGAGGAGTAAGGACTTCCGACACGCTTTCCGGAGC. The mutant cDNAs were sequenced to confirm the presence of the desired substitution.

Cell Transfection—HEK293 cells were grown and stably transfected with expression plasmids containing WT or mutant hCB₁ receptor sequences as described previously (19).

Immunochemistry—Selected colonies were expanded and tested for receptor expression by immunochemistry as described previously (19). Immunofluorescence was visualized with a laser-scanning confocal fluorescence microscope (Nikon). In control experiments, no labeling was observed with the secondary antibody alone or when the primary antibody was incubated with CB₁ receptor peptide at 100 µg/ml. For internalization studies, cells were preincubated with 1 µM CP-55,940 for 30 min or 1 h at 37 °C and then labeled (19).

Radioligand Binding—As described previously (19), binding was initiated by the addition of 50 or 60 µg of membrane protein into tubes containing [³H]CP-55,940 (158 Ci/mmol) or [³H]rimonabant (16.9 Ci/mmol). Nonspecific binding was assessed by the addition of 1 µM unlabeled CP-55,940 or rimonabant to the tubes.

[³⁵S]GTPS Binding Assay—Cells were harvested in phosphate-buffered saline containing 1 mM EDTA and centrifuged at 500 x g for 5 min, as described previously (20). The cell pellet was homogenized and centrifuged at 50,000 x g, for 10 min at 4 °C. Binding was initiated by the addition of 10 µg (for agonist experiments) or 20 µg (for antagonist experiments) of membrane protein into glass tubes containing 0.1 nM [³⁵S]GTPS, 10 µM GDP in GTPS binding buffer (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.

Immunoprecipitation and Western Blot Analysis—HEK293 cells expressing WT, L7.60F, or L7.60I receptors were homogenized in a glass homogenizer in ice-cold buffer composed of 20 mM Na-HEPES (pH 8.0), 2 mM MgCl₂, 1 mM EDTA, containing a protease inhibitor mixture (1.04 mM 4-(2-aminoethyl)benzenesulfonyl fluoride, 0.8 µM aprotinin, 20 µM leupeptin). After sedimentation at 1000 x g for 5 min at 4 °C to remove nuclear debris, the supernatant was collected and sedimented at 17,000 x g for 20 min at 4 °C. The pellet (P2 membrane fraction) was then solubilized using CHAPS hydrate detergent according to the method described by Houston and Howlett (2). The immunoprecipitation of the CB₁ receptor and associated proteins from the detergent-solubilized extract was performed as described previously (4, 5).

Immunoprecipitated samples were subjected to PAGE on 0.1% SDS, 10% polyacrylamide, 6 M urea gels. Electrophoretic transfer of proteins from the gel to polyvinylidene difluoride membranes was carried out in 10 mM CAPS buffer with 0.01% SDS (pH 11), for 12 h (0–4 °C) at 20 V using a Bio-Rad Trans-Blot cell. Blots were incubated with affinity-purified anti-CB₁-1–14) combined with the indicated anti-G_i antibody in blocking buffer for 3 h at room temperature (4, 5).

Neuron Preparation and Electrophysiological Recording of Ca²⁺ Currents—Superior cervical ganglion (SCG) neurons were isolated from adult male Wistar rats as described previously (21) with minor modifications. Briefly, isolated superior cervical ganglia were enzymatically digested and plated onto poly-L-lysine precoated 35-mm dishes in minimum essential medium with 10% fetal calf serum, 1% glutamine, and 1% penicillin/streptomycin. After 3–4 h at 37 °C in 5% CO₂ plasmid cDNA (100 ng/µl) encoding hCB₁, hCB₁-L7.60I or hCB₁-L7.60F was microinjected directly into the nucleus of single SCG neurons. The pEGFP-N1 plasmid (10 ng/µl) encoding the enhanced green fluorescent protein was used as a co-injection marker (21). For G-protein reconstitution experiments, pertussis toxin-insensitive G_oA(C351G) and G_i3(C351G) subunits were each microinjected with ₁ and ₂ at a ratio (mass) of 0.5:1.25:1.25. Neurons were treated overnight with 500 ng/ml pertussis toxin (PTX). Ca²⁺ currents were recorded from neurons at room temperature 16–20 h after injection using the whole-cell patch clamp technique (21). Ca²⁺ currents were elicited by voltage steps from -80 mV to +5 mV. A double-pulse protocol consisting of two 25-ms steps to +5 mV was used to elicit Ca²⁺ currents. The second voltage step to +5 mV was preceded by a 50-ms voltage step to +80 mV in order to remove the voltage-dependent, G-protein-mediated inhibition of the Ca²⁺ channels. Current amplitudes were measured isochronally as the average current between 10 and 11.8 ms after the voltage step to +5 mV. To isolate Ca²⁺ currents for whole-cell recording, cells were bathed in external and internal solutions as described previously (21). Neurons were superfused with external solution or with external solution containing D-Trp⁸ somatosatin-14 or WIN-55,212-2 using the SF-77B Perfusion Fast-Step device (Warner Instruments).

Data and Statistical Analysis—Results are presented as means ± S.E. Data from radioligand binding and GTPS assays were analyzed, and curves were generated through nonlinear regression analyses (or nonlinear analysis, when appropriate) with Prism software (Prism GraphPad, San Diego). K_d, B_max, E_max, and EC₅₀ values were calculated. Statistical analysis of variance (one-way analysis of variance) and the Bonferroni multiple comparison post hoc tests were performed for B_max and E_max with GraphPad Prism. Two-tailed Student's t tests were performed on the log K_d values, log EC₅₀ values ±95% confidence limits and for analyses of Ca²⁺ currents. p values <0.05 were defined as statistically significant.

Molecular Modeling, Receptor Model Construction
Amino Acid Numbering System—The amino acid numbering scheme proposed by Ballesteros and co-workers (14) was used. Sequence numbers used are human CB₁ sequence numbers unless otherwise noted (14). Residues in the intracellular extension of TMH7 (H8) are numbered here as if they are part of TMH7 following the literature precedent set by Prioleau et al. (13).

WT CB₁ Receptor Model Construction—Using the Loopy program within the protein structure modeling suite, Jackal 1.5 (J. Z. Xiang and B. Honig, Columbia University), extracellular (EC-1 His¹⁸¹–Ser¹⁸⁵, EC-2 Cys²⁵⁷–Glu²⁷³, and EC-3 Gly³⁶⁹–Lys³⁷⁶) and intracellular loops (IC-1 Ser¹⁴⁶–Arg¹⁵⁰, IC-2 Pro²²¹–Val²²⁸, and IC-3 Ala³⁰¹–Pro³³²) as well as portions of the N and C termini were added to our previously built transmembrane helix (TMH) bundle model of the CB1 receptor (65). The Modeler program was then used to refine loop structures (22, 23). The chosen loop configurations for the final bundle model were those that produced a low value of the modeler objective function. Moreover, because of their close spatial proximity, the EC1 and EC3 loops were run concurrently.

EC-2 Loop—One of the significant sequence divergences between rhodopsin and CB₁ is in the second extracellular (EC-2) loop region. This loop in CB₁ is shorter than in rhodopsin and is missing the conserved disulfide bridge between the cysteine in EC-2 and C3.25 in TMH 3 of rhodopsin. Instead, there is a Cys residue at the extracellular end of TMH4 in CB₁ and a Cys near the middle of the EC-2 loop that experiments suggest may form a disulfide bridge (24). Consequently, the position of the EC-2 loop with respect to the binding site crevice in CB₁ around TMHs 3–4–5 is likely to be quite different from that in rhodopsin. For this reason, the refined EC-2 loop (Cys²⁵⁷–Glu²⁷³) structure built by Modeler was removed, and the conformation of this loop was calculated using the Biased Scaled Collective Variable in Monte Carlo method (25, 26). The aqueous environment of the EC-2 loop was modeled during these calculations with a recently developed implicit solvent model that is based on a screened Coulomb potential formulation (the SCP-ISM) (27, 28). The EC-2 loop was modeled with an internal C4.66(257)-Cys²⁶⁴ disulfide bridge based upon mutation results from the Farrens laboratory (see Ref. 24), which show that these two cysteines are required for high level expression and receptor function.

IC-3 Loop—The CB₁ IC-3 loop is much longer than the corresponding sequence in rhodopsin. NMR experiments have been performed on a peptide fragment comprised of the CB1 sequence span from the intracellular end of TMH5 to the intracellular end of TMH6 in micelles (29). This study suggested that part of the IC-3 loop is -helical. This region occurs after the intracellular end of TMH5 (K5.64(300)) and consists of a short -helical segment from Ala³⁰¹ to Arg³⁰⁷, followed by an elbow region (Arg³⁰⁷–Ile³⁰⁹) and an -helical segment (Gln³¹⁰–Ser³¹⁶) up to a Ile–Ile–Ile (Ile³¹⁷–Ile³¹⁹) in IC-3. Based on these results, we replaced the initial Modeler-built IC-3 loop with this -helix-elbow--helix region, and then the rest of IC-3 loop (Ile³¹⁷–Pro³³²) was re-built and optimized using Modeler.

N and C Termini—The N and C termini were added last to the model. The N terminus of CB₁ is 111 residues in length. A shorter portion of this terminus from Asn⁹⁵ to Asn¹¹² (NIQC-GENFMDIECFMVLN) was added to the CB₁ model using Modeler. A C-terminal fragment Ser⁴¹⁴–Gly⁴²⁷ (SCEGTAQ-PLDNS-MG), which contains a putative palmitoylation site at Cys⁴¹⁵ (24), was also added using Modeler, and Cys⁴¹⁵ was palmitoylated.

Embedding the CB₁ Receptor Model in a Lipid Bilayer—To build a receptor plus phospholipid bilayer system, we used a snapshot from a previous simulation of 1-palmitoyl-2-oleoylphosphatidylcholine (POPC). In these previous calculations, a patch of hydrated POPC was equilibrated for 3 ns at 310 K. The system was constructed using the VMD membrane and solvate plug-ins. The constant area/pressure NPAT ensemble was employed using a value of 68.0 Ų as set via the VMD plug-in. This compares to earlier calculated values of 65.5 Ų (30) to 72.2 Ų (31) at 300 K. Experimental values ranged from 63–66 Ų and depended on the temperature (32–34). The CB₁ receptor model (including all extracellular and intracellular loops) was positioned in the bilayer such that TMH4 was perpendicular to the bilayer surface, and the ends of the helices were placed approximately at the water-lipid interface. Water and POPC molecules that overlapped with the protein were removed. The remaining system contained 240 POPC molecules, 19,737 water molecules, the CB₁ receptor, and 11 chloride ions, the latter of which were added to achieve charge neutrality for this system, for a total of 96,074 atoms.

Molecular Dynamics Simulations—The NAMD2 (35) molecular simulation package along with the CHARMM27 parameter set (36–38) was employed for the molecular minimizations and dynamics calculations using a TIP3P water model. The system, as described above, was conjugate gradient energy-minimized (to a gradient of less than 0.05 kcal/mol) to first remove unfavorable contacts and then allow the protein itself to relax. The protein and lipid were first fixed, and the waters were energy-minimized for 5000 steps; then the protein and waters were fixed, and the lipids were energy-minimized for 5000 steps. The CB₁ heavy atoms were then fixed, and the system was energy-minimized for 5000 steps. The helix backbone atoms of CB₁ were then fixed and energy-minimized for 5000 steps. Finally, the system was energy-minimized for 20,000 steps without constraints.

Keeping the minimized CB₁ fixed, the rest of the system was warmed slowly in 10 K increments for 20 ps per increment to 310 K at constant volume (NVT ensemble), using Langevin coupling to a heat bath (39), running 20 ps per interval. Then 1 ns of NPT dynamics was performed to pack the lipids near the protein. The system was re-minimized and again slowly warmed in 10 K increments for 20 ps per increment to 310 K. Next, during an additional 0.1 ns of 310K NVT dynamics, the heavy atoms in CB₁ were constrained by a harmonic force (k = 0.5 kcal/mol Ų). Using the NVT ensemble at 310 K, the heavy atom constraints on the receptor were released in five steps (k = 0.4, 0.3, 0.2, 0.1, and 0.05 kcal/mol/Ų) of 0.1 ns each. The ensemble was switched to NPAT (310 K/1 atm) and run for 1 ns with strong coupling to the pressure bath. From this starting point, the strong coupling of the temperature and pressure bath was reduced, and 9 ns of production was performed.

L7.60F and L7.60I Mutants—Using the CHARMM (37) modeling package, L7.60F and L7.60I mutations were generated from the starting CB₁ WT receptor model. Identical system minimization, warming, and productions runs to that described above for the WT CB₁ were performed.

Analysis of Molecular Dynamics Trajectories—Analysis was performed with the VMD package and associated plug-ins (39) and with analysis scripts developed in-house. For the 9 ns of production runs, the root mean square deviation from the initial structure was calculated. The Hbond analysis facility of VMD was used to analyze the trajectories for the incidence of hydrogen bonding. Interactions for which the heteroatom to heteroatom distance was 3.5 Å or less, with heteroatom-H-heteroatom angles between 180 to 130° were considered hydrogen bonds. The secondary structure analysis facility in VMD, STRIDE (40), was used to analyze the structure of H8 in WT CB₁ and each of the mutants.

	RESULTS

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Radioligand Binding of WT and L.760 Mutant CB₁ Receptors—HEK293 cell lines expressing the WT hCB₁ receptor or L7.60I or L7.60F mutant receptors were generated, and cell surface receptors were visualized with immunocytochemistry (Fig. 2). The WT hCB₁ and mutant receptors displayed comparable levels of expression. Saturation binding analysis with [³H]rimonabant showed that the WT hCB₁ and L7.60I and L7.60F mutant receptors possess similar receptor levels with no change in affinity for rimonabant (Fig. 3A). The B_max values (fmol/mg membrane protein) of 1700 ± 80 (n = 4), 1502 ± 118 (n = 4), and 1209 ± 80 (n = 3) and the K_d values (nM) of 1.9 ± 0.27, 1.8 ± 0.38, and 1.2 ± 0.17 for WT hCB₁, L7.60I, and L7.60F mutants, respectively, were not significantly different. Similar results were obtained with [³H]CP-55,940 binding analysis (Fig. 3B). The receptor levels (fmol/mg membrane protein) of 1320 ± 93 (n = 3), 1431 ± 63 (n = 3), and 1417 ± 158 (n = 3) and the K_d values (nM) 1.5 ± 0.35, 1.4 ± 0.18, and 2.1 ± 0.58 of the WT hCB₁, L7.60I, and L7.60F mutant receptors, respectively, were not significantly different from each other nor from binding levels obtained with [³H]rimonabant.

View larger version (78K): [in this window] [in a new window]   FIGURE 2. Immunolabeling of HEK293 cells stably expressing WT hCB1 receptor (A), L7. 60I (B), or L7.60F (C) mutant hCB1 receptors are shown. Living cells were labeled with a rabbit anti-CB1 receptor (1–77) antibody and visualized with Alexa Fluor 488-conjugated goat anti-rabbit. Cell surface labeling observed on cells expressing mutant receptors was similar to that of cells expressing WT CB1 receptors. D, immunolabeling was blocked by preincubation with 100 µg/ml of the peptide against which the antibody was raised. Scale bars are 10 µm.

View larger version (20K): [in this window] [in a new window]   FIGURE 3. Saturation binding analysis with [3H]rimonabant (A) and [3H]CP-55,940 (B) of the WT human CB1 receptor (), L7. 60F (), or L7.60(404)I () mutant hCB1 receptors. Experiments were performed with membranes of stably transfected HEK293 cells. Nonspecific binding was determined in the presence of excess nonradiolabeled ligands. No significant difference was found in binding affinities and relative levels of receptor expression between the WT receptor and the mutant CB1 receptors. Data are means ± S.E. of 3–4 independent experiments each performed in triplicate.

H8 Mutations Impair Agonist-stimulated Efficacy—[³⁵S]GTPS binding to G-proteins was used to determine the ability of structurally distinct cannabinoid agonists (WIN-55,212-2, CP-55,940, and HU-210) to activate G-proteins in cells expressing the WT hCB₁, L7.60I, or L7.60F mutant receptors (Table 1 and supplemental Fig. 10). In experiments with membranes isolated from untransfected HEK293 cells, no stimulation by WIN-55,212-2, CP-55,940, or inhibition by rimonabant was detected (data not shown) (20). Compared with the [³⁵S]GTPS binding activities (dpm) of WT hCB₁ (809.67 ± 83.34 (n = 18)) and the L7.60F mutant (694.45 ± 65.32 (n = 11)), the L7.60I mutant receptor appeared to exhibit a somewhat greater (931.40 ± 146.82 (n = 10)) basal binding activity. Nevertheless, agonist-independent basal constitutive activity levels for the three receptors were not significantly different from each other. Using this same method, differences in basal activity could be detected between mutants of the mouse CB₁ receptor (20). The maximal stimulation of [³⁵S]GTPS binding (E_max) values obtained for WT hCB₁ receptor were 95.8 ± 1.8% (n = 3) with the potent cannabinoid analog, HU-210, 144 ± 3.1% (n = 3) with the bicyclic CP-55,940, and 132 ± 1.0% (n = 7) with the aminoalkylindole WIN-55,212-2 (Table 1). Compared with the WT hCB₁ receptor, both L7.60I and L7.60F mutations caused a significant reduction (p < 0.001, n = 3) in the maximal stimulation by CP-55,940 and WIN-55,212-2 (Table 1 and supplemental Fig. 10, B and C). The L7.60I mutation also caused a significant reduction (p < 0.001, n = 4) in the maximal stimulation by HU-210 (Table 1 and supplemental Fig. 10A). The potencies of CP-55,940 and HU-210 for both mutations were not significantly different from WT, whereas a small but significant increase was observed in the potencies of WIN-55,212-2.

View this table: [in this window] [in a new window]   TABLE 1 Cannabinoid agonist efficacies and potencies for stimulation of [35S]GTPS binding Maximal stimulation and potencies were determined in [35S]GTPS binding experiments using membranes from stably transfected HEK293 cells expressing WT hCB1 or mutant receptors. Data are means and corresponding 95% confidence limits (in parentheses) of at least three independent experiments each performed in triplicate. Statistical analysis is as follows: for Emax one-way analysis of variance followed by Bonferroni's post hoc test; for EC50, two-tailed Student's t test on log values.

View this table: [in this window] [in a new window]   TABLE 2 Effect of rimonabant on [35S]GTPS binding Minimal stimulation and potencies were determined in [35S]GTPS binding experiments using membranes from stably transfected HEK293 cells expressing WT hCB1 or mutant receptors. Data are means and corresponding 95% confidence limits of at least three independent experiments each performed in triplicate. Statistical analysis is as follows: for Emax one-way analysis of variance followed by Bonferroni's post hoc test; for EC50, two-tailed Student's t test on log values.

View larger version (46K): [in this window] [in a new window]   FIGURE 4. Receptor-G-protein association demonstrated by co-immunoprecipitation of G subtypes with the hCB1 receptor from CHAPS-solubilized HEK293 membranes. Each lane was loaded with immunoprecipitated protein from CHAPS extract of cell membranes as described (100 µg of protein), and Western blot analysis was performed. A representative immunoblot stained with anti-CB1-(1–14) plus anti-Gi1 (A), anti-Gi2 (B), or anti-Gi3 (C) antibodies. A and B, 1st and 3rd lanes, WT; 2nd lane, L7.60F (F); 4th lane, L7.60I (I); C, 1st lane, WT; 2nd lane, L7.60F (F); 3rd lane, L7.60I (I). D, Western blots were scanned, and the ratio of the density of the Gi band to the CB1 receptor band was calculated. The Gi/CB1 ratios for the mutant receptors were normalized to the WT value as 100%, and the mean ± S.E. was calculated from three independent experiments.

View larger version (95K): [in this window] [in a new window]   FIGURE 5. Agonist-induced internalization of hCB1 receptor expressed in stably transfected HEK293 cells. WT hCB1 receptor (A–C) or L7.60I mutant receptor (D and E) or L7.60F mutant receptor (F and G) were incubated in the absence (A, D, and F) or presence of 1 µM CP-55,940 for 30 min (B, E, and G) or 1 h (C). Surface hCB1 receptors were labeled with CB1-(1–77) polyclonal antibody. This figure is representative of three individual cell culture experiments. Scale bars are 10 µm.

View larger version (14K): [in this window] [in a new window]   FIGURE 6. CB1 receptor-mediated sequestration of G-proteins leading to the inhibition of somatostatin signaling by hCB1 WT and L7.60I and L7.60(404)F mutants. 1st bar, somatostatin (0.1 µM) inhibited the Ca2+ current in uninjected SCG neurons through endogenous somatostatin receptors. In neurons injected with the WT hCB1 receptor (2nd bar), somatostatin could not efficiently inhibit Ca2+ current, a measure of G-protein sequestration by the hCB1 receptor. 3rd and 4th bars depict the inhibition of somatostatin signaling by L7.60I and L7.60F mutants, respectively. No significant differences were found between the hCB1 WT and mutant receptors. The data are expressed as means ± S.E. from 9 recordings.

H8 Mutations Shorten the Time Course and Reduce the Magnitude of Ca²⁺ Channel Inhibition—When expressed in SCG neurons, agonist-activated WT hCB₁ receptors efficiently couple to PTX-sensitive G_i/o proteins and inhibit voltage-dependent, N-type Ca²⁺ channels. We tested whether the L7.60I and L7.60F mutations in helix 8 would alter Ca²⁺ channel inhibition when expressed in SCG neurons. Activation of WT hCB₁ receptors by 1 µM WIN-55,212-2 resulted in a 67.8 ± 2.8% inhibition of the Ca²⁺ current in SCG neurons (Table 3). The Ca²⁺ current inhibition by WIN 55,212-2 was significantly reduced by the L7.60F and L7.60I mutant receptors, 59.9 ± 1.8 and 54.3 ± 4.0%, respectively (Table 3, no PTX). Thus, a good correlation was seen between G-protein activation in HEK293 cells (Table 1 and supplemental Fig. 10) and Ca²⁺ current measurements in SCG neurons (Table 3). In addition, the time to reach 90% inhibition of the maximal Ca²⁺ current (seconds) after the application of WIN-55,212-2 was significantly faster for the L7.60F (45.6 ± 5.5) and L7.60I (51.4 ± 4.0) mutant receptors compared with the WT hCB₁ receptor (70.4 ± 5.9) (Table 4, no PTX). Taken together, these results suggest that the H8 mutations alter both the magnitude and time course of Ca²⁺ channel effector coupling.

View this table: [in this window] [in a new window]   TABLE 3 Ca2+ current inhibition (%) by WIN 55,212-2 in SCG neurons expressing wild type hCB1 or H8 mutant receptors reconstituted with PTX-insensitive GoA(C351G)/G12 or Gi3(C351G)/G12 N-type Ca2+ channel inhibition and reconstitution of hCB1 coupling with PTX-insensitive G proteins were performed. Each receptor WT hCB1, L7.60F, or L7.60I was expressed in SCG neurons with PTX treatment and expression of PTX-insensitive G proteins GoA(C351G)12 or Gi3(C351G)12. N-type Ca2+ channel inhibition by WIN-55,212-2 was significantly lower in L7.60F and L7.60I mutants compared with WT hCB1. Inhibition of Ca2+ channels by the WT hCB1 receptor was abolished by overnight treatment with PTX. PTX-insensitive G proteins GoA or Gi3 reconstituted the WIN55,212-2-induced inhibition of Ca2+ channels by WT hCB1 receptors in SCG neurons treated overnight with PTX. Inhibition of N-type Ca2+ channels by the L7.60F mutant receptor was reconstituted by GoA or Gi3 in PTX-treated SCG neurons. However, WIN55,212-2-induced inhibition of Ca2+ channels was smaller for L7.60I mutant receptor reconstituted with Gi3 compared with reconstitution with GoA.

View this table: [in this window] [in a new window]   TABLE 4 Time to 90% Ca2+ current inhibition (seconds) by WIN 55,212-2 in SCG neurons expressing wild type hCB1 or H8 mutant receptors reconstituted with PTX-insensitive GoA(C351G)/G12 and Gi3(C351G)/G12 The time course of WIN55,212-2-induced inhibition of N-type of Ca2+ channels was significantly faster by both the L7.60F and L7.60I mutant receptors (No PTX column). Reconstitution with GoA greatly prolonged the time to 90% inhibition of the Ca2+ current by WT and mutant receptors. Reconstitution with Gi3 mimicked the time course for WT hCB1 by the reconstituted WT hCB1 and the L7.60IF mutant but not the L7.60I mutant. The data are expressed as means ± S.E., and significance was tested using a two-tailed Student's t test to compare the groups.

CB₁-mediated Ca²⁺ current inhibition was reconstituted in PTX-treated SCG neurons by co-injection of WT hCB₁ and PTX-insensitive G_oA(C351G)/G₁₂ or G_i3(C351G)/G₁₂ cDNAs. G₁₂ has been shown previously to provide optimal coupling between other GPCRs and N-type Ca²⁺ channels (47). Ca²⁺ currents were inhibited 65.5 ± 3.7% by WIN-55,212-2 in PTX-treated SCG neurons reconstituted with G_oA(C351G) (Table 3, column 2), which was no different from WT hCB₁ using endogenous G_i/o proteins (Table 3, column 1). However, the magnitude of Ca²⁺ current inhibition of WT hCB1 reconstituted with G_i3(C351G) (56.3 ± 4.8%) was significantly lower (Table 3, column 3) than WT hCB₁ using endogenous G_i/o proteins. G_oA(C351G)/G₁₂ or G_i3(C351G)/G₁₂ reconstitution of Ca²⁺ current inhibition by WIN-55,212-2 was not different between WT hCB₁ (65.5 ± 3.7% for G_oA and 56.3 ± 4.8 for G_i3) and the two mutant receptors L7.60F (63.8 ± 3.5% for G_oA and 56.9 ± 5.4% for G_i3) and L7.60I (63.7 ± 5.7%, for G_oA and 43.6 ± 5.8%, for G_i3) (Table 3). However, WIN-55,212-2 inhibition of the Ca²⁺ current was significantly smaller for the L7.60I mutant when reconstituted with G_i3(C351G) (43.6 ± 5.8%) compared with reconstitution with G_oA(C351G) (63.7 ± 5.7%) (Table 3).

The most striking differences were in the time required for WIN-55,212-2 to inhibit the Ca²⁺ current. The time to 90% inhibition of the Ca²⁺ current (seconds) by WIN-55,212-2 was significantly faster for both the L7.60F (45.6 ± 5.5) and L7.60I (51.4 ± 4.0) mutant receptors compared with the WT receptor (70.4 ± 5.9) (Table 4). Unexpectedly, a significantly longer time was needed for Ca²⁺ current inhibition by both mutant and WT hCB₁ receptors when reconstituted with G_oA(C351G)/G₁₂ (Table 4). Thus, the time to 90% inhibition of the Ca²⁺ current (seconds) by WIN-55,212-2 for WT hCB₁ receptors coupled to endogenous G_i/o proteins was 70.4 ± 5.9 versus 130.5 ± 11.0 for WT hCB₁ receptors when reconstituted with G_oA(C351G)/G₁₂. For the L7.60F mutant coupled to endogenous G_i/o proteins, the time to 90% inhibition of the Ca²⁺ current (seconds) by WIN-55,212-2 was 45.6 ± 5.5 versus 109.5 ± 6.1 for L7.60F when reconstituted with G_oA(C351G)/G₁₂ (Table 4). For the L7.60I mutant coupled to endogenous G_i/o proteins, the time to 90% inhibition of the Ca²⁺ current (seconds) by WIN-55,212-2 was 51.4 ± 4.0 versus 96.7 ± 10.9 for L7.60I when reconstituted with G_oA(C351G)/G₁₂ (Table 4). Reconstitution with G_i3(C351G) mimicked the time course of Ca²⁺ current inhibition by both the WT hCB₁ receptor and the L7.60F mutant (Table 4). For WT hCB₁ the time course of Ca²⁺ current inhibition was 79.4 ± 8.4 for G_i3 reconstitution versus 70.4 ± 5.9, for endogenous G_i/o proteins. For the L7.60F mutant the time course of Ca²⁺ current inhibition was 57.6 ± 7.4 for G_i3 reconstitution versus 45.6 ± 5.5% for endogenous G_i/o proteins. However, neither G_oA(C351G) (96.7 ± 10.9) nor G_i3(C351G) (79.3 ± 12.4) reconstitution mimicked the time course of Ca²⁺ current inhibition by the L7.60I mutant (51.4 ± 4.0) (Table 4).

We also found a surprising difference between G_i3 and G_oA in the reconstitution of WT hCB₁ constitutive activity as measured by the basal facilitation ratio (Table 5). The basal facilitation ratio is the amplitude of the Ca²⁺ current in response to the second voltage step divided by the amplitude of the Ca²⁺ current in response to the first voltage step of the double-pulse protocol. The second voltage step to +5 mV is preceded by a voltage step to +80 mV that reverses the voltage-dependent, G-mediated inhibition of the Ca²⁺ channels. Constitutive activation of G-proteins by GPCRs will result in tonic inhibition of Ca²⁺ channels and consequently produce a basal facilitation ratio >1. The more constitutively active a GPCR is the larger the basal facilitation ratio will be and vice versa. The basal facilitation ratios of neurons expressing the WT hCB₁ (1.21 ± 0.03) or L7.60F (1.22 ± 0.02) or L7.60I (1.28 ± 0.06) mutant receptors were not significantly different (Table 5). These results are well correlated to our findings in HEK293 cells. In neurons reconstituted with G_oA the basal facilitation ratio was significantly smaller for both WT hCB₁ (1.07 ± 0.03) and L7.60F mutant (1.12 ± 0.04) but not for the L7.60I mutant receptor (1.26 ± 0.09) (Table 5). In contrast, reconstitution with G_i3 produced a significantly enhanced basal facilitation ratio (1.49 ± 0.10, 1.91 ± 0.23, and 1.91 ± 0.25) for all three receptors (WT, L7.60F, and L7.60I, respectively), (Table 5). These results suggest that hCB₁ coupling to G_i3 is responsible for the constitutive activity of hCB₁ receptors and consequently the tonic inhibition of voltage-dependent, N-type Ca²⁺ channels in SCG neurons. These results also suggest that hCB₁ coupling to G_oA will not result in tonic Ca²⁺ channel inhibition but will support inhibition of Ca²⁺ channels in the presence of an agonist.

View this table: [in this window] [in a new window]   TABLE 5 Differences in constitutive activity as measured by the basal facilitation ratio Reconstitution with Gi3 significantly enhanced the basal facilitation ratio for WT hCB1, L7.60F, and L7.60I. Each receptor hCB1, L7.60F, or L7.60I was expressed in SCG neurons without PTX treatment or with PTX and expression of PTX-insensitive G proteins GaoA(351G)12 or Gi3(C351G)12. The basal facilitation ratio was similar between WT hCB1, L7.60F, and L7.60I receptors. However, reconstitution with GoA significantly reduced the basal facilitation ratio for both WT CB1 and L7.60F mutant but not for the L7.60I mutant compared to without PTX treatment. Reconstitution with Gi3 significantly enhanced the basal facilitation ratio for WT hCB1, L7.60F, and L7.60I receptors.

Comparison of WT CB₁ H8 Conformation in POPC with Mutant CB₁ H8 Conformation in POPC—As both mutant receptors showed differences in G protein coupling versus WT CB₁, we tested the effect of a membrane bilayer environment on H8 by incorporating the models of WT CB₁ versus each L7.60 mutant into a pre-equilibrated bilayer model. Fig. 7 illustrates simulation results for the position of WT CB₁ H8 in a POPC bilayer. This simulation suggests that the amphipathic helix, H8 (which is situated approximately parallel to the membrane plane), lies in the phosphate/glycerol region of the bilayer and that water penetrates into the phospholipid head group region. This extent of water penetration has been noted for other lipid bilayers, such as 1-stearoyl-2-docosahexaenoyl-sn-glycero-3-phosphocholine (18:0/22:6 PC) (48). The distribution of side chains in H8 of rhodopsin exhibits an amphipathic pattern, where charged polar groups cluster on one side and hydrophobic ones cluster on the other side of the helix (9). This same pattern is seen in Fig. 7 for CB₁ H8. Hydrophobic residues, L7.60, F7.64, and F7.68 lie in the phosphate/glycerol region of the bilayer facing up toward the helical bundle and the acyl chain region of bilayer. Palmitoylated C-terminal residue Cys⁴¹⁵ also resides in this same region, whereas the palmitate side chain extends into the acyl chain region of bilayer. Polar residues, R7.61, H7.62, and R7.65 each extend from the phosphate/glycerol region toward the aqueous region of the bilayer (i.e. the cytoplasm). Residues S7.66(410) and D7.59(403) are on the same side of H8, pointing toward TMH1–2.

View larger version (70K): [in this window] [in a new window]   FIGURE 7. Simulation results for the position of WT hCB1 H8 in a POPC bilayer. For the intracellular extension of TMH7, H8 is shown here in magenta. A palmitoylation site at Cys415 is indicated, with its palmitic acid chain shown in cyan. Residues near and along the elbow region between TMH7 and H8 as well as H8 residues are shown here in yellow. The residue mutated in the present study, L7.60, is highlighted in green. POPC choline nitrogens are shown in blue contoured at their van der Waals radii. Phosphorus atoms of the POPC phosphate region are shown in gold, and the POPC glycerol oxygen atoms are shown in red with both contoured at their van der Waals radii. Water molecules are shown in stick representation. Lipid acyl chains have been removed here for simplicity. This figure shows that the amphipathic helix, H8 (which is situated essentially parallel to the membrane plane), lies in the phosphate/glycerol region of the bilayer and that water penetrates into the phospholipid head group region.

View larger version (58K): [in this window] [in a new window]   FIGURE 8. Results for the position of H8 relative to TMH1 (green) in the L7.60I and L7.60F mutants versus WT CB1. The IC-1 loop is shown in purple. Here the average structure from the last 5 ns of the trajectory is superimposed onto the lower eight residues of TMH7(N7.49–R7.56) in the WT bundle. It is clear here that the position of H8 is different for each system. Both the L7.60F (yellow) and L7.60I (red) mutant H8s lie lower than WT H8 (white), but the L7.60F mutant (yellow) is clearly packed closer to TMH1 here.

The NPXXY(X)_5,6F motif in rhodopsin, which links by aromatic interaction, Y7.53(306) in the NPXXY motif of TMH7 to F7.60(313) in H8, has been proposed to provide structural constraints in rhodopsin that rearrange in response to photoisomerization (12). This same interaction has been shown to be important for the switching of the 5HT-2C receptor among multiple active and inactive conformations (13). The CB₁ receptor lacks a NPXXY(X)_5,6F motif, as residue 7.60(405) is a Leu in CB₁. However, the L7.60F mutation results in the introduction of this motif into CB₁. An analysis over the last 5 ns of the L7.60F trajectory revealed that the distance between aromatic ring centers of Y7.53 and F7.60 met the criterion for aromatic stacking distance (4.5 Å < « 7.0 Å) over greater than 90% of this trajectory (51). The average energy of interaction between residue 7.60 and Y7.53 over the last 5 ns of the trajectories for WT CB₁, L7.60F, and L7.60I was found to be -1.59 (0.64), -4.26 (0.84), and -2.28 (0.47) kcal/mol, respectively. The larger energy of interaction for the L7.60F mutant is likely a reflection of the aromatic stacking in which it is engaged with Y7.53. Taken together, the enhanced interaction of the L7.60F mutant with the V1.53/V1.56 groove and the aromatic stacking interaction with Y7.53 that can be maintained by the L7.60F mutant are likely to result in a better packing of H8 in the L7.60F against TMH1 as suggested in Fig. 8.

Further analysis of the geometries of H8 in WT CB₁, L7.60I, and L7.60F mutants revealed that these helices also differ in their geometries surrounding the elbow region (R7.56 to K7.58) and in their hydrogen bond patterns along H8. Results of STRIDE secondary structure analysis of the region from L7.55 to D7.59 over the last 5 ns (1000 frames) of the WT, L7.60F and L7.60I trajectories revealed that L7.55 was in a helical region in all three receptors (992, 998, and 1000 frames, respectively). At R7.56, WT CB₁ differed from the L7.60F and L7.60I mutants because structures were helical for WT CB₁ in only 69 of 1000 frames, whereas for the L7.60F and L7.60I mutants this residue was in a helical region for 831 and 983 out of 1000 frames, respectively. S7.57 was found to be in a coil region for all three receptors, whereas K7.58 was helical in 751 out of 1000 frames for WT CB₁ and helical for all 1000 frames for the L7.60F and L7.60I mutants. D7.59 and residue 7.60 were in a helical region for all three receptors. As Mukhopadhyay et al. (15) have shown the WT CB₁ residue of primary importance in determining apparent affinity for G-protein is R7.56, the fact that R7.56 is not in a helical region for WT CB₁ on average compared with the 7.60 mutants may contribute to the differential interaction with G-proteins reported here for WT CB₁ and the 7.60 mutants. Table 6 illustrates the backbone and values for the elbow region in each receptor. Analysis of the distribution of backbone and angles over the last 5 ns of the trajectory also revealed several important differences in elbow region geometry between WT CB₁ and the two L7.60 mutants. The major difference in values occurs for WT residue R7.56 (WT -107.3° (14.2°); L7.60F -81.1° (10.6°); L7.60I -86.0° (17.6°)). Although the value for WT K7.58 appears to be similar to those values for the mutants, we found that this angle actually is made up of two populations ( = -50° and -105°), whereas the angles of the mutants clustered around the single value reported in Table 3. There are several differences to be noted in values as well for the elbow region. The angle for WT CB₁ L7.55 is smaller than in either mutant (WT -6.06° (15.5°), L7.60F -48.6° (11.3°), and L7.60I -35.4° (14.5°)), whereas the WT values for residues S7.57 and K7.58 are higher than in the mutants (S7.57, WT 159.1° (37.7°), L7.60F 122.7° (15.2°), and L7.60I 114.2° (11.8°); K7.58, WT -70.3° (18.2°), L7.60F -42.7° (10.6°), and L7.60I -43.1° (11.8°)). For WT S7.57, as the large standard deviation suggests, we found that this angle actually is made up of two populations ( = -165°and +120°). Although the +120° value is very close to the corresponding values for the mutants, the = -165° angle is quite different from the mutant S7.57 values.

View larger version (34K): [in this window] [in a new window]   FIGURE 9. The hydrogen bonding patterns along the H8 backbone for WT hCB1 and the L7.60I and L7.60F mutant structures averaged over the last 5 ns of each run. Hydrogen bonds are shown as green dotted lines. It is clear that each H8 has a distinct hydrogen bonding pattern. Of note is the result that hydrogen bonds that link H8 to the elbow region (R7.56(400)–K7.58(402)) occur in WT hCB1 less frequently than in either the L7.60I or L7.60F mutants.

Taken together, these molecular dynamics results suggest that WT CB₁, L7.60F, and L7.60I mutants each have distinct locations for H8 and distinct flexibilities in their elbow regions. Given the role that H8 appears to play in activation and G-protein coupling, our modeling studies suggest that it is likely that WT CB₁, L7.60F, and L7.60I structural differences in the H8 region may underlie functional differences.

Comparison of WT CB₁ H8 Conformation in POPC Results with H8 Peptide Studies in Micelles—In the rhodopsin-derived model of CB₁ reported here, the D7.59(403) to P7.69(413) region forms an amphipathic helix that is packed against TMH1 at its intracellular end. This structure is consistent with our early Fourier transform analysis of the CB₁ primary sequence that predicted the CB₁ 403–413 segment to be an intracellular helical domain (14) and with the crystal structure of rhodopsin (9). Significant helicity has been demonstrated when a CB₁ 400–416 peptide (401–417 rat CB₁ numbering) was present in an anionic micelle environment (SDS or phosphatidic acid), with a 3₁₀ helical structure being favored over an -helical structure (15). NMR measurements at 285 K of a short peptide comprising a portion of the predicted TMH7 region and amphipathic H8 region of the CB₁ receptor (I7.52(397)-G7.73(418), rat CB₁ sequence numbers) have been reported in DPC (dodecylphosphocholine) micelles (16). The ¹H nuclear Overhauser effect spectroscopy pattern in DPC micelles indicated a high degree of -helical structure, in particular from S7.57(402) to -M7.67(412), with a break in the helix at L7.55(400),R7.56(401). Xie and Chen (17) examined the same peptide as well as a related mutant peptide. In this study, a salt bridge was found between D7.59(404) and H7.62(407). In the modeling studies reported here, we found that this interaction did not occur possibly because H8 is packed against the CB₁ TMH bundle, and this constraint may not permit some of the interactions possible in the isolated peptide (17). Finally, it is important to note that the CB₁ 400–416 (401–417 rat) peptide has not been found to maintain helicity in an aqueous environment (15). It has been hypothesized that an environmentally driven shift in H8 conformation may occur upon receptor activation (52).

	DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS DISCUSSION REFERENCES

 
Recent studies have underscored the significance of the H8 domain of GPCRs in ligand binding, switching between the active and inactive conformations, and activation of G-proteins (13, 53, 54). Okuna and co-workers (55) recently reported that out of a set of 180 class A GPCRs, residue 7.60 is a Phe in 135 of these receptors. Mutation studies of the NPXXY(X)_5,6F motif, which links the TMH7 and H8 domain, examined the involvement of this motif in ligand recognition and signal transduction, including G-protein coupling (12, 13). Prioleau et al. (13) demonstrated that Y7.53 in the NPXXY domain of the 5HT-2C receptor influences the receptor ability to switch among multiple active and inactive conformations. Fritze et al. (12) showed that an F7.60(313)A mutation in rhodopsin causes dramatic inhibition of G_t-protein activation. In contrast, substitutions of F7.60 to alanine in the melanin-concentrating hormone type 1 (56) and angiotensin II_1A (57) receptors resulted in normal activation. Zhong et al. (58) reported a similar finding for an L7.60A/F7.61A double mutation in the oxytocin receptor. Nevertheless, in both the Tetsuka et al. (56) and Zhong et al. (58) studies, hydrophilic residues in H8 were demonstrated to play a major role in G-protein activation. Moreover, substitution of F7.60(383) in the ₁-adrenergic receptor with a positively charged histidine resulted in improper receptor biosynthesis (50).

This study demonstrates for the first time that the divergence from NPXXY(X)_5,6F to NPXXY(X)_5,6L in the CB₁ receptor contributes to proper association with G_i3 and maximal efficacy in signal transduction. This residue is also implicated in regulating the rate of CB₁ internalization process but not G_i/o protein sequestration. These results indicate that although the general consensus is that H8 plays a major role in the structure and function of its respective GPCR, the ultimate effect is dependent on the specific sequence (receptor) under study and perhaps the specific G-protein as well (50, 58). Furthermore, given the variety of effects observed in these studies, as well as an earlier set of experiments on the prostaglandin EP3 receptor (59), which demonstrated that different splicing of the C terminus produced coupling to different G-proteins, one can conclude that H8 is a region, perhaps in cooperation with certain intracellular loops, that governs G-protein specificity (50, 56, 60).

With the H8 mutant receptors, we demonstrate that this selective coupling to G-protein subtypes by chemically different cannabinoid ligands affects downstream signal transduction of CB₁ receptor. Structurally different cannabinoid agonists CP-55,940, WIN-55,212-2, and HU-210 could activate the L7.60I and L7.60F mutant receptors in [³⁵S]GTPS binding assays. However, compared with the WT hCB₁, the L7.60F mutation resulted in a reduction in cannabinoid agonist efficacies. Hence, restoring the NPXXY(X)_5,6F motif resulted in lower receptor stimulation. The L7.60I mutation exhibited severely impaired maximal receptor activation in response to CP-55,940. It is likely that CP-55,940 and WIN-55,212-2 may signal through G_i3 in addition to other G-proteins in WT CB₁, as their signaling was reduced in a ligand-dependent fashion in both the L7.60F and L7.60I mutant receptors.

Our study supports and expands upon three previous studies of the CB₁ cannabinoid receptor. Glass and Northup (3) showed that structurally different cannabinoid ligands affect maximal activation of G_i and G_o proteins in Sf9 cells, but this study did not distinguish between G-protein subtypes. Mukhopadhyay and Howlett (5) demonstrated selective coupling of CB₁ receptor peptides to G-protein subtypes and later (61) that receptor-G-protein complexes for the different subtypes are dependent on the cannabinoid ligand structure. However, these studies did not show if this selective coupling by the different ligands affects downstream signal transduction of the CB₁ receptor.

Our co-immunoprecipitation studies in HEK293 cells demonstrated that although WT CB₁ associates with G_i1, G_i2, and G_i3, the L7.60F and L7.60I mutants predominantly associate with G_i1 and G_i2. This physical association between the CB₁ receptor and the G-protein can be readily dissociated by agonists, as occurs in the typical trimeric G-protein kinetic activation model. As spontaneous dissociation might be the mechanism for constitutive activity, and dissociation is also produced by an abundance of GTPS, the observation that the mutant CB₁ receptors are readily dissociated from G_i3 suggests altered kinetics of the G-protein activation cycle for G_i3. In SCG neurons, reconstitution with G_i3 resulted in Ca²⁺ current inhibition and significantly higher basal facilitation. This suggests that the interaction between G_i3 and the mutant receptors may be transient. Taken together, the F7.60 mutations may not only interfere with recognition of G_i3 but enhance the rate of dissociation of G_i3 from subunits. This reduction in coupling to G_i3 seen here is likely to be the result of an alteration in the "elbow" region of CB₁.

The inverse agonist, rimonabant, has been shown to promote and/or stabilize an inactive state of the receptor-G-protein complex, in which nucleotide exchange is prohibited (7). Mimicking the NPXXY(X)_5,6F motif by the L760F, but not by the L7.60I, mutation maintained maximal inhibition by rimonabant. Our modeling studies suggested that residue 7.60 in the L7.60F mutant has higher interaction with the V1.53/V1.56 groove on TMH1 compared with the L7.60I mutant. This interaction may cause H8 and TMH1 to come closer together in the L7.60F mutant thus restricting the movement of NPXXY-H8-IC1 axis to move outward toward the G-proteins, which in return may result in reduced nucleotide exchange. Alterations in H-bond and ionic interactions between NPXXY-H8 and IC1 have been previously shown to control G-protein signaling (60). However, it is difficult to interpret the events related to rimonabant as due to inverse agonist activity because of the ability of most cells to synthesize anandamide or 2-arachidonylglycerol under certain experimental conditions (62). Thus, endocannabinoids present in the vicinity of the CB₁ receptors may provide a level of stimulation that can be competitively antagonized by rimonabant.

Following agonist binding, the mutant receptors could still be internalized, supporting previous studies that demonstrated that CB₁ internalization is pertussis toxin-insensitive and therefore does not require a prior association with G-proteins (63). However, a time course internalization study indicated that a greater number of L7.60F and L7.60I mutant receptors were internalized during a 30-min time period than WT hCB₁, suggesting that L7.60 slows the rate of CB₁ receptor internalization. As the positions and geometries of H8 in the mutant receptors have been altered (see Figs. 8 and 9), this topographical change may also influence the interactions between the mutant receptors and G-protein-coupled receptor kinases and/or -arrestins. Both the bovine rhodopsin crystal structure and results from our modeling studies indicate an intimate relationship between residue 7.60(404) and the NPXXY motif. Internalization defects have been reported for the ₂-adrenergic receptor upon mutation of Y7.53(397), whereas the ability of the mutant to stimulate maximal adenylyl cyclase activity and be normally desensitized through phosphorylation remained the same as WT (64). Although total internalization of CB₁ was previously reported to be G-protein-independent (63), it appears that coupling to G-proteins delicately regulates specific mechanisms involved in this process.

Sequestration of G-proteins is a process that requires efficient coupling by GPCRs. Previously, Nie and Lewis (21) have underlined the importance of the distal C-terminal tail of the CB₁ receptor in the sequestration of G-proteins by deleting residues 418–472 in CB₁. In SCG neurons, both the L7.60F and L7.60I mutant receptors were able to sequester G-proteins from the endogenously expressed somatostatin receptor in a similar manner to the WT. These results suggest that the NPXXY(X)_5,6F motif is not part of the structural basis for G-protein sequestration.

In summary, the present study demonstrates for the first time that the evolutionary change from NPXXY(X)_5,6F to NPXXY(X)_5,6L in the CB₁ receptor is required for association with G_i3 in its GDP-bound inactive state in which constitutive signaling is minimized and agonist efficacy is maximized. The reduction in maximal activation seen upon mutation at position 7.60 is shown to be dependent on ligand structure and appears to be profound for the bicyclic cannabinoid CP-55,940. Certain mutations at position 7.60 also have a profound effect on the CB1 inverse agonist rimonabant ability to inactivate CB1. With the G-protein reconstitution experiments, we also demonstrate for the first time that coupling of CB₁ to G_i3 is responsible for tonic inhibition of voltage-dependent, N-type Ca²⁺ channels, whereas G_o slows the rate of agonist-induced Ca²⁺ current inhibition. The L7.60 residue is also implicated in regulating the rate of CB₁ internalization process but not G_i/o protein sequestration. Our results suggest that the evolutionary modification in CB₁ to L7.60, compared with the conserved Phe residue at the same position of rhodopsin is significant for the functional coupling to G-proteins for the endocannabinoid system.

	FOOTNOTES

 
^* This work was supported by National Institute on Drug Abuse Grants DA09978, DA05274 (to M. E. A.), DA00489, DA039434 (to P. H. R.), DA03690, and DA12385 (to A. C. H.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Figs. 10 and 11.

¹ Present address: Dept. of Physiology and Pharmacology, Wake Forest University, Winston-Salem, NC 27157.

² To whom correspondence should be addressed: CA Pacific Medical Center Research Institute, 475 Brannan St., Ste. 220, San Francisco, CA 94107. Tel.: 415-600-3607; Fax: 415-600-1725; E-mail: aboodm{at}cpmcri.org.

³ The abbreviations used are: GPCR, G-protein-coupled receptor; CB, cannabinoid; WT, wild type; TMH, transmembrane helix; H8, helix 8; IC, intracellular loop; HEK293, human embryonic kidney 293; SCG, superior cervical ganglion; GTPS, guanosine 5'-3-O-(thio)triphosphate; POPC, 1-palmitoyl-2-oleoylphosphatidylcholine; PTX, pertussis toxin; CHAPS, 3-[(3-chlamidopropyl)dimethylammonio]-1-propanesulfonic acid; CAPS, 3-(cyclohexylamino)propanesulfonic acid; VMD, visual molecular dynamics; h, human.

	ACKNOWLEDGMENTS

 
We thank Dr. Ken Mackie (University of Washington) for providing CB₁ receptor antibody and blocking peptide (DA11322 and DA00286).

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