The Third Intracellular Loop Stabilizes the Inactive State of the Neuropeptide Y1 Receptor

Abstract

Constitutively active G-protein-coupled receptors (GPCRs) can signal even in the absence of ligand binding. Most Class I GPCRs are stabilized in the resting conformation by intramolecular interactions involving transmembrane domain (TM) 3 and TM6, particularly at loci 6.30 and 6.34 of TM6. Signaling by G_i/G_o-coupled receptors such as the Neuropeptide Y1 receptor decreases already low basal metabolite levels. Thus, we examined constitutive activity using a biochemical assay mediated by a G_i/G_q chimeric protein and a more direct electrophysiological assay. Wild-type (WT-Y1) receptors express no measurable, agonist-independent activation, while μ-opioid receptors (MOR) and P2Y₁₂ purinoceptors showed clear evidence of constitutive activation, especially in the electrophysiological assay. Neither point mutations at TM6 (T^6.30A or N^6.34A) nor substitution of the entire TM3 and TM6 regions from the MOR into the Y1 receptor increased basal WT-Y1 activation. By contrast, chimeric substitution of the third intracellular loop (ICL3) generated a constitutively active, Y1-ICL3-MOR chimera. Furthermore, the loss of stabilizing interactions from the native ICL3 enhanced the role of surrounding residues to permit basal receptor activation; because constitutive activity of the Y1-ICL3-MOR chimera was further increased by point mutation at locus 6.34, which did not alter WT-Y1 receptor activity. Our results indicate that the ICL3 stabilizes the Y1 receptor in the inactive state and confers structural properties critical for regulating Y receptor activation and signal transduction. These studies reveal the active participation of the ICL3 in the stabilization and activation of Class I GPCRs.

Neuropeptide Y (NPY)2 is one of the most abundant peptides in the nervous system. NPY and its receptors have been implicated in various biological functions and neuronal disorders, such as epilepsy, obesity, and anxiety. There are 5 confirmed subtypes of NPY receptors (Y1, Y2, Y4, Y5, and y6), which are class I G-protein-coupled receptors (GPCRs) coupling to G_i/G_o proteins (1). Recent evidence indicates that an unexpectedly large number of the GPCRs demonstrate constitutive activity, so they can signal even in the absence of ligand binding (2, 3, 4). However, little is known about the activation properties of Y receptors or their propensity for constitutive activation.

GPCR activation is determined by the flexibility of its transmembrane (TM) segments to move relative to each other (5). GPCRs are stabilized in the inactive state by a network of non-covalent, intramolecular interactions that restricts TM mobility (6, 7). The interactions between the (D/E)RY and X₁BBX₂X₃B motif on TM3 and TM6, respectively, are considered to be critical for GPCR activation (8). Single point mutation at X ^6.30₁ and/or X ^6.34₃ is sufficient in some GPCRs to destabilize this covalent network, thus allowing the GPCR to adopt a conformation favorable for intracellular G-protein binding. In effect, point mutation(s) at loci 6.30 and/or 6.34 frequently induce constitutive activity in class I GPCRs (6, 9–11). However, some GPCRs are resistant to constitutive activation in this way, suggesting the participation of alternative regions that regulate GPCR activation.

Y receptors are coupled to the activation of G_i/G_o proteins, hence detection of their receptor activation is complicated by the nature of G_i/G_o-protein signaling that primarily inhibits adenylyl cyclase, thereby suppressing the production of readily detectable metabolites. We circumvented this by employing a biochemical assay in which a chimeric Gα_Δ6qi4myr protein positively coupled Y1 receptor activation to the G_q-signaling pathway, increasing inositol phosphate (IP) production (12, 13). Secondly, we developed an electrophysiological assay whereby GPCR activation was measured as an increase in basal prepulse facilitation (BPF) of a co-expressed N-type voltage-dependent calcium channel (14–16). The use of both approaches maximized the detection and estimation of constitutive activity in this class of G_i/G_o-coupled receptors. However, the WT-Y1 receptor did not express significant constitutive activity, nor was it susceptible to constitutive activation by single point mutations at position 6.30 (T^6.30A) or 6.34 (N^6.34A). By chimeric substitution of functional regions, such as a complete TM or intracellular loop (ICL), into the Y1 receptor from analogous regions in the constitutively active μ-opioid receptor (MOR), we identified the structural region that stabilizes the Y1 receptor. The loss of the region that imparts the most relevant stabilizing interactions would result in the generation of a constitutively active chimeric Y1 receptor. Our findings indicate that the Y1 receptor is stabilized in the inactive state by the ICL3 region and suggest the relative importance of this atypical region for regulating the activation of other Y receptors and class I GPCRs.

EXPERIMENTAL PROCEDURES

Generation of Y1 Receptors

The subcloning of the human Y1 receptor cDNA into the eukaryotic expression plasmid pEGFP-N1 (Clontech, Heidelberg, Germany) has been described previously (17). The C-terminal fusion to the enhanced green fluorescent protein (EGFP) or yellow fluorescent protein (EYFP) allowed for fluorescence detection of the heterologously expressed receptor and does not alter the function of Y receptors (17). The human (h) Y1 receptor EYFP fusion gene was additionally subcloned into the eukaryotic expression plasmid pVitro2-mcs(hygro) (Invivo-Gen, Toulouse, France) via a BamHI and an AvrII site. Mutations were introduced with the QuikChange™ site-directed mutagenesis method (Stratagene). The entire coding region was sequenced to ensure the accuracy of all constructs. In the Y1-ICL3-MOR construct, the nucleotides coding for amino acids 232–264 of the hY1 receptor were replaced by nucleotides coding for amino acids 255–283 of the hMOR. The Y1-ICL3-MOR construct was additionally mutated at position 281 (originally from the hMOR construct) by point mutation from Thr to Lys to generate the Y1-ICL3(T^6.34K)-MOR chimera. In the Y1-TM3-TM6-MOR chimera, nucleotides coding for amino acids 116–139 and 255–286 of the hY1 receptor were replaced by nucleotides coding for amino acids 145–168 and 274–305 of the hMOR, respectively; this Y1-TM3-TM6-MOR also contains an additional spontaneous Lys to Arg point mutation at amino acid 252. The hMOR cDNA in pcDNA3.1 (+) was kindly provided by Dr. M. Statnick (Lilly Research Laboratories, Indianapolis, IN). The purinergic hP2Y₁₂ receptor cDNA in pcDps was kindly provided by Dr. T. Schoneberg (University of Leipzig, Leipzig, Germany).

Amino acid sequences of several GPCRs at the X₁BBX₂X₃B motif were applied for a multiple sequence alignment with ClustalX1.81 (18). The Gonnet scoring matrix was selected with the penalty value for opening a gap setting at 10 and the penalty for extending a gap was set at 0.20.

Inositol Phosphate Accumulation Assay

COS-7 Cell Transfection—African green monkey kidney COS-7 cells were maintained in standard Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% heat-inactivated fetal calf serum and 0.01% penicillin-streptomycin. Cells were grown as monolayers at 37 °C in a humidified atmosphere of 5% CO₂ and 95% air. All cell culture agents were obtained from Invitrogen.

The phospholipase Cβ pathway signal, IP accumulation, was used to determine the activation of G_i/G_o-GPCRs by co-transfection of the chimeric G-protein Gα_Δ6qi4myr (kindly provided by Dr. E. Kostenis, 7TM Pharma A/S, Hoersholm, Denmark). A G_i/G_o-GPCR can couple to the G_q pathway via this chimeric protein, which has a Gα_q backbone with an altered C terminus comprised of residues from Gα_i and an N-terminal myristoylation consensus site (12, 13). COS-7 cells were seeded into 24-well plates (1.0 × 10⁵ cells/well) and transiently transfected with cDNA plasmids to be tested (0.20 μg), plus the Gα_Δ6qi4myr (0.05 μg) using 0.75 μl of metafectene (Biontex, München, Germany). As a positive control, cells were transiently co-transfected with the G_i/G_o-protein-coupled P2Y₁₂ receptor (0.20 μg) and Gα_Δ6qi4myr (0.05 μg). As a negative control, cells were transiently co-transfected with the cDNA plasmid encoding EGFP (0.20 μg) and Gα_Δ6qi4myr (0.05 μg).

Determination of IP Accumulation—IP formation was measured 2 days after transfection. Cells were incubated with 2 μCi/ml of myo-[³H]-inositol (25.0 Ci/mmol; Amersham Biosciences, Freiburg, Germany) 16 h prior to measurement. The cells were washed once and stimulated in DMEM containing 10 mm LiCl. Cells were incubated for 1, 3, or6hat37°Cinthe absence, or in differing concentrations (1 nm to 10μm) of an agonist, either pNPY (synthesized by automated solid phase peptide synthesis with the 9-fluorenylmethoxycarbonyl-tert-butyl (Fmoc/tBu) strategy (19); or 10 μm ADP (Sigma-Aldrich), as appropriate. The reactions were stopped by the aspiration of DMEM and cell lysis with 300 μl of 0.1 m NaOH. After adding 100 μl of 0.2 m formic acid, intracellular IP levels were determined by anion-exchange chromatography as described (20, 21). Ligand potency of generated chimeric GPCRs was determined (Table 1).

TABLE 1.

Agonist binding and signal transduction for the WT-Y1 receptor and Y1 chimera Data are given as the mean ± S.E. for specific binding studies (n = 3) and signal transduction assays (n = 2).

	Specific binding	Ligand potency EC50
	% relative to WT-Y1	nm
WT-Y1	100	1.97 ± 0.18
T6.30A-Y1	102 ± 5	1.49 ± 0.12
N6.34A-Y1	78 ± 5	1.69 ± 0.21
Y1-ICL3-MOR	75 ± 9	5.4 ± –0.34
Y1-TM3-TM6-MOR	20 ± 3	N/Aa
Y1-ICL3(T6.34K)-MOR	50 ± 5	N/Aa

^aCapacity for agonist binding was retained, but receptor could not be activated

Radioligand Binding Studies—Competitive radioligand binding studies were performed with N-[propionyl-³H]pNPY (GE Healthcare Europe GmbH, Braunschweig, Germany) in the absence (total binding) and presence (nonspecific binding) of 1 μm unlabeled NPY as described previously (22; Table 1).

Receptor Localization and Quantification

To estimate plasma membrane localization, a cell surface, enzyme-linked immunosorbent assay (ELISA; 23) was performed with the WT-Y1 receptor and its mutants carrying an N-terminal hemagglutinin (HA) tag. The N-terminal nine amino acid residue epitope (YPYDVPDYA) was derived from the influenza virus HA protein and inserted downstream of the start codon. The entire coding region was sequenced to ensure the accuracy of all constructs. COS-7 cells were seeded into 48-well plates (35,000 cells/well) and transfected (0.20 μg of plasmid DNA and 0.60 μl of metafectene/well). Three days after transfection, the cells were fixed with formaldehyde without disrupting the cell membrane and probed with a biotin-labeled anti-HA antibody (1 μg of 12CA5 with biotin/ml; Sigma-Aldrich). Bound anti-HA antibody was detected by a peroxidase-labeled streptavidin conjugate (0.29 μg/ml; Sigma-Aldrich). Following removal of excess unbound conjugate, H₂O₂ and o-phenylenediamine (2.5 mm each in 0.1 m phosphate-citrate buffer; pH 5) were added to serve as substrate and chromogen, respectively. After 10–30 min, the enzyme reaction was stopped by addition of 1 m HCl containing 50 mm Na₂SO₃. Color development was measured bichromatically at 492 nm and 620 nm using an ELISA reader (Sunrise, Tecan).

Immunofluorescence studies were carried out to determine the subcellular distribution of P2Y₁₂, MOR, WT-Y1, T^6.30A-Y1 (data not shown), N^6.34A-Y1 (data not shown), Y1-ICL3-MOR, Y1-TM3-TM6-MOR, and Y1-ICL3(T^6.34K)-MOR receptors (Fig. 1). COS-7 cells were transferred into 24-well plates that contained sterilized glass cover slips and then transfected. After 48 h, the cells were fixed with formaldehyde. They were treated with an anti-HA monoclonal antibody (10 μg 12CA5/ml; Sigma-Aldrich) and then incubated with a tetramethylrhodamine isothiocyanate (TRITC)-linked anti-mouse IgG secondary antibody (Sigma-Aldrich). The nuclei were visualized with Hoechst 33342 (1 μg/ml; Sigma-Aldrich). Fluorescence images were obtained with a confocal laser-scanning microscope (Leica TCS SP2; Heidelberg, Germany).

FIGURE 1.

Surface expression of all WT and chimeric GPCRs tested. A, schematic of structural mutations generated in this study and illustrations of the resulting mutant or chimeric GPCRs. B, representative confocal microscopic images of COS-7 cells that were transiently transfected with GPCRs tagged with hemagglutinin at the N terminus (TRITC image) and fused to EGFP at the C terminus (EGFP image). As cells were not permeabilized, the TRITC-tagged reagent bound only to GPCRs that were correctly trafficked and inserted into the plasma membrane. Merged images show the membrane localization of the GPCR. Cells were fixed in 4% paraformaldehyde and processed for immunofluorescence as described under “Experimental Procedures.” Images are representative of three independently performed experiments. C, representative confocal EGFP/EYFP images of transiently transfected cells expressing the GPCR fusion proteins. Images are fused with those for the membrane-permeant Hoechst 33342 nuclear stain. Cells were not permeabilized for these images. Images show membrane localization to varying degrees as fluorescent GPCRs are caught in different stages of their synthesis, transport, and insertion. There were no systematic differences in the distribution of the GPCR constructs. The Y1-TM3-TM6-MOR and Y1-ICL3(T^6.34K)-MOR chimera were slightly less well integrated in the membrane compared with the WT-Y1 receptor. This correlates well with the agonist binding data, as specific binding only reached 50% for the Y1-ICL3(T^6.34K)-MOR and 20% for the Y1-TM3-TM6-MOR chimera, relative to the WT-Y1 receptor (Table 1). Scale bar, 10 μm.

Electrophysiological Assay: Prepulse Facilitation

tsA-201 Cell Transfection—Human embryonic kidney-derived tsA-201 cells were maintained in standard DMEM supplemented with 10% heat-inactivated fetal calf serum and 1% penicillin-streptomycin. Cells were grown and maintained as monolayers at 37 °C in a humidified atmosphere of 5% CO₂ and 95% O₂. All cell culture agents were obtained from Invitrogen.

When the tsA-201 cells reached 90% confluence, they were detached with trypsin-EDTA (0.25% trypsin with EDTA 4 mm Na) and plated onto glass coverslips at 50% confluence 7 h prior to transfection. Using a standard calcium phosphate precipitation protocol (24), the plated cells were always transiently transfected with cDNA constructs encoding for the N-type voltage-dependent calcium channel (VDCC) subunits (α_1B, α₂δ, and β_1b; 3:2:2 ratio with a total of 7.0 μg/ml of DMEM). In most cases, proteins of interest were intrinsically tagged with a fluorescent protein that does not affect their functions; otherwise, a plasmid encoding EGFP (0.05 μg/ml) was additionally co-expressed. Where indicated, N-type VDCC subunits were co-expressed with the (1.0 μg/ml each) WT-Y1-EGFP, T^6.30A-Y1-EGFP, N^6.34A-Y1-EGFP, WT-MOR, T^6.34K-MOR-EYFP, Y1-ICL3-MOR-EYFP, Y1-ICL3(T^6.34K)-MOR-EYFP, or Y1-TM3-TM6-MOR-EYFP receptor construct. Where indicated, the chimeric Gα-protein (Gα_qα_o-ECFP; 0.6 μg/ml; kindly provided by Dr. N. Gautam, Washington University, St. Louis, MO) was also transiently co-expressed in the tsA-201 cells as scavengers of Gβγ. The cDNA constructs were first added to a transfection mixture containing (in mm): 640 NaCl, 25 HEPES, 120 CaCl₂, 0.75 Na₂HPO₄·H₂O before being added to the cells. Following a 16-h incubation at 37 °C in the calcium phosphate/cDNA mixture, the cells were washed with fresh DMEM and allowed to recover at 37 °C for 6 h. The cells were then moved to a 30 °C incubator to prevent further cell division.

Patch Clamp Recordings—A coverslip with the transfected cells attached was placed in a recording chamber, attached to a fixed stage upright microscope (Axioskop FS-2, Carl Zeiss) that was continuously perfused with external recording solution composed of (in mm): 20 BaCl₂, 1 MgCl₂, 10 HEPES, 40 tetraethylammonium chloride, 10 glucose, 65 CsCl (adjusted to pH 7.20 with TEA-OH; 256–260 mOsm/liter). The microscope was equipped with epifluorescence optics (excitation 450–490 nm; emission 515 nm; Chroma, Burlington, VT), and only those cells exhibiting fluorescence were considered for patching. Patch pipettes (borosilicate glass Sutter Instruments, Novato, CA) were pulled using a Sutter P-97 microelectrode puller, then in some cases were fire-polished with a Micro-Forge 200 (World Precision Instruments, Sarasota, FL) to yield tip resistances of 5–7 MΩ when back-filled with internal solution composed of (in mm): 108 Cs-methanesulfonate, 4 MgCl₂, 9 EGTA, 9 HEPES, 1 Na-GTP (adjusted to pH 7.20 with CsOH; 238–243 mOsm/liter). All electrophysiological recordings were conducted at room temperature (22 °C). Patch pipettes were connected to the headstage of either an Axopatch 200B or Axopatch 1D amplifier (Axon Instruments, Foster City, CA) used in voltage-clamp mode and linked via a Digidata 1322 interface (Axon) to a computer equipped with Clampex 8.2 software (Axon). The whole cell currents were filtered at 1 kHz (-3 dB) and the cell membrane capacitance and series resistance were manually compensated. Current traces were digitized at 10 kHz and analyzed offline. An online leak-subtraction protocol was used, where 8 pulses of opposite polarity and 1/8th amplitude were acquired immediately prior to the test protocol.

The barium current (I_Ba) elicited by each test pulse (P1 or P2) consisted of a 105 mV, 14 ms depolarizing voltage step from a holding potential (V_h) of -90 mV. In the prepulse (PP) protocol, a 15-ms depolarizing voltage step from V_h to +105 mV (a 195-mV depolarization) was applied immediately following the first test pulse (P1); the second test pulse (P2) was then preceded by a brief (4 ms) interpulse interval at the holding potential (24). In the absence of the prepulse, the two test pulses were separated from each other by 19 ms. The parameters of the voltage protocols used here: V_h; amplitude and duration of P1, P2, and PP; and duration of the interpulse interval, were optimized to maximize the effect of prepulse facilitation. Only cells showing peak inward I_Ba amplitudes between 150 pA and 2000 pA in response to P1 (I_P1) were included in the data set.

Drug Applications—NPY (Peptidec Technologies, Pierrefonds, QC, Canada) and [d-Ala²,N-MePhe⁴,Gly-ol⁵]enkephalin (DAMGO; Bachem, Torrance, CA) were diluted into the external recording solution from concentrated stock solutions to their final concentrations immediately prior to use. Stock solutions of 10 mm ADP, 100 μm NPY, and 100 μm DAMGO was prepared in HPLC grade water (Fisher Scientific) and stored at -20 °C. During drug application, 1 mm ADP, 50 nm NPY, or 1 μm DAMGO was perfused into the recording chamber for 3–5 min. Following drug application, the cells were washed out with bath perfusion solution for at least 8 min. In experiments involving guanosine 5′-[β-thio]diphosphate (GDP-β-S; Sigma-Aldrich), it was added directly to the internal pipette solution to a final concentration of 1 mm immediately prior to use.

Data Analysis

Current traces were analyzed using Clampfit 9.2 (Axon Instruments). Sample traces were reproduced using Axum 5.0 (MathSoft, Miami Beach, FL). Statistics and graphs were produced using GraphPad Prism v 4.03 (GraphPad Software, San Diego, CA). Statistical significance for mean comparisons involving only 2 groups and 3 or more groups was determined using the unpaired Student's t test and Tukey's Multiple Comparison Test, respectively, unless indicated otherwise. Statistical significance for single means was compared with 0 with a one-sample Student's t test. Statistical significance for slope comparisons between lines of best-fit was determined using a comparison of Linear Regression. The differences were considered to be significant at p < 0.05. Results were presented as means ± S.E. where appropriate.

RESULTS

WT-Y1 Receptor Is Not Constitutively Active—Basal activation of the WT-Y1 receptor was determined in the absence of agonist, with a biochemical assay measuring IP accumulation and an electrophysiological assay measuring basal prepulse facilitation (BPF). Activation of the G_i/G_o-coupled Y1 receptor typically suppresses the production of cAMP so that a direct measure of Y1 receptor activation is complex. In the IP assay, co-transfection of the chimeric Gα_Δ6qi4myr subunit (12) coupled the activation of all G_i/G_o-protein-coupled receptors tested to the normally G_q-mediated stimulation of the phospholipase Cβ pathway and subsequent accumulation of IP.

In this functional biochemical assay, basal activity of the highly constitutively active P2Y₁₂ receptor (25, 26) was readily detected, but not that of the native MOR (27). There was a significant increase in intracellular IP accumulation following the co-expression of the constitutively active P2Y₁₂ receptor but not of the demonstrably constitutively active MOR relative to the EGFP control (Fig. 2). The IP assay is therefore not sensitive to subtle levels of GPCR constitutive activity. Furthermore, this assay did not reveal significant basal receptor activation by the WT-Y1 receptor (Fig. 2).

FIGURE 2.

Expression of constitutive GPCR activation measured as basal IP accumulation. COS-7 cells transiently co-transfected with the chimeric G-protein (Gα_Δ6qi4myr) and either EGFP only, the purinergic P2Y₁₂ receptor, MOR, or WT-Y1. The IP level accumulated was determined after a 1-h incubation in the absence of an agonist and reported as the relative fold increase relative to the EGFP condition where GPCR expression is absent (EGFP: 1.00; P2Y₁₂: 13.0 ± 1.6; MOR: 2.3 ± 0.3; WT-Y1: 1.2 ± 0.1). ^**, p < 0.01.

The electrophysiological assay measuring basal prepulse facilitation (BPF) can detect constitutive activity of G_i/G_o-coupled receptors expressed in individual cells (14–16). Whole cell I_Ba was elicited using a double-pulse protocol, with or without an intervening depolarizing prepulse (PP) and results were reported as the ratio (I_P2/P1) of the peak current elicited by the second test pulse (P2) relative to that of the first (P1; Fig. 3A). BPF refers to the percent increase in I_P2/P1 elicited by the PP in the absence of an agonist (Fig. 3A).

FIGURE 3.

Expression of constitutive GPCR activation measured as BPF. A, left upper traces, voltage double-pulse protocol (black) consisted of two +105 mV (14 ms) depolarizing voltage steps (P1 and P2, respectively) separated by a 19-ms interval resting at V_h = -90 mV. Where applicable, the +195 mV prepulse (PP, gray) lasting 15 ms immediately follows P1 and precedes P2 by 4 ms. Left lower traces, overlay of sample traces of barium current (I_Ba) elicited by the double-pulse protocol in the absence (black) or presence (gray) ofthe PP. Increase in I_P2/I_P1 ratio produced by the prepulse was reported as BPF. Right upper traces, voltage conditions of the prepulse protocol were as described in the upper left panel. Right lower traces, overlay of samples traces of I_Ba produced in the absence (black) and presence (gray) of 50 nm NPY for the agonist-mediated activation of Y1 receptors. This is representative of the MOR and P2Y₁₂ receptor response to 1 μm DAMGO and 1 mm ADP as well. B, N-type calcium channel subunits (α_1B, α₂δ, β_1b) were co-expressed in all conditions, in the absence (EGFP control) or presence of the GPCR indicated. Constitutive GPCR activation was indicated by a significant increase in BPF relative to the EGFP control. Percent BPF increase refers to the relative increase in I_P2/P1 ratio in the presence of PP (EGFP: 12.4 ± 7.0%; P2Y₁₂: 52.3 ± 7.1%; MOR: 52.4 ± 8.2%; WT-Y1: 25.4 ± 5.4%). ^**, p < 0.01.

Constitutive activity of both the P2Y₁₂ receptor and MOR were revealed by the electrophysiological BPF assay. Co-expression of either the P2Y₁₂ receptor or the MOR resulted in a significant increase in BPF relative to control cells in which no GPCRs were co-expressed with the N-type VDCCs. In contrast, co-expression of the WT-Y1 receptor did not cause a significant increase in BPF over control, indicating that it is quiescent in the absence of an agonist (Fig. 3B). WT-Y1 receptor activation and intracellular signaling remained intact in this preparation, as 50 nm NPY produced a significant inhibition of I_P1 and increased prepulse facilitation (Table 2).

TABLE 2.

Inhibition of barium currents at the first test pulse (I_P1) and increase in PF following agonist-mediated receptor activation Data are given as the mean ± S.E. and determined if significantly different from zero (one sample t test).

Agonist	Receptor	n	IP1 inhibition	p value	PF increase	p value
			%		%
50 nm NPY	WT-Y1	21	73.8 ± 3.6	***a	94.7 ± 11.6	***
	T6.30A-Y1	7	84.5 ± 4.1	***	188.4 ± 65.0	*b
	N6.34A-Y1	12	82.1 ± 2.5	***	197.5 ± 56.0	***
	Y1-TM3-TM6-MOR	4	7.8 ± 9.0	nsc	N/A	N/A
	Y1-ICL3-MOR	17	60.6 ± 2.8	***	26.7 ± 9.5	*
	Y1-ICL3(T6.34K)-MOR	9	62.4 ± 1.3	***	11.6 ± 6.9	ns
1 μm DAMGO	WT-MOR	4	61.2 ± 4.2	***	34.4 ± 5.7	**d
	T6.34K-MOR	4	36.0 ± 6.2	**	–13.3 ± 9.8	ns

^a***, p < 0.0001

^b*, p < 0.05

^cns, not significant; one-sample t test

^d**, p < 0.01

Even in the absence of any co-expressed GPCRs, a significant level of BPF was observed (Fig. 4A). We tested the hypothesis that the BPF produced in the absence of any co-expressed GPCRs was due to the actions of endogenous Gβγ, expressed in excess of the Gα-subunit within the cell. A scavenger of Gβγ, the chimeric Gα subunit Gα_qα_o that is activated by a G_q-protein-coupled GPCR but transduces via the G_o pathway (28), was introduced to the cell. Co-expression of Gα_qα_o completely abolished the BPF observed in the absence of any co-expressed GPCRs, as well as that observed in the presence of the WT-Y1 or even the known constitutively active MOR (Fig. 4A).

FIGURE 4.

Modulation of BPF by GPCR-mediated turnover of G-proteins. A, co-expression of a chimeric Gα (Gα_qα_o) subunit (gray) abolished BPF in all conditions (absence of Gα_qα_o-EGFP: 25.4 ± 6.0%; WT-Y1: 25.4 ± 5.4%; MOR: 46.0 ± 8.0%; Presence of Gα_qα_o-EGFP: -4.9 ± 1.7%; WT-Y1: -2.3 ± 6.6%; MOR: -10.0 ± 3.6%). ^***, p < 0.0001. B, comparison of GPCR-mediated BPF decline following an intracellular application of 1 mm GDP-β-S (dashed line, open circle). In the absence of GDP-β-S (solid line, solid circle), a gradual decline in BPF (slope) was observed in all conditions of GPCR expression (EGFP: 2.4 ± 0.7% min^-1, thin black; MOR: 3.4 ± 1.7% min^-1, bold black; WT-Y1: 3.2 ± 1.4% min^-1, bold gray). GDP-β-S accelerated the gradual decline of BPF over time only in MOR-expressing cells (10.5 ± 2.2% min^-1, p < 0.05; EGFP: 2.8 ± 0.5% min^-1; WT-Y1: 1.6 ± 1.1% min^-1). Slopes of best-fit lines were generated using linear regression. C, comparison of BPF produced at 1 min (black; EGFP: 18.3 ± 6.8%; MOR: 49.5 ± 16.8%; P2Y₁₂: 32.6 ± 11.0%; WT-Y1: 28.2 ± 6.0%) and 6 min (gray; EGFP: 17.0 ± 6.9%; MOR: 17.6 ± 14.6%; P2Y₁₂: 5.5 ± 2.5%; WT-Y1: 18.5 ± 10.8%) following the application of intracellular GDP-β-S. ^*, p < 0.05; ^**, p < 0.001 (paired t test).

To test the possibility that the endogenous expression of excess Gβγ could impede the detection of possible constitutive WT-Y1 receptor activation, we examined the effects of GDP-β-S, an inhibitor of G-protein activation, on BPF. We hypothesized that GDP-β-S will determine whether or not the BPF produced by the WT-Y1 receptor was caused by receptor-mediated turnover of Gβγ. In all our experiments, we observed a gradual decline of BPF with time that was independent of GPCR expression (Fig. 4B). When 1 mm GDP-β-S was applied intracellularly via the pipette to inhibit basal GPCR-mediated activation of BPF, it did not affect the rate of BPF decline in cells expressing the WT-Y1 receptor, or in cells that do not co-express any GPCRs (Fig. 4B). However, GDP-β-S accelerated the rate of BPF decline in MOR-expressing cells (Fig. 4B). The BPF produced by MOR-expressing cells was significantly reduced after a 6 min GDP-β-S application (Fig. 4C). The sensitivity of BPF to inhibition by GDP-β-S was observed in cells expressing the highly constitutively active P2Y₁₂ receptor as well (Fig. 4C). This is consistent with the hypothesis that the basal activity of the MOR and P2Y₁₂ receptors resulted in an accelerated rate of GDP-β-S binding to Gα-subunits, thus reducing the Gβγ subunits available for binding to the N-type VDCCs. In contrast, intracellular GDP-β-S did not decrease BPF produced by cells expressing the WT-Y1 receptor (Fig. 4C). This suggested that the BPF observed in WT-Y1 receptor-expressing cells was not attributed to basal WT-Y1 receptor activation.

WT-Y1 Receptor Is Resistant to Constitutive Activation by Single Point Mutation at Loci 6.30 and 6.34—Constitutive GPCR activation is governed by a loss of intramolecular interactions that constrain the receptor in an inactivated state (6, 7, 29). The involvement of the cytoplasmic end of TM 6, specifically at locus 6.30 and 6.34, has been implicated in such restraining intramolecular interactions (30). A single point mutation at locus 6.30 or 6.34 was introduced to the WT-Y1 receptor to test the hypothesis that the quiescence of the WT-Y1 receptor is regulated at these specific loci. The mutant T^6.30A-Y1 and N^6.34A-Y1 receptors were normally trafficked and expressed at the plasma membrane, as revealed by the ELISA cell surface imaging assay (data not shown), and these mutant receptors retained agonist binding capacity similar to that of the WT-Y1 receptor (Table 1).

Agonist-mediated activation of both mutant Y1 receptors resulted in the robust inhibition of I_P1 and produced a significant increase in NPY-mediated prepulse facilitation (Table 2). Comparison between the BPF observed with the WT-Y1 receptor and the T^6.30A-Y1 and N^6.34A-Y1 mutant receptors showed that neither the T^6.30A-Y1 nor the N^6.34A-Y1 mutants produced significantly greater levels of BPF relative to the WT-Y1 receptor (Fig. 5A). This suggested that the T^6.30A and N^6.34A point mutations neither impaired receptor function, nor conferred constitutive activity upon the WT-Y1 receptor.

FIGURE 5.

T^6.30A-Y1 and N^6.34A-Y1 receptors do not express constitutive activity. A, neither the T^6.30A -Y1 (27.6 ± 7.2%) nor N^6.34A-Y1 (27.8 ± 6.4%) receptor demonstrated a significant increase in BPF relative to the WT-Y1 receptor (25.4 ± 5.4%). B, substitution of TM3 and TM6 from the MOR into the WT-Y1 receptor resulted in the generation of the Y1-TM3-TM6-MOR chimera, which did not produce a significant increase in BPF (EGFP: 19.2 ± 2.6%; MOR: 47.1 ± 3.6%; WT-Y1: 23.4 ± 4.8%; Y1-TM3-TM6-MOR: 41.8 ± 4.5%). ^*, p < 0.05; ^**, p < 0.01.

To examine the potential contribution of any stabilizing interactions between residues along TM3 and TM6, these two transmembrane segments of the WT-Y1 receptor were replaced with those from the constitutively active WT-MOR. Relative to the WT-Y1 receptor, the resulting Y1-TM3-TM6-MOR chimera (Fig. 1A) did not demonstrate a significant increase in BPF over the WT-Y1 receptor (Fig. 5B). Furthermore, the Y1-TM3-TM6-MOR chimera did not display agonist sensitivity (Table 2) though it was able to bind NPY (Table 1). This indicated that the Y1-TM3-TM6-MOR chimeric receptor does not express constitutive activity and suggested that exchange of the TM3 and TM6 regions from the WT-Y1 receptor interfered with agonist-mediated receptor activation and signal transduction despite retaining the capacity for agonist binding.

ICL3 Domain Governs WT-Y1 Receptor Activation—As seen above, the WT-Y1 receptor was not susceptible to constitutive activation by single point mutation at specific loci on TM6. The loci 6.30 and 6.34 on the WT-Y1 receptor lie on the interface between TM6 and the ICL3 regions of the receptor. To test the hypothesis that the ICL3 region contributes residues that stabilize the inactive state of the WT-Y1 receptor (31, 32), the ICL3 region of the WT-Y1 receptor was replaced with that from the MOR. The resulting Y1-ICL3-MOR chimera is a functional receptor that responded to 50 nm NPY agonist application. In the absence of the agonist, the Y1-ICL3-MOR chimera also demonstrated a robust BPF response. The BPF observed with expression of the Y1-ICL3-MOR chimera was significantly greater than that of the WT-Y1 receptor (Fig. 6). Furthermore, following a 5-min intracellular application of 1 mm GDP-β-S, the BPF produced in cells expressing the Y1-ICL3-MOR chimera was significantly reduced by 87.0 ± 14.9% (n = 6; p < 0.005, paired t test). This suggested that the BPF observed in cells expressing the chimeric Y1-ICL3-MOR receptor was in part attributable to basal activation of the receptor. Thus, exchange of the ICL3 region of the WT-Y1 receptor with that of the MOR resulted in the generation of a mainly Y1 receptor that is constitutively active.

FIGURE 6.

Increased basal Y1 receptor activation following mutagenesis at ICL3. Substitution of ICL3 from the Y1 receptor with that from the MOR produced a constitutively active Y1-ICL3-MOR chimeric receptor. BPF produced by the Y1 chimera (41.8 ± 4.5%) was greater than that produced by the positive EGFP control (19.2 ± 2.6%) or WT-Y1 receptor (23.6 ± 3.1%) and was similar to that produced by the MOR (41.1 ± 4.0%). ^*, p < 0.05.

We then tested the hypothesis that following the ICL3 substitution, the constitutive activity of the WT-Y1 receptor is susceptible to further modulation. In the MOR, basal activity was increased following a single Thr (T) to Lys (K) point mutation at locus 6.34 of the WT to result in the T^6.34K-MOR receptor mutant (33). We examined the constitutive activity of the Y1-ICL3-MOR chimera following an additional T^6.34K point mutation at the junction between the TM6 and ICL3 domain. If the hypothesis is correct, then a comparison of the increase in constitutive activity between the Y1-ICL3-MOR and Y1-ICL3(T^6.34K)-MOR chimerae should be similar to that between the WT-MOR and T^6.34K-MOR. Both the T^6.34K-MOR and Y1-ICL3(T^6.34K)-MOR responded to agonist application (Table 2). The Y1-ICL3(T^6.34K)-MOR chimeric mutant produced a similar inhibition of I_P1 as the Y1-ICL3-MOR, but the T^6.34K-MOR mutant was less responsive to the agonist than the WT-MOR.

The T^6.34K point mutation produced a similar increase in the BPF produced by the WT-MOR (74.9 ± 26.0%; n = 15; p < 0.01) and the Y1-ICL3-MOR (77.7 ± 24.3%; n = 17; p < 0.01; Fig. 7). Despite the robust elevation in constitutive activity observed in the BPF assay for both mutant and chimeric receptor containing the T⁶³⁴K point mutation, the IP assay revealed evidence of constitutive activity for the T^6.34K-MOR (IP accumulation: 3.27 ± 0.29-fold over WT-MOR; n = 30; p < 0.05) only. Our results showed that following the exchange of the native Y1 ICL3 region with that from the WT-MOR, the Y1 receptor demonstrated a similar susceptibility to structural changes that also increased basal receptor activation in the WT-MOR.

FIGURE 7.

T^6.34K point mutation increased the basal activity of constitutively active MOR and Y1 receptor chimera. The T^6.34K point mutation within the ICL3 region (gray, T^6.34K-MOR: 95.0 ± 14.2%; Y1-ICL3(T^6.34K)-MOR: 78.8 ± 9.2%) further increased the BPF produced by both the WT-MOR (52.4 ± 8.2%) and Y1-ICL3-MOR receptor (41.8 ± 4.5%). ^*, p < 0.05.

DISCUSSION

Detection of constitutive activity in G_i/G_o-protein-coupled GPCRs is more complex than with GPCRs coupling to G-proteins such as G_s or G_q that naturally stimulate the production of readily examined second messengers. In the biochemical assay here, the coexpression of a chimeric Gα_Δ6qi4myr-protein converted the inhibitory G_i/G_o-signaling pathway into a stimulatory G_q signal that was identified by the accumulation of IP (12, 25, 34, 35). The IP assay was insensitive to low levels of GPCR activation but responded with a wider dynamic range; thus may be particularly useful to study GPCRs that express strong constitutive activity. While a potentially sensitive biochemical method of measuring activity in potassium channels activated by G_i/G_o-coupled GPCRs has been described recently (36), its suitability and sensitivity to constitutive activity is unknown. In contrast, the electrophysiological detection of BPF can track even subtle changes in constitutive activity over a short period of time, including the gradual loss of GPCR-mediated Gβγ turnover in the presence of GDP-β-S in an individual cell. The sensitivity of the BPF assay can be largely attributed to the presence of an internal control, because the expression of BPF produced by the activated and inactivated GPCR state was evaluated during one continuous current recording.

The WT-Y1 receptor differs from many class I GPCRs in that it expresses neither any detectable constitutive activity nor sensitivity to simple structural changes that enhance constitutive activity in other similar receptors. By substituting a complete TM or ICL3 region from the MOR into the Y1 receptor, we identified which functional region is most important for the activation of the Y1 receptor. Though the activation of most class I GPCRs involves the TM3 and TM6 regions (37), they do not appear to be critical for stabilizing the inactive state of the Y1 receptor. Replacement of the ICL3 but not the TM3 and TM6 region of the WT-Y1 receptor generated a constitutively active, chimeric Y1 receptor. This suggests that the ICL3 region of the Y1 receptor contributes significant intramolecular interactions that stabilize the Y1 receptor in the inactive state. Interestingly, following the loss of constraining interactions from the WT-ICL3, the residues at the cytoplasmic end of TM6 appear to become more important for stabilizing the inactive conformation of the Y1 receptor.

Contributions of TM6 to WT-Y1 Receptor Activation—Others have indicated that the Y receptor family displays little or no basal activation (38). The class I GPCRs, which include the Y receptors, is stabilized in the inactive state by an ionic lock (5, 8, 39, 40). The ionic lock comprises a network of hydrogen bonds and charge interactions between the conserved (E/D)RY motif on TM3 and specific residues at locus 6.30 and 6.34 along TM6 (5). Constitutively active mutants can be generated in several receptors by point mutation(s) at locus 6.30 and/or 6.34. However, the applicability of this mutagenesis depends on the presence of a conserved D/E^6.30 residue. Although D/E^6.30 is nearly 100% conserved among rhodopsin and monoamine GPCRs (10), it is not universally conserved among all class I GPCRs (i.e. MOR: 33, 41; gonadotrophin releasing hormone receptor (GnRHR): 42; angiotensin I receptor (AT₁R): 43, 44), and it is not present in the Y1 receptor (45, 46).

Two different phenotypes of GPCR activation can be identified based on the presence or absence of D/E^6.30 in the native GPCR. When D/E^6.30 is present in the wild-type GPCR, a constitutively active mutant receptor could be generated by point mutation at either position 6.30 or 6.34 (α_1B-AR: 6, 9; β₂-AR: 10, 11; 5-HT_2A receptor 47, 48; luteinizing-stimulating hormone receptor (LSHR): 49, 50; thyrotropin-stimulating hormone receptor (TSHR): 51–53). This corresponded with the rhodopsin model, which suggested that both residues at loci 6.30 and 6.34 are necessary to regulate GPCR activation (10, 47). The second phenotype arose in the absence of D/E^6.30 in the wild-type receptor. Constitutive activity was observed only following site-directed mutagenesis at locus 6.34 (33) but not 6.30 (41). This suggested that only one constraining intramolecular interaction existed at TM6 for GPCRs lacking a D/E^6.30 residue (41).

Our studies on the Y1 receptor provide evidence of a third phenotype for GPCR activation where neither point mutation at locus 6.30 nor 6.34 can elicit constitutive, agonist-independent GPCR activation in the absence of a native D/E^6.30. A similar insensitivity to constitutive activity was also observed in GnRHR (42) and AT₁R (43, 44). In contrast to the distinctive role of TM3 and TM6 for receptor stabilization in many Class I GPCRs, our studies revealed that neither of these TM regions contains residues that play a significant role in constitutive activation of the WT-Y1 receptor.

Role of ICL3 in Stabilizing the WT-Y1 Receptor—As with the α₁-adrenergic receptor (α₁-AR; 31, 32), the ICL3 of the Y1 receptor confers properties important for activation and signal transduction. GPCR activation becomes a physiologically relevant event when it affects an intracellular signal. Several residues within the ICL3 (54–58) and at the TM6/ICL3 junction (59, 60) are essential for the formation of a G-protein-binding pocket (61–63). Because the ICL3 mediates G-protein recognition and activation (61), structural changes involving the ICL3 can mimic conformational changes of agonist-mediated GPCR activation even in the agonist-unbound state. It is interesting to note that, despite the change in basal activity conferred upon the Y1 receptor with a chimeric ICL3, the ability to activate G_i/G_o-proteins appeared unimpaired. This implies that the functional aspects of the region are at least partly independent of the effect of basal activity. Although identification of those specific ICL3 residues that have the most significant impact on Y1 receptor activation can be addressed by single point mutations along the ICL3, it is beyond the scope of the present study.

The ICL3 substitution also caused structural changes to nearby regions of the Y1 receptor. Point mutation at locus 6.34 of the quiescent WT-Y1 receptor had no effect on its basal activity. By contrast, the basal activity of the Y1-ICL3-MOR chimera was substantially increased by the T^6.34K point mutation at this previously disengaged locus. This indicates that the structural change in the Y1-ICL3-MOR chimera unmasked the previously unseen participation of residues along TM6 in stabilizing the inactive conformation of the Y1 receptor.

The NPY system appears to be relatively quiescent during basal physiological conditions in vivo. Thus, there is little or no effect of germline knock-out of NPY or its receptors on basal physiological responses (64). Similarly, there are only modest effects in vivo following the administration of NPY antagonists (e.g. 65, 66), except under circumstances where the NPY system is strongly activated (e.g. 67). Activation of G_i/G_o-coupled GPCRs tends to inhibit neuronal activity via a number of mechanisms (68), and signaling via G_i/G_o-coupled GPCRs appears to interact with that of G_s- or G_q/11-coupled receptors (69–71). Given this, there may be significant adaptive advantages to agonist/receptor systems where there is no signaling in the absence of any agonist.

CONCLUSION

Mutant GPCRs that demonstrate the activation of their G-proteins even in the absence of agonist-binding have advanced the understanding of the molecular mechanisms involved in GPCR activation. Constitutively active mutant GPCRs are useful tools for identifying specific regions or residues that comprise the structural machinery conferring GPCR activation. Our experiments identified the relative quiescence of the WT-Y1 receptor, which is resistant to constitutive activation by disturbing specific intramolecular interactions along TM6. Instead, we observed that the ICL3 region plays an important role for its constitutive activation, which is highly unusual relative to most class I GPCRs.