Article31 July 2008free access Dopamine D2 receptors form higher order oligomers at physiological expression levels Wen Guo Wen Guo Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Eneko Urizar Eneko Urizar Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Michaela Kralikova Michaela Kralikova Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Juan Carlos Mobarec Juan Carlos Mobarec Department of Structural and Chemical Biology, Mount Sinai School of Medicine, New York, NY, USA Search for more papers by this author Lei Shi Lei Shi Department of Physiology and Biophysics and the Institute for Computational Biomedicine, Weill Medical College of Cornell University, New York, NY, USA Search for more papers by this author Marta Filizola Marta Filizola Department of Structural and Chemical Biology, Mount Sinai School of Medicine, New York, NY, USA Search for more papers by this author Jonathan A Javitch Corresponding Author Jonathan A Javitch Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Pharmacology, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Wen Guo Wen Guo Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Eneko Urizar Eneko Urizar Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Michaela Kralikova Michaela Kralikova Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Juan Carlos Mobarec Juan Carlos Mobarec Department of Structural and Chemical Biology, Mount Sinai School of Medicine, New York, NY, USA Search for more papers by this author Lei Shi Lei Shi Department of Physiology and Biophysics and the Institute for Computational Biomedicine, Weill Medical College of Cornell University, New York, NY, USA Search for more papers by this author Marta Filizola Marta Filizola Department of Structural and Chemical Biology, Mount Sinai School of Medicine, New York, NY, USA Search for more papers by this author Jonathan A Javitch Corresponding Author Jonathan A Javitch Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Pharmacology, College of Physicians and Surgeons, Columbia University, New York, NY, USA Search for more papers by this author Author Information Wen Guo1,2,‡, Eneko Urizar1,2,‡, Michaela Kralikova1,2, Juan Carlos Mobarec4, Lei Shi5, Marta Filizola4 and Jonathan A Javitch 1,2,3 1Center for Molecular Recognition, College of Physicians and Surgeons, Columbia University, New York, NY, USA 2Department of Psychiatry, College of Physicians and Surgeons, Columbia University, New York, NY, USA 3Department of Pharmacology, College of Physicians and Surgeons, Columbia University, New York, NY, USA 4Department of Structural and Chemical Biology, Mount Sinai School of Medicine, New York, NY, USA 5Department of Physiology and Biophysics and the Institute for Computational Biomedicine, Weill Medical College of Cornell University, New York, NY, USA ‡These authors contributed equally to this work *Corresponding author. Center for Molecular Recognition and Departments of Psychiatry and Pharmacology, College of Physicians and Surgeons, Columbia University, 630W 168th street, P&S, Room 11-401, NY 10032, USA. Tel.: +1 21 230 573 08; Fax: +1 21 230 555 94; E-mail: [email protected] The EMBO Journal (2008)27:2293-2304https://doi.org/10.1038/emboj.2008.153 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info G-protein-coupled receptors are generally thought to be organized as dimers; whether they form higher order oligomers is a topic of much controversy. We combined bioluminescence/fluorescence complementation and energy transfer to demonstrate that at least four dopamine D2 receptors are located in close molecular proximity in living mammalian cells, consistent with their organization as higher order oligomers at the plasma membrane. This implies the existence of multiple receptor interfaces. In addition to the symmetrical interface in the fourth transmembrane segment (TM4) we identified previously by cysteine (Cys) crosslinking, we now show that a patch of residues at the extracellular end of TM1 forms a second symmetrical interface. Crosslinking of D2 receptor with Cys substituted simultaneously into both TM1 and TM4 led to higher order species, consistent with our novel biophysical results. Remarkably, the rate and extent of crosslinking at both interfaces were unaltered over a 100-fold range of receptor expression. Thus, at physiological levels of expression, the receptor is organized in the plasma membrane into a higher order oligomeric structure. Introduction G-protein-coupled receptors (GPCRs) comprise a diverse, well-studied system for transducing signals from the extracellular milieu to a variety of intracellular signalling molecules (Bartfai et al, 2004). GPCRs have been inferred to be dimers in the plasma membrane (Pin et al, 2007; but see Chabre et al, 2003; Chabre and le Maire, 2005; James et al, 2006; Meyer et al, 2006). Class C receptors form homo- and heterodimers but have been inferred not to be organized as higher order oligomers (Brock et al, 2007). In contrast, several lines of evidence have suggested that class A receptors might exist as higher order oligomers, although this remains controversial (Chabre et al, 2003). Despite an explosion of recent interest, our understanding of the structural details and functional role of GPCR oligomerization is still limited. Most importantly, it has not been established whether activation of class A rhodopsin-like GPCRs is affected by their organization in a particular quaternary structure. Recently, both rhodopsin (Bayburt et al, 2007) and β2-adrenergic receptor (B2AR) (Whorton et al, 2007) have been shown to signal efficiently to G proteins when reconstituted into lipid nanodiscs containing only a single receptor. Thus, after solubilization and reconstitution, these GPCRs can function alone. However, whether in fact they do function alone in vivo cannot be addressed by such studies and requires an exploration of their native organization. Much evidence indicates that class C receptors exist and function as homo- or heterodimers (reviewed in Pin et al, 2003). Findings in class A glycoprotein hormone receptors support the existence of trans- as well as cis-activation (Ji et al, 2002), which requires them to be organized at least as dimers. In addition, ligand binding to one protomer in a GPCR dimer appears to be sufficient to cause an activation-like conformational change in the unoccupied protomer (Damian et al, 2006; Brock et al, 2007), consistent with transfer of information between the protomers and thus with a functional role of dimerization. An understanding of the structural basis of crosstalk between receptors must include their interfaces, but information about these interactions is still limited. In various class A receptors, cysteine (Cys) crosslinking studies have supported a contribution of transmembrane 4 (TM4) (Guo et al, 2003, 2005; Klco et al, 2003; Kota et al, 2006) or TM1 and TM2 (Klco et al, 2003) to a symmetrical interface. FRET studies performed in the alpha-factor yeast GPCR supported a contribution of TM1 to the dimer interface (Overton and Blumer, 2002). Work with peptides and receptor fragments has supported a contribution of TM1, TM4 and/or TM6 to interaction surfaces (Hebert et al, 1996; Ng et al, 1996; Baneres and Parello, 2003; Carrillo et al, 2004). Recently, a role for TM4 as part of a dimerization interface has been extended to the class B secretin-like class of GPCRs (Harikumar et al, 2007). Given the placement of TM1 and TM4 in the high-resolution structures of rhodopsin (Palczewski et al, 2000) and the B2AR (Cherezov et al, 2007), it is not possible for these segments to contribute to the same dimer interface. Although not without substantial controversy (Chabre et al, 2003), higher order packing of native rhodopsin into rows of well-organized protomers has been visualized by atomic force microscopy (AFM) (Fotiadis et al, 2003), and biochemical and biophysical findings in other GPCRs also suggest the possibility of higher order organization (Park and Wells, 2004; Lopez-Gimenez et al, 2007; Philip et al, 2007). A higher order organization of GPCRs could simultaneously provide for symmetrical TM1 and TM4 interfaces (Liang et al, 2003). Here, we use protein complementation assays combined with resonance energy transfer to demonstrate that in living mammalian cells, the dopamine D2 receptor (D2R) is organized in a unit comprised of at least four protomers. In addition to the symmetrical interface in TM4 we established previously by Cys crosslinking, in the present work, we identified a patch of residues at a second symmetrical interface at the extracellular end of TM1. As predicted by our biophysical data, crosslinking of receptors with Cys substituted simultaneously into both TM1 and TM4 led to higher order species, consistent with a higher order organization of D2R in the plasma membrane of intact cells. The rate and extent of crosslinking were unaltered over a 100-fold range of receptor expression, suggesting that the receptor is organized into a higher order structure at physiological levels of expression. Results Higher order oligomerization: resonance energy transfer evidence Given the controversy regarding the possibility that class A GPCRs are organized as higher order oligomers, we developed a biophysical approach using a combination of luminescence and fluorescence complementation and energy transfer approaches to explore the higher order structure of the D2R in living cells. First, we used bimolecular fluorescence (BiFC) (Hu et al, 2002) and luminescence (BiLC) (Paulmurugan et al, 2004) complementation assays to study the D2R dimer. Split fluorescent (or luminescent) proteins are not fluorescent (or luminescent) when expressed alone, but when fused to proteins that are located in close proximity, they can be assembled early in biosynthesis and thereby complement fluorescence (or luminescence) and report on molecular proximity (Kerppola, 2006). Coexpression of C-terminally tagged D2R with either split monomeric Venus (mVenus) (Zacharias et al, 2002) (D2–V1 and D2–V2) or split Renilla luciferase 8 (RLuc8) (Loening et al, 2006) (D2–L1 and D2–L2) resulted in efficient complementation of fluorescence or luminescence, respectively (Figure 1A and C). Evidence for specificity of the interactions came from experiments using the thyrotropin receptor (TSHr) fused at its C terminus to split Venus or RLuc8. TSHr readily complemented with TSHr (data not shown), but much lower levels of heterocomplementation were observed between D2R and TSHr with either combination of constructs (Figure 1B and D). Figure 1.D2R ‘dimerization’ as assessed by protein complementation. HEK 293T cells transiently coexpressing D2R split Venus (A) or RLuc8 (C) or D2R and the corresponding TSHr splits (B, D) were harvested 48 h post-transfection, washed with PBS, centrifuged and resuspended in PBS. Fluorescence was recorded for 1 s using 500 nm excitation and 540 nm emission filters (Polarstar, BMG). Unfiltered luminescence was recorded for 1 s (Gain 3900). Background was determined with cells expressing only one of the receptor probes and the signal to background ratio was plotted against the FACS ratio (A–D). For the FACS ratio, cells transfected in parallel were labelled with primary and secondary antibodies (Abs) as described previously (Costagliola et al, 1998) and in the Methods. Relative staining for each receptor was determined independently in the same cells with the same secondary anti-mouse Ab to determine the FACS ratio. Representative results from at least three independent experiments are shown. Inset in (A) illustrates HTRF experiments performed in cells expressing identical amounts of D2–V1 and D2–V2 as compared with SF–D2 and SM–D2, respectively. Download figure Download PowerPoint Homogeneous time-resolved fluorescence (HTRF) experiments using probes directed against distinct N-terminal epitope tags in the split D2R constructs confirmed their interaction at the cell surface. Complementation of the split probes did not affect energy transfer (Figure 1A inset), indicating that complementation does not enhance D2R–D2R interactions. Thus, the proximity of receptors in a dimeric or oligomeric complex early in biosynthesis (Kerppola, 2006) is maintained on the plasma membrane, with or without complementation. To confirm that complementation is without impact on plasma membrane expression, we performed confocal microscopy analysis of various combinations of the transfected receptor constructs. As expected, Venus complemented by D2–V1 and D2–V2 is almost exclusively on the plasma membrane (Supplementary Figure 1c). In striking contrast, the lower levels of complementation of TSHr and D2R originate exclusively from receptor complexes that are retained intracellularly (Supplementary Figure 1c). Similar intracellular retention was observed upon coexpression of D2R–V1 and the single membrane-spanning protein CD8 fused to V2, which was tested as an additional control (Supplementary Figure 1c). We also carried out cell surface fluorescence-activated cell sorting (FACS) analysis, which indicated that any combination of TSHr and D2R that can complement (V1–V2 or L1–L2 in both orientations) leads to complete intracellular retention of TSHr (Supplementary Figure 1d inset). Thus, the low nonspecific complementation appears to result from receptors that become inappropriately attached by cotranslational folding of Venus or RLuc8, but the quality control mechanism of the cell does not allow these species to reach the cell surface. In parallel, we also used bioluminescence resonance energy transfer (BRET) (Xu et al, 1999) to study receptor interactions. We performed BRET titration experiments with cells coexpressing constant amounts of D2–RLuc8 and increasing concentrations of D2–Venus (full-length versions of the previously split proteins). A specific and saturable BRET signal was observed for D2R–D2R (Figure 2A), as well as for the TSHr–TSHr (data not shown), which is known to homodimerize (Urizar et al, 2005). The level of BRET between D2R–RLuc8 and TSHr–Venus was low, but the expression levels of the TSHr–Venus were not sufficient to titrate fully the D2–RLuc8. To insure that the direction and the orientation of the BRET between the receptors did not affect the results and to allow full titration between TSHr and D2R, we swapped the probes and still observed very low and right-shifted BRET, which in this case was titrated fully (Figure 2A). In contrast to the hetero-complementation studies described above, TSHr–RLuc8 was seen on the plasma membrane in the presence of D2–Venus (data not shown). Figure 2.Biophysical evidence of higher order oligomerization. (A) Two-protomer BRET: increasing amounts of D2–Venus were coexpressed with constant amounts of either D2–RLuc8 or TSHr–RLuc8 in HEK 293T cells. At 48 h post-transfection, BRET was performed and the BRET signals were plotted against the relative expression levels of each tagged receptor. Results were analysed by nonlinear regression assuming a model with one site binding (GraphPad Prism 4.0) on a pooled data set from four independent experiments. (B) BRET was performed as described above in cells coexpressing constant amounts of D2–RLuc8 and increasing amounts of D2–Venus in the absence (▪ and solid line), or presence of three different class A GPCRs (D2R: □ and dashed line; TSHr: • and dotted line; CXCR4: × and dotted line). Cell surface expression of each untagged receptor used as a competitor was monitored by FACS (not shown). As above, pooled BRET signals from three independent experiments were plotted against relative expression ratios assuming a one-binding site model. (C) BRET assay was performed in cells coexpressing constant D2–RLuc8 and increasing D2–Venus with increasing amounts of untagged D2R as described above. The increasing level of surface expression of the competing D2R and surface expression of TSH and CXCR4 was confirmed by FACS (data not shown). A representative experiment performed three times is shown. (D and inset) Three-protomer BRET: cells coexpressing constant amounts of D2–L1 and D2–L2 as a BRET donor (complemented RLuc8) and increasing amounts of either D2–Venus or TSHr–Venus as the acceptors were treated and analysed identically as described in (A). Data from a pooled data set from four independent experiments are shown. (E) Three-protomer BRET: cells coexpressing increasing amounts of D2–V1 and D2–V2 as the BRET acceptor (complemented mVenus) and either D2–RLuc8 or TSHr–RLuc8 as donors were treated and analysed identically as described in (A). Data from a pooled data set from four independent experiments are shown. (F and inset) Four-protomer BRET: Cells coexpressing increasing amounts of D2–V1 and D2–V2 as the BRET acceptor (complemented mVenus) and constant amounts of either D2–L1 and D2–L2 or TSHr–L1 and TSHr–L2 as donors (complemented RLuc8) were treated and analysed identically as described in (A). Data from a pooled data set from four independent experiments are shown. Cartoons represent different receptor species: D2 depicted as a 7TM alone and TSHr as the 7TM with the large extracellular domain. mVenus and the splits are represented as elliptical and RLuc8 and the splits as rectangular. In the legends, + separates donor (first) from the acceptor (second) and _ indicates the complemented pairs. Download figure Download PowerPoint To confirm the specificity of these interactions, to rule out a role of the biosensor orientation and to avoid biased comparisons between different BRET couples, we performed BRET experiments in the presence of different concentrations of untagged receptor, which would be expected to inhibit the BRET signal by competing for dimerization with the receptors fused to the probes. To ensure that the signal was consistent over the entire range of coexpression, we performed these experiments over a wide range of donor concentrations. Coexpression of untagged D2R decreased the BRET signal, whereas coexpression of TSHr or CXCR4 was without effect (Figure 2B). Competition over the entire titration range rules out changes in the absolute or relative levels of expression of the BRET probes as the cause of the decreased BRET signal (Figure 2B and C). The decrease in maximal BRET, without an associated increase in BRET50, suggests caution in the use of BRET50 analysis in studying receptor interactions that might be stable. Thus, BiFC, BiLC, HTRF and BRET assays all were consistent with the robust formation of D2R ‘dimers’, although, one can only infer from these parallel approaches that the receptor forms at least a dimer. Once we established the basic ‘dimeric’ unit, we combined the different constructs to probe the existence of higher order oligomeric complexes in living cells. Complementing RLuc8 with coexpression of D2–L1 and D2–L2 resulted in a very efficient donor for the full-length D2–Venus but not for the TSHr–Venus (Figure 2D). As for the full-length BRET (see above), only the complemented RLuc8 generated by coexpressing D2–L1 and D2–L2 was a donor for D2–Venus and none of the other combinations assayed substituting L1, L2 or both with TSHr–splits gave significant BRET signals (Supplementary Figure 2a and b). Similar results were obtained expressing constant amounts of D2–RLuc8 full-length construct as donor with the split acceptor constructs, mVenus (D2–V1 and D2–V2) (Figure 2E; Supplementary Figure 2c). All the tested probes were expressed and functional (Supplementary Figure 1a and b) and transfections were adapted so that the relative expression levels would be comparable. Although the TSHr constructs could not be combined with the D2R constructs to produce ‘three-protomer’ BRET, under similar conditions, BRET was readily observed when all the probes were attached to TSHr (Supplementary Figure 2b). In summary, when the three probes were all from D2R or all from TSHr, regardless of which reporter was split, BRET was most efficient, whereas the titration curves were right shifted dramatically when one of the protomers was substituted by the other receptor. This set of experiments confirms that D2R can form higher order complexes containing at least three receptors. To establish that the minimum oligomeric unit is formed by at least four receptors, we carried out experiments in which both the donor and the acceptor were split and complemented. We coexpressed constant amounts of split RLuc8 (D2–L1 and D2–L2) with increasing amounts of split mVenus (D2–V1 and D2–V2), and we detected specific and saturable energy transfer between the complemented probes used as donor and acceptor, respectively (Figure 2F). Again, when any of the four probes was substituted by a homologous TSHr probe, the BRET curves were dramatically right shifted. It is important to note that complementation of either luminescence or fluorescence requires that the RLuc8 or Venus be assembled early in biosynthesis. Therefore, given the enormous number of ‘non-productive’ receptor combinations that would fail to show fluorescence or luminescence, or would show luminescence and/or fluorescence but not BRET (Supplementary Figure 5), our observation of a substantial ‘four-protomer BRET’ signal is remarkable. Our demonstration of resonance energy transfer between the complemented RLuc8 and the complemented Venus shows that at least four receptors are located in close molecular proximity in living cells. A patch of residues at the extracellular end of TM1 forms a second symmetric dimerization interface We had previously identified a site of symmetrical interaction in TM4 of D2R (Guo et al, 2003, 2005), but the energy transfer data described above imply the existence of another interface. A molecular model proposed based on the AFM data of native murine rhodopsin (Fotiadis et al, 2003; Liang et al, 2003) placed a second symmetrical interface in TM1, and we therefore first focused on this region. In our background D2R construct (see Methods), the endogenous Cys1684.58 at the TM4 interface is mutated to Ser to eliminate crosslinking (Guo et al, 2003). In this background, we mutated each residue in TM1 to Cys, one at a time, from the extracellular end (P321.30) to the beginning of the putative first intracellular loop (R611.59) and stably expressed these mutants in HEK 293 cells. On the basis of immunoblotting, 28 of the mutant receptors were well expressed and maturely glycosylated, whereas mutation to Cys of the highly conserved residue N521.50 led to loss of immunoreactivity (Figure 3A and B). Of the 28 Cys mutants that were expressed, treatment with 1 mM copper phenanthroline (CuP) led to oxidative crosslinking of four mutants (Y361.34C, Y371.35C, L401.38C and L431.41C) to a species with a mobility on SDS–PAGE consistent with that of a D2R dimer (Figure 3A and B), as we observed previously with selected Cys in TM4 (Guo et al, 2005). To estimate the susceptibilities to crosslinking, we treated the mutants with a range of CuP concentrations. As shown in Figure 3, the crosslinking was saturable at all four positions, with different susceptibilities to crosslinking and different maximal crosslinking (Figure 3C–J). The mutant Y371.35C was the most susceptible to crosslinking (Figure 3E and F). Figure 3.TM1 forms a symmetrical ‘dimerization’ interface in D2R. (A, B) Intact cells stably expressing substituted Cys mutants in TM1 from P321.30C to R611.59C (denoted using the indexing system described in Methods) (except for non-expressed N521.50C and the endogenous C561.54, which is present in the background construct and all the mutants) were treated with 1 mM CuP in a 1:2 molar ratio at 25°C for 10 min, washed with PBS buffer, treated with NEM and analysed by immunoblotting. (C–J) Crosslinking was performed as described above with increasing concentrations of CuP for the four Cys mutants of interest Y361.34C (C, D), Y371.35C (E, F), L401.38C (G, H) and L431.41C (I, J). The dimer fraction is plotted against CuP concentration to determine the apparent crosslinking rates and efficiency. (K, L). Intact cells were treated with 1 mM CuP for Y361.34C, L401.38C and L431.41C and 5 μM for Y371.35C in the absence or presence of 10 μM sulpiride or 10 μM quinpirole. Quantification of crosslinking fractions was performed as described in Methods. All experiments were repeated ⩾3 times, and one representative experiment is shown. Download figure Download PowerPoint To explore the potential effect of ligand binding on the TM1 interface, we treated each of the four mutants with a non-saturating concentration of CuP in the presence and absence of either agonist or antagonist. In contrast to our findings in TM4 (Guo et al, 2005), we did not see a significant effect of these ligands on crosslinking of any of the TM1 Cys mutants (Figure 3K and L). These data suggest that a patch of residues near the extracellular end of TM1 contributes to a symmetrical interface that does not appear to undergo major conformational changes upon ligand binding. Consistent with our predictions (Shi et al, 2001), the recent high-resolution structure of the B2AR (Cherezov et al, 2007), which has much greater sequence similarity with the D2R, showed that the greatest divergence between rhodopsin and the B2AR structure is the presence of a much straighter TM1 in B2AR. We built a homology model of the D2R based on the B2AR structure (see Materials and methods), and packed the TM1 interface to satisfy our crosslinking data (Figure 4). The same packing is not possible in the rhodopsin model, as the TM1 helices would clash in such a configuration (Figure 4D and E). Figure 4.Molecular model of the TM1 interface and of a D2R oligomeric arrangement. Ribbon representations of (A) vertical, (B) extracellular and (C) cytoplasmic views of the proposed TM1–TM1 interacting regions of the D2R homodimer. The TM1 and H8 helices at the interface are highlighted by thicker traces. The Cβ atoms of residues that crosslink when mutated to Cys are shown in CPK representations in different colours. (D) Vertical and (E) extracellular views of the best fit between rhodopsin-based models of the D2R protomers (only TM1 helices are shown; orange and yellow colours) and the proposed B2AR-based model of the TM1–TM1 homodimer of D2R (cyan and grey colours). (F) Schematic representation of a possible D2R oligomeric organization based on inferences from our crosslinking studies (see Methods). Eight protomers are shown. The four protomers with red in the interior can be crosslinked in the 1.35C/4.58C construct. The four protomers contained in the yellow contour can be crosslinked in the 1.35C/5.41C construct (see Discussion). Download figure Download PowerPoint The configuration that was most compatible with our observed TM1 crosslinking places helix 8 (H8) in direct interaction with H8 from the adjacent protomer (see Discussion). In an attempt to validate this prediction, we created a series of Cys mutations (F4377.64C, L4387.65C and K4397.66C) in H8 and stably expressed these in HEK 293 cells. In intact cells, no crosslinking of the H8 Cys mutants was observed with CuP treatment (data not shown). In contrast, treatment with the membrane permeant crosslinker HgCl2 led to crosslinking of L4387.65C, consistent with the prediction of our model of the TM1 interface (Figures 4A–C and 5C). Thus, based on the biochemical and computational data, we infer that H8 is part of the symmetrical TM1 interface. Figure 5.TM1 and TM4 contribute to symmetrical interfaces in higher order D2R complexes. (A) Intact cells stably expressing a D2R TM1–TM4 double Cys mutant with both TM1 Y371.35C and the endogenous TM4 C1684.58 were treated with increasing concentrations of CuP as described in Figure 3. A representative blot is shown with the different species as determined by the mass molecular weight standards. (B) Fractions of the different species at the indicated CuP concentration from three independent experiments (mean±s.d.). (C) Intact cells stably expressing F4377.64C, L4387.65C or K4397.66C in H8 an
