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G-protein-gated K+ channels (Kir3.1–Kir3.4) control electrical excitability in many different cells. Among their functions relevant to human physiology and disease, they regulate the heart rate and govern a wide range of neuronal activities. Here we present the first crystal structures of a G-protein-gated K+ channel. By comparing the wild-type structure to that of a constitutively active mutant, we identify a global conformational change through which G-proteins could open a G-loop gate in the cytoplasmic domain. The structures of both channels in the absence and presence of PIP2 show that G-proteins open only the G-loop gate in the absence of PIP2, but in the presence of PIP2 the G-loop gate and a second inner helix gate become coupled, so that both gates open. We also identify a strategically located Na+ ion-binding site, which would allow intracellular Na+ to modulate GIRK channel activity. These data provide a mechanistic description of multi-ligand regulation of GIRK channel gating.
G-protein-gated K+ (GIRK) channels are members of the inward rectifier (Kir) channel family, so named because the outward flow of K+ ions is inhibited by intracellular polyamines and Mg2+, which block the pore in a voltage-dependent manner. Kir channels play an essential role in many physiological processes including neuronal signaling, kidney function, insulin secretion and heart rate control. Mutations of Kir channels underlie numerous diseases including primary aldosteronism, Andersen syndrome, Bartter syndrome, and congenital hyperinsulinism (Choi et al., 2011; Hibino et al., 2010).
All Kir channels share the same basic topology: four subunits combine to form a canonical K+ pore-forming transmembrane domain (TMD) and a large cytoplasmic domain (CTD) (Fig. 1A). It is thought that ion conduction may be regulated by two gates in series: one is formed by the inner helices of the TMD (Doyle et al., 1998; Jiang et al., 2002), and the other by the G loop at the apex of the CTD (Nishida et al., 2007; Pegan et al., 2005). Various regulatory molecules are thought to control these gates, but the control mechanisms are still unknown.
The anionic lipid phosphatidylinositol 4,5-bisphosphate (PIP2) is essential for the activation of all Kir channels (Hibino et al., 2010). GIRK channels (Kir3.x) are unique in that they also require G proteins, in combination with PIP2, for activation (Huang et al., 1998; Logothetis et al., 1987; Reuveny et al., 1994; Sui et al., 1998; Wickman et al., 1994). Certain GIRK channel subtypes are also modulated by intracellular Na+ ions (Ho and Murrell-Lagnado, 1999a, b; Lesage et al., 1995; Sui et al., 1996; Sui et al., 1998). GIRK channel activation elicits the flow of K+ ions across the cell membrane and thus drives the membrane voltage towards the Nernst potential for K+. Near the K+ Nernst potential, voltage-dependent Na+ and Ca2+ channels tend to be silenced and therefore electrical excitation is diminished. This is an important signaling mechanism by which hormone and neurotransmitter stimulation of G protein coupled receptors (GPCRs) regulates many essential physiological processes (Luscher and Slesinger, 2010). For example, acetylcholine secreted by the vagus nerve controls heart rate through stimulation of muscarinic GPCRs in cardiac pacemaker cells (Logothetis et al., 1987; Pfaffinger et al., 1985).
Electrophysiological studies have sought to understand how the various ligands – G proteins, PIP2, and Na+ – interact simultaneously with GIRK channels to regulate their gating. These studies suggest that G proteins and Na+ function in a co-dependent manner with PIP2 to open GIRK channels (Huang et al., 1998; Sui et al., 1998; Zhang et al., 1999). Here we present crystal structures of a quiescent, closed GIRK2 channel and of a point mutant that is constitutively active, independent of G protein stimulation. Further, both structures are determined in the absence and presence of PIP2. These structures render a molecular mechanistic description of multi-ligand regulation of GIRK channels.
Four GIRK channel isoforms (Kir3.1 – Kir3.4) associate into various homo/hetero-tetrameric complexes. GIRK2 (Kir3.2) forms functional homotetramers and is thus a good candidate for crystallographic structure determination (Kofuji et al., 1995). To obtain suitably diffracting crystals, we modified the mouse GIRK2 cDNA to remove unstructured regions of the N- and C-termini (Nishida et al., 2007; Nishida and MacKinnon, 2002; Pegan et al., 2005; Tao et al., 2009). The resulting channel differs from the corresponding human ortholog by only one amino acid near the structured end of the C-terminus (Asn377 is Ser in human GIRK2) (Figure S2). This channel with unstructured regions of the N- and C-termini removed, which we refer to as wild type in this study, exhibits the fundamental characteristics of the full-length GIRK2: G protein activation, inhibition by tertiapin-Q, and a strongly rectifying current-voltage curve (Figure 1C,D).
Wild type GIRK2 crystals diffracted X-rays to 3.6 Å resolution. Initial phases were determined by molecular replacement using a GIRK2 CTD structure and a model was built and refined to working and free residuals (Rw/Rf) of 26.0/27.3% (Figure 1A,B and Table S1) (Inanobe et al., 2007). The overall architecture of GIRK2 is similar to the G protein-independent ‘classical inward rectifier’ channel Kir2.2 (Tao et al., 2009), but has two significant differences. First, the turrets surrounding the extracellular entryway to the pore form a wider, more open vestibule in GIRK2 (Figure S1B,C). This structural difference may provide a simple explanation for pharmacological differences between classical inward rectifiers and GIRK channels. Many GIRK channels, including GIRK2, are inhibited by certain pore-blocking toxins such as tertiapin, as shown in Figure 1C, whereas classical inward rectifier channels are not (Jin and Lu, 1999). The more open turrets in GIRK2 would allow tertiapin to fit into the vestibule, whereas the more restrictive turrets in classical inward rectifiers appear to prevent toxin binding. The second structural difference occurs at the interface between the TMD and CTD. In Kir2.2 the CTD is extended away from the TMD, whereas in GIRK2 the two components are tightly juxtaposed (Figure 1A,B and Figure S1A).
The TMD-CTD interface in GIRK2 is mediated by both hydrophilic and hydrophobic interactions between the interfacial helices of the TMD, the TM-CTD linker, and the βC-βD loop of the CTD (Figure 1B). These interactions were absent in the more extended Kir2.2 structure (Tao et al., 2009). It seems likely that they play an important role in the control of GIRK2 channel activity because they are in close proximity to the two constrictions along the ion pathway that have been hypothesized to function as gates. One gate - the inner helix gate - is formed by the inner helices of the TMD, just inside the membrane, above the level of the interfacial helix (Figure 1A and Figure S1A). Another gate - the G-loop gate - is formed by the G-loop at the apex of the CTD, just outside the membrane, below the level of the interfacial helix (Figure 1A and Figure S1A). In this structure of GIRK2, both gates are tightly closed.
Near the TMD-CTD interface, immediately beneath the βC-βD loop, there is electron density that cannot be attributed to protein atoms (Figure 2A). Given the surrounding protein chemical groups, this density most likely represents either a metal ion, for example Na+, or a water molecule. To distinguish between these possibilities we crystallized the channel in the presence of Tl+, a monovalent metal ion identifiable by its X-ray anomalous signal, which has been used previously to analyze the Na+ binding site in the transport protein LeuT (Boudker et al., 2007). An anomalous peak, the third strongest in the anomalous difference electron density map (4.6 σ, after sites in the selectivity filter and at the intracellular pore entryway), identifies this extra density as a metal cation (Figure 2B). In the native structure, the density is undoubtedly due to a Na+ ion coming from the crystallization solution of 1M NaCl.
The discovery of a Na+ binding site is interesting because GIRK channels that contain either Kir3.2 or Kir3.4 subunits are known to be activated by elevated levels of intracellular Na+ with an EC50 of 30–40 mM (Ho and Murrell-Lagnado, 1999a, b; Lesage et al., 1995; Sui et al., 1996; Sui et al., 1998; Zhang et al., 1999). Sodium activation of GIRK channels is thought to serve an important physiological function by producing negative feedback on excessive electrical excitability: during excitation, Na+ entry can elevate intracellular Na+ concentrations above the normal range of 5–15 mM, enough to activate GIRK channels and drive the membrane potential negative again. Mutagenesis studies have pinpointed an aspartic acid in the βC-βD loop as a critical determinant of Na+ activation (Asp228 in Kir3.2), and the presence of aspartic acid rather than asparagine accounts for the Na+ sensitivity of GIRK channels with Kir3.2 and Kir3.4 subunits (Ho and Murrell-Lagnado, 1999a, b; Zhang et al., 1999) (Figure S2). Asp228 is one of the coordinating residues of the Na+ site in the crystal structure and when Asp228 is mutated to asparagine, the Na+ density is no longer observed (Figure 2C). This direct correlation between structure and function observed when mutating Asp228 suggests that we have identified the regulatory Na+ binding site. In the refined model, Na+ is coordinated not only by the side chain carboxylate of Asp 228, but also by main chain carbonyl oxygen atoms from Arg230, Asn231 and Ser232 (Figure 2A). Main chain carbonyls from Leu275 and Val276 in the βE-βG loop may also participate, as well as the flanking histidines (His69 and His233), which may help to coordinate a water molecule near this site. As we present additional data below it will become clear why this Na+ ion is strategically located in the channel to modulate gating.
We suspected that the TMD-CTD interface likely plays an important role in controlling the channel’s gates, and in particular in transmitting conformational changes that allow G proteins to regulate the gates. Using the closed structure of GIRK2 to guide our experiments, we introduced point mutations into the TMD-CTD interface and assessed their effects using a functional assay developed by Jan and coworkers (Kubo et al., 1993). In this assay, the GIRK2 channel was co-expressed in Xenopus oocytes with the M2 muscarinic G protein coupled receptor (Kubo et al., 1993). Stimulation of the M2 receptor by acetylcholine (ACh) causes GIRK2 channel opening, mediated by endogenous G proteins in the oocyte. When the wild type GIRK2 channel is expressed, acetylcholine application stimulates large K+ currents (Figure 1C). Subsequent application of tertiapin-Q (TPN-Q) distinguishes GIRK2 currents from endogenous oocyte channel currents. For the wild type channel, the magnitude of current stimulated by acetylcholine nearly equals that inhibited by TPN-Q, which means there is little or no G protein-independent GIRK2 current (Figure 1C).
Mutations in the TMD-CTD interface were introduced at many of the most conserved positions and in a number of cases the mutations we chose corresponded to naturally occurring mutants that underlie channelopathies (Decher et al., 2007; Donaldson et al., 2003; Lin et al., 2006; Plaster et al., 2001; Zhang et al., 1999) (Figure S2). Not surprisingly, the majority of these mutations yielded nonfunctional GIRK2 channels (listed in Figure 3 legend). The mutations that did express are shown in Figure 3A. Grey bars show that functional expression levels in each case are comparable to wild type, while black bars show the fraction of GIRK2 current that is stimulated by the addition of acetylcholine. This fraction is near unity for wild type and four of the mutants (Figure 3A). In contrast, the R201A mutation stands out because its acetylcholine-stimulated fraction is less than 0.2, which means that its activity is largely independent of acetylcholine (Figure 3B).
To analyze the structural alterations underlying constitutive activation in the R201A mutant, we determined its crystal structure at 3.1 Å resolution. Compared to wild type, this channel shows a large rearrangement of the βC-βD loop, movement of the His233 side chain to fill the void left by deletion of the Arg201 side chain, and a shifting of the strand comprising residues 235–237 to interact more closely with residues 272–275 of the βE-βG loop (Figure 3C). These conformational changes are associated with a displacement and rotation of the G loop and a change in the positions of three amino acids that form the gate’s constriction: Met313, Gly318 and Met319 (Figure 3D, E Supplementary Movie 1). The net effect of these concerted changes is a widening of the G loop gate from about 6.0 Å to nearly 12.0 Å, and the appearance of oxygen atoms from the Gly318 main-chain carbonyl and the Thr320 side chain to face the pore (Figure 3D,E). These conformational changes effectively create a 9 Å diameter hydrophilic pore (as delimited by the van der Waals surfaces of the pore-lining oxygen atoms), which should be sufficient to allow passage of an 8 Å diameter hydrated K+ ion.
The R201A mutant also shows a propagated conformational change that extends to the perimeter of the CTD, a distance of approximately 30 Å from the site of the R201A mutation, and encompasses the Na+ binding site (Figure 4, Supplementary Movie 1). This propagated conformational change is mediated by a domino-like displacement of β strands, which in turn leads to a reorganization of several hydrophobic amino acids (Val67, Leu257, and Val276) and a rotamer switch in the conformation of the Tyr58 side chain near the CTD perimeter. Several independent lines of evidence suggest that this propagated conformational change is similar to the conformational change that occurs when G proteins bind to the channel.
First, site-directed mutagenesis studies of GIRK/Kir2.1 chimeric channels have identified several buried and surface resides as being essential in mediating G protein activation of GIRK channels. The critical surface residues are located around the βL-βM loop of the CTD, especially residue Leu344, thus implicating this region as a binding site for G proteins (Finley et al., 2004; He et al., 1999). Therefore it is reasonable to think that G protein binding to this region could regulate this conformational change, leading to G loop gate opening. It was also shown that mutation of the buried residue Leu273 to isoleucine eliminated G protein activation (He et al., 2002). Until now it was unclear how this would affect channel function, but here we can see that Leu273 is located on the βE-βG loop next to Val276 and that it undergoes a significant conformational change in the R201A mutant. It is conceivable that a mutation at this position could alter the propagation of a conformational change from the surface of the protein to the G loop.
Second, NMR studies of the interaction between G protein subunits and the GIRK1 CTD using transferred cross saturation and chemical shift perturbation experiments have identified regions on the CTD that either directly contact a G protein or undergo a conformational change in response to G protein binding (Yokogawa et al., 2011). These studies concluded that G proteins bind to a surface mainly comprised of the βL-βM and βD-βE loops (Figure S3B). They also concluded that G protein binding elicits conformational changes near this surface as well as at locations within the N-terminus, especially Tyr58 (Figure S3C,D). The NMR data match the conformational changes that we observe in the βL-βM loop, N-terminus, and in particular Tyr58, which is the residue that undergoes a rotamer switch in the R201A mutant (Figure S3A and Figure 4). We will show below that the R201A mutant in the presence of PIP2 reveals an additional conformational change that is consistent with the NMR data on the βD-βE loop (Figure S3E,F).
Taken together, the mutagenesis and NMR data support the hypothesis that constitutive activation in the R201A mutant channel results because this mutant favors a conformation similar to that induced by G protein binding. The G loop gate is open in the R201A mutant, but the inner helix gate remains closed. We note, however, that G protein stimulation alone is insufficient to achieve ion conduction in GIRK channels. In electrophysiological assays, the signaling lipid PIP2 is also required, in addition to G proteins, for channel activation.
We next determined the crystal structure of wild type GIRK2 in the presence of C8-PIP2 (8 carbon acyl chains) at 3.0 Å resolution. The structure shows one ordered PIP2 lipid molecule per subunit bound near the TMD-CTD interface. The negatively charged phosphates of the PIP2 molecule are coordinated by several positively charged residues: Lys64, Lys194, Lys199, and Lys200, along with backbone amides at the junction of the interfacial and outer helices (Figure 5, Figure S4A,B). The side chains for residues Lys90 and Arg92 did not show any appreciable electron density, but may still contribute to an overall positive electrostatic potential of the binding site (Figure 5, Figure S4A,B). The presence of PIP2 induced a modest displacement of the protein main chain near the PIP2 binding site, where the interfacial helix turns into the outer helix (Figure S4C). PIP2 also produced a slight rotation of the inner helices, accompanied by a weakening of electron density for the Phe192 side chain, which forms the most constricted region of the inner helix gate (Figure S4D,E,F). Clearly, however, both the G loop gate and inner helix gate remain closed in the presence of PIP2, consistent with the observation that PIP2 alone is insufficient to open GIRK channels in electrophysiological experiments.
In contrast to the small effect of PIP2 on the conformation of the wild type channel, PIP2 had a profound effect on the R201A mutant. A crystal structure in the presence of C8-PIP2, determined at 3.5 Å resolution, shows a large change in both the G loop and the inner helix gates (Figure 6A,B, Supplementary Movie 2). Because of the way in which channel molecules are packed against each other in this orthorhombic crystal, only two of the four subunits are free to bind PIP2 and undergo the conformational change (Figure S5). In the PIP2 bound subunits, we observe conformational changes similar to those observed in the R201A mutant. We also observe a rigid body rotation of the CTDs, which necessitates a movement of the βD-βE loop (Figure 6C,D, Figure S3E,F). The rotation of the CTDs is propagated across the TMD-CTD interface and is associated with a rotation and splaying apart of the inner helices. The net effect of PIP2 in combination with the R201A mutation is an opening of the G loop gate to 15 Å and the inner helix gate to 11 Å (Figure 6A,B, Supplementary Movie 2).
Figure 7A shows a surface rendering of the entire pore lining for the wild type channel and for the R201A mutant in the presence of PIP2. Although we do not know for sure whether in a cell two or four PIP2 molecules bind to the channel in its G protein-activated state, this rendering shows the case in which we allow four subunits to bind PIP2 and undergo the conformational change, which we anticipate would likely occur in the unconstrained environment of the membrane. It is evident that in the setting of dual activation by G protein subunits (mimicked by the R201A mutant) and PIP2, the highly constricted pore of the quiescent wild type channel opens wide enough to allow a hydrated K+ ion passage from the cytoplasm to the selectivity filter.
This study presents the first molecular structures of a G protein gated K+ channel, GIRK2. Crystallization of the wild type channel and of a constitutively active mutant, both in the absence and presence of the signaling lipid PIP2, reveal four distinct structures that we believe represent physiologically relevant conformations that underlie GIRK channel gating. In contrast to voltage-dependent and other ligand-gated K+ channels, GIRK channels clearly contain two functional gates – an inner helix gate and a G loop gate – that are regulated by a combination of cytoplasmic and membrane stimuli.
Until this study, all previous structures of Kir family channels including several bacterial family members, a chimera with a bacterial TM and eukaryotic CTD, and a eukaryotic ‘classical inward rectifier’, exhibited a tightly closed inner helix gate (Clarke et al., 2010; Kuo et al., 2003; Nishida et al., 2007; Tao et al., 2009). The conformation of the inner helix gate in three structures presented here – wild type without PIP2, wild type with PIP2, and the R201A mutant without PIP2 – have a closed inner helix gate conformation similar to previous structures. In the R201A mutant in the presence of PIP2 we observe an open inner helix gate conformation. As discussed above, we think it is likely that crystal lattice contacts prevented PIP2 binding and opening in two of the subunits, but that in a membrane all four subunits would bind PIP2 and undergo the conformational change. If four subunits undergo the change, then the inner helix gate will open wide enough to permit passage of a hydrated K+ ion, but not as wide as in voltage-gated K+ and MthK Ca2+ gated K+ channels (Jiang et al., 2002; Jiang et al., 2003; Long et al., 2005; Long et al., 2007). A narrower passageway from the cytoplasm to the selectivity filter in inward rectifier K+ channels might be functionally significant because strong voltage-dependent block by intracellular polyamines and Mg2+ give rise to their namesake conduction property – inward rectification (Lopatin et al., 1994; Matsuda et al., 1987). The strong voltage dependence of these blocking ions is in part due to the coupled movement of blocking and permeant ions in the pore (Spassova and Lu, 1998). A very wide ion pathway would allow blocking and permeant ions to interchange their positions along the pathway (i.e. slip by each other), but a narrower pathway will force the ions to move in a queue, and thus the movement of K+ will impart excess voltage dependence onto the blocking ion and give rise to stronger rectification (Spassova and Lu, 1998).
We discovered the constitutively active R201A mutant by evaluating the initial wild type closed structure and then perturbing the TM-CTD interface because it appeared as if it ought to be involved in regulating the gates. Not surprisingly, most mutations in this region abolished channel function. Fortuitously, in the R201A mutant we observe constitutive activation associated with an open G loop gate and a propagated conformational change across the CTD. The extent and distribution of this conformational change matches very well previous mutational studies implicating the βLβM loop as a site for binding G protein subunits, and NMR studies on the effect of G protein binding to isolated CTD structures (Finley et al., 2004; He et al., 2002; He et al., 1999; Yokogawa et al., 2011). We thus hypothesize that the R201A mutant stabilizes a conformation that is similar to the G protein activated state.
Electrophysiological experiments have shown that elevated levels of Na+, along with the presence of PIP2, can activate certain GIRK channels that have an aspartic acid residue at position 228 (Ho and Murrell-Lagnado, 1999a, b; Lesage et al., 1995; Sui et al., 1996; Sui et al., 1998; Zhang et al., 1999). Our data may start to address the mechanism of this mode of activation. We show that a Na+ ion binds in a pocket between the βC-βD and βE-βG loops. This lies directly on the pathway of the propagated conformational changes that we observed in the R201A structure, which we believe to mimic the effects of a bound G protein. Thus, Na+ may work to influence the conformation of this pathway. One possible mechanism by which the Na+ ion could do this is by binding to Asp228 and thereby weakening the ionized hydrogen bond that it forms with Arg201. Through such an interaction we imagine that Na+ could in part mimic the effect of the R201A mutation and, by extension, the effect of G proteins as well.
There is a very good correlation between the four crystal structures and past electrophysiological studies showing a dual requirement for both G proteins and PIP2 to open GIRK channels (Huang et al., 1998; Sui et al., 1998; Zhang et al., 1999). We can now view this dual requirement through a thermodynamic cycle of the four structures: PIP2 alone binds but does not open either gate, G proteins (R201A) alone open the G loop gate but not the inner helix gate, but G proteins in addition to PIP2 open both gates (Figure 7B, Figure S6). In words, the relations suggest that PIP2 functions to couple tightly the two gates so that upon G protein stimulation both gates open to activate the channel.
One well established role of the CTD is to form an extended pore that is important for producing rectification (Nishida and MacKinnon, 2002; Tao et al., 2009; Yang et al., 1995). In this study we observe how the CTD also has the ability to undergo conformational changes that allow cytoplasmic ligands to allosterically control gates in the pore. In realizing that G proteins apparently regulate the gates through a pathway of conformational change that includes a modulatory Na+ binding site, we have begun to see the structural underpinnings of a very complex form of gating regulation.
A truncated GIRK2 cDNA (consisting of residues 52–380 – hereafter referred to as wild type) was subcloned by PCR into pPICZ and pGEM vectors for expression in Pichia pastoris, or Xenopus laevis oocytes, respectively. The ORF in the pPICZ vector contained a C-terminal PreScission protease site, followed by green fluorescent protein (GFP), and a His10 tag. The pGEM vector contained a C-terminal FCYENE tag (Ma et al., 2001)) and then a c-Myc tag.
Protein expression was induced in stable P. pastoris cell-lines by the addition of methanol for 20–24 hrs at 24 °C. Cells were harvested by centrifugation, frozen in liquid N2, and stored at −80 °C until needed. Frozen cells were lysed in a mixer mill, then solubilized for 1 hour at room temperature (RT) by resuspending in 50 mM Hepes pH 7.35, 150 mM KCl, 4 % (w/v) n-decyl-β-D-maltopyranoside (DM), and protease inhibitor cocktail. Clarified supernatant was then incubated with Talon metal affinity resin for 1 hr at RT with gentle mixing. The resin was washed in batch with 5 column volumes (cv) of Buffer A (50 mM Hepes pH 7.0, 150 mM KCl, 0.4 % (w/v) DM, then loaded onto a column and further washed with 5 cv Buffer A + 40 mM imidazole, then 2 cv Buffer A + 80 mM. The column was then eluted with Buffer A + 300 mM imidazole. Peak fractions were pooled and 20 mM DTT, 3 mM TCEP, 1 mM EDTA were added, and then cut with PreScission protease for either 2 hrs at RT or overnight at 4C. The cleaved protein was then concentrated to run on a Superdex-200 gel filtration column in 20 mM TRIS-HCl pH 7.5, 150 mM KCl, 0.2 % (w/v) DM (anagrade), 20 mM DTT, and 1 mM EDTA.
Purified protein representing the peak tetramer fractions was pooled and concentrated in a 50 K MWCO concentrator to 6–7 mg/mL, mixed 1:1 with crystallization solution, then crystallized using the hanging drop vapor diffusion method. For the PIP2 complexes, 1,2-dioctanoyl-sn-glycero-3-phospho-(1′-myo-inositol-4′,5′-bisphosphate) (C8-PIP2, Avanti Polar Lipids) was added to the concentrated protein at a final concentration of 2 mM right before setting up the drops. For R201A + PIP2, 10 mM spermine was also added, although it was not seen in the electron density maps. Crystallization drops were incubated at 20 °C and crystals (D-shaped plates) usually appeared after 1–3 days. The wild type and R201A mutant crystals grew in 50 mM Na citrate pH 6.0, 1 M NaCl, 30–35 % PEG 400. The D228N mutant crystals grew in 50 mM Na citrate pH 6.0, 1 M NaNO3, 24–26 % PEG 400. The wild type + PIP2 crystals grew in 50 mM Na citrate pH 6.0, 1 M NaCl, 20 % PEG 400. The R201A mutant + PIP2 crystals grew in 50 mM Na Hepes pH 7.25, 0.5 M NaCl, 25 % PEG 400. For collecting Tl+ anomalous diffraction data, wild type protein was crystallized in the presence of KNO3 instead of NaCl. These crystals were then transferred to solutions containing TlNO3. For data collection, the crystals were cryoprotected and flash-frozen in liquid N2. The structures were solved by molecular replacement with the MOLREP (Vagin and Teplyakov, 2000) program, using the GIRK2 cytoplasmic domain structure as a search model (Inanobe et al., 2007). The models were built using COOT (Emsley and Cowtan, 2004) and refined with REFMAC (Murshudov et al., 1997). Data collection and refinement statistics are shown in Table S1.
X. laevis oocytes were injected with 50 nL cRNA and incubated for 1–3 days before recording currents. All experiments were performed at room temperature using two-electrode voltage-clamp. For gap-free recordings, oocytes were held at −80 mV in ND96 (96 mM NaCl, 2 mM KCl, 0.3 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, pH 7.6 with KOH), then perfused with a highK solution (98 mM KCl, 0.3 mM CaCl2, 1 mM MgCl2, 10 mM HEPES, pH 7.6 with KOH) that also contained either 10 μM acetylcholine (ACh) or 1 μM tertiapin-Q (TPN-Q). GIRK2-specific current was calculated by subtracting the current remaining after TPN-Q blockage (background oocyte currents) from the maximum current elicited with highK + ACh. Basal GIRK2 current was calculated by subtracting the current remaining after TPN-Q blockage from the maximum current elicited with highK only. For measuring rectification, the membrane potential was ramped from −80 mV to +80 mV over 100 ms in the presence of either highK buffer + 1 μM ACh to measure total currents, or highK + 1 μM ACh + 100 nM TPN-Q to measure background oocyte currents. GIRK2-specific currents were calculated by subtracting the background currents from the total currents.
We thank P. Hoff and members of D. Gadsby’s laboratory(Rockefeller University) for assistance with oocyte preparation; K. R. Rajashankar and K. Perry at beamline 24ID-C (Advanced Photon Source, Argonne National Laboratory), H. Robinson at beamline X29 (National Synchrotron Light Source, Brookhaven National Laboratory), and M. Becker at beamline 23ID-B (Advanced Photon Source, Argonne National Laboratory) for assistance at the synchrotron; members of the MacKinnon laboratory for assistance; and A. Banerjee, J. Butterwick, X. Tao, and P. Yuan for comments on the manuscript. R.M. is an Investigator in the Howard Hughes Medical Institute.
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