We began by characterizing the expression of RGS complex subunits in the mouse hippocampus. We detected robust expression of RGS6, RGS7, R7BP and Gβ5 by western blotting (Figure 1A). To understand the contribution of individual subunits to GABA_BR-GIRK signaling, we studied the effects of selective knockout of Rgs6, Rgs7, and R7bp in mice. Elimination of RGS7 dramatically reduced levels of Gβ5 and R7BP in the hippocampus (Figure 1A,B). In contrast, elimination of RGS6 had no significant effect on the expression of R7BP or Gβ5. Similarly, loss of one RGS protein did not affect the expression of the other, or the GIRK channel subunit GIRK2. Given the interdependence of subunit expression in R7 RGS complexes (Chen et al., 2003; Anderson et al., 2007; Grabowska et al., 2008), these results suggested that RGS7 was likely the dominant catalytic subunit in the hippocampus.

Figure 1

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We next compared GABA_BR-GIRK responses in hippocampal pyramidal neurons from Rgs6, Rgs7, and R7bp knockout (^−/−) and wild-type (WT) mice (Figure 1C–F). Application of a saturating concentration (100 μM) of the GABA_BR agonist baclofen elicited currents with similar maximal amplitudes in neurons of all genotypes (Figure 1D). We also measured the activation and deactivation kinetics of the baclofen-induced currents. At this saturating concentration of baclofen, we observed no significant differences between genotypes in the activation phase of the response (Figure 1C,F). Desensitization of current during the timeframe of agonist application was negligible and similar across genotypes, suggesting that the response at the steady-state is not compounded by the GIRK channel inactivation (Figure 1C,E). While no change in the current deactivation kinetics was observed in Rgs6^−/− neurons, elimination of RGS7 markedly slowed response deactivation (Figure 1C,G). The rate of GIRK current deactivation in R7bp^–/– neurons was also slower than in wild-type neurons, but the effect was substantially smaller than seen in neurons from Rgs7^−/− mice (Figure 1C,G). Importantly, there were no detectable changes in any of the measured response parameters in Rgs6^−/− neurons as compared to wild-type, arguing that GABA_BR-GIRK signaling in hippocampal neurons is modulated by RGS7 and R7BP, but not RGS6.

Given the observed changes in the kinetics of GIRK channel modulation by GABA_BR, we next sought to determine the impact of ablating Rgs7 and R7bp on the sensitivity of the GIRK response to GABA_BR activation. Increasing baclofen concentrations caused a progressive enhancement in GIRK-mediated currents in all genotypes (Figure 2A). However, pronounced differences in the concentration-response relationship were evident (Figure 2B). First, in both Rgs7^−/− and R7bp^−/− neurons, curves were shifted to the left relative to wild-type neurons, indicating that ablation of RGS7 or R7BP increased GABA_BR-GIRK coupling sensitivity. Second, while RGS7 ablation resulted in a largely parallel leftward shift of the curve, its shape in R7bp^−/− neurons was markedly steeper. Indeed, response amplitudes resembled those seen in wild-type neurons at lower agonist concentrations, but at higher concentrations, the responses more resembled those seen in Rgs7^−/− neurons.

Figure 2

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We next tested whether GABA_BR-GIRK current kinetics exhibited a similar dependence on agonist concentration. Comparing activation rates across different concentrations of baclofen revealed that onset kinetics were affected by RGS7 ablation only at low baclofen concentrations, whereas no effect of R7BP elimination was seen at any level of GABA_BR stimulation (Figure 2C). Since GIRK activation kinetics could be influenced by the G protein deactivation cycle (Doupnik et al., 1997; Lambert et al., 2010), we did not put significant emphasis on the analysis of the differences in the raising phases of the response. The impact of RGS7 ablation on deactivation rates was consistent at different agonist concentrations (Figure 2D). In contrast, R7BP ablation significantly affected deactivation kinetics only at higher agonist concentrations, consistent with the effects on response sensitivity (Figure 2D).

Analysis of the responses also revealed that elimination of RGS7 and R7BP resulted in a significant delay between agonist removal and the beginning of current deactivation (Figure 2E). This lag time showed an exponential dependence on agonist concentration that continued to develop past the saturation point for the maximal GIRK response (∼10 μM baclofen) in all genotypes (Figure 2F). While there was no difference in lag time between genotypes at non-saturating baclofen concentrations (∼EC₆₀), it became pronounced as the current response reached saturation (∼EC₉₀) (Figure 2G). Combined with the observations that GABA_BR continues to increase the amount of activated G proteins past the saturation point of the GIRK channel response (Hensler et al., 2012), these data suggest that the lag time reflects the clearance of free βγ subunit produced above the stoichiometric level relative to the GIRK channel.

To provide an independent evidence that increase in the lag time reflects changes in stoichiometry of Gβγ subunits relative to their effector molecule, we utilized a bioluminescence resonance energy transfer (BRET)-based approach that monitors interactions of Gβγ with a reporter derived from an effector (GRK3) upon reconstitution in transfected cells (Figure 3A). In this assay, we induced production of free Gβγ subunits via GABA_BR activation and then measured delay time between antagonizing GABA_BR and the onset of signal decay, while changing the Gβγ to GRK3 effector ratio (Figure 3B). In agreement with the electrophysiological recordings of GIRK channel activity, BRET experiments showed that the increase in Gβγ stoichiometry over an effector results in prolongation of the response deactivation lag (Figure 3C,D).

Figure 3

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In transfected cells, R7BP is essential for the membrane localization of RGS7 (Drenan et al., 2005; Narayanan et al., 2007). Furthermore, we previously reported that knockout of R7bp resulted in a reduction of the total membrane-bound RGS7 protein in hippocampal tissue (Panicker et al., 2010). To analyze the effect of R7BP on RGS7 localization in hippocampal pyramidal neurons, we performed high-resolution immunoelectron microscopy. Consistent with the earlier findings, immunoparticles for RGS7 were abundant on the extrasynaptic plasma membrane of dendritic spines and dendritic shafts of pyramidal cells, as well as at intracellular sites (Fajardo-Serrano et al., 2013). In the hippocampus of R7bp^−/− animals, immunoparticles for RGS7 were distributed similarly to the wild-type, but they were more frequently observed just beneath the plasma membrane and more broadly distributed in somata (Figure 4). Indeed, quantitative analysis indicated that in R7bp^−/− mice, RGS7 was less frequently detected in the plasma membrane and tended to accumulate within 100 nm of the plasma membrane. We also detected a significant increase in RGS7 labelling in the rough endoplasmic reticulum (rER) in the soma of R7bp^−/− pyramidal neurons (2489 vs 3371 immunoparticles in wild-type and R7bp^−/− neurons, respectively). These findings indicate that knockout of R7BP caused a modest but significant redistribution of RGS7 away from the plasma membrane.

Figure 4

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The slower kinetics and increased sensitivity of GIRK channels to GABA_BR activation suggested that there was a net up-regulation of GIRK-dependent inhibition in hippocampal neurons lacking RGS7. Since GIRK channels significantly contribute to setting neuronal excitability (Chen and Johnston, 2005), we next determined how deletion of RGS7 influences the excitability of CA1 pyramidal neurons. To characterize intrinsic electrophysiological properties of different genotypes, the responses of CA1 neurons to somatic current injections ranging from −150 pA to +300 pA were measured (Figure 5A). While membrane resistance was similar in both genotypes (Rin = 120 ± 10 MΩ vs 105 ± 11 MΩ, p=0.31, in wild-type and Rgs7^−/− cells, respectively), resting membrane potential (RMP) was significantly hyperpolarized in Rgs7^−/− neurons (−64.7 ± 0.5 mV vs −68.5 ± 0.7 mV in wild-type and Rgs7^−/−, respectively; Figure 5B). In addition, the current required to elicit action potentials (APs) was significantly higher in RGS7^−/− neurons (I = 104.3 ± 8.8 pA vs 146.4 ± 17.0, in WT and Rgs7^−/− cells, respectively; Figure 5C). Finally, Rgs7^−/− neurons fired significantly fewer action potentials in response to depolarizing current as compared to wild-type controls (Figure 5D). Collectively, these observations argued that hippocampal pyramidal neurons from Rgs7^−/− mice were less excitable than wild-type counterparts.

Figure 5

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We further examined whether decreased intrinsic excitability of Rgs7^−/− neurons is caused by dysregulated net excitatory transmission. For this purpose, we measured spontaneous EPSCs (sEPSC) in CA1 pyramidal neurons (Figure 5E–G). There were no significant differences in either amplitudes or frequencies of sEPSC events between Rgs7^−/− and wild-type neurons (Figure 5F–G). These observations indicate that excitatory input into CA1 pyramidal neurons is unchanged by elimination of RGS7 and suggest that the observed decrease in neuronal excitability is caused by changes in the intrinsic membrane properties of CA1 neurons, likely stemming from enhanced activity of the postsynaptic GIRK channel.

Previous studies have implicated GIRK channels in several forms of hippocampal synaptic plasticity, including long-term potentiation (LTP) (Cramer et al., 2010), long-term depression (LTD) (Cooper et al., 2012) and depotentiation (DP) (Chung et al., 2009; Cooper et al., 2012). Because GABA_B receptors play key roles in these processes and since GABA_B-GIRK signaling is severely dysregulated in Rgs7^−/− neurons, we next examined the impact of RGS7 ablation on hippocampal synaptic plasticity.

A high-frequency stimulation protocol of 2 tetanized stimuli (TS, 100 Hz for 1s each) produced robust LTP in both genotypes (Figure 6A,D). The extent of the potentiation in Rgs7^−/− slices was not significantly different from that measured in wild-type slices, arguing that elimination of RGS7 has no effect on LTP. A low-frequency stimulation (LFS, 2 Hz for 10 min, 1200 pulses) elicited LTD in both wild-type and Rgs7^−/− hippocampal slices (Figure 6B,D). However, the extent of inhibition was significantly smaller in Rgs7^−/− slices. Interestingly, the magnitude of the first fEPSP after the train of LFS stimulation was significantly reduced in knockout mice (51 ± 6% and only 72 ± 6% in wild-type and Rgs7^−/− slices correspondingly, *p=0.03, t test, n = 6), suggesting that the LTD impairment in Rgs7^−/− is due to a deficit in an induction mechanism(s). The depotentiation produced by consecutive application of TS and LFS was also significantly impaired in Rgs7^−/− mice (Figure 6C,D).

Figure 6

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We further investigated the impact of RGS7 ablation on paired-pulse facilitation (PPF), a form of short-term plasticity that results from the enhancement of presynaptic vesicle release in response to two closely-spaced stimuli (Zucker, 1989; Dobrunz and Stevens, 1997). We found that fEPSP facilitation was similar between genotypes across the examined range of 10–1000 ms interpulse intervals (Figure 6E), indicating that presynaptic mechanisms are not likely to be the cause of the observed alterations in synaptic plasticity. We next compared the dependence of fEPSP slopes (Figure 6F) and presynaptic fibre volley (FV) amplitudes (Figure 6G) on stimulus intensity. Analysis of the relationship between FV amplitudes and fEPSP slopes by linear fitting reveals their nearly perfect correspondence between wild-type and Rgs7^−/− slices, indicating preservation of basal synaptic transmission properties (Figure 6H). Together, these results suggest that RGS7 elimination selectively impaired LTD and DP forms of synaptic plasticity, likely by a post-synaptic mechanism.

Changes in the intrinsic excitability of CA1 pyramidal neurons, together with deficits in hippocampal synaptic plasticity, suggested that RGS7 complexes play a role in spatial learning and memory. To test this possibility, we first studied the impact of RGS or R7BP ablation in contextual fear conditioning, a test that requires hippocampal processing for memory formation (Maren, 2001). We found significant deficits in context recognition 24-hr after training in Rgs7^−/− mice, but not in Rgs6^−/− or R7bp^−/− mice (Figure 7A–C). Importantly, we observed no difference in baseline freezing behavior before associative training between the genotypes. Furthermore, there were no significant differences between wild-type mice and mutant mice in amygdala-dependent cue recognition.

Figure 7

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To clarify the impairment of hippocampus dependent learning and memory in Rgs7^−/− mice, we conducted other tests that rely on hippocampal function. In the Morris water maze test, Rgs7^−/− mice showed delayed escape latencies and reduced success rates during acquisition trials (Figure 7D,E). In the probe trial, when the platform was removed 24 hr after training, it took Rgs7^−/− mice significantly longer to reach the target area where the platform was located during training as compared to their wild-type littermates (Figure 7F). Mice lacking RGS7 also made fewer crossings over the target area during the probe trial. Importantly, despite a pronounced spatial impairment, Rgs7^−/− mice performed normally in the visible platform version of water maze.

In the novel object recognition test, wild-type littermates spent significantly more time exploring novel objects over familiar objects (Figure 7G). However, Rgs7^−/− mice spent approximately equal time exploring familiar and novel objects. Furthermore, there was a significant difference in time exploring novel objects between Rgs7^−/− mice and their wild-type littermates. No genotype differences in object exploratory behavior were observed during the training day, when the mice were first introduced to identical objects. Taken together, these results indicate that elimination of RGS7 leads to a disruption of spatial learning and memory in mice.

The results of this study, together with prior investigations, establish the RGS7/Gβ5/R7BP complex as an essential regulator of GABA_BR-GIRK signaling in the hippocampus. In hippocampal neurons, RGS7 is closely co-localized with both GABA_BR and GIRK2-containing channels (Fajardo-Serrano et al., 2013). Furthermore, in transfected cells, RGS7 can directly interact with GIRK channel subunits as demonstrated by both co-immunoprecipitation and bioluminescence energy transfer approaches (Xie et al., 2010; Zhou et al., 2012). The complex may further involve GABA_BR as well as G protein subunits (Kovoor and Lester, 2002; Fajardo-Serrano et al., 2013), supporting the contention that components of the entire pathway are scaffolded into a larger macromolecular assembly.

The interaction between RGS7 with GIRK channels is mediated by the Gβ5 subunit, which mimics signal-transducing Gβγ subunits in binding to the channel (Xie et al., 2010; Zhou et al., 2012). Knockout of Gβ5 greatly slows GABA_BR-GIRK response deactivation rates and dramatically increases agonist sensitivity of the signaling pathway (Xie et al., 2010). In addition to mediating interactions with the channel, Gβ5 is essential for the expression of all four R7 RGS proteins (Chen et al., 2003). With several R7 RGS proteins expressed in hippocampus, the identity of the exact isoform that participates in GIRK channel regulation in the region, through complex formation with Gβ5, remained unknown. The elimination of RGS7, reported in this study, largely phenocopies the loss of Gβ5, both in terms of kinetics and agonist sensitivity. This argues that in hippocampal pyramidal neurons, GIRK signaling is regulated by RGS7 and not by other R7 RGS proteins. Indeed, while RGS6 regulates GIRK signaling in cerebellar neurons (Maity et al., 2012) and sinoatrial pacemaking cells (Posokhova et al., 2010; Yang et al., 2010), RGS6 ablation had no effect on any GIRK response parameters in hippocampal pyramidal neurons.

The function of R7 RGS proteins is regulated by the membrane anchor R7BP, which augments the ability of R7 RGS proteins to deactivate G protein signaling in reconstituted systems (Drenan et al., 2006; Masuho et al., 2013). Consistent with these observations, we show that knockout of R7BP reduces plasma membrane localization of RGS7 and slows GABA_BR-GIRK current deactivation in hippocampal pyramidal neurons. Our results also agree well with a recent study reporting similar deceleration of GIRK deactivation kinetics in R7bp^−/− hippocampal neurons (Zhou et al., 2012). However, the consequences of RGS7 and R7BP elimination are different in two important ways. First, relative to the loss of RGS7 or Gβ5, elimination of R7BP had a very modest effect on GABA_BR-GIRK deactivation kinetics, and was seen only at high GABA_BR agonist concentrations. This suggests that the RGS7/Gβ5 complex can regulate GABA_BR-GIRK signaling without R7BP. Secondly, loss of R7BP and RGS7 has different effects on GABA_BR-GIRK coupling sensitivity. While elimination of RGS7 (or Gβ5) results in a parallel leftward shift in the concentration-response relationship, the concentration-response measured in R7bp^−/− neurons shows markedly greater cooperativity. Thus, at low levels of GABA_BR stimulation, R7BP is dispensable and its elimination fails to affect response amplitude. As the response reaches saturation, R7BP becomes indispensable and its elimination has the same effect on response amplitude as loss of RGS7. These observations are inconsistent with the proposed role for R7BP as a critical factor in the assembly of RGS7-GIRK complexes (Zhou et al., 2012), which postulates the equivalence of R7BP effects across agonist concentrations and a similar impact of R7BP and RGS7 elimination on GIRK channel kinetics, and certainly not an increase in cooperativity of GIRK channel activation.

Based on our findings, we propose an alternative model, where R7BP sets GABA_BR-GIRK coupling efficiency. In this model, at low levels of GIRK activation, when released Gβγ subunits do not saturate the GIRK channel, deactivation kinetics are primarily mediated by RGS7/Gβ5 directly associated with the GIRK subunits. Under these conditions, there are virtually no genotype-dependent differences in the lag time of GIRK deactivation upon agonist removal. With higher agonist concentrations, as GABA_BR produces more Gβγ subunits, the amount of available free Gβγ exceeds that of activatable GIRK channels, evidenced by a delay that precedes GIRK current deactivation upon agonist removal. Under these conditions, R7BP elimination affects response sensitivity as much as elimination of RGS7. While response deactivation kinetics reflect Gβγ inactivation after it had a chance to interact with the channel, response sensitivity likely relates to the efficiency of the Gβγ reaching the GIRK channel. Considering that R7BP affects both of these parameters only when free Gβγ is produced in excess, we think that the RGS7/Gβ5 complex exists in two states: (1) anchored to the GIRK complex where it affects deactivation kinetics, and (2) anchored via R7BP to the plasma membrane outside of GIRK complex where it primarily determines response sensitivity.

In agreement with this model, we observe an increase in cooperativity of GIRK channel activation upon R7BP loss that suggests that RGS7/Gβ5 alone, but not in complex with R7BP, promotes more productive association of Gβγ with the channel. On an intuitive level, this may reflect larger proportion of Gβγ subunits that reach GIRK while being deactivated in its vicinity by RGS7/Gβ5. In contrast, complexes of RGS7/Gβ5 with R7BP may predominantly act elsewhere on the plasma membrane, thereby decreasing the number of Gβγ subunits capable of reaching the channel. By physically binding to the GIRK channel, RGS7/Gβ5 may further act to promote productive interactions of Gβγ (and/or Gα) with the channel, thus increasing cooperativity of its activation. This model is also consistent with the observed decrease in RGS7 on the plasma membrane but largely preserved localization at specific postsynaptic sites where it might be anchored via complex formation with GIRK channels. Thus, the main role of R7BP appears to be in tuning the sensitivity of the response by endowing RGS7/Gβ5 complexes with an ability to deactivate G proteins before they reach the GIRK channel.

RGS proteins are potent negative regulators of both the extent and duration of neurotransmitter signaling via G protein-coupled receptors (Sjogren, 2011; Xie and Martemyanov, 2011). As a result, knockout of individual RGS genes in mice is frequently associated with augmented GPCR signaling. This makes the analysis of changes in sensitivity of behavioral or cellular reactions to neurotransmitter actions associated with elimination of individual RGS proteins a powerful strategy that allows for establishing physiological receptor-RGS pairings. Using this strategy, two RGS proteins have been previously implicated in controlling GABA_BR signaling in vivo: RGS6 in cerebellar neurons (Maity et al., 2012) and RGS2 in the ventral tegmental area (Labouebe et al., 2007). Now, our findings establish that in hippocampal neurons, GABA_BR signaling is negatively regulated by RGS7. These observations complement earlier findings with RGS14, the only other RGS protein implicated in hippocampal synaptic plasticity and spatial learning (Lee et al., 2010). However, in contrast to RGS7 that acts in the CA1 region, RGS14 functions in CA2 synapses and its ablation augments LTP and improves spatial learning in mice.

The major forms of synaptic plasticity (LTP, LTD and depotentiation) in the hippocampus have been implicated in learning and memory (Martin et al., 2000). Recent models propose that all three forms cooperate to affect distinct aspects of spatial information storage (Kemp and Manahan-Vaughan, 2007). In relevance to this thinking, we found a selective disruption of LTD and depotentiation in RGS7^−/− mice, but normal LTP. There is a mounting evidence suggesting an active and selective role of LTD in object recognition and creation of spatial representation of memory (Kemp and Manahan-Vaughan, 2007). For example, LTD but not LTP was found to be important for spatial memory consolidation and memory enhancement for novelty acquisition (Ge et al., 2010; Dong et al., 2012). Consistent with this, electrophysiological recordings in behaving animals show facilitation of LTD during novelty exploration (Xu et al., 1997, 1998; Manahan-Vaughan and Braunewell, 1999; Dong et al., 2012). Disruption of LTD via genetic mutations often results in memory deficits, particularly affecting behavioral flexibility (Nicholls et al., 2008), while LTD enhancement can lead to improvement in spatial reversal learning (Duffy et al., 2008). In particular, our findings resemble the phenotype of SRF^−/− mice, which exhibited a selective disruption of LTD in hippocampal CA1 neurons paralleled by an inability to learn hippocampus-dependent tasks (Etkin et al., 2006). While specific knowledge on how depotentiation contributes to memory is still lacking, it has been observed that exposure to novel environment and spatial exploration depotentiates previously-induced LTP (Xu et al., 1998). Together, our observations in mice lacking RGS7 reinforce the idea that normal inhibitory synaptic plasticity (LTD and depotentiation) is required for hippocampal-dependent learning and memory.

Although, it has never been systematically addressed in a single study, GIRK channel elimination was reported to enhance LTP (Cramer et al., 2010). Given that the Girk2^−/− neurons are relatively depolarized, the enhanced LTP likely results from a general increase in excitability. Conversely, GIRK2 over-expression in transgenic models of Down syndrome leads to reduced LTP (Kleschevnikov et al., 2004; Siarey et al., 2005), but enhanced LTD (Siarey et al., 2005; Cooper et al., 2012). Interestingly, GIRK2 over-expression and ablation both resulted in decreased depotentiation (Chung et al., 2009; Cooper et al., 2012).

Similar to Down syndrome mouse models, Rgs7^−/− mice also exhibited augmented GABA_BR-GIRK signaling. However, in the case of Rgs7^−/− mice, this augmentation resulted in yet another distinct phenotype: selective deficits in LTD and depotentiation, with normal LTP. Several lines of evidence argue that these changes in synaptic plasticity are driven by a postsynaptic mechanism whereby RGS7 controls inhibitory GIRK-dependent signaling in the CA1 pyramidal neurons. First, CA1 pyramidal neurons lacking RGS7 are hyperpolarized and less excitable relative to wild-type neurons, a property that is significantly shaped by the GIRK channels. Second, basal excitatory transmission is unaltered in Rgs7^−/− slices. Third, direct measurements of events that reflect excitatory presynaptic function (e.g., FV and PPR) reveal no changes caused by the elimination of RGS7. Given these observations, we propose that GIRK signaling sets the general excitability of postsynaptic pyramidal neurons, which in turn determines a sliding scale window for the induction of different forms of synaptic plasticity. By adjusting signaling strength in the hippocampal GABA_BR-GIRK pathway, the RGS7/Gβ5/R7BP complex influences the range of neuronal responses necessary for memory formation. While dysregulation of GABA_BR-GIRK signaling in hippocampus may be sufficient for explaining the effects of RGS7 on learning an memory, at this point we cannot rule out a contribution of other brain regions and/or signaling pathways in the process. In any event, we believe that the results of this study establish RGS7 complex as an important molecule for understanding and/or correcting the pathology of neuropsychiatric disorders associated with disruptions in synaptic plasticity and imbalances in inhibitory signaling.

All studies were carried out in accordance with the National Institute of Health guidelines and were granted formal approval by the Institutional Animal Care and Use Committee of the Scripps Research Institute. The generation of Rgs6^−/− (Posokhova et al., 2010), Rgs7^−/− (Cao et al., 2012), R7bp^−/− (Anderson et al., 2007) mice were described earlier. All animals used for comparing genotypes were littermates derived from heterozygous breeding pairs. Mice were housed in groups on a 12 hr light–dark cycle with food and water available ad libitum. Males and females (2–5 months) were used for all experiments.

Lysates were prepared by homogenizing hippocampal tissue from age-matched littermates by sonication in the lysis buffer (1 × PBS, 150 mm NaCl, 1% Triton X-100, protease inhibitors) followed centrifugation at 14,000×g for 10 min. The resulting extract was used for protein concentration determination by the BCA protein assay (Pierce, Rockford, IL). The lysates were adjusted to equalize total protein content by adding lysis buffer and 2x SDS sample buffer. Samples were boiled for 5 min, resolved on SDS-PAGE gels, transferred onto PVDF membrane and subjected to western blot analysis using HRP conjugated secondary antibodies and ECL West Pico (Pierce) detection system. Signals were captured on film and scanned by densitometer, and band intensities were determined using NIH ImageJ software. Rabbit anti-R7BP (TRS) and rabbit Gβ5 (ADTG) were generous gifts from Dr William Simonds (NIDDK/NIH). Anti-RGS6 was generated and used previously (Posokhova et al., 2010). Chicken IgY anti-RGS7 antibody was from Thermo Scientific (Waltham, MA), anti-GIRK2 was purchased from Alomone labs (Jerusalem, Israel) and anti-β-actin was from Sigma (St. Lois, MO).

Primary cultures of hippocampal neurons were prepared using a modified version of a published protocol (Xie et al., 2010). Briefly, hippocampi were extracted from neonatal (P1-3) pups and placed into an ice-cold HBSS/FBS solution: Hank’s Balanced Salt Solution (Sigma; St. Louis, MO), 4.2 mM NaHCO₃, 1 mM HEPES, and 20% FBS. The tissue was washed twice with HBSS/FBS, and then three times with HBSS alone. Hippocampi were digested at room temperature for 5 min with 10 mg/ml Trypsin Type XI (Sigma; St. Louis, MO) in a solution that contained (in mM): 137 NaCl, 5 KCl, 7 Na₂HPO₄, and 25 HEPES (pH 7.2). The tissue was washed twice with HBSS/FBS and three times with HBSS alone, and then hippocampi were mechanically-dissociated in HBSS (supplemented with 12 mM MgSO₄) using Pasteur pipettes of decreasing diameter. The neurons were pelleted by centrifugation (600×g for 10 min at 4°C) and plated onto 8-mm glass coverslips pre-treated with Matrigel (BD Biosciences; San Jose, CA) in 48-well plate. Neurons were allowed to adhere for 30 min prior to adding 0.3 ml of pre-warmed culture medium consisting of Neurobasal A (Life Technologies; Carlsbad, CA), 2 mM GlutaMAX-I (Life Technologies, Carlsbad, CA), 2% B-27 supplement, and 5% FBS. After 4–12 hr, culture media was completely replaced with the same media without FBS. Neurons were incubated at 37°C/5% CO₂, and half of the medium was replaced with fresh medium on each of the first 3 days of culture. Neurons were kept in culture for 10–14 days prior to experiments.

Coverslips containing neurons were transferred to a chamber containing a low-K⁺ bath solution (in mM): 145 NaCl, 4 KCl, 1.8 CaCl₂, 1 MgCl₂, 5.5 D-glucose, 5 HEPES/NaOH (pH 7.4). Borosilicate patch pipettes (3–5 MΩ) were filled with (in mM): 130 KCl, 10 NaCl, 1 EGTA/KOH (pH 7.2), 0.5 MgCl₂, 10 HEPES/KOH (pH 7.2), 2 Na₂ATP, 5 phosphocreatine, 0.3 GTP. Baclofen (R-(+)-b-(aminomethyl)-4-chlorobenzenepropanoic acid hydrochloride) was purchased from Sigma (St. Louis, MO). Baclofen-induced currents were measured at room temperature using a high-K⁺ bath solution (in mM): 120 NaCl, 25 KCl, 1.8 CaCl₂, 1 MgCl₂, 5.5 D-glucose, 5 HEPES/NaOH (pH 7.4). The high-K⁺ bath solution (+/− baclofen) was applied directly to the soma and proximal dendrites with an SF-77B rapid perfusion system (Warner Instruments, Inc.; Hamden, CT).

Current responses to the application of the high-K⁺ solution (+/− baclofen) were measured at a holding potential of −80 mV. Membrane potentials and whole-cell currents were measured in large neurons (>75 pF) with hardware (Axopatch-700B amplifier, Digidata 1440A) and software (pCLAMP v. 10.3) from Molecular Devices (Sunnyvale, CA). All currents were low-pass filtered at 2 kHz, sampled at 5 kHz, and stored on computer hard disk for subsequent analysis. Peak and steady-state current amplitudes were measured for each experiment. Current activation rates were extracted from a standard exponential fit of the current trace corresponding to the onset of drug effect and the peak evoked current, while deactivation rates were extracted from an exponential fit of the trace corresponding to the return of current to baseline following removal of drug (Clampfit v. 10.3 software). Current desensitization was defined as % change in steady state current from the maximal baclofen-evoked response amplitude during 10 s of continuous drug application. Only experiments where access resistances were stable and low (<20 MΩ) were included in the analysis.

Immunohistochemical reactions were carried out using the pre-embedding immunogold method as described earlier (Lujan et al., 1996). Briefly, after blocking with 10% serum for 1 hr at room temperature free-floating sections were incubated for 48 hr with anti-RGS7 antibodies (1–2 mg/ml). Sections were washed and incubated for 3 hr with goat anti-rabbit IgG coupled to 1.4 nm gold (Nanoprobes Inc) at 1:100 dilution. Sections were washed, postfixed in 1% glutaraldehyde and processed for silver enhancement of the gold particles with an HQ Silver kit (Nanoprobes Inc.). The reacted sections were treated with osmium tetraoxide (1% in 0.1 M PB), block-stained with uranyl acetate, dehydrated in graded series of ethanol and flat-embedded on glass slides in Durcupan (Fluka) resin. Regions of interest were cut at 70–90 nm on an ultramicrotome (Reichert Ultracut E; Leica). Staining was performed on drops of 1% aqueous uranyl acetate followed by Reynolds’s lead citrate. Ultrastructural analyses were performed in a Jeol-1010 electron microscope.

To establish the relative the abundance of RGS7 immunoreactivity along the plasma membrane of pyramidal cells, we used 60-µm coronal slices processed for pre-embedding immunogold immunohistochemistry. The procedure was similar to that used previously (Lujan et al., 1996). Briefly, for each of three animals from different postnatal ages and adult, three samples of tissue were obtained for preparation of embedding blocks (totalling nine blocks for each age). To minimize false negatives, electron microscopic serial ultrathin sections were cut close to the surface of each block, as immunoreactivity decreased with depth. We estimated the quality of immunolabelling by always selecting areas with optimal gold labelling at approximately the same distance from the cutting surface. Randomly selected areas were then photographed from the selected ultrathin sections and printed with a final magnification of 45 000X. Quantification of immunogold labelling was carried out in reference areas totalling approx. 1,800 µm² for each age. Immunoparticles identified in each reference area and present in different subcellular compartments (dendritic spines, dendritic shafts and somata) were counted. We measured the radial distance of each immunoparticle to the plasma membrane, being 0 for those just located in the plasma membrane. The data was expressed as percentage of immunoparticles along the radial distance from the plasma membrane expressed in nanometers.

Mice were sacrificed under isoflurane anesthesia, and brains were rapidly removed and placed in ice-cold artificial cerebrospinal fluid (aCSF) without CaCl₂, composed of (mM): 124 NaCl, 3 KCl, 24 NaHCO₃, 1.25 NaH₂PO₄, 1 MgSO₄, and 10 D-Glucose, equilibrated with 95% O₂ and 5% CO₂. The tissue was cut in 350–400 µm thick sections with a Vibrating microtome (Leica VT1200S, Germany). The slices were warmed to 35°C for 25–45 min in aCSF supplemented with 2 mM CaCl₂, and equilibrated with 95% O₂ and 5% CO₂. Then slices were maintained in gassed aCSF at room temperature until being transferred to submerged-type recording chambers of volume ∼1.5 ml. Here, the slices were constantly superfused (1–2 ml/min) with warmed (30–31°C), gassed aCSF. All measurements were performed by an experimenter blind to genotype.

CA1 neurons were visually identified in the hippocampal transverse slices of 350 µM thickness using Scientifica SliceScope system. Glass microelectrodes with an open-tip resistance of 3.5–6.5 MΩ were used. The following internal solution was used (mM): 130 K-Gluconate, 20 KCl, 10 K-HEPES, 0.2 EGTA, 0.3 Na-GTP and 4 Mg-ATP (pH 7.3). To determine intrinsic cellular properties such as resting membrane potential, input resistance and spike numbers, 500 ms, 50 pA, 10-step hyperpolarizing and depolarizing current injections were delivered every 10 s. Cells with series resistance >20 MΩ or resting membrane potentials > −55 mV were excluded from analysis. Liquid junction potential was −14 mV. Spontaneous EPSCs (sEPSCs) were measured by holding the cells at −70 mV in normal aCSF. At least 100 events, which are above 4 pA, were obtained during sEPSC measurements in each cell.

Field excitatory postsynaptic potentials (fEPSPs) were elicited by a concentric bipolar stimulating electrode (inner diameter [ID]: 25 µm; outer diameter [OD]: 125 µm, FHC Inc., Bowdoin, ME) connected to a constant current isolated stimulator unit (A-M Systems; Carlsborg, WA) and recorded with low resistance (3–5 MΩ) glass pipettes (ID: 1.16 mm, OD: 1.5 mm, Harvard Apparatus, Holliston, MA) filled with aCSF. The electrodes were placed in the stratum radiatum of the CA1 area of the dorsal hippocampus slices. Stimulation frequency was set to 0.05 Hz. Input-output curves were generated by adjusting the stimulus intensity in increments of 10 µA, from 0 to 100 µA. Paired pulse ratio (PPR) was assessed using a succession of paired pulses separated by time intervals ranging from 25 to 1000 ms, delivered every 20 s. The degree of facilitation was determined by taking the ratio of the initial slope of the second fEPSP to the initial slope of the first fEPSP. A PPR >1 was considered to reflect facilitation. For synaptic plasticity experiments, a stable baseline for at least 30 min was achieved prior to induction. LTP and LTD were recorded for 1 hr after HFS (2 tetanized stimuli (TS) of 100 Hz for 1s each) or LFS (2 Hz for 10 min, 1200 pulses) was applied. Depotentiation was achieved by applying HFS followed by LFS after 1-2 min interval.

Spatial learning and memory were evaluated in the Morris water maze using a video tracking systems (EthoVision XT, Noldus Information Technology, Wageningen, Netherlands). Mice first received 4 cued trials (visible but variable platform location) on the first day to determine if non-associative impairment in other behavioral responses affecting performance in this task, such as exploratory activity, motor coordination, vision and motivation. After completing the cued trials, spatial learning acquisition was evaluated during the place condition (hidden platform, constant location; 4 trials/day, 6 consecutive days). Escape latency and success rates in finding hidden platform were calculated for all place trials. Retention performance was evaluated during probe trials (platform removed), which were conducted 24 hr after the last cued trial on the sixth day. Latency to the first target platform area crossing and the number of crossings in the probe trial (1 min) served as the dependent variables.

For fear conditioning experiments, mice were habituated in individual conditioning chambers to obtain freezing baseline on the first day. On the second day, mice were trained in individual conditioning chambers (Med Associates, St. Albans, VT). Video images were recorded via video tracking systems (EthoVision XT, Noldus Information Technology, Wageningen, Netherlands). In context A, visible light was turned on, a stainless steel grid floor inserted, and the chambers were cleaned with 70% ethanol prior to conditioning. In context B, white plastic inserts were placed inside the chamber to change the shape, size and texture of the wall and the floor. A small weight boat with orange extract is placed behind the wall inserts to provide a novel smell for the chambers. The chambers were cleaned with 70% isopropanol. During the training sessions, mice were placed in context A and allowed to explore for 3 min prior to the delivery shocks (0.75 mA, 1 s). We used auditory tone (85 dB, 30 s duration) as CS and an electric foot shock (delivered through the grid floor, 0.75 mA AC current, 1 s duration) as US. Mice received 3 tone-shock stimuli (with 1–2 min interval) during the training day. In every session, tone always co-terminated with electric shock. Mice were removed from the conditioning chamber 30 s after the last shock. Contextual test (5 min) was done in context A 24 hr after training. Cue test was performed 1 hr after contextual test in context B. The animals were brought into the test room in a different covered mover with different bedding inside to avoid contextual reminders. The animal was immediately placed in the contextual B chamber and allowed to explore for 3 min prior to the delivery the auditory tone (85 dB) for another 3 min. The freezing response was measured with the automated tracking and analyzed offline.

For the novel object recognition test, mice were habituated to the arena for two consecutive days, where they were placed in a white plastic open-box of dimensions 43.2 cm2 by 30.5 cm height for 10 min each. The acquisition phase of the object recognition assay involved placing each individual mouse in the test arena for 8 min in the presence of two identical objects (either silver or black objects in different shape, size and texture, the order of which alternated between mice). The video was recorded via a camera. The time spent investigating both identical objects was recorded by automatic tracking system (EthoVision XT, Noldus Information Technology, Wageningen, Netherlands). 24 hr after the acquisition phase, each mouse was reintroduced to the test arena for 8 min. During this phase (retention testing phase) the mouse was presented with one copy of the object that it was exposed to during the acquisition phase (familiar object) and one novel object (either silver or black objects, depending on which was presented during the acquisition phase). The time spent investigating each object was recorded. All objects and the testing box were thoroughly cleaned with 70% methanol between mice to remove any odor cues.

Agonist-dependent cellular measurements of bioluminescence resonance energy transfer (BRET) between Venus-Gβ1γ2 and its effector fragment masGRK3ct-Nluc were performed upon reconstitution in living cells as previously described with slight modification (Hollins et al., 2009; Masuho et al., 2013). The masGRK3ct-Nluc construct contained amino acids G495-L688 of bovine GRK3 (NP_776925), preceded by a myristic acid attachment peptide (mas; MGSSKSKTSNS). The stop codon of GRK3 was replaced with a GGGS linker, which was followed by the NanoLuc (Nluc) (Hall et al., 2012). Briefly, GABA_B1R, GABA_B2R, GαoA, Venus156-239-Gβ1, and Venus1-155-Gγ2 constructs were transfected into HEK293T/17 cells at a 1:1:2:1:1 ratio with increasing masGRK3ct-Nluc from 0.125 to 6 in ratio. 5 µg total DNA was delivered per 4 × 10⁶ cells in a 6-cm-dish. 16–24 hr post transfection cells were stimulated with 100 µM GABA followed by treatment with 100 µM CGP 54626. The BRET signal is determined by calculating the ration of the light emitted by the Venus-Gβ1γ2 (535 nm) over the light emitted by the masGRKct-Rluc8 (475 nm). The average baseline value recorded prior to agonist stimulation was subtracted from BRET signal values, and the resulting difference (ΔBRET) was obtained.

Statistical analyses were performed using Prism (GraphPad Software, Inc.; La Jolla, CA). Data are presented throughout as the mean ± SEM. Student t test, one-way or two-way ANOVA, followed by Bonferroni’s post hoc test were used as appropriate. The minimal level of significance was set at p<0.05.

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

We thank Dr Orlandi for his help with preparing samples for the EM studies, Drs Clement (Chelliah), Kourrich and Ozkan for technical advice and discussions and Mrs Martemyanova for help with animal breeding and genotyping. This work was supported by NIH grants DA021743 (KAM), DA026405 (KAM), MH061933 (KW), DA034696 (KW), HL105550 (KW and KAM) and by grants from the Spanish Ministry of Education and Science (BFU-2012-38348) and CONSOLIDER (CSD2008-00005) to RL.

Animal experimentation: This study was performed in strict accordance with the recommendations in the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. All of the animals were handled according to approved institutional animal care and use committee (IACUC) protocols (#08-133) of the Scripps Research Institute.

© 2014, Ostrovskaya et al.

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