To investigate physiological functions of RNG105 in adult mice, we generated RNG105 conditional deletion mice by crossing floxed Rng105 mice and Camk2a-Cre transgenic mice for gene deletion in the central nervous system (Figure 1A–C). In Camk2a-Cre;Rng105^f/f mice, exons 5–6 were flanked by loxP sequences and deleted by Cre expression (Figure 1A). Because frame shift occurs in the exon 5–6-deleted mRNA, most of the functional domains of RNG105 ranging from the N-terminal coiled-coil domain to the C-terminal RG-rich domain (a.a. 123–707) are deleted (Shiina et al., 2005). In addition, most of the exon 5–6-deleted mRNA appeared to be degraded by nonsense-mediated mRNA decay (NMD): in the hippocampus where Cre was highly expressed, expression of Rng105 transcripts from all exons was reduced to the comparable level to that of exon 5–6 transcripts as judged by RNA-seq analysis (Figure 1—figure supplement 1A).

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Camk2a-Cre;Rng105^f/f mice were born in the Mendelian ratio and showed no apparent abnormalities just after birth. Although their growth was retarded during the lactation period, their body weight recovered thereafter to 80–90% of that of control (Rng105^f/f) mice (Figure 1D). In addition to growth retardation, Camk2a-Cre;Rng105^f/f mice were susceptible to death. However, death was spontaneous, and more than 40% of Camk2a-Cre;Rng105^f/f mice survived for more than 4 months (Figure 1E). Thus, RNG105 conditional deletion overcame the neonatal lethality of RNG105 conventional KO.

Western blotting revealed that the expression of RNG105 protein was markedly reduced in the cerebral cortex and hippocampus, but not so much in the cerebellum of Camk2a-Cre;Rng105^f/f mice (Figure 1F). Although the anti-RNG105 antibody can recognize truncated RNG105 protein (a.a. 1–122) encoded by the exon 5–6-deleted mRNA, it did not detect any truncated form of RNG105 in the cerebrum of Camk2a-Cre;Rng105^f/f mice (Figure 1—figure supplement 1B). This supported the notion that the exon 5–6-deleted mRNAs were degraded by NMD and hardly any truncated RNG105 protein was expressed in Camk2a-Cre;Rng105^f/f mice. Immunostaining of hippocampal slices showed that RNG105 was markedly reduced in the somatic layer (stratum pyramidale [SP]) and dendritic layer (stratum radiatum [SR]) of pyramidal neurons, confirming the reduction of RNG105 expression in neurons of Camk2a-Cre;Rng105^f/f mice (Figure 1G).

To investigate the impact of RNG105 deficiency on synaptic function, we first examined the morphology of pyramidal neurons and dendritic spines in the hippocampal CA1 region. The density of hippocampal neurons, as judged from nuclear staining, was not affected in Camk2a-Cre;Rng105^f/f mice (Figures 1G and 2A). To trace the morphology of pyramidal neurons, Camk2a-Cre;Rng105^f/f mice were crossed with Thy1-GFP transgenic mice. Fluorescence imaging revealed that the length and branching of dendrites were comparable between Rng105^f/f and Camk2a-Cre;Rng105^f/f mice (Figure 2B−D). The density of spines on dendrites was also equivalent between the genotypes, but the size of spines was smaller in Camk2a-Cre;Rng105^f/f mice (Figure 2E−G). Furthermore, the number of mushroom spines was significantly reduced, which suggested that synaptic strength and/or stimulation-dependent plasticity were attenuated in Camk2a-Cre;Rng105^f/f mice (Figure 2E,H).

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We then analyzed structural plasticity of dendritic spines using a two-photon glutamate uncaging technique. Hippocampal neurons from floxed RNG105 (Rng105^f/f) mice were cultured in dishes and co-transfected with CMV-Cre and mCherry in order to delete the Rng105 gene and trace the morphology of the neurons, respectively. Immunostaining of the neurons indicated that the expression of RNG105 was significantly reduced in mCherry-positive neurons compared with neighboring mCherry-negative neurons (Figure 3A,B). RNG105 deletion did not affect spine density on dendrites, but reduced the size of spines (Figure 3C,D), which was similar to the results in the hippocampal slices.

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In response to stimulation by glutamate uncaging, single spines close to the uncaging spots were transiently increased in their volume to ~3 fold and sustained an increased state of ~2.5 fold over 60 min in Rng105^f/f neurons (Figure 3E,F). The increase in spine volume during the sustained phase is translation-dependent (Nishiyama and Yasuda, 2015; Tanaka et al., 2008), which was confirmed by cycloheximide addition (Figure 3F,G). In contrast to Rng105^f/f neurons, CMV-Cre;Rng105^f/f neurons significantly reduced the spine volume during the sustained phase (Figure 3E−G). These results indicated that RNG105 was required for the stimulation-induced structural plasticity of spines in the translation-dependent late-phase LTP.

To examine whether RNG105 regulates the function of synapses, we measured electrophysiological responses of hippocampal CA1 neurons to stimulation. First, basal synaptic transmission at CA3-CA1 synapses was recorded. The relationship between the magnitude of input stimulation and the postsynaptic responses (amplitude and slope of field excitatory postsynaptic potential [fEPSP]) indicated that RNG105 deficiency reduced both the amplitude and slope of fEPSP to about half of those of control mice (Figure 4A,B). Paired-pulse ratio (PPR) was significantly decreased after LTP induction in Camk2a-Cre;Rng105^f/f mice, but it was not significantly different between the genotypes (Figure 4C,D). These results indicated that postsynaptic responses to stimulation were markedly reduced in Camk2a-Cre;Rng105^f/f mice, suggesting downregulation of AMPARs by RNG105 deficiency.

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We also measured LTP at CA3-CA1 synapses. Theta-burst-induced LTP in CA1 neurons revealed that fEPSP was increased from baseline by ~100%, which was comparable between Rng105^f/f and Camk2a-Cre;Rng105^f/f mice (Figure 4E,F). However, because the absolute amplitude and slope of basal fEPSP in Camk2a-Cre;Rng105^f/f mice were about half of those in Rng105^f/f mice, fEPSP after LTP induction in Camk2a-Cre;Rng105^f/f mice was also about half of that in Rng105^f/f mice (Figure 4E,F). As a result, the absolute amplitude and slope of fEPSP in Camk2a-Cre;Rng105^f/f mice reached, even after LTP induction, only the same level as the basal fEPSP in Rng105^f/f mice (Figure 4E,F). After the induction of LTP, we often observed epileptic-like repetitive patterns in fEPSP waveforms, which may be a back-propagation of repetitive action potentials, suggesting aberrant neuronal excitation after LTP induction in Camk2a-Cre;Rng105^f/f mice (Figure 4C, inset). Together, in Camk2a-Cre;Rng105^f/f mice, fEPSP amplitude was reduced both in the steady state and after LTP induction, which may be related to abnormal neuronal excitation and impaired structural plasticity of spines.

Next, we examined whether RNG105 is required for normal behavior in learning and memory tasks. In open-field test, there were no differences in exploratory horizontal locomotion among Rng105^f/f, RNG105 hetero-deletion (Camk2a-Cre;Rng105^f/+) and Camk2a-Cre;Rng105^f/f mice in the initial trial (Figure 5A). The exploratory activity of Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice was decreased over 3 days, suggesting that the mice were habituated to the novel place with trials. In contrast, Camk2a-Cre;Rng105^f/f mice did not show such experience-dependent reduction in exploratory activity, suggesting that Camk2a-Cre;Rng105^f/f mice had problems in becoming habituated to a novel environment (Figure 5A).

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Habituation is a complex behavior, which is impaired by several factors such as memory deficits, incomplete initial exploration of the entire area because of high level of anxiety, and low level of locomotor activity (Bolivar, 2009). Among them, the latter two factors were not likely reasons for the impaired habituation in Camk2a-Cre;Rng105^f/f mice, because the initial exploratory activity in the open field test was normal and anxiety-like behavior in the light/dark transition test was also normal in Camk2a-Cre;Rng105^f/f mice (Figure 5A; Figure 5—figure supplement 1A).

We further conducted a novel object recognition test, which is another test to assess habituation (Figure 5—figure supplement 1B). In the first session, mice were habituated to two identical objects and showed no biased preference for either object. In the second session in which one of the objects was replaced by a novel one, Rng105^f/f mice showed an increased preference for the novel object. In contrast, Camk2a-Cre;Rng105^f/f mice did not show such an increased preference for the novel object (Figure 5—figure supplement 1B). These results supported the notion that habituation, a form of learning and memory, was impaired in Camk2a-Cre;Rng105^f/f mice.

In the rotarod test, Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice showed increasing retention time on the rod and decreasing number of falls from the rod over 3 days, indicating that the mice learned the rotarod skill day by day (Figure 5B). Camk2a-Cre;Rng105^f/f mice showed indistinguishable performance from the other genotypes, indicating that RNG105 conditional deletion did not affect motor skill learning. This was consistent with a mild reduction in RNG105 proteins in the cerebellum of Camk2a-Cre;Rng105^f/f mice, as well as a notion that RNG105 may be simply not required for this cerebellum-dependent form of learning and memory.

To test spatial learning and memory, Morris water maze was conducted. Mice were first subjected to a visible platform test. Escape latency of Camk2a-Cre;Rng105^f/f mice was longer than that of Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice, but significantly reduced over trials (Figure 6A). These results indicated that Camk2a-Cre;Rng105^f/f mice required more training than the other genotypes, but they had escape motivation, vision, and motor skills sufficient to accomplish the task.

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The mice were then subjected to a hidden platform test, which necessitates hippocampus-dependent long-term spatial memory (Figure 6B). Repeated trials enabled Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice to learn the platform location and escape on the platform faster than before the trials. In contrast, the escape latency of Camk2a-Cre;Rng105^f/f mice did not shorten at all over the trials (Figure 6B). Following the last trial, a probe test was conducted without the platform. Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice intensively searched around the target place (Figure 6C). In contrast, Camk2a-Cre;Rng105^f/f mice showed a circular swimming path along the wall and reduced the search time around the target place (Figure 6C). The time that Rng105^f/f and Camk2a-Cre;Rng105^f/+ mice spent in the target quadrant was significantly longer than in the other quadrants, whereas Camk2a-Cre;Rng105^f/f mice markedly reduced the time in the target quadrant compared to the other genotypes (Figure 6D). These severe phenotypes raised a concern that Camk2a-Cre;Rng105^f/f mice might be incapable of learning the task. However, if outliers were eliminated (two Camk2a-Cre;Rng105^f/f mice showing maximum values in L and O quadrants [Figure 6D], p<0.01 using Smirnov-Grubbs test), statistical significance was detected in the quadrant effect in Camk2a-Cre;Rng105^f/f mice (one-way repeated measures ANOVA, F[3,45]=3.234, p=0.0309). Consistently, the density plot of swim path showed weak preference of Camk2a-Cre;Rng105^f/f mice for the target quadrant (Figure 6C), suggesting Camk2a-Cre;Rng105^f/f mice were able to learn the task. Together, these results indicated that RNG105 was critical for the formation of spatial long-term memory.

Finally, a contextual fear conditioning test (passive avoidance) was conducted. Before receiving foot shock in a dark chamber, all genotypes spent 60–70% of the test time in the dark chamber (Figure 6E). At 5 min after the foot shock, none of the genotypes stayed in the dark chamber, indicating that fear-conditioned learning had took place. However, at 1 day after the foot shock, Camk2a-Cre;Rng105^f/f mice spent a significantly longer time in the dark chamber than the other genotypes. Furthermore, at 1 week after the foot shock, Camk2a-Cre;Rng105^f/f mice spent as long a time in the dark chamber as before the foot shock (Figure 6E). Because Camk2a-Cre;Rng105^f/f mice spent a comparable time to the other genotypes in the dark chamber before the foot shock, as well as showing normal anxiety-like behavior in the light/dark transition test (Figure 5—figure supplement 1A), the increased time in the dark chamber at 1 day and 1 week was considered not to be due to increased anxiety, but reduced long-term memory in Camk2a-Cre;Rng105^f/f mice. In another contextual fear conditioning test, in which mice received foot shock in a single chamber and their freezing responses were measured in the same chamber at 5 days after the foot shock, Camk2a-Cre;Rng105^f/f mice showed less freezing behavior than Rng105^f/f mice (Figure 6—figure supplement 1). These results indicated that RNG105 was required for long-term fear conditioning memory formation.

Besides the impairment in memory formation, Camk2a-Cre;Rng105^f/f mice sometimes exhibited seizures during and just after the Morris water maze and contextual fear conditioning tests (Video 1). This phenotype was reminiscent of the epileptic-like fEPSP appeared after relatively intense stimulation of LTP (Figure 4C). Taken together, behavioral analyses demonstrated an essential role of RNG105 in the formation of long-term memory.

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To understand the underlying mechanisms by which RNG105 regulates synaptic strength, synaptic structural plasticity and long-term memory formation, we investigated the impact of RNG105 deficiency on mRNA localization in neurons. Although RNG105 is involved in mRNA transport to dendrites in vitro (Shiina et al., 2010), it is unknown whether RNG105 regulates somato-dendritic mRNA localization in vivo, and if so, which mRNAs change the dendritic localization in RNG105-deficient mice.

First, we extracted somatic and dendritic mRNAs from the hippocampal CA1 region in adult mice. The somas of the CA1 pyramidal neurons are aligned in the SP, from which apical dendrites elongate in the SR (Figures 1G and 7A). We microdissected and isolated each layer for the preparation of soma and dendrites of pyramidal neurons (Figure 7B,C), as demonstrated in previous studies (Cajigas et al., 2012; Ainsley et al., 2014). Next, mRNAs extracted from SP and SR of Rng105^f/f mice and Camk2a-Cre;Rng105^f/f mice were subjected to RNA-seq analysis (Supplementary file 1). For each mRNA, relative concentration (FPKM: read counts normalized by transcript length) in SR to SP, which we termed the ‘dendritic accumulation index (DAI)", was calculated. Statistical analysis identified SP-enriched (low DAI) mRNAs and SR-enriched (high DAI) mRNAs. Because the hippocampus contains not only pyramidal neurons but also other types of cells such as interneurons and glial cells, mRNAs also expressed in these cell types were eliminated from the mRNA lists as in the previous study (Cajigas et al., 2012) (Supplementary file 2A,B). As a result, we identified 1122 dendritically enriched mRNAs (D-mRNAs) and 2106 somatically enriched mRNAs (S-mRNAs) in control mice (Figure 7D; Supplementary file 3A−C). The D-mRNAs included already known dendritic mRNAs such as Camk2a, Eef1a1, Dlg4, Iptr1, Arc, Shank2, Homer2, and Limk1 (Figure 7D; Supplementary file 3A), suggesting that the strategy was appropriate to detect somato-dendritic mRNA distribution pattern.

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Comparison between Rng105^f/f and Camk2a-Cre;Rng105^f/f mice revealed that the somato-dendritic distribution pattern of mRNAs was different between the genotypes; the variance of DAI of mRNAs was smaller in Camk2a-Cre;Rng105^f/f mice than in Rng105^f/f mice, and the gap in DAI between D- and S-mRNAs was narrower in Camk2a-Cre;Rng105^f/f mice (Figure 8A). Variance values (s²) of the DAI of D- and S-mRNAs were 1.945 for Rng105^f/f mice and 1.092 for Camk2a-Cre;Rng105^f/f mice, which were significantly different (n = 3,228, F₀ = 1.781, p<0.005). These results indicated an in vivo role of RNG105 to establish the asymmetric localization of mRNAs in the soma and dendrites.

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We further analyzed whether changes in the somato-dendritic localization of mRNAs in Camk2a-Cre;Rng105^f/f mice were mRNA-selective. For each mRNA, the ratio of DAI in Camk2a-Cre;Rng105^f/f mice to Rng105^f/f mice was calculated (Figure 8B). The ratio was lower than one for majority of, but not all, the D-mRNAs, indicating that RNG105 deficiency reduced the dendritic localization of many, but specific D-mRNAs. To further address mRNA-selectivity, relation between DAI (Rng105^f/f) and the ratio of DAI (Camk2a-Cre;Rng105^f/f/Rng105^f/f) for D- and S-mRNAs was analyzed (Figure 8C). D-mRNAs showed large variance and low correlation (|r| = 0.3455). In contrast, S-mRNAs showed smaller variance and high correlation (|r| = 0.7269) (Figure 8C). These results indicated that the effect of RNG105 deficiency on dendritic localization of mRNAs varied among D-mRNAs, suggesting RNG105 targets selective D-mRNAs. On the other hand, the relatively uniform effect of RNG105 deficiency on S-mRNAs indicated that the effect was mRNA-non-selective. The apparent increase in DAI of S-mRNAs in Camk2a-Cre;Rng105^f/f mice may be mainly a secondary effect of the decrease in DAI of D-mRNAs because the sum of total mRNAs' log(DAI) should be nearly zero.

To identify the biological categories in which the D-mRNAs are involved, gene ontology (GO) enrichment analysis was conducted. First, D- and S-mRNAs in control mice were classified into GO categories (Figure 8D; Supplementary file 4A−I). Major categories of D-mRNAs were ‘regulation of Arf protein signal transduction’, which included GTPase-activating proteins (GAPs) and guanine nucleotide exchange factors (GEFs) of small G protein ADP-ribosylation factor (Arf), and ‘structural constituent of ribosomes’ which included ribosomal subunit proteins (Figure 8D; Supplementary file 4A,B). D-mRNAs included in these categories were plotted on MA plots, which indicated that the dendritic accumulation of these mRNAs was comparable to that of well-known dendritic mRNAs (Figure 9, cf. Figure 7D). Arf is known to regulate membrane trafficking between the cell surface and endosomes, and in particular, Arf6 participates in the surface expression of AMPARs (D'Souza-Schorey and Chavrier, 2006; Jaworski, 2007; Oku and Huganir, 2013; Zheng et al., 2015). Arf is also known as a regulator of actin dynamics and dendritic spine formation via Rac1 activation (D'Souza-Schorey and Chavrier, 2006; Jaworski, 2007). There are various Arf GAPs and GEFs possessing or not possessing a membrane-associated pleckstrin-homology (PH) domain, among which the Arf GAPs and GEFs identified here were mostly the PH domain-possessing types and classified in ‘pleckstrin homology’ category with other PH domain-containing proteins (Figure 8D; Supplementary file 4E). The Arf GAPs and GEFs were also classified in ‘GTPase regulator activity’, which included regulators of other small G proteins such as Ras, Rho, and Rac (Figure 8D; Supplementary file 4G), known to be involved in actin reorganization and spine morphogenesis (Nishiyama and Yasuda, 2015). ‘Fc gamma receptor-mediated phagocytosis’, ‘leukocyte activation’, and ‘chemotaxis’ categories also had high-fold enrichment scores, which contained several overlapping proteins such as PI3 kinase pathway proteins and Rac pathway proteins involved in actin regulation, and extracellular membrane proteins (Figure 8D; Supplementary file 4D,F,H).

Figure 9

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GO enrichment analysis was further conducted on D-mRNAs whose dendritic localization was reduced in Camk2a-Cre;Rng105^f/f mice compared with Rng105^f/f mice (the ratio of DAI < 0.8). Major categories with high-fold enrichment scores were the same as above, for example, ‘regulation of Arf protein signal transduction’, ‘structural constituent of ribosomes’, and ‘GTPase regulator activity’, suggesting that RNG105 was responsible for the dendritic localization of mRNAs classified in the major categories (Figures 8E and 9; Supplementary file 4A−I).

Reduction in the DAI of Arf regulator mRNAs in Camk2a-Cre;Rng105^f/f mice was reminiscent of the results that fEPSP, which is mediated by AMPARs, was reduced in Camk2a-Cre;Rng105^f/f mice (Figure 4). We then examined whether the DAIs of mRNAs encoding other regulatory proteins of AMPARs (Bassani et al., 2013; Henley and Wilkinson, 2013) were also reduced in Camk2a-Cre;Rng105^f/f mice. mRNAs such as Cnih2, Arc, Dlg4, Pik3r2, Camk2a, and Cplx2 were identified as D-mRNAs. Furthermore, their DAIs were reduced in Camk2a-Cre;Rng105^f/f mice compared with Rng105^f/f mice (Supplementary file 5A). These results indicated that various mRNAs, encoding AMPAR regulators involved in AMPAR surface expression and retention to postsynapses, were localized to dendrites in an RNG105-dependent manner.

By contrast, mRNAs encoding AMPAR subunits themselves (Gria1-4) were somatically enriched and their DAIs were not reduced in Camk2a-Cre;Rng105^f/f mice (Supplementary file 5A). Notably, the DAI of Gria2 mRNA was markedly increased, rather than decreased, in Camk2a-Cre;Rng105^f/f mice (Supplementary file 5A), suggesting that RNG105 influences specifically, if not directly, the dendritic localization of Gria2 mRNA.

RNG105 deficiency also influenced, if not directly, the total expression level of some mRNAs, as judged from the S-mRNA concentration (FPKM). mRNAs whose expression was reduced in Camk2a-Cre;Rng105^f/f mice included Rng105 itself, and notably, a considerable number of immediate early genes (IEGs) such as Fos, Btg2, Egr1, Egr4, Dusp1, and Arc (Supplementary file 6). Because the expression of these IEGs was reportedly upregulated by neuronal activation (Saha et al., 2011; Iacono et al., 2013), these results suggested a reduction in neuronal activity by RNG105 deficiency. In addition, the drastic increase in Gria2 mRNA localization to dendrites in Camk2a-Cre;Rng105^f/f mice may be also attributed to reduced neuronal activity in RNG105-deficient mice (Grooms et al., 2006).

The reduction in fEPSP amplitude and dendritic localization of mRNAs for AMPAR regulators suggested that RNG105 regulates AMPAR scaling. In particular, given the reduction in steady-state fEPSP, we hypothesized that RNG105 may be important for homeostatic scaling of AMPARs. To test this, we analyzed GluR1 and GluR2 cell surface expression in response to activity deprivation with TTX and APV in primary cultured neurons from wild-type (Rng105^+/+) and RNG105 knockout (Rng105^−/−) mice. Surface and total GluR1 and GluR2 were immunostained before and after cell permeabilization, and quantified by counting the number and measuring the fluorescence intensity of their puncta in dendrites (Figure 10). After activity deprivation, the number of surface GluR1 puncta normalized to that of total GluR1 was significantly increased, and the intensity of surface GluR1 tended to be increased, in dendrites of Rng105^+/+ neurons. By contrast, the activity deprivation-dependent increase in surface GluR1 expression was not observed in Rng105^−/− dendrites (Figure 10A,C). Although homeostatic scaling of GluR2 is controversial (Isaac et al., 2007; Wierenga et al., 2005; Gainey et al., 2009), our results indicated that the number of GluR2 surface puncta was increased by activity deprivation in dendrites of Rng105^+/+ neurons (Figure 10B,D). By contrast, this increase in surface GluR2 puncta was not observed in Rng105^−/− neurons, similarly to GluR1.

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Comparison of Rng105^+/+ and Rng105^−/− neurons indicated that the number of surface GluR1 in dendrites was significantly reduced in Rng105^−/− neurons (Figure 10A,C), confirming the previous results (Ohashi et al., 2016). By contrast, GluR2 surface expression in dendrites was not much reduced by RNG105 deficiency (Figure 10B,D), which could be due to the increase in Gria2 mRNA localization to dendrites by RNG105 deficiency (Supplementary file 5A) and/or different surface expression pathways of a fraction of GluR2 from that of GluR1 (Tanaka and Hirano, 2012).

We further conducted biotin labeling of cell surface proteins of cultured neurons followed by immunoblot measurement for GluR1 and GluR2 (Figure 10—figure supplement 1). GluR1/2 in total cell lysates was detected as a major band of ~100 kDa, which was reduced in amount after biotin labeling, accompanied by an increase in the amount of an upper band (Figure 10—figure supplement 1A). The upper band, but not the lower band, bound to avidin agarose beads, indicating that the upper band was biotin-labeled and mobility-shifted surface GluR1/2, whereas the lower band was non-labeled intracellular GluR1/2 (Figure 10—figure supplement 1A−C). Because the upper band was also detected in control lysates without biotinylation, the band was considered to contain a non-specific protein(s) as well as biotinylated GluR1/2.

The most obvious difference between TTX/APV-treated and untreated neurons was the less amount of the lower GluR1 band in TTX/APV-treated neurons from Rng105^+/+ mice, but not from Rng105^−/− mice, which suggested that TTX/APV treatment reduced intracellular GluR1 in Rng105^+/+ neurons, but not in Rng105^−/− neurons (Figure 10—figure supplement 1B). Then we quantified the intensity of the lower band in the cell lysate and the upper band in the avidin agarose-bound fraction, and calculated the ratio of surface to intracellular GluR1 and GluR2 (Figure 10—figure supplement 1D,E). The ratio of surface/intracellular GluR1 was increased in TTX/APV-treated neurons compared to untreated neurons from Rng105^+/+ mice. Compared to Rng105^+/+ neurons, the ratio of surface/intracellular GluR1 was lower and was not significantly increased by TTX/APV treatment in Rng105^−/− neurons (Figure 10—figure supplement 1D). We noted that the GluR1 band intensity was lower both in the total lysates and avidin-bound fractions in Rng105^−/− neurons than in Rng105^+/+ neurons (Figure 10—figure supplement 1B, upper panel), which may be because of the reduced density of neural networks in Rng105^−/− primary cultured neurons (Shiina et al., 2010). This weaker band intensity in Rng105^−/− neurons made it difficult to identify differences in the surface/intracellular ratio between the genotypes at a glance, but when twice the volume of samples from Rng105^−/− neurons was loaded on gels, the difference in the surface/intracellular ratio between the genotypes was obvious: the intensity of the lower GluR1 bands was higher, whereas that of the upper GluR1 bands was lower, in Rng105^−/− neurons than in Rng105^+/+ neurons (Figure 10—figure supplement 1B, bottom panel). As for GluR2, although statistical significance was not detected, the surface/intracellular ratio of GluR2 tended to be increased by TTX/APV treatment (Figure 10—figure supplement 1E). These results were consistent with the results of the immunofluorescence imaging of GluR1 and GluR2. Taken together, these results indicated that RNG105 impacts on homeostatic scaling of AMPARs in dendrites, which is coupled to basal synaptic function and also influences synaptic strength in activated state.

All animal care, experiments and behavioral testing procedures were approved by the Institutional Animal Care and Use Committee of the National Institutes of Natural Sciences, and performed in accordance with the guidelines from the National Institutes of Natural Sciences, Niigata University and the Science Council of Japan.

To generate a loxP-flanked (floxed) Rng105 construct, we isolated the Rng105/caprin1 gene by PCR from a genomic DNA library of C57BL/6 mice. A DNA fragment, which carried a 34 bp loxP sequence and a neomycin resistance gene (Neo) flanked by two frt sites, was inserted into the site 227 bp upstream of exon 5. The other loxP site was introduced into the site 282 bp downstream of exon 6. The targeting vector contained exons 5 and 6 of the Rng105 gene flanked by loxP sequences, 7.33 kb upstream and 5.8 kb downstream homologous genomic DNA fragments, and the diphtheria toxin (DT) gene for negative selection (Figure 1A). The targeting vector was introduced into C57BL/6 ES cells (RENKA), and homologous recombinants were selected by G418 resistance and identified by Southern blotting analysis of genomic DNA after EcoRI digestion. ES cell clones with the correct recombination were injected into eight-cell stage embryos of CD-1 mice to generate chimeric mice, which were mated with C57BL/6 mice to obtain heterozygous offspring (Rng105^f/+). Homozygous floxed Rng105 mice (Rng105^f/f) were obtained by mating heterozygotes. Floxed Rng105 mice were further crossed with Camk2a-Cre transgenic mice (C57BL/6-TgN[a-CaMKII-nlCre]/20, RIKEN RBRC00254) to obtain heterozygous conditional deletion mice (Camk2a-Cre;Rng105^f/+). Homozygous conditional deletion mice (Camk2a-Cre;Rng105^f/f) and control mice (Rng105^f/f) were obtained by crossing Camk2a-Cre;Rng105^f/+ female and Rng105^f/f male mice because Cre is expressed in male germ cells as well as in the central nervous system under the control of the Camk2a promoter. Genotyping was performed by PCR with primers 5'-AGATGGCTTTTCTTCTGCCA-3' and 5'-CTGGAAAACACGCTCAACAA-3', which amplified a 918 bp product from the wild-type Rng105 allele and a 978 bp product from the floxed Rng105 allele; and primers 5'-GTCGATGCAACGAGTGATGA-3' and 5'-AGCATTGCTGTCACTTGGTC-3', which amplified a 291 bp product from the Cre transgene.

The Thy1-GFP transgenic mice (Tg[Thy1-EGFP]MJrs/J) were purchased from Jackson Laboratory (Bar Harbor, ME, USA). We crossed Camk2a-Cre;Rng105^f/+ female and Thy1-GFP;Rng105^f/f male mice to obtain Thy1-GFP;Rng105^f/f control mice and Thy1-GFP;Camk2a-Cre;Rng105^f/f RNG105 conditional deletion mice.

In the glutamate uncaging experiments, dissociated hippocampal neurons were prepared from Rng105^f/f embryos at embryonic day 17–18 (E17–18). Neurons were plated at a density of 1.6 × 10⁶ cells/cm² onto poly-D-lysine-coated coverslips in glass-bottomed dishes (MatTek, Ashland, MA, USA) in Neurobasal-A medium (Thermo Fisher Scientific, Waltham, MA, USA) containing B-27 supplement (Thermo Fisher Scientific), 0.5 mM glutamine and 25% Neuron culture medium (Wako Pure Chemical Industries, Osaka, Japan). Cultures were incubated at 37°C in a 5% CO₂ incubator. The neurons were transfected with plasmids at 6 days in vitro (DIV) using conventional calcium-phosphate transfection method.

In the GluR1 and GluR2 immunostaining and biotinylation experiments, dissociated cerebral cortical neurons were prepared from individual littermates at E17.5. Neurons were cultured in the same way as above at a density of 6.4 × 10⁴ cells/cm².

CHO-K1 cells (RCB0285, RIKEN BRC, Tsukuba, Japan) were cultured in HAM's F-12 (Wako Pure Chemical Industries) containing 5% fetal calf serum (FCS) at 37°C in the 5% CO₂ incubator. The cells were transfected with plasmids using Lipofectamine 2000 (Thermo Fisher Scientific) in accordance with the manufacturer's protocol. CHO-K1 cells were not included in the list of commonly misidentified cell lines maintained by the International Cell Line Authentication Committee. The origin of the cells (Chinese hamster) was confirmed by PCR in RIKEN BRC (link of datasheet is http://www2.brc.riken.jp/lab/cell/detail.cgi?cell_no=RCB0285). The cells were negative for mycoplasma by both PCR and nuclear staining, which were performed based on protocols by RIKEN BRC (http://cell.brc.riken.jp/ja/quality/myco_kensa).

To construct the expression vector for Cre, Cre cDNA was obtained using RT-PCR from RNA isolated from the cerebral cortex of Camk2a-Cre transgenic mouse with primers 5'-GGGGAATTCATGTCCAATTTACTGACC-3' and 5'-CTCGAATTCCTAATCGCCATCTTCCAGC-3'. The product was cloned into the EcoRI site of pEGFP-N1 (Clontech, Mountain View, CA, USA) whose GFP coding sequence was deleted by BamHI/NotI digestion. To construct the expression vector for RNG105 (a.a. 1–122)-GFP, Rng105 cDNA was obtained by RT-PCR from RNA isolated from mouse cerebral cortex with primers 5'-GTCGACATGCCCTCGGCCACCAGCCACAG-3' and 5'-GAATTCGGAGCTTTATATCTTGACTTAATG-3'. The product was cloned into the XhoI/EcoRI sites of pEGFP-N1 (Clontech). An expression vector for mCherry (pCS2-mCherry) was kindly donated by Dr. N. Kinoshita.

Extracts of mouse brains were prepared by homogenization in 50 mM Tris (pH 8.0), 150 mM NaCl, 1% NP-40, protease inhibitors (1 mM PMSF, 10 µg/ml leupeptin, pepstatin, and aprotinin) and 1 mM dithiothreitol. After centrifugation for 10 min at 10,000 × g at 4°C, the supernatant was added to Laemmli sample buffer and boiled. Extracts from cultured CHO cells were prepared in the same way. The extracts were separated by SDS-PAGE, transferred to polyvinylidene fluoride membranes (Merck Millipore, Billerica, MA, USA) and probed with an anti-RNG105 polyclonal antibody (Shiina et al., 2010), anti-α-tubulin monoclonal antibody (1:2,000, DM1A, Sigma-Aldrich, St. Louis, MO, USA), or anti-GFP monoclonal antibody (1:500, GF200, Nacalai Tesque, Kyoto, Japan). Biotinylated secondary antibodies (GE Healthcare, Chicago, IL, USA) and alkaline phosphatase-conjugated streptavidin (GE Healthcare) were used for the detection with a bromochloroindolyl phosphate/nitro blue tetrazolium solution.

To immunostain brain slices, adult mouse brains were infused with Tissue-Tek (Sakura Finetek, Tokyo, Japan), frozen in liquid nitrogen and sectioned at 10 µm using a cryostat (HM500-OM, Carl Zeiss, Oberkochen, Germany). The sections were mounted on silane-coated coverslips and dried for ~30 min at room temperature. The samples were fixed with 3.7% formaldehyde in phosphate-buffered saline (PBS; 137 mM NaCl, 8.1 mM Na₂HPO₄, 1.5 mM KH₂PO₄, and 2.7 mM KCl, pH 7.4) for 10 min at room temperature and permeabilized with 0.5% Triton X-100 in PBS. After blocking with 10% FCS in PBS, the samples were incubated with the anti-RNG105 antibody over night at 4°C. After washing with PBS, the samples were incubated with Alexa488-conjugated anti-rabbit IgG (1:400, Jackson ImmunoResearch, West Grove, PA, USA) and 1 µg/ml 4',6-diamidino-2-phenylindole (DAPI) (Wako Pure Chemical Industries) for 1 hr at room temperature to label RNG105 and nuclei, respectively. To immunostain cultured neurons, neurons at 12 DIV were fixed and stained in the same way. Fluorescence images were acquired using an A1 confocal laser microscope equipped with a Ti-E inverted microscope (Nikon, Tokyo, Japan) with a 10 × objective lens or a PlanApo VC60 × oil objective lens.

Brains were removed from the Thy1-GFP expressing Rng105^f/f and Camk2a-Cre;Rng105^f/f mice and fixed with 3.7% formaldehyde in PBS for 2 hr at room temperature. The brains were sectioned at 100 µm using a vibratome VT1200S (LEICA, Wetzlar, Germany), and the sections were mounted in Mowiol (Merck Millipore). Fluorescence images were acquired using the A1 confocal microscope with a 20 × objective lens to measure the length and branching of dendrites, and a 100 × objective lens to measure the size and morphology of spines. The images were analyzed using ImageJ software. Spines were classified as the mushroom type when W_neck/W_head < 0.5 and W_head > 0.4 µm, where W_neck and W_head are the width of spine neck and spine head, respectively.

Time-lapse two-photon imaging of dendritic spines was performed using an FVMPE-RS two-photon laser scanning microscope (Olympus, Tokyo, Japan) equipped with an Insight DS Dual-line laser system (Spectra Physics, Santa Clara, CA, USA) and a 95% O₂/5% CO₂ incubator (Tokai Hit, Shizuoka, Japan) with a water immersion objective lens XLPLN25XWMP2 (Olympus). The culture medium of dissociated hippocampal neurons (12–15 DIV) was exchanged with modified artificial cerebrospinal fluid (ACSF) (125 mM NaCl, 2.5 mM KCl, 3 mM CaCl₂, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, 20 mM glucose, 1 μM tetrodotoxin [TTX] [Wako Pure Chemical Industries], and 50 µM picrotoxin [Sigma-Aldrich], gassed with 95% O₂/5% CO₂ before use) containing 2 mM MNI-caged L-glutamate (Tocris, Ellisville, MO, USA). MNI-caged L-glutamate was uncaged locally near single spine heads by 120 pluses (2 ms pulse duration at 2 Hz) of 740 nm laser illumination with laser power of 5 mW. For imaging of spines with mCherry, a 1,040 nm laser was used. 20 images at 0.5 µm focus step were projected by summation, and the fluorescence intensity of spine (sum of pixel intensity in the spine area), which reflects spine volume, was measured using ImageJ software as described previously (Matsuzaki et al., 2004).

Hippocampal slice preparation and electrophysiology were performed as described previously (Shinoda et al., 2011). Briefly, postnatal 8–10 (P8–10) weeks old Camk2a-Cre;Rng105^f/f mice or Rng105^f/f littermates were deeply anesthetized and decapitated, brains were isolated and cooled rapidly to 4°C. Transverse 400 µm thickness hippocampal slices were prepared using a vibratome RPO7 (D.S.K, Kyoto, Japan) and high-sucrose cutting solution (the formulations of all solutions are described below), and maintained in ACSF at room temperature for at least 2 hr. A bipolar stimulation electrode was placed in the CA1 SR region to stimulate CA3 Schaffer collateral fibers. Field excitatory postsynaptic potentials (fEPSPs) were recorded using a capillary glass electrode (Harvard Apparatus, Massachusetts, MA, USA) filled with ACSF placed in the CA1 SR after a 0.05 Hz test pulse generated by a pulse generator Master-8 (A.M.P.I., Jerusalem, Israel) equipped with an isolator ISO-Flex (A.M.P.I.). For LTP recording and theta-burst stimulation, 50% maximum stimulus intensity was used. LTP was induced by theta-burst stimulation (Four trains with 10 s intervals between trains; each train had five bursts separated by 200 ms and included four pulses delivered at 100 Hz). Data were amplified using a MultiClamp 700A (Molecular Devices, Sunnyvale, CA, USA), digitized at 10 kHz and filtered at 2 kHz using a Digidata 1440 system (Molecular Devices) with pCLAMP9 software (Molecular Devices). The formulation of the solutions were (in mM): High-sucrose cutting solution; 234 sucrose, 2.5 KCl, 1.25 NaH₂PO₄, 0.5 CaCl₂, 10 MgSO₄, 26 NaHCO₃, and 11 D-glucose, gassed with 95% O₂/5% CO₂. ACSF; 125 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 2 CaCl₂, 1 MgCl₂, 26 NaHCO₃, and 11 D-glucose, gassed with 95% O₂/5% CO₂.

In all behavioral tests, male mice (3–4 months old; 2–4 mice were co-housed in a 12 hr dark/light cycle) were used during the light cycle. The open field test was conducted in a round apparatus (85 cm in diameter with a 40 cm wall) with concentric circles and radial lines drawn on the floor, which divided the floor into 25 blocks. A mouse, naive to the apparatus, was placed in the center of the field and allowed to explore for 5 min. The total number of line crossings was recorded as an index of exploratory activity. The mouse was subjected to the test on three consecutive days to assess habituation to the novel place.

The test was conducted with an apparatus consisted of a light chamber and a dark chamber separated by a sliding door (LDK-M, Melquest, Toyama, Japan). A mouse was automatically monitored using a SCANET system (Melquest). The mouse was placed into the light chamber, and 5 s later, the door was opened. The mouse was allowed to move freely between the two chambers for 5 min. Distance traveled, time spent in each chamber, and the number of transition between the chambers were measured.

To habituate to the test environment, mice were placed in a chamber (43 × 43 × 29.5 cm) and allowed to explore freely for 1.5 hr on four consecutive days. In the first session, two identical objects (object 1 and object 2, cell culture flasks) were placed at the opposite corners of the chamber, and the mouse was allowed to explore the chamber for 5 min. In the second session, object two was replaced by a different type of object (object 3, a wooden prism), and the mouse was allowed to explore the chamber for 5 min. The interval between the sessions was 2.5 hr. The number of mouse interactions with the objects was counted, where object interaction was defined as the nose directing toward the object at a distance ≤2 cm.

The rotarod test was conducted using ROTA-ROD for mice 7650 (Ugo Basile, Varese, Italy). A mouse was placed on a rod rotating at a fixed speed of 24 rpm. The duration of a trial was 3 min, and if the mouse fell from the rod within 3 min, the mouse was re-placed on the rod. The number of falls and the longest latency without falling within a trial were measured. The mouse was subjected to the test on three consecutive days to assess motor skill learning.

The Morris water maze was conducted in a round water tank with a diameter of 110 cm filled with ~24°C water. First, the mouse was subjected to visible platform test. In this test, a black platform (6 cm in diameter) was placed 0.5 cm above the water surface. A mouse was placed in the pool and the escape latency to the platform was measured. If the mouse found the platform within a 1 min time limit, the mouse was allowed to stay on the platform for 30 s. If not, the mouse was guided to the platform before staying on the platform for 30 s. Mice were given six trials per day for five consecutive days. The location of the platform and the start position were changed randomly in each trial.

Next, the mice were subjected to a hidden platform test, in which a clear platform (6 cm in diameter) was placed 1 cm below the water surface. The water was clouded with non-toxic white paint (Sakura Color Products, Osaka, Japan). Spatial cues (four different plane figures) were attached on the interior wall of the tank above the water surface. The test was conducted in the same way as in the visible platform test with a 2 min time limit. The start position was changed randomly, but the location of the platform was fixed. The mice were given six trials per day for 10 consecutive days.

One day after the last trial of the hidden platform task, the mice were subjected to a probe test. The platform was removed from the pool and the swimming path of the mouse was tracked for 1 min using a computer-based video tracking system ANY-Maze (Stoeling, Wood Dale, IL, USA). Time spent in the target quadrant, where the hidden platform had been placed, and in the other quadrants was also measured.

The test was conducted using the LDK-M chamber (Melquest) with the SCANET system (Melquest). One day before the training day, to habituate to the test environment, the mouse was allowed to freely explore both chambers for 30 min with the door opened. On the training day, the mouse was placed in the light chamber and allowed to freely explore the chambers for 15 min after the door opened, and the time spent in the dark chamber during the first 5 min was measured (Pre-FS). After that, when the mouse entered the dark chamber, the door was closed and foot shock (0.5 mA, 1 s duration, 4 times with 1 min intervals) was delivered. After the foot shock, the mouse was returned to its home cage. The mouse was replaced in the light chamber at 5 min, 1 day, and 1 week after the foot shock and allowed to freely explore the chambers for 5 min after the door opened, and the time spent in the dark chamber was measured.

A mouse was placed in a test chamber (15 × 15 × 15 cm, the dark chamber of LDK-M with the door closed) and allowed to explore freely for 30 s. Then the mouse received a foot shock (0.5 mA, 2 s) and was returned to its home cage. Five days after the conditioning, the mouse was replaced in the test chamber and in a control chamber (24.5 × 15.5 × 14.8 cm, CL-0112–3, CLEA Japan, Tokyo, Japan) for 1 min each. Freezing time in the chambers was measured by ANY-Maze.

Mouse brains (P12 weeks old) were removed in ice cold sterile PBS, and embedded in a 4.5% low-melting point agarose (Agarose XP, Nippon Gene, Tokyo, Japan). Coronal sections (500 µm thick) were sliced in ice-cold sterile PBS using the vibratome VT1200S, and transferred into ice-cold sterile PBS. Brain slices were stained with 1 µg/mL DAPI for 10 min at 4°C. Stratum pyramidale (SP) and stratum radiatum (SR) in the hippocampal CA1 region were microdissected manually using glass capillaries (G-1, Narishige, Tokyo, Japan) with the use of a stereomicroscope. Glass capillaries were made using a needle puller (model PB-7, Narishige). The borders between the stratum oriens (SO) and SP, and between the SP and SR were cut to isolate the SP, and the border between SP and SR, and between SR and stratum lacnosum-moleculare (SLM) were cut to isolate the SR. To avoid contamination of SR with the soma of pyramidal neurons, the isolated SR was checked to confirm it did not contain a high-density DAPI-positive nucleic layer (SP) with the use of an inverted fluorescence microscope (IX83, Olympus) with a 40 × objective lens. To obtain sufficient amount of RNA for RNA-seq, left and right hippocampi from three mice were dissected and collected into one sample tube. Triplicate samples were prepared for RNA-seq analysis. The samples were stored at −80°C until RNA extraction.

Total RNA was extracted from the tissues (SP and SR) using ISOGEN (Nippon gene) in accordance with the manufacturer's protocols with minor modifications. The isolated tissue was homogenized in 400 µL ISOGEN by pipetting. After the homogenate was stored at room temperature for 5 min, 100 µL chloroform was added and the sample was shaken vigorously for 30 s. After being stored on ice for 5 min, the sample was centrifuged for 15 min (21,900 × g, 4°C), and about 240 µL of the aqueous phase containing RNA was collected in a new tube. The sample was centrifuged for 15 min (21,900 × g, 4°C) again and the aqueous phase was collected into a new tube. To remove phenol and chloroform from the sample, 240 µL diethyl ether was added to the sample and vortexed for 1 min. After the sample was centrifuged for 1 min (21,900 × g, 4°C), the upper phase was removed. This diethyl ether treatment was performed three times. Then, 25 µL 3 M sodium acetate and 250 µL 2-propanol were added to the sample and mixed, and then RNA was precipitated on ice for 30 min. After the sample was centrifuged for 15 min (21,900 × g, 4°C), the supernatant was removed. The RNA precipitate was washed with 500 µL 70% ethanol, dried using a vacuum desiccator and dissolved in 22 µL RNase-free water (TaKaRa, Shiga, Japan).

Next, DNA was removed from the sample by DNase treatment. 0.5 µL DNase (RT grade) (Nippon Gene) and 10 × buffer were added to the 22 µL RNA solution, and the sample was incubated for 15 min at 37°C. Then, 25 µL phenol-chloroform mixture was added, and the sample was vortexed for 30 s. After the sample was centrifuged for 10 min (21,900 × g, 4°C), upper phase was collected. Phenol and chloroform were removed from the sample by diethyl ether treatment as described above. Then, 2.5 µL 3 M sodium acetate and 62.5 µL 100% ethanol were added to the sample and mixed, and then RNA was precipitated at −20°C for 30 min. After the sample was centrifuged for 20 min (21,900 × g, 4°C), the supernatant was removed. The RNA precipitate was washed with 150 µL 70% ethanol, dried, and dissolved in 10 µL RNase-free water. RNA solution was stored at −80°C until use for the preparation of cDNA libraries.

Before preparation of cDNA libraries, the quality of the extracted RNA was checked. RNA integrity was measured using an Agilent 2100 bioanalyzer (Agilent Technologies, Santa Clara, CA, USA) with RNA 6000 Nano Kit and RNA 6000 Pico Kit (Agilent Technologies) in accordance with the manufacturer's protocols. The RNA integrity number was over seven for all samples, which was sufficient quality for RNA-seq.

Total RNA (200 ng per sample) was used as the starting material to prepare cDNA using TruSeq RNA sample Preparation Kit v2 (Illumina, San Diego, CA, USA) at a half scale compared with the manufacturer's protocols, as follows. Poly-A-containing mRNAs were purified using Oligo dT magnetic beads. After the mRNAs were denatured, they were fragmented at 94°C for 4 min. First-strand cDNA was synthesized using SuperScript II Reverse Transcriptase (Thermo Fisher Scientific), and then second-strand cDNA was synthesized. After the ends of the fragments were blunted, an adenine nucleotide was added to the 3'-end at 37°C for 30 min. RNA adapter index was ligated to the 5'- and 3'-ends of the ds cDNA, and the cDNA was amplified with 10 PCR cycles. cDNA quality was validated using the bioanalyzer with a High Sensitivity DNA Kit (Agilent Technologies). The cDNA libraries were quantified with quantitative PCR (qPCR) using a 7500 real-time PCR system (Thermo Fisher Scientific) and adjusted to 2 nM.

In each experiment, an equal amount of cDNA from SP and SR was analyzed, and three independent experiments were conducted. Twelve cDNA libraries (four kinds of samples × three biological replicates) were sequenced with 101 bp paired-end sequencing using a HiSeq1500 (Illumina). The resulting reads were mapped to the mouse genome (Mus musculus Ensemble NCBIM37) using TopHat (v 2.0.11). The mapped reads were assembled using Cufflinks (v 2.2.1), and differential gene enrichment analysis was conducted using Cuffdiff.

S-mRNAs and D-mRNAs in control (Rng105^f/f) mice were identified as follows. First, mRNAs from SP and SR of Rng105^f/f mice were analyzed using Cuffdiff, as described above, which identified mRNAs whose concentration (FPKM value) was significantly different between SP and SR, designated as ‘yes’, and not significantly different between the layers designated as ‘no’ in the ‘all mRNAs’ list (Supplementary file 1). After this, mRNAs, whose FPKM values were more than zero both in SP and SR, were selected and subjected to subsequent data analysis. ‘Yes’ mRNAs were divided into two groups, candidates for dendritic mRNAs whose DAIs (relative FPKM value in SR to SP) were more than 1, and candidates for somatic mRNAs whose DAIs were less than 1. Next, to identify mRNAs specifically expressed in pyramidal neurons, we eliminated mRNAs also expressed in other types of cells such as glial cells, interneurons, and endothelial cells (Cajigas et al., 2012; Doyle et al., 2008; Daneman et al., 2010; Cahoy et al., 2008) from each of the lists (Supplementary file 2A,B). Finally, we identified 1122 dendritic mRNAs, 2106 somatic mRNAs, and 2814 non-significant mRNAs in Rng105^f/f mice (Supplementary file 3A−C). Furthermore, to compare the dendritic localization of mRNAs between Rng105^f/f and Camk2a-Cre;Rng105^f/f mice, the ratio of DAI in Camk2a-Cre;Rng105^f/f mice to Rng105^f/f mice was calculated for each mRNA and indicated in the ‘DAI (Camk2a-Cre;Rng105^f/f)/DAI (Rng105^f/f)’ columns (Supplementary file 3A−C).

Gene ontology enrichment analysis was performed using DAVID functional annotation tools. Significance of overrepresentation of GO terms was assessed using the Benjamini-Hochberg false discovery rate (FDR) criterion at p<0.05.

Immunostaining of cultured cortical neurons (9 DIV) for GluR1 and GluR2 was conducted as described previously (Ohashi et al., 2016). To block neuronal activity, 0.7 µM TTX and 20 µM D-2-amino-5-phosphonovaleric acid (APV) (Sigma-Aldrich) were added to the medium for 24 hr at 37°C in a 5% CO₂ incubator. Live neurons were incubated with an anti-GluR1 (1:15, PC246, Merck Millipore) or an anti-GluR2 antibody (1:100, MAB397, Merck Millipore) at 37°C in a 5% CO₂ incubator for 1 hr. After the neurons were washed in Neurobasal-A medium, they were fixed with 3.7% paraformaldehyde in PBS for 20 min at 25°C. Fixed neurons were blocked for 30 min in 10% FCS in DMEM (Sigma-Aldrich), and incubated with an Alexa Fluor 488-conjugated anti-rabbit and mouse IgG antibodies (1:400, Thermo Fisher Scientific) in 10% FCS in DMEM for 3 hr at 25°C to label cell surface GluR1 and GluR2. After neurons were washed in PBS, they were fixed again and permeabilized with 0.25% Triton X-100 in PBS for 10 min. After the neurons were blocked, they were incubated with the anti-GluR1 antibody (1:50) or anti-GluR2 antibody (1:100) for 12 hr at 4°C and then with a Cy3-conjugated anti-rabbit and mouse IgG antibodies (1:400, Jackson Immuno Research) for 3 hr at 25°C to label intracellular and residual surface GluR1 and GluR2.

Neurons were imaged using the IX83 inverted fluorescence microscope (Olympus) with a 40 × objective lens and an ORCA-R2 digital CCD camera (Hamamatsu Photonics, Hamamatsu, Japan). Because GluR1 and GluR2 were detected in a punctate manner, they were quantified by counting the number and measuring the fluorescence intensity of the puncta in dendrites. To count the number of GluR1 and GluR2 puncta, the images were converted into binary images by selecting dendritic regions and using the MaxEntropy threshold algorithm in ImageJ software. The number of GluR1 and GluR2 puncta in dendrites was counted using the magic wand tool and analysis tool in Adobe Photoshop software. To count total GluR1 and GluR2 puncta, the binary images of before and after permeabilization were merged and used. To measure the fluorescence intensity of GluR1 and GluR2 puncta in dendrites, the binary image and original image were layered in Adobe Photoshop software. ROIs were selected in the binary layer using the magic wand tool, and fluorescence intensity in the ROIs in the original layer was calculated by multiplying mean pixel intensity by area of the ROIs. The sum of fluorescence intensity was normalized by dendrite length and by the intensity of GluR1 and GluR2 in the soma. GluR1 and GluR2 intensity in the soma was calculated using the layered images by multiplying mean pixel intensity by area of ROIs in the soma and normalized by the area of the soma. Fluorescence intensity of GluR1 and GluR2 puncta immunostained before permeabilization was normalized by that immunostained after permeabilization.

Biotinylation assay was conducted as described previously (Chung et al., 2000; Snyder et al., 2001; Aoto et al., 2008). Primary cultured cortical neurons (9 DIV) in two 30 mm dishes were treated with or without 1 μM TTX and 100 μM APV in the culture medium for 24 hr prior to surface biotinylation (Aoto et al., 2008). The dishes were placed on ice and washed three times with ice-cold ACSF (124 mM NaCl, 5 mM KCl, 1.25 mM NaH₂PO₄, 26 mM NaHCO₃, 0.8 mM MgCl₂, 1.8 mM CaCl₂, and 10 mM D-glucose, gassed with 95% O₂/5% CO₂). Then the neurons were incubated with ACSF containing 1 mg/ml sulfo-NHS-LC biotin (Thermo Fisher Scientific) for 30 min on ice. After washing once with ice-cold 100 mM glycine in ACSF and three times with ice-cold TBS (50 mM Tris, 150 mM NaCl, pH 7.5), the neurons were lysed in 100 μl of modified RIPA buffer (1% Triton X-100, 0.1% SDS, 0.5% deoxycholic acid, 50 mM NaHPO₄ [pH 7.2], 150 mM NaCl, 2 mM EDTA, 25 mM β-glycerophosphate, 1 mM PMSF, 10 µg/ml leupeptin). The lysate was centrifuged at 14,000 × g for 15 min at 4°C, and 85 μl of supernatant was incubated with 40 μl of NeutraAvidin Agarose beads (Thermo Fisher Scientific) for 3 hr at 4°C with gentle rocking. After washing three times with modified RIPA buffer, biotinylated proteins were eluted from the beads with 80 μl of SDS sample buffer and boiled for 5 min. The total lysate and biotinylated eluate were analyzed by western blotting with the anti-GluR1 (1:50, PC246, Merck Millipore), and anti-GluR2 (1:1000, MAB397, Merck Millipore) antibodies. Alkaline phosphatase-conjugated secondary antibodies (1:5000, 711-055-152 and 115-055-146, Jackson Immuno Research) and Can Get Immunoreaction Enhancer Solution (TOYOBO, Osaka, Japan) were used for the detection with a bromochloroindolyl phosphate/nitro blue tetrazolium solution. Quantification of the band intensity was conducted as previously described (Ohashi et al., 2016). A standard dilution series of the lysate from cultured neurons was loaded on the same gel, and used to generate a standard curve and calculate the relative intensity of upper and lower bands of GluR1 and GluR2. The band intensity was measured using ImageJ software. Triplicate from three mice (nine samples) were analyzed for each group.

Sample numbers and experimental repeats are indicated in figure legends. Statistical significance was determined using Student's t-test, paired t-test, one-way ANOVA, one-way repeated measures ANOVA, two-way ANOVA, two-way repeated measures ANOVA, post-hoc Tukey-Kramer test, and Bonferroni post-hoc t-test as indicated in the figure legends. Statistical analysis was performed in R, Excel, or an Excel add-in software Statcel (The Publisher OMS Ltd., Saitama, Japan). Exact F, t, and p-values are indicated in Statistical reporting table (Supplementary file 7). Post-hoc power analysis was performed with G*Power (http://www.gpower.hhu.de/) and statistical power is indicated in the Statistical reporting table. No blinding method was used in this study. In behavioral tests, animals that died before the experimental endpoint were excluded from the data analysis. The Gene ontology term enrichment was analyzed using DAVID functional annotation tools (https://david.ncifcrf.gov/).

Raw and processed data files for the RNA-seq analysis have been deposited in the NCBI Gene Expression Omnibus (GEO) under series accession number GSE96552 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE96552).

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

We thank Dr. M Matsuzaki for advice on glutamate uncaging experiments; C Matsuda for technical assistance; Spectrography and Bioimaging Facility and Functional Genomic Facility, NIBB Core Research Facilities for technical supports. This work was supported by Grant-in-Aid for Scientific Research on Priority Areas (Molecular Brain Science) from the MEXT of Japan, JSPS KAKENHI Grant Number 16K07361 to NS, and JSPS KAKENHI Grant Number 16J07985 to RO (RO is a JSPS Research Fellow). The work was also supported by the Uehara Memorial, the DAIKO, and the Toyoaki foundations to NS.

Animal experimentation: All animal care, experiments and behavioral testing procedures were approved by the Institutional Animal Care and Use Committee of the National Institutes of Natural Sciences (Permit Number: 17A059), and performed in accordance with the guidelines from the National Institutes of Natural Sciences, Niigata University and the Science Council of Japan.

1. Received: June 21, 2017
2. Accepted: October 22, 2017
3. Version of Record published: November 21, 2017 (version 1)

© 2017, Nakayama et al.

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